Hair Cell Micromechanics & Hearing (Singular Audiology Text.)

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Author: Charles I Berlin | Richard P. Bobbin

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Hair Cells Micromechanics and Hearing

A Singular Audiology Textbook Jeffrey L. Danhauer, Ph.D. Audiology Editor

Hair Cells Micromechanics and Hearing Edited by

Charles I. Berlin, Ph.D. Kenneth and Frances Barnes Bullington Professor of Hearing Science Director, Kresge Hearing Research Laboratory of the South Louisiana State University Health Sciences Center Department of Otolaryngology and Biocommunication New Orleans, Louisiana

Richard P. Bobbin, Ph.D. Professor Department of Otorhinolaryngology and Biocommunication Kresge Hearing Research Laboratory of the South Louisiana State University Health Sciences Center New Orleans, Louisiana

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Hair Cells: Micromechanics and Hearing by Charles I. Berlin, Ph.D. and Richard P. Bobbin, Ph.D.

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Executive Production Editor: Karen Leet Production Editor: Brad Bielawski

Editorial Assistant: Kristin Banach

COPYRIGHT © 2001 by Singular, an imprint of Delmar, a division of Thomson Learning, Inc. Thomson Learning™ is a trademark used herein under license Printed in Canada 1 2 3 4 5 XXX 06 05 04 02 01 For more information contact Singular, 401 West “A” Street, Suite 325 San Diego, CA 92101-7904 Or find us on the World Wide Web at http://www.singpub.com

ALL RIGHTS RESERVED.No part of this work covered by the copyright here-on may be reproduced or used in any form or by any means—graphic, electronic, or mechanical, including photocopying, recording, taping, Web distribution or information storage and retrieval systems—without written permission of the publisher.

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NOTICE TO THE READER Publisher does not warrant or guarantee any of the products described herein or perform any independent analysis in connection with any of the product information contained herein. Publisher does not assume, and expressly disclaims, any obligation to obtain and include information other than that provided to it by the manufacturer. The reader is expressly warned to consider and adopt all safety precautions that might be indicated by the activities herein and to avoid all potential hazards. By following the instructions contained herein, the reader willingly assumes all risks in connection with such instructions. The Publisher makes no representation or warranties of any kind, including but not limited to, the warranties of fitness for particular purpose or merchantability, nor are any such representations implied with respect to the material set forth herein, and the publisher takes no responsibility with respect to such material. The publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or part, from the readers’ use of, or reliance upon, this material.

Contents

Preface by Charles I. Berlin, Ph.D. and Richard P. Bobbin, Ph.D. Contributors Chapter 1 Transduction and Adaptation by Vertebrate Hair Cells

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David P. Corey, Ph.D. Chapter 2 How a Living Hair cell Repairs Itself: Involvement of Purinoceptors In the Repair of Hair Bundle Mechanoreceptors of Sea Anemones

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Glen M. Watson, Ph.D. Chapter 3 Fast Transducer Adaptation, Physiological Implications and Underlying Mechanisms

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Anthony J. Ricci, Ph.D. Chapter 4 Hearing in a Primitive Animal

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Patricia Mire, Ph.D. Chapter 5 Brn-3.1 is Required for Development and Survival of the Hair Cell

93

Allen F. Ryan, Ph.D. Chapter 6 Enhancing Signal Discrimination by Means of a Metabotropic Glutamate Receptor

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Adam W. Hendricson, Grace Athas, Ph.D., Paul S. Guth, Ph.D. Chapter 7 Additional Studies on the Role of ATP as a Neuromulator in the Organ of Corti

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Richard P. Bobbin, Ph.D., Christopher S. LeBlanc, Manisha Mandhare, Margarett S. Parker Index

155 v

Preface

On October 8th, 1999, Dr. David Corey of Harvard University became the sixth winner (recipient) of the annual Kresge-Mirmelstein Award. This award was jointly conceived by the late Rona Mirmelstein and Dr. Charles I. Berlin to promote hearing research in general, and tangibly reward the world’s best auditory scientists for their successful efforts. The winner is now chosen by the previous winners purely on the basis of the power and importance of their scientific contribution and this selection is made with no other consideration. Once he or she accepts the award, each winner is asked to exchange a chapter describing his or her work for the cash award. At the meeting, related other invited investigators give papers and the manuscripts are then collated into a publication. In fact, the resulting manuscripts become what, in German, is an ideal word…a Festschrift, meaning a collection of writings to celebrate either a person or an event. In this case we celebrate both the person and the event. Each of the previous winners contributed the leading chapter to the following books all published by Singular Publishing Group: Winner (Recipient)

Year Awarded

Title of Book

Year Published

William Brownell, Ph.D.,

1994

Hair Cells and Hearing Aids

1996

Robert Wenthold, Ph.D.,

1995

Neurotransmission and Hearing Loss

1997

David Kemp, Ph.D.

1996

Otoacoustic Emissions

1998

M.C.Liberman, Ph.D.

1997

The Efferent Auditory System

1999

Karen Steele, Ph.D.

1998

Genetics and Hearing Loss

2000

David Corey, Ph.D.

1999

Hair Cells: Micomechanics and Hearing

2000

The seventh recipient of this award is Peter Dallos, Ph.D. for his work cloning the hair cell motor about which Brownell wrote. We expect Dr. Dallos’ award to be given on October 20th, 2000. vii

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HAIR CELLS MICROMECHANICS AND HEARING

The meeting was held at the New Orleans Marriott Hotel and called to order by our redoubtable Dr. Douglas Webster, who has in the past been an exemplary Master of Ceremonies, and returned from his retirement in Arizona to serve in this special capacity. Our colleague, Nicolas Bazan, M.D., Ph.D., co-sponsored this meeting and also gave introductory remarks to set the stage for what turned out to be a thrilling day in auditory science. Our Department Head, Dr. Daniel W. Nuss greeted the group, and remained throughout the day in testimony to his interest in these important scientific issues. The first talk was by Dr. David Corey and entitled “Dancing on the Head of a Pin: Hair Cell Micromechanics and the Cochlea”. The frog vestibular hair cell was the model here for an outstanding talk on the importance of the transduction apparatus for adaptation. He reviewed the two general mechanism proposed for adaptation, one involving a myosinpowered motor that maintains tension on the channels, and another involving calcium entering through the transduction channels. The second talk of the day was presented by Dr. Glen Watson on sea anemones, that are essentially living hair cells. Dr. Watson presented evidence that showed how living hair bundles on hair cells are repaired following trauma by secretory proteins called “repair proteins”. The recovery induced by repair proteins is enhanced by the chemical messenger called adenosine triphosphate (ATP) acting on ATP receptors and using calcium as a second messenger. Because ATP receptors are found on the hair bundles of mammalian hair cells, this research by Dr. Watson may have direct implications on how human hearing recovers following noise exposure. Dr. Anthony Ricci, who was among the first to suggest that adaptation of the transduction apparatus may impart a mechanical tuning to the hair bundles, amplified the topic introduced by Dr Corey. Dr. Ricci reviewed existing data regarding fast adaptation with emphasis on the evidence that adaptation may be a mechanical tuning mechanism. He also presented evidence that there may be multiple mechanisms of adaptation. Dr. Patricia Mire, who studies the hair bundles on tentacles of sea anemones as a model of hair bundles on hair cells of vertebrates, discussed the similarities and differences of these two systems. Dr. Mire presented data demonstrating that the hair bundles on the tentacles are modulated by chemoreceptors and possess self repair mechanisms. A tip link model of transduction with modifications for functioning in prey detection in a noisy environment was presented. Dr. Allen Ryan, discussed the development of the ear and the role of transcription factors which regulate gene expression by binding to specific DNA sequences. Dr. Ryan presented evidence that the transcription factor Brn-3.1 is a master gene regulatory factor required for the differentiation of hair cells from their precursors. Studies of Brn-3.1 may lead to discoveries of genetic mutations that underlie various forms of genetic deafness. Dr. Paul Guth, together with colleagues Drs. Grace Athas and Adam W. Hendricson discussed the evidence for a role of metabotropic glutamate

PREFACE

ix

receptors present on vestibular hair cells in transmitter release from these cells. The hypothesis being tested is whether these receptors are recruited under conditions of mechanically-evoked activity, but not at rest. Finally, Dr. Richard Bobbin, and colleagues, presented a continuation of their ATP story published in the previous book of this series. Additional immunohistochemical and electrophysiological data are presented examining the role of ATP and ATP receptors on cells exposed to perilymph in modulation of cochlear mechanics and a control of ion flow. These receptors appear to have a different function than those exposed to endolymph which act in repair of the hair bundles as discussed in an earlier chapter by Dr. Watson. Dr. Nuss closed our meeting expressing his appreciation and his admiration for the science presented, and urged us to continue this exciting and useful series. The careful reader will note that the current text was finished and published in record time. This is due in no small part to the extraordinary efforts, diligence and talents of Lillie Long, our Editorial Assistant. Ms. Long not only reminded the busy authors of their obligations on a weekly basis, but she personally revised and restructured references and formats to make the book as uniform and precise as possible. She showed a genuine talent for redaction and document preparation, and met publisher’s deadline with remarkably satisfying regularity. Our special thanks to her for her performance. Finally, we must acknowledge the far-sighted, thoughtful and committed munificence of Frances Barnes Bullington. Mrs. Bullington, who is a retired speech pathologist, funded the Regents-supplemented Professorship which Dr. Berlin now holds, and has made a bequest to fund a Chair as well. At this time our Board of Regents accepts petitions to supplement donated funds of $600,000 so that they reach the $1Million needed to support a Chair in perpetuity. It is this generosity of purse as well as spirit and commitment to our work that will ultimately leave our Otolaryngology Department with both a Professorship and a Chair funded in the names of Kenneth and Frances Barnes Bullington. We will be forever indebted to her. Charles I. Berlin, Ph.D. Kenneth and Frances Barnes Bullington Professor of Hearing Science Director, Kresge Hearing Research Laboratory of the South LSU Medical Center, Department of Otolaryngology and Biocommunication Richard P. Bobbin, Ph.D. Professor, LSU Medical Center, Department of Otolaryngology and Biocommunication

Contributors

Grace B. Athas, Ph.D. Pharmacology Tulane University School of Medicine New Orleans, Louisiana Richard P. Bobbin, Ph.D. Professor Department of Otorhinolaryngology and Biocommunication Kresge Hearing Research Laboratory of the South Louisiana State University Health Sciences Center New Orleans, Louisiana David P. Corey, Ph.D. Professor Howard Hughes Medical Institute Department of Neurobiology Harvard Medical School and Massachusetts General Hospital Boston, Massachusetts

Christopher S. LeBlanc Neuroscience Graduate Student Louisiana State University Health Science Center New Orleans, Louisiana Manisha S. Mandhare Technician in Otorhinolaryngology Department Louisiana State University New Orleans, Louisiana Patricia Mire, Ph.D. Research Assistant Professor of Biology The University of Louisiana Lafayette Lafayette, Louisiana

Paul S. Guth, Ph.D. Professor of Pharmacology Tulane University School of Medicine New Orleans, Louisiana

Margarett S. Parker, Ph.D. Department of Otorhinolaryngology and Biocommunication Kresge Hearing Research Laboratory of the South Louisiana State University Health Sciences Center New Orleans, Louisiana

Adam W. Hendricson Department of Pharmacology Tulane University School of Medicine New Orleans, Louisiana

Anthony Ricci, Ph.D. Assistant Professor Neuroscience Center of Excellence Kresge Hearing Research Laboratory of the South xi

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Louisisana State University Health Sciences Center Department of Neuroscience New Orleans, Louisiana Allen F. Ryan, Ph.D. University of California, San Diego School of Medicine Department of Otolaryngology – Head and Neck Surgery La Jolla, California

Glen M. Watson, Ph.D. Associate Professor Department of Biology University of Louisiana at Lafayette Lafayette, Louisiana

1 Transduction and Adaptation by Vertebrate Hair Cells David P. Corey, PhD

Two of the great mysteries facing auditory physiologists of a quarter century ago were how the mechanical stimulus of sound initiated a neural signal in the hair cells of the cochlea, and how the organ of Corti could amplify the vibration of the basilar membrane in a frequency-specific way to create the remarkably sharp tuning that characterizes auditory nerve fibers. Since then, the first question has been largely answered, at least in terms of a structural and biophysical model for transduction. Many details remain to be elucidated, especially in identifying the proteins that constitute the transduction apparatus, but there is strong evidence supporting the tip-links model for transduction and essentially none in conflict with it. There is less understanding of mechanical amplification by the organ of Corti. In part, this is because the most successful hair-cell organs for physiology have come from amphibians and reptiles, which were thought not to have amplification of the sort found in the mammalian cochlea. That view may be changing, with the recognition that most vertebrate ears can generate otoacoustic emissions, and that emissions may be a by-product of a cellular amplifier that has gone into oscillation. In addition, the amazingly large and fast electromotility of the mammalian outer hair cell has attracted much attention—deservedly so—but while it seems like the obvious motor to drive a cochlear amplifier, there are certain difficulties in working it into a high-frequency amplifier. Hair cells adapt to maintained stimuli. Several mechanisms mediate adaptation, including mechanical relaxations in the stimulus and changes in transmitter release, but an important one happens at the level of the transduction apparatus, where the mechanically-activated conductance tends to return to a resting level of 10 to 20% open, even if the mechanical deflection of the hair bundle is maintained. Two general mechanisms have been proposed for adaptation, one involving a myosin-powered motor 1

2


that maintains tension on the channels, and another involving calcium entering though the transduction channels and binding just inside to close the channels. Recent evidence suggests that both mechanisms may be operative, perhaps in different proportions in different cells. Moreover, the calcium-closure mechanism has characteristics that could power a mechanical amplification in the organ of Corti. It may be that a component of adaptation is in fact the long-sought cochlear amplifier.

TRANSDUCTION Bundle Structure Hair cells are epithelial cells, and have evolved their specialized structure from components of more generic epithelia. The mechanosensory part of the hair cell is, of course, the hair bundle, composed of two types of cilia. Stereocilia are elongated microvilli, and like microvilli have cores of crosslinked actin filaments (Flock, Cheung, Flock, and Utter, 1981; Tilney, DeRosier, and Muloy, 1980). They number between 20 and 300 in hair cells from different vertebrate organs; an example from the bullfrog saccule is shown in Figure 1–1, where the bundles contain about 60 stereocilia. Stereocilia can range in length from a few micrometers in the high-frequency regions of mammalian cochleas, to 80 m or more in the semicircular canals. In all cases, their heights rise in a regular staircase from the short edge of the bundle (the negative side) to the tall edge (positive). Each stereocilium contains hundreds of actin filaments, perhaps 1200 in the medium-sized stereocilia of Figure 1–1A, and these are cross linked by fimbrin to create a stiff rod (Sobin and Flock, 1983; Shepherd, Barres, and Corey, 1989). The actin filaments decrease in number as the stereocilium tapers near its bottom end, so that only a few dozen filaments coalesce into a rootlet that enters the cuticular plate to anchor the stereocilium. Thus deflections of the bundle cause stereocilia to pivot at their insertions in the cuticular plate, but not bend. The hair bundle also includes a single kinocilium in all hair cells but those of avian and mammalian cochleas. In those organs, the kinocilium grows out normally when the bundle grows or regenerates, but is lost in the mature hair cell. The placement of the kinocilium at the tall edge of the bundle, and its attachment to the tallest stereocilia (Jacobs and Hudspeth, 1990), enable it to transmit mechanical force from an overlying accessory structure such as the otolithic membrane or cupula to the stereocilia. In contrast to stereocilia, kinocilia have the internal structure of motile cilia, containing a 9+2 arrangement of microtubules. Both stereocilia and kinocilia are enclosed by the cell membrane, like fingers in a glove, so that the actin or microtubule cores are intracellular and in electrical continuity with the cell body.

TRANSDUCTION AND ADAPTATION BY VERTEBRATE HAIR CELLS

3

Figure 1–1. A, Three hair cells in the sensory macula of the bullfrog saccule. The bundle has a morphological axis of polarity defined by the graded heights of the stereocilia (seen on the middle bundle) and the eccentric placement of the kinocilium (seen on the left bundle), such that the tallest stereocilia are at the positive edge. These bundles are about 8 m tall, and contain 50-60 stereocilia. B, A tip link (arrow) extending between two adjacent stereocilia. At each end are electrondense attachment plaques between the membrane and the actin cores. The tip link is 150-200 nm in length; stereocilia are ~400 nm in diameter.

Adjacent stereocilia are connected by four sets of extracellular links: basal connectors near the taper region, shaft connectors often extending the length of stereocilia, horizontal tip connectors that seem to hold the tips of stereocilia together while allowing slipping, and tip links (Pickles, Comis, and Osborne, 1984; Goodyear and Richardson, 1992). While the first three links extend laterally and symmetrically between stereocilia, the tip links extend obliquely upwards from the tip of each stereocilium to the side of the adjacent taller stereocilium, along the axis from short to tall (Figure 1–1B). They are about ~10 nm in thickness and ~150 nm long in nearly all vertebrate hair cells, and at each end terminate at electron-dense plaques situated between the membrane and the outermost actin filaments (Pickles et al., 1984, Assad, Shepherd, and Corey, 1991; Jacobs and Hudspeth, 1990).

Physiology Deflection of the hair bundle towards the tallest stereocilia (a positive stimulus) causes a depolarizing receptor potential of up to ~30 mV (Hudspeth and Corey, 1977; Crawford and Fettiplace, 1978). The depolarization

4


is associated with a drop in resistance of the cell membrane, indicating that positive deflection opens ion channels. If the hair cell is voltage-clamped, opening of these “transduction” channels is manifest as an increase in inward current, at negative holding potentials, of several hundred picoamperes amplitude (Figure 1–2A) (Corey and Hudspeth, 1979a; Assad, Hacohen, and Corey, 1989). In bullfrog hair cells, positive deflection of ~0.4 (m (around the diameter of one stereocilium) is sufficient to open all the channels, while a negative deflection of ~0.1 m closes the fraction of channels that are open at the resting bundle position. The I(X) curve generated by plotting receptor current against deflection (Figure 1–2B), shows the range over which deflection can change channel open probability, and shows the resting current of 10–20%. Most small cations will pass through the channel, including Na+, K+, Li+, Cs+, tetramethylammonium, and even Tris, but the channel is particularly permeable to Ca2+ (Corey and Hudspeth, 1979a; Lumpkin, Marquis, and Hudspeth, 1997; Ricci and Fettiplace, 1998). Because the bundles are normally bathed in a high-K+ and low-Ca2+ endolymph, K+ carries the bulk of the receptor current in vivo. However, even when Ca2+ is less than 0.1% of the cation composition, it carries as much as 10% of the current, and plays an important role in regulation and feedback for the transduction apparatus (Ricci and Fettiplace, 1998). The single-channel conductance of the transduction channels is about 100 pS in normal saline, so that each open channel carries a current of 6–7 pA at the resting potential (Crawford, Evans, and Fettiplace, 1991). In tur-

Figure 1–2. A, Receptor current (lower panel) evoked by movement of the hair bundle (upper panel) in a dissociated bullfrog hair cell. Positive deflection of the bundle increases a cation conductance and allows inward current of up to several hundred picoamperes in these cells. B, The I(X) curve generated by plotting current against the deflection of the tip of the hair bundle. Positive deflections of ~0.5 m open all channels; negative deflections of ~0.1 m close those channels open at the resting position. Modified from Assad et al., 1989.


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tle cochlea, healthy cells can have well over 1000 pA of receptor current, indicating that each hair cell may have >200 functional channels, or >4 channels per stereocilium (Wu, Ricci, and Fettiplace, 1999). Transduction channels can be blocked by a variety of larger cations, and the block is usually voltage-dependent, indicating that these compounds bind within the pore of the channel. The best known of the blocking drugs are aminoglycoside antibiotics such as streptomycin, which block with a Ki of ~20 µM at the resting potential (Kroese, Das, and Hudspeth, 1989). Perhaps surprisingly, curare will also block the channel at a concentration of 2–3 µM (Glowatzki, Ruppersberg, Zenner, and Rusch, 1997). Amiloride and its analogs such as benzamil also reduce the receptor current, with Ki near 5 M, but the voltage dependence and Hill coefficient suggest a mechanism different from pore block (Rusch, Kros, and Richardson, 1994). Transduction channels are located at the tips of stereocilia, as shown by mapping the location of current sinks around the bundle (Hudspeth, 1982). Although surprising and perhaps controversial (Ohmori, 1988), the result has been confirmed by iontophoretic application of a channel blocker and by calcium imaging of individual stereocilia (Jaramillo and Hudspeth, 1991; Denk, Holt, Shepard, and Corey, 1995; Lumpkin and Hudspeth, 1995). Thus, current entering the stereocilia must pass down their lengths to depolarize the cell body, a process which takes just microseconds in stereocilia such as those in Figure 1–1A. Following deflection of the hair bundle, transduction channels open in ~40 µs at room temperature, and ~10 µs at mammalian temperatures (Corey and Hudspeth, 1979b; 1983a). Larger positive deflections cause more rapid opening than smaller deflections. These argue against activation of channels by a diffusible second-messenger, and suggest instead that the deflection directs a force directly onto a mechanically-sensitive transduction channel protein. If deflection were to move one end of an elastic “gating spring”, the other end of which pulls on the channel, then a simple biophysical model can describe the kinetics and force sensitivity (Corey and Hudspeth, 1983a). Support for this model has come from very fine measurements of the movement of hair bundles when force is applied to them. Howard and Hudspeth (1988) found that application of force deflected the bundle by an amount that could be attributed to the stiffnesses of the stereocilia and the gating springs acting in parallel. However, if the deflection moved the bundle to a range where channels opened, then the bundle moved a bit more. The extra movement could be attributed to the slight relaxation in the gating springs that occurred as channels opened, and was calculated to correspond to a movement of the channel protein upon opening of ~4 nm. The extra movement was blocked by drugs that block the pore, suggesting the channel’s gate can’t close if a blocker is in the pore. Hair cells respond only to deflections directed along their morphological axis, from the short to the tall stereocilia, while deflections perpendicu-

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lar to this axis cause little if any response (Shotwell, Jacobs, and Hudspeth, 1981). Thus, a transduction apparatus must be at the tips of the stereocilia, must involve some sort of direct linkage to the channel, and must be anisotropic—having some characteristic appropriate to sense directional stimuli.

Transduction Model The discovery of tip links provided both a structure and a model for transduction (Figure 1–3) (Pickles et al., 1984). They are oriented only along the sensitive axis, and not from side to side. Moreover, their oblique orientation, unique among interstereociliary links, confers a directional sensitivity: deflection of the bundle in the positive direction will stretch the links, and deflection in the negative direction will relax them. The “tip links model” supposes that these links pull directly on the transduction channels to open them, and that the tip links either are the gating springs themselves or are mechanically in series with other elastic elements. Two experiments support the importance of the tip links. Assad et al. (1991) found that reduction of the extracellular Ca2+ concentration to well below 1 µM abolished the mechanical sensitivity within seconds, in hair cells from the bullfrog. Even when the normal Ca2+ was restored, the me-

Figure 1–3. The tip-links model for transduction. A, Stereocilia pivot at their bases and remain touching at their tips, so that positive deflection of the bundle would stretch the tip links. B, The tip link is imagined to be directly connected to ion channels at each end. Stretch of the tip link (or of a different elastic element in series with it) would pull on the channels to open them.


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chanical sensitivity did not return during the duration of the experiment (many minutes). They also found that a few seconds of low Ca2+ eliminated the tip links observed with electron microscopy, and restoration of normal Ca2+ did not restore the links. A similar correlation of tip links with mechanical sensitivity was observed in bird hair cells (Zhao, Yamoah, and Gillespie, 1996), but in this case the hair cell epithelia were kept in culture for many hours after cutting tip links with low Ca2+, to allow recovery. Both tip links and mechanical sensitivity returned over 7-10 hours, indicating a dynamic regulation and repair of the transduction apparatus.

Extensions of the Model Although the tip-links model seems fundamentally sound, several refinements have been made in recent years. First, the model gives no way to determine whether the transduction channels are at the upper or lower ends of the tip links; either would work, as current through the channels could flow down stereocilia to depolarize the hair cell. With calcium imaging of individual stereocilia of a bundle, Denk et al. (1995) were able to suggest an answer. They loaded hair cells with a calcium indicator dye, and reasoned that any stereocilium that has a functional transduction channel would increase its fluorescence when calcium entered through the open channel. If the channels are only at the lower end, then the tallest stereocilia could never show a fluorescence increase upon bundle stimulation; if channels are at the upper end, then the shortest would remain dim. They found that all stereocilia could increase their fluorescence upon stimulation, indicating that transduction channels can be at both lower and upper ends of tip links, and probably at both ends. Moreover, the notion that channels are at both ends requires a revision of Howard and Hudspeth’s (1988) estimate of a 4-nm movement associated with channel opening. The earlier data could be refitted by a model with two channels each moving ~2.2 nm (Denk et al., 1995). Second, the largest transduction currents seen in bullfrog hair cells (~400 pA) could be accounted for by about 50 channels on about 25 intact tip links. Indeed, the calcium imaging showed many stereocilia that remained dim during stimulation, as if a fraction of the 50 or so possible tip links were missing (Denk et al., 1995). On the other hand, more recent recordings of much larger transduction currents from turtle cochlear hair cells can only be accounted for by supposing 5 or more channels per tip link (Ricci and Fettiplace, 1998), a number clearly incompatible with the idea of one at each end. Third, a branching of the tip link at each end, long observed in transmission electron micrographs, has been documented much more clearly by quick-freeze, deep-etch electron micrographs of bullfrog hair cells (Kachar, 1999). The tip link appears to be a helical filament with at least

8


two strands, two or three distinct branches appear to connect the upper end of the tip link with the taller stereocilium’s membrane, and three or more finer strands appear where the lower end contacts the tip of the shorter stereocilium. Perhaps then, there is a channel at each of the fine strands at each end of the tip link, amounting to 6 or 8 channels per tip link. Then 3 or 4 channels at one end would be mechanically in parallel, and together these would be mechanically in series with the 3 or 4 at the other end. Such an arrangement makes models of channel gating quite complex, but, then, evolution did not create hair cells with concern for the biophysicist.

Ca2+ Metabolism in Stereocilia Patch-clamp experiments show that calcium carries a significant fraction of the receptor current in hair cells, and indicator dyes show that calcium concentration rises inside stereocilia when channels open. As we shall see, the intracellular calcium concentration near the transduction channels has a pronounced feedback effect on the open probability of the channels, so calcium metabolism within the stereocilia is critical in regulation of transduction. Thus, we need to know how high the concentration gets near the channel, and where the calcium goes. Two groups have used calcium imaging together with diffusion models to determine the calcium concentration within stereocilia (Ricci and Fettiplace, 1998; Ricci, Wu, and Fettiplace, 1998; Lumpkin and Hudspeth, 1998). These indicate that Ca2+ entering through channels can be sequestered and removed by four mechanisms: simple diffusion down the stereocilia, buffered diffusion of Ca2+ bound to a mobile endogenous buffer, binding of Ca2+ to fixed endogenous buffer, and extrusion of Ca2+ by a calcium pump in the stereociliary membrane. Modeling of responses to bundle deflections indicates that all four mechanisms must be active to account for the fluorescence transients observed (Lumpkin and Hudspeth, 1998). Peak calcium levels depend both on the extracellular Ca2+ concentration and on buffers such as EGTA or BAPTA added to the patch recording solution. Under physiological conditions, the Ca2+ concentration at the tips of the stereocilia can reach a few tenths of a micromolar, but is much lower further from the tips. These values, however, are average concentrations across the whole diameter of a stereocilium; very close to transduction channels the concentration can be much higher. For instance, 10 nm from the channel, where Ca2+ may be acting to regulate transduction, the concentration may reach a steady-state value of 5 to 50 µM within a few tens of microseconds following channel opening (Ricci et al., 1998; Lumpkin and Hudspeth, 1998). Calcium buffers in the hair cell body have been studied, and appear to be at a concentration of several millimolar. The buffer may be a protein


9

such as calbindin or calretinin (Roberts, 1993; 1994). It is not clear whether this same buffer is in the stereocilia, but the effective endogenous mobile buffer concentration is equivalent to 0.1-0.4 mM BAPTA (Ricci et al., 1998). Calcium must also be extruded from the cytoplasm. Since there is little Na+ in endolymph to drive a Na+/Ca2+ exchanger, attention has been focused on Ca2+ ATPases. Antibodies to plasma membrane calcium ATPases (PMCA) label hair cells, especially the stereocilia, and especially the tips of stereocilia (Crouch and Schulte, 1995; Apicella et al., 1997; Yamoah et al., 1998). Quantitative immunoblots of purified hair bundles indicated that each stereocilium may contain as many as 20,000 molecules of PMCA, or 2,000 molecules per µm2 (Yamoah et al., 1998). During a maintained deflection, PMCA pumps Ca2+ from the stereocilia, creating an outward current of several picoamperes. Dialysis of hair cells with PMCA inhibitors blocks this current, and also leads to a pronounced rise in Ca2+ concentration within stereocilia (Yamoah et al., 1998). There are at least four different isoforms of PMCA, but the evidence is that PMCA2 is the isoform in stereocilia (Furuta et al., 1998). Indeed, two alleles of the mouse mutant deafwaddler, which is deaf and shows vestibular abnormalities, have either a point mutation or a frameshift in their PMCA2 genes (Street, McKee-Johnson, Fonseca, Temple, and Noben-Trauth, 1998). Similarly, a mouse with a targeted deletion of PMCA2 is deaf, has balance problems, and has structural defects in the organ of Corti (Kozel et al., 1998).

ADAPTATION Most sensory systems adapt, decreasing their response even though a stimulus is maintained (Adrian, 1928). Auditory nerve fibers decrease their firing rate during a sustained tone (Smith, Brachman, and Goodman, 1983), and vestibular nerve fibers decrease their firing rate during a sustained acceleration (Goldberg and Fernandez, 1971; Eatock, Corey, and Hudspeth, 1987). While adaptation may occur at a variety of levels between a stimulus to the hair cell and the firing of a nerve, an important— perhaps the dominant—part of adaptation occurs in the transduction apparatus (Eatock et al., 1987; Eatock, 2000). A sustained positive deflection of the hair bundle causes an initial increase in receptor current (channels open), followed by a decline in current and conductance (channels close) (Figure 1–4A,B). Expressed in terms of the I(X) curve, the decline in current could occur in three ways: by a broadening of the curve, so that a larger range of deflection is needed to open or close all the channels; by a vertical compression of the curve, as if inactivation reduces the number of channels available to be opened; or by a shift of the curve along the deflection axis,

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Figure 1–4. Models for adaptation of the transduction apparatus in hair cells. A-C, Physiology of adaptation. The open probability (Popen) of channels is increased by a step deflection of the bundle, but then declines with a time course of 1-30 ms (B). When the bundle is brought back to the rest position, Popen goes to zero, and then recovers to ~15% over the next 10-50 ms. Adaptation can also be seen as the Popen(X) curve moving along the deflection axis, so that the same deflection causes a progressively smaller Popen (C). D-E, Two models for adaptation. D, The slipping/climbing model. Channels (and other associated proteins) would be tethered to the actin cores by a few dozen motor proteins (perhaps myosin-I), which are always trying to climb up the actin. Increased tip-link tension caused by bundle deflection would pull the channels down the side of the taller stereocilium, so as to relax the tension on them. If the bundle is deflected negatively to relax the tip links, then the motors would be able to climb and restore resting tension. E, The calcium closure model. When a channel opens, calcium passing through it would bind to a site on or near the channel, shifting the relation between tension and open probability so the channel tends to close. If channels close, the local calcium concentration rapidly dissipates and shifts the relation back so channels tend to open. The models are not mutually exclusive.

as if the cell is resetting the stimulus needed to open a certain proportion of channels. In fact, adaptation is primarily associated with a shift of the I(X) curve, whereby the hair cell moves the curve towards the sustained bundle position, bringing the channel open probability back towards the resting value (Figure 1– 4C) (Corey and Hudspeth, 1983b; Eatock et al., 1987).


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Adaptation is not complete: the full shift of the I(X) curve is not as large as the sustained bundle deflection, so the receptor current does not return completely to the resting level. In bullfrog hair cells with 4 mM Ca2+ bathing the bundles, the extent of adaptation is about 80%, that is, adaptation shifts the I(X) curve by about 80% of the deflection (Shepherd and Corey, 1994). At lower Ca2+ concentrations, it can be less complete (Ricci and Fettiplace, 1997). Thus the hair cell responds mostly to the change in bundle position, but there is a residual response to the absolute bundle position. Calcium inside the tips of the stereocilia speeds adaptation considerably. The Ca2+ concentration inside the tips can be affected in three ways, and each affects adaptation: Lowering the extracellular Ca2+ decreases Ca2+ flux through the transduction channels, and slows the rate of adaptation, measured either as the rate of shift of the I(X) curve or the change in receptor current (Corey and Hudspeth, 1983b; Eatock et al., 1987; Hacohen, Assad, Smith, and Corey, 1989; Assad et al., 1989; Crawford, Evans, and Fettiplace, 1989). Raising the calcium buffer concentration within the hair cell causes Ca2+ to be more rapidly bound once it enters, and slows the rate of adaptation (Crawford et al., 1989; Ricci and Fettiplace, 1997). Depolarizing the hair cell reduces the driving force for Ca2+ influx, reduces Ca2+ entry, and reduces or abolishes adaptation (Assad et al., 1989; Denk et al., 1995). Ca2+ appears to affect the rate of adaptation by acting at a site very near the transduction channel. If a hair cell is depolarized to block Ca2+ entry and eliminate adaptation, then repolarized to allow Ca2+ entry while the bundle is still deflected, adaptation starts again within 1-2 ms of the repolarization. Since Ca2+ could only diffuse about 1 µm in that time, it must find the adaptation control site very near the transduction channel in the tip of the stereocilium (Assad et al., 1989). In addition, slow calcium buffers like EGTA are not effective in slowing adaptation, whereas fast buffers like BAPTA are, suggesting that the buffer must capture Ca2+ before it diffuses very far, if it is to prevent Ca2+ from reaching the adaptation site (Ricci et al., 1998). Initial studies characterized adaptation as a single exponential process with a time constant of 10-30 ms (Eatock et al., 1987; Assad et al., 1989). More recently, it has become clear that adaptation may have two different components, with one as fast as 0.3 ms (Ricci et al., 1998). The fast process tends to dominate with small deflections, but saturates with larger deflections, leaving the slower process. Moreover, some pharmacological manipulations have shown differential effects on the time course of adaptation and on the resting position of the I(X) curve, which adaptation is supposed to regulate (Ricci and Fettiplace, 1997). Two fundamentally different models for adaptation have been proposed, one involving a mechanical adjustment of tension on the gating spring (Howard and Hudspeth, 1987), and one involving Ca2+ binding di-

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rectly to a site at or near the channel to change the relation between tension and open probability (Figure 1–4 D,E) (Crawford et al., 1989; Wu et al., 1999). It is interesting to consider the evidence for each of these, and to ask whether both could be operating simultaneously in the hair cell.

The Active Motor Model Howard and Hudspeth (1987) used a flexible glass stimulus probe to measure the mechanical correlates of adaptation in bullfrog saccular hair cells. They found that a force applied to the bundle caused a deflection of a certain amount, followed by an additional, slower deflection with the same time course as adaptation (~30 ms). The slow deflection occurred for forces directed in either the positive or negative direction along a cell’s physiological axis, but not for forces directed perpendicular to the axis. When adaptation of the receptor current slowed or disappeared during prolonged recording, the slow deflection did as well. Because adaptation was associated with a mechanical relaxation, Howard and Hudspeth (1987) speculated that adaptation might come about by a movement of the upper tip-link attachment point along the side of the stereocilium. In this model, an active motor complex in each stereocilium is continuously trying to “climb” the stereocilium, pulling up the attachment point to increase tension in the tip link. When a positive deflection tightens the tip link (and opens channels) the attachment slips down to relax the tension and allow channels to close. When a negative deflection loosens the tip link (and closes channels), the motor can climb to restore tension. At steady state, the motor slips as fast as it climbs, and the steady state tension is just sufficient to open 10-20% of the channels. The shift of the I(X) curve can then be seen as a shift of the upper tip-link attachment point, adjusted for the geometrical gain of the bundle. Thus, a positive deflection of 200 nm would stretch the gating spring by about 24 nm (for bundle dimensions in bullfrog saccule), and a subsequent slippage of 20 nm would allow most of the opened channels to reclose. Howard and Hudspeth went on to suggest that myosin would be an attractive mechanoenzyme to power the motor, as it moved with the right speed for adaptation and in the right direction relative to the actin polarity in stereocilia. This short, provocative model really comprises three different ideas. First, it supposes that adaptation is fundamentally a mechanical adjustment process, moving one end of the physiologically-defined gating spring to change tension. Second, by equating the gating spring with the tip link, it makes the morphological prediction that the electron-dense attachment of the tip link moves during adaptation. Third, it suggests a protein of the myosin family as the molecular basis for adaptation. All three ideas have been tested in the last decade. The mechanical basis of adaptation was studied by systematically measuring adaptation rates for positive and negative deflections, and de-


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veloping a mechanical model for adaptation based on an active, force-producing motor and on the previously measured stiffness of hair bundles (Assad and Corey, 1992). Reducing calcium entry (by depolarizing the hair cell) reduced both climbing and slipping rates, but reduced slipping more. The model predicted that depolarization would therefore increase tension on the channels, increasing resting open probability to ~80% and shifting the I(X) curve by ~120 nm; this was observed experimentally. Because of the upward slant of tip links, an increase in tip-link tension should pull an unrestrained bundle in the negative direction. The predicted position change with depolarization was about 100 nm, which could be measured with high-resolution video microscopy (Assad and Corey, 1992). Finally, the bundle movement caused by depolarization was abolished when tip links were cut with BAPTA, suggesting that tip links convey the tension for bundle movement (Assad et al., 1991). Thus a quantitative model based on adaptation rates accurately predicted movement of the bundle, supporting a mechanical basis for adaptation. Morphological correlates of adaptation have been more difficult to obtain. Video microscopy with a resolution of 30-40 nm showed no gross movement of the stereocilia or cuticular plate during adaptation, shifting attention to smaller-scale molecular rearrangements (Shepherd, Assad, Parakkal, Kachar, and Corey, 1991). Thus far, it has not been possible to see the predicted changes in attachment plaque position, which are just tens of nanometers, above the natural variability in plaque position. On the other hand, cutting tip links with BAPTA should relieve tension and allow motors to climb even in an undeflected bundle. Indeed, BAPTA treatment was followed by an upward movement of the attachments of 50-70 nm, as measured from transmission electron micrographs. (Shepherd et al., 1991). There is broad, but as yet circumstantial, support for myosin’s involvement in adaptation. First, it has been shown that the actin cores of stereocilia can support myosin motility. Beads coated with chicken muscle myosin move along demembranated bullfrog stereocilia, always moving towards the tips at 1-2 m/s (Shepherd et al., 1990). Second, blockers of the ATPase cycle such as ADPS, which should arrest myosin while strongly bound to actin, block adaptation when dialyzed into the hair-cell cytoplasm (Gillespie and Hudspeth, 1993). Similarly, phosphate analogs that arrest myosin while weakly bound should inhibit myosin force production, and apparently cause release of resting tension on channels (Yamoah and Gillespie, 1996). The myosin gene superfamily contains dozens of members, organized into fifteen classes. At least eight of these classes occur in vertebrates. A search for hair-cell myosins has followed two paths. Gillespie, Wagner, and Hudspeth (1993) used vanadate trapping of adenine nucleotides to identify three putative myosins within stereocilia, of molecular weights 120, 160, and 230 kDa. The 120 kDa band was labeled with an antibody to

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a myosin type I, and this antibody particularly labeled the tips of stereocilia. Solc, Derfler, Duyk, and Corey (1994) used degenerate PCR to amplify fragments of most myosins expressed in the hair-cell epithelium, and found ten different myosins from six different classes (I, II, V, VI, VII and X). Interestingly, two of these myosins (VI and VIIa) cause inherited deafness in mice and/or humans when mutated (Avraham et al., 1995; Gibson et al., 1995). The myosin-I was cloned in full, and an antibody raised against the tail domain (Solc et al., 1994). An extensive antibody study found that myosins-I, -VI and –VIIa are all expressed by the hair cells, and all are in the stereocilia as well as elsewhere in the cells (Hasson et al., 1997). These have molecular weights of approximately 120, 160, and 230 kDa, respectively. Recently, a myosin-XV has also been discovered in stereocilia (Liang et al., 1999). Of all these myosins, however, only myosin-I is concentrated in the tips of stereocilia (Figure 1–5A,B) (Hasson et al., 1997). Because localization with light microscopy cannot determine the relation of a myosin to the tip-link attachments, and because myosins might naturally climb to the tips of stereocilia unless otherwise prevented, the location of myosin-I was also determined with immunogold electron microscopy. This showed that myosin-I immunoreactivity was indeed associated with both end of the tip links, where it may link the channels to the actin cores (Figure 1–5C) (Garcia, Yee, Gillespie, and Corey, 1998). Like most myosins, myosin-I has binding sites for regulatory light chains such as calmodulin. Three calmodulin molecules bind to myosin-I, and confer a calcium dependence to the myosin activity (Zhu, Sata, and Ikebe, 1996; Burlacu, Tap, Lumpkin, and Hudspeth, 1997). Calmodulin is in stereocilia, especially concentrated at the tips (Shepherd et al., 1989; Walker et al., 1993), and antagonists of calmodulin block adaptation (Corey, Smith, Barres, and Koroshetz, 1987; Walker and Hudspeth, 1996). It may be that calmodulin mediates the Ca2+ sensitivity of adaptation (Walker and Hudspeth, 1996; Gillespie and Corey, 1997). While all these studies are consistent with a mechanoenzyme such as myosin mediating adaptation, none specifically proves the involvement of myosin-I. We now need physiological inhibitors or activators based on the specific molecular sequence of myosin-I, as a definitive test of this motor. A mutant form of myosin-I (Gillespie, Gillespie, Mercer, Shah, and Shokat, 1999), which is more sensitive than the wild-type to a certain inhibitor, could provide such a test.

Ca2+-Dependent Closure Model As mentioned, the rate of adaptation depends on the Ca2+ concentration inside the tips of stereocilia. With high extracellular Ca2+, and low or slow internal calcium buffer, adaptation can become quite fast, with a time constant as short as 0.3 ms (Ricci and Fettiplace, 1997). This is probably too


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Figure 1–5. Myosin-I immunoreactivity in bullfrog hair cells. A-B, Immunofluorescence of single dissociated cells shows myosin-I in the cell body, and also in the hair bundle where it is especially concentrated at the tips of stereocilia. The kinocilium also shows fluorescent label. Scale bar = 2 µm. C, Summary of myosinI distribution in the tips as seen with immunogold electron microscopy. Gold particles marking the antibody were counted and averaged over many stereocilia in six bundles, and their density is indicated as particles/m2 of surface membrane. The greatest density is within ~200 nm of either end of the tip link. Reprinted from Garcia et al. (1998), Figures 1 and 7.

fast to be mediated by a tension adjustment system that requires an ATPase cycle of myosin. For instance, the calculated 1-2 m/s climbing rate of the adaptation motor and a myosin step size of 8 nm suggest an ATPase cycle time of 4 to 8 ms. A fast time constant is still compatible with a myosin motor if Ca2+ rapidly causes the myosin to release tension, for instance, if Ca2+ reversibly causes the lever domain of myosin to become more compliant (Gillespie and Corey, 1997). However, such a mechanism could account

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only for a limited shift of the I(X) curve—perhaps 0.1 m given the myosin lever dimensions and the bundle geometry—and fast adaptation can cause more shift than that. As an alternative, Howard and Hudspeth (1988) and Crawford et al. (1989, 1991) proposed that Ca2+ binding directly to an intracellular site on the channel could shift the energetics of channel opening, such that the channel with calcium bound required more force to reach the same open probability; that is, calcium would tend to close the channel. The original model, in which the energy of one of the closed states is affected, cannot explain adaptation to large deflections, but a later model, in which calcium changes the “set point” of the channel, accounts for adaptation of up to 0.8 m (Wu et al., 1999). These two models—one in which Ca2+ relaxes tension and allows channel closure, and the other in which Ca2+ causes the channel to shut even with the same tension—have quite different predictions for the mechanical behavior of a hair bundle during adaptation. In the motor model, a force that opens channels would quickly move the bundle a certain amount determined by the force and the stiffness of the bundle. Bundle stiffness is determined by two springs in parallel: the stereocilia pivot stiffness and the gating spring stiffness. If myosin motors slip to allow channel closure, the gating spring relaxes, which could be seen as a decrease in stiffness (chord stiffness but not slope stiffness). The bundle would move forward with the time course of adaptation. On the other hand, if adaptation happens because Ca2+ causes the channel to shut, the small movement of the channel’s gate slamming shut would tighten the gating spring (e.g., Figure 1–3), in essence increasing the stiffness and pulling the whole bundle backward. Which behavior is seen with adaptation? In fact, both positive and negative going movements have been observed, with different time courses (Howard and Hudspeth, 1987; Jaramillo and Hudspeth, 1993; Benser, Marquis, and Hudspeth, 1996). Figure 1–6 shows a rapid forward movement (arrowhead), followed by a quick hook back—the “twitch”—and then a slower and larger forward relaxation. Both the twitch and the slower relaxation depend on Ca2+, becoming slower and smaller when Ca2+ is reduced. The termination of force allows the bundle to move back, but no twitch is seen on the return, at least following large deflections. On the other hand, a negative force evokes a slow negative relaxation of the bundle, and then a twitch is seen when termination of force allows the bundle to move forward towards the rest position These two mechanical behaviors, which are separable by time course and stimulus polarity, suggest that both proposed mechanisms of adaptation may be occurring in hair cells. The calcium closure might be fast (1-5 ms), and cause the quick twitch; a myosin slipping would be slower (10-50


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Figure 1–6. The “twitch”, and subsequent slow relaxations of the hair bundle during maintained force steps. A, The positive force stimulus in this experiment caused a quick forward deflection of ~50 nm (arrowhead), a rapid twitch back of ~10 nm, and then a relaxation forward of ~25 nm. The receptor potential recorded simultaneously (middle trace) showed a transient depolarization, and then an oscillation during and following the twitch. B, A negative force evoked a quick negative deflection, and slower negative relaxation. The twitch was not observed until the bundle was returned to the rest position (arrowhead), and was associated with an oscillation. Reprinted from Benzer et al. (1996), Figure 2.

ms) and may cause the slow relaxation. A careful analysis of the time course of adaptation in turtle hair cells has found that two time constants are needed to fit the transduction current. Moreover, they depend differently on external Ca2+ and internal buffer concentration, further indicating their separability (Wu et al., 1999). Comparison with a model of Ca2+ diffusion in stereocilia suggests that the fast phase is controlled at a Ca2+ binding site that is 20-50 nm from the site of Ca2+ entry, and may be the channel itself (Wu et al., 1999). The slow phase is controlled at a more distant site, 150 to 200 nm from the site of entry, which corresponds well with the location of myosin-I. The authors were careful not to speculate about molecular mechanisms of the two components, but it will be interesting to find which parts of the transduction complex correspond to each phase. Needed also is a careful temporal correlation of the two phases of transduction current with the two phases of mechanical response.

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Frequency Tuning Ringing in a system can be caused by negative feedback with a delay. If the feedback has a gain greater than one, the system can go into spontaneous oscillation, at a frequency determined in part by the delay. The calcium closure mechanism could provide such a feedback (Choe, Magnasco, and Hudspeth, 1998; Wu et al., 1999), which can be seen by supposing a sinusoidal force stimulus to the bundle. The forward phase of force would cause the bundle to move forward, opening channels. Ca2+ entry and the subsequent closure of channels would pull back on the gating springs, pulling the bundle back with a slight delay. If the delay was such that the channel closure was coincident with the negative-going phase of the force stimulus, then the negative bundle movement would be amplified by the pullback. During the next few moments, channels would close, Ca2+ would unbind and diffuse away from the binding site, and then the next positive-going phase of the stimulus could more easily open channels and move the bundle forward. If the stimulus were sinusoidal, the movement, and thus the stimulus to the channels, would be amplified by feedback like this; if the stimulus were a step, the movement would be characterized by ringing following the initial deflection; and if there were no stimulus, but the feedback gain was large enough, the bundle could undergo a continuous oscillation in position. Ringing in bundle position was first observed in turtle hair cells, when recording the receptor potential with a single microelectrode (Crawford and Fettiplace, 1985). Similarly, spontaneous twitches, oscillatory bursts, and continuous oscillation were seen in the movement of unstimulated bullfrog hair bundles (Howard and Hudspeth, 1988; Benser et al., 1996; Martin and Hudspeth, 1999). Turtle hair cells have, in addition, a well-characterized oscillation of membrane potential that is distinct from the transduction and is generated by Ca2+ and K+ channels in the basolateral membrane, so it was not clear whether the bundle oscillation was generated within the transduction apparatus or was some sort of voltage feedback on bundle position. However, oscillations in transducer current have been observed when the hair cell was voltage-clamped and voltage-gated conductances could not influence the transient current (Figure 1–7) (Ricci et al., 1998). Moreover, ringing in transduction current was observed with a stiff stimulus probe that would not allow mechanical feedback on bundle position (Figure 1–7), so that some part of the ringing might be generated simply by delays in unbinding and binding of Ca2+ to the transduction channel. Two models have been developed that describe the ringing based on calcium closure of the transduction channel. Choe et al. (1998) suppose two binding sites on the channel for Ca2+, which steepens the relation between Ca2+ concentration and open probability. In this model, the bundle is allowed to move, and within a realistic range of parameters, can show spontaneous oscillations. Close to that range, it can amplify oscillatory


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Figure 1–7. Damped oscillations in transducer currents from turtle cochlear hair cells. Bundles were deflected with a stiff stimulus probe by ±50 nm (top panel). Positive deflections evoked inward receptor currents, which oscillated during the maintained deflection. Negative deflections stopped the inward receptor current, but there was sometimes an oscillation when the bundle was brought back to the rest position. The frequency of oscillation varied among different cells, and with different amounts of calcium buffer in the cytoplasm; two examples are shown. Reprinted from Ricci et al. (1998) figure 9

stimuli, and shows the most amplification for the smallest stimuli. The characteristic frequency can range from 20 Hz to 20 kHz. A similar model was developed by Wu et al. (1999) to describe the two phases of adaptation, but this model also shows ringing responses to step deflections of the

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bundle. Although the model incorporates two mechanisms for adaptation at different distances from the channel, the ringing seems to depend on the closer, fast mechanism. Perhaps because the channel closure has first order rather than second order dependence on Ca2+, the model does not show spontaneous oscillation. The interesting point is that two models that are similar only in basic outline both predict ringing responses to small stimuli, suggesting that this mechanism would work to amplify within a fairly broad range of parameters. At least one of the models can produce ringing at frequencies of tens of kilohertz, so that it might work for the highest frequencies of mammalian hearing. On the other hand, the oscillations that have been actually observed in lower vertebrates are much slower. Ringing in turtle occurs at several hundred Hertz (Ricci et al., 1998), and frog bundles oscillate at tens of Hertz (Martin and Hudspeth, 1999). It will be important to see if such behavior occurs in animals that hear at much higher frequencies. It is apparent that a hair bundle spontaneously oscillating in the absence of mechanical stimulus is putting energy into a mechanical system, and that such energy could be used to amplify vibrations at the characteristic frequency (Crawford and Fettiplace, 1985). This result has recently been explored in more depth by estimating the viscous drag of the bundle and a stimulus probe, to calculate the work done in each cycle of the oscillation (Martin and Hudspeth, 1999). Where does the energy come from? If driven by the Ca2+-closure mechanism (which is still hypothetical), the energy comes from the Ca2+ gradient across the stereocilium membrane, which is in turn established by the Ca2+ ATPase in the membrane. However, the energy in this model comes not from Ca2+ passing through the channel and down the standing gradient, but from Ca2+ being driven onto the binding site when it is at high concentration near an open channel, and then unbinding when the channel closes and the concentration is low. That is, the gradient is really temporal rather than spatial (Choe et al., 1998). Fast intracellular buffers reduce the temporal differences, and tend to eliminate the oscillations (Ricci et al., 1998). Regardless of the mechanism, it seems likely that the fast component of adaptation is associated with a negative feedback force on the bundle, and that this may lead to mechanical ringing of the bundle. Because the bundle puts energy into the ringing, it can amplify the vibration of a sinusoidal stimulus near the characteristic frequency, and nearby hair cells of the cochlea might act together to amplify the vibration of the basilar membrane. Perhaps the cochlear amplifier is in the transduction channel itself.

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Eatock, R. A. (2000). Adaptation in hair cells. Annual Review of Neuroscience 23, 285–314. Eatock, R. A., Corey, D. P., & Hudspeth, A. J. (1987). Adaptation of mechanoelectrical transduction in hair cells of the bullfrog’s sacculus. Journal of Neuroscience 7, 2821–2836. Flock, A., Cheung, H. C., Flock, B., & Utter, G. (1981). Three sets of actin filaments in sensory cells of the inner ear. Identification and functional orientation determined by gel electrophoresis, immunofluorescence, and electron microscopy. Journal of Neurocytology 10, 133–147. Furuta, H., Luo, L., Hepler, K., & Ryan, A. F. (1998). Evidence for differential regulation of calcium by outer versus inner hair cells: Plasma membrane Ca-ATPase gene expression. Hearing Research 123, 10–26. Garcia, J. A., Yee, A. G., Gillespie, P. G., Corey, D. P. (1998). Localization of myosinIbeta near both ends of tip links in frog saccular hair cells. Journal of Neuroscience 18, 8637–8647. Gibson, F., Walsh, J., Mburu, P., Varela, A., Brown, K. A., Antonio, M., Beisel, K. W., Steel, K. P., & Brown, S. D. M. (1995). A type VII myosin encoded by the mouse deafness gene Shaker-1. Nature 374, 62–64. Gillespie, P. G., & Corey, D. P. (1997). Myosin and adaptation by hair cells. Neuron 19, 955–958. Gillespie, P. G., Gillespie, S. K., Mercer, J. A., Shah, K., & Shokat, K. M. (1999). Engineering of the myosin-I nucleotide-binding pocket to create selective sensitivity to n(6)-modified ADP analogs. Journal of Biological Chemistry 274, 31373–31381. Gillespie, P. G., & Hudspeth, A. J. (1993). Adenine nucleoside diphosphates block adaptation of mechanoelectrical transduction in hair cells. Proeedings of the National Academy of Sciences USA 90, 2710–2714. Gillespie, P. G., Wagner, M. C., & Hudspeth, A. J. (1993). Identification of a 120 kD hair-bundle myosin located near stereociliary tips. Neuron 11, 581–594. Glowatzki, E., Ruppersberg, J. P., Zenner, H. P., & Rusch, A. (1997). Mechanically and ATP-induced currents of mouse outer hair cells are independent and differentially blocked by d-tubocurarine. Neuropharmacology 36, 1269–1275. Goldberg, J. M., & Fernandez, C. (1971). Physiology of peripheral neurons innervating semicircular canals of the squirrel monkey. I. Resting discharge and response to constant angular accelerations. Journal of Neurophysiology 34, 635–660. Goodyear, R., & Richardson, G. (1992). Distribution of the 275 kD hair cell antigen and cell surface specialisations on auditory and vestibular hair bundles in the chicken inner ear. Journal of Computational Neurology 325, 243–256. Hacohen, N., Assad, J. A., Smith, W. J., & Corey, D. P. (1989). Regulation of tension on hair-cell transduction channels: Displacement and calcium dependence. Journal of Neuroscience 9, 3988–3997. Hasson, T., Gillespie, P. G., Garcia, J. A., MacDonald, R. B., Zhao, Y., Yee, A. G., Mooseker, M. S., & Corey, D. P. (1997). Unconventional myosins in inner-ear sensory epithelia. Journal of Cell Biology 137, 1287–1307. Howard, J., & Hudspeth, A. J. (1987). Mechanical relaxation of the hair bundle mediates adaptation in mechanoelectrical transduction by the bullfrog’s saccular hair cell. Proeedings of the National Academy of Sciences, USA 84, 3064–3068. Howard, J., & Hudspeth, A. J. (1988). Compliance of the hair bundle associated with gating of mechanoelectrical transduction channels in the bullfrog’s saccular hair cell. Neuron 1, 189–199.


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Hudspeth, A. J. (1982). Extracellular current flow and the site of transduction by vertebrate hair cells. Journal of Neuroscience 2, 1–10. Hudspeth, A. J. and Corey, D. P. (1977). Sensitivity, polarity, and conductance change in the response of vertebrate hair cells to controlled mechanical stimuli. Proceedings of the National Acedemy of Sciences, USA 74, 2407–2411. Jacobs, R. A., & Hudspeth, A. J. (1990). Ultrastructural correlates of mechanoelectrical transduction in hair cells of the bullfrog’s internal ear. Cold Spring Harbor Symposium on Quantitative Biology 55, 547–561. Jaramillo, F., & Hudspeth, A. J. (1991). Localization of the hair cell’s transduction channels at the hair bundle’s top by iontophoretic application of a channel blocker. Neuron 7, 409–420. Jaramillo, F., & Hudspeth, A. J. (1993). Displacement-clamp measurement of the forces exerted by gating springs in the hair bundle. Proeedings of the National Academy of Sciences, USA 90, 1330–1334. Kachar, B., Mammano, F., & Kruc, M. (1999). Structural analysis of the tip-link molecular complex. Association for Research in Otolaryngology Abstracts, #16. Kozel, P. J., Friedman, R. A., Erway, L. C., Yamoah, E. N., Liu, L. H., Riddle, T., Duffy, J. J., Doetschman, T., Miller, M. L., Cardell, E. L., & Shull, G. E. (1998). Balance and hearing deficits in mice with a null mutation in the gene encoding plasma membrane Ca2+-ATPase isoform 2. Journal of Biological Chemistry 273, 18693–18696. Kroese, A. B., Das, A., & Hudspeth, A. J. (1989). Blockage of the transduction channels of hair cells in the bullfrog’s sacculus by aminoglycoside antibiotics. Hearing Research 37, 203–217. Liang, Y., Wang, A., Belyantseva, I. A., Anderson, D. W., Probst, F. J., Barber, T. D., Miller, W., Touchman, J. W., Jin, L., Sullivan, S. L., Sellers, J. R., Camper, S. A., Lloyd, R. V., Kachar, B., Friedman, T. B., & Fridell, R. A. (1999). Characterization of the human and mouse unconventional myosin XV genes responsible for hereditary deafness DFNB3 and Shaker 2. Genomics 61, 243–258. Lumpkin, E. A., & Hudspeth, A. J. (1995). Detection of Ca2+ entry through mechanosensitive channels localizes the site of mechanoelectrical transduction in hair cells. Proeedings of the National Academy of Sciences, USA 92, 10297–10301. Lumpkin, E. A., & Hudspeth, A. J. (1998). Regulation of free Ca2+ concentration in hair-cell stereocilia. Journal of Neuroscience 18, 6300–6318. Lumpkin, E. A., Marquis, R. E., & Hudspeth, A. J. (1997). The selectivity of the hair cell’s mechanoelectrical-transduction channel promotes Ca2+ flux at low Ca2+ concentrations. Proeedings of the National Academy of Sciences, USA 94, 10997–11002. Martin, P., & Hudspeth, A. J. (1999). Active hair-bundle movements can amplify a hair cell’s response to oscillatory mechanical stimuli. Proeedings of the National Academy of Sciences, USA 96, 14306–14311. Ohmori, H. (1988). Mechanical stimulation and fura-2 fluorescence in the hair bundle of dissociated hair cells of the chick. Journal of Physiology 399, 115–137. Pickles, J. O., Comis, S. D., & Osborne, M. P. (1984). Cross-links between stereocilia in the guinea pig organ of Corti, and their possible relation to sensory transduction. Hearing Research 15, 103–112. Ricci, A. J., Fettiplace, R. (1997). The effects of calcium buffering and cyclic AMP on mechano-electrical transduction in turtle auditory hair cells. Journal of Physiology (London) 501, 111–124.

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Ricci, A. J., & Fettiplace, R. (1998). Calcium permeation of the turtle hair cell mechanotransducer channel and its relation to the composition of endolymph Journal of Physiology (London) 506, 159–173. Ricci, A. J., Wu, Y. C., & Fettiplace, R. (1998). The endogenous calcium buffer and the time course of transducer adaptation in auditory hair cells. Journal of Neuroscience 18, 8261–8277. Roberts, W. M. (1993). Spatial calcium buffering in saccular hair cells. Nature 363, 74–76. Roberts, W. M. (1994). Localization of calcium signals by a mobile calcium buffer in frog saccular hair cells. Journal of Neuroscience 14, 3246–3262. Rusch, A., Kros, C. J., & Richardson, G. P. (1994). Block by amiloride and its derivatives of mechano-electrical transduction in outer hair cells of mouse cochlear cultures. Journal of Physiology (London) 474, 75–86. Shepherd, G. M. G., Assad, J. A., Parakkal, M., Kachar, B., & Corey, D. P. (1991). Movement of the tip-link attachment is correlated with adaptation in bullfrog saccular hair cells. Journal of General Physiology 95, 25a. Shepherd, G. M. G., Barres, B. A., & Corey, D. P. (1989). “Bundle Blot” purification and initial protein characterization of hair-cell stereocilia. Proceedings of the National Academy of Sciences, USA 86, 4973–4977. Shepherd, G. M. G., & Corey, D. P. (1994). The extent of adaptation in bullfrog saccular hair cells. Journal of Neuroscience 14, 6217–6229. Shepherd, G. M. G., Corey, D. P., & Block, S. M. (1990). Actin cores of hair-cell stereocilia support myosin motility. Proeedings of the National Academy of Sciences, USA 87, 8627–8631. Shotwell, S. L., Jacobs, R., & Hudspeth, A. J. (1981). Directional sensitivity of individual vertebrate hair cells to controlled deflection of their hair bundles. Annual New York Academy of Science 374, 1–10. Smith, R. L., Brachman, M. L., & Goodman, D. A. (1983). Adaptation in the auditory periphery. Annual New York Academy of Science 405, 79–93. Sobin, A., & Flock, A. (1983). Immunohistochemical identification and localization of actin and fimbrin in vestibular hair cells in the normal guinea pig and in a strain of the waltzing guinea pig. Acta Otolaryngology (Stockholm) 96, 407–412. Solc, C. K., Derfler, B. H., Duyk, G. M., & Corey, D. P. (1994). Molecular cloning of myosins from the bullfrog saccular macula: A candidate for the adaptation motor. Auditory Neuroscience 1, 63–75. Street, V. A., McKee-Johnson, J. W., Fonseca, R. C., Tempel, B. L., & Noben-Trauth, K. (1998) Mutations in a plasma membrane Ca2+-ATPase gene cause deafness in deafwaddler mice. Nature Genetics 19, 390–394. Tilney, L. G., DeRosier, D. J., & Mulroy, M. J. (1980). The organization of actin filaments in the stereocilia of cochlear hair cells. Journal of Cell Biology 86, 244–259. Walker, R. G., & Hudspeth, A. J. (1996). Calmodulin controls adaptation of mechanoelectrical transduction by hair cells of the bullfrog’s sacculus Proeedings of the National Academy of Sciences, USA 93, 2203–2207. Walker, R. G., Hudspeth, A. J., & Gillespie, P. G. (1993). Calmodulin and calmodulin-binding proteins in hair bundles. Proeedings of the National Academy of Sciences, USA 90, 2807–2811. Wu, Y. C., Ricci, A. J., & Fettiplace, R. (1999). Two components of transducer adaptation in auditory hair cells. Journal of Neurophysiology 82, 2171–2181.


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Yamoah, E. N., & Gillespie, P. G. (1996). Phosphate analogs block adaptation in hair cells by inhibiting adaptation-motor force production. Neuron 17, 523–533. Yamoah, E. N., Lumpkin, E. A., Dumont, R. A., Smith, P. J., Hudspeth, A. J., & Gillespie, P. G. (1998). Plasma membrane Ca2+-ATPase extrudes Ca2+ from hair cell stereocilia. Journal of Neuroscience 18, 610–624. Zhao, Y., Yamoah, E. N., & Gillespie, P. G. (1996). Regeneration of broken tip links and restoration of mechanical transduction in hair cells. Proeedings of the National Academy of Sciences,USA 93, 15469–15474. Zhu, T., Sata, M., & Ikebe, M. (1996) Functional expression of mammalian myosin 1: Analysis of its motor activity. Biochemistry 35, 513–522. Note: This chapter is expanded from a review in press in Proceedings of the National Academy of Sciences, USA by J. R. Holt and D. P. Corey.

2 How A Living Hair Cell Repairs Itself: Involvement of Purinoceptors in the Repair of Hair Bundle Mechanoreceptors of Sea Anemones Glen M. Watson, Ph.D.

In sea anemones, hair bundle mechanoreceptors regulate discharge of nematocysts into vibrating targets. Consequently, proper functioning of hair bundles can be inferred from a bioassay based on counting nematocysts discharged into vibrating targets. Following trauma caused by exposure to calcium-free seawater, hair bundle mechanoreceptors are repaired by secretory proteins called “repair proteins” (RP). Exogenously supplied RP speeds the restoration of vibration dependent discharge from 4 hr in seawater alone to 7–8 min in RP. In addition, the recovery is further enhanced to 2 min by including extracellular ATP. ATPase activity is confirmed in isolated repair proteins at the TEM level. I here present evidence that the repair process includes an activation of purinoceptors. At modest levels of repair proteins, PPADS, a known inhibitor of purinoceptors, blocks or delays the recovery of vibration dependent discharge. Calcium imaging suggests that activated purinoceptors induce calcium transients in the tentacle epidermis. The calcium transients are blocked in the presence of PPADS, or in the presence of anti-P2X antibodies. Furthermore, RP induces calcium transients in tissue previously exposed to calcium-free seawater. Recovery of vibration dependent discharge of nematocysts is inhibited in animals preloaded with W7, an inhibitor of calcium/calmodulin, or with nifedipine, an inhibitor of L-type calcium channels. In control animals with healthy hair bundles, exposure to ATP causes a loss of vibration de27

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pendent discharge that requires approximately 6 hr to recover. This detrimental effect of extracellular ATP on vibration sensitivity is blocked in the presence of PPADS, W7, or nifedipine. Taken together, these results suggest that extracellular ATP may function in repair of hair bundles both by serving as an energy source for ecto-ATPases in repair proteins and by serving as a ligand for purinoceptors. Apparently, the purinoceptors are regulated by components of RP to enhance repair without allowing damage to the hair bundles. Activated purinoceptors induce a calcium influx through cell surface channels sensitive to dihydropyridines. Elevated calcium initiates a calcium/calmodulin second messenger pathway.

INTRODUCTION Sea anemones are predominantly sessile, marine invertebrates that use hair bundles to detect swimming movements of potential prey (Watson and Mire, 1999). The hair bundles are strikingly similar to those of the acousticolateralis system of vertebrates in terms of fine structure. Anemone hair bundles consist of actin based stereocilia interconnected by numerous linkages including tip links (Watson and Roberts, 1995; Watson, Mire, and Hudson, 1997). Upon stimulation by vibrations at specific, key frequencies, the hair bundles somehow predispose anemones to discharge nematocysts into targets subsequently touched to their tentacles (Watson, Mire, and Hudson, 1998a; see Chapter 4 by Dr. Mire for recent developments on the physiology of anemone hair bundles). Discharge of nematocysts involves the forceful and rapid eversion of a tubule from the nematocyst capsule into contact with the target organism (Skaer and Picken, 1965; Holstein and Tardent, 1984). Depending on the type of nematocyst, the everting tubule may adhere to the surface of the target or penetrate its integument to inject potent toxins (Mariscal, 1984). In the laboratory, we developed a bioassay in which vibrating test probes coated with a thin layer of gelatin are touched to tentacles and then withdrawn. Discharged nematocysts penetrating the gelatin coating are counted using phase contrast microscopy. The bioassay consistently detects a doubling of nematocysts discharged into test probes vibrating at specific, key frequencies as compared to probes vibrating at other frequencies or not vibrating at all. The enhancement in discharge above baseline levels is referred to as “vibration dependent discharge” (Watson, Mire, and Hudson,1998a; Watson and Mire, 1999). It is interesting to note that vibration dependent discharge is inhibited by compounds known to inhibit signal transduction in hair cells of the acousticolateralis system of vertebrates. Among these compounds are aminoglycoside antibiotics including streptomycin. Streptomycin inhibits mechanotransduction of anemone hair cells (Mire and Watson, 1997) and vibration dependent discharge (Watson, Mire, and Hudson, 1997) at

HOW A LIVING HAIR CELL REPAIRS ITSELF

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doses known to inhibit signal transduction in vertebrate hair cells (Kroese, Das, and Hudspeth, 1989). Such inhibition is completely reversible both in anemones and in vertebrates. Furthermore, agents thought to attack tip links (among other linkages interconnecting stereocilia) abolish mechanosensitivity in vertebrate hair cells and vibration dependent discharge in anemones. Among these are elastase (Osborne and Comis, 1990; Preyer, Hemmert, Zenner, and Gummer, 1995) and calcium depleted buffers containing such calcium chelators as EGTA or BAPTA (Assad, Shepherd, and Corey, 1991; Crawford, Evans, and Fettiplace, 1991; Zhao, Yamoah, and Gillespie, 1996). In anemones, the loss of vibration dependent discharge caused by exposure to calcium-free seawater is temporary. The recovery period increases with the duration of exposure to calcium-free seawater (Watson, Mire, and Hudson, 1998b). After 1hr in calcium-free seawater, the hair bundles are completely disorganized and vibration dependent discharge is abolished. However, vibration dependent discharge and normal morphology of the hair bundles are restored within 4 hr after the animals are returned to normal seawater. Such a rapid recovery suggests that hair bundles of anemones are repaired rather than replaced by cell division and differentiation of new hair cells. Assuming that exposure to calcium-free seawater damages or destroys linkages interconnecting stereocilia while otherwise leaving the hair bundle intact, we reasoned that the repair process involves replacing lost or damaged linkages. Even so, the process of repair is difficult to understand because linkages are extracellular in distribution and interconnect stereocilia membranes separated by substantial distances. Nevertheless, repair of pre-existing hair bundles was confirmed by additional experiments in which it was shown that specific secretory proteins called, “repair proteins” restore vibration dependent discharge and normal morphology of the hair bundles (Watson, Mire, and Hudson, 1998b). Exogenously supplied repair proteins restore vibration dependent discharge within 7–8 min to anemones previously exposed to calcium-free seawater for 1 hr. This process is further enhanced by adding ATP with the repair proteins. In the combined presence of RP and 10–6 M ATP, vibration dependent discharge is restored within 2 min. ATPase activity is confirmed at the TEM level for isolated repair proteins. Thus, it appears likely that exogenously supplied ATP facilitates biochemical reactions carried out by enzymatic domains of the repair proteins (Watson, Venable, Hudson, and Repass, 1999). These biochemical reactions are likely to be component parts of the repair process. On the other hand, effects of ATP need not be limited to enzymatic activity within repair proteins. It is conceivable that ATP may initiate intracellular processes involved in repair by activating purinoceptors. Purinoceptors are known to be present in vertebrate hair cells (e.g., Housley, Raybould, and Thorne, 1998; see chapter 7 by Dr. Bobbin for additional information concerning purinoceptors in au-

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ditory systems of mammals). In the present paper, I will present data implicating purinoceptors in repair of hair bundles, then discuss some details of the second messenger pathways initiated by activated purinoceptors.

MATERIALS AND METHODS The bioassay for vibration sensitivity based on nematocyst discharge into vibrating test probes was performed as described previously (Watson and Hudson, 1994). Calcium imaging was performed on excised tentacles threaded with a fine, human hair. The ends of the hair were glued to the glass slide. Tentacles were loaded with 5 µM fluo-3 AM according to the manufacturer’s instructions (Molecular Probes, Eugene, OR). Specimens were anesthetized in potassium seawater (formulated as described previously, Watson et al., 1997). Excised tentacles were perfused with potassium seawater which then was exchanged with the experimental solution in potassium seawater by switching a valve in the supply tubing. The tissue was examined using a LOMO Multiscope light microscope (LOMO America, Prospect Heights, IL ) equipped with fluorescence and a 70X water immersion objective (n.a. = 1.23) (GEK, Ltd., Charlottesville, VA). Digital images were obtained using a cooled CCD camera (SBIG, Santa Barbara, CA) and subsequently analyzed using Image Pro Plus software (Media Cybernetics, Silver Springs, MD). Video microscopy of hair bundles during the repair process was performed with an IMT-2 inverted microscope (Olympus, Tokyo) equipped with transmitted and reflected DIC optics.

RESULTS Evidence that Purinoceptors are Involved in Repair In the presence of exogenously supplied RP at 3.33 µl/ml, full recovery of vibration dependent discharge is achieved within 10 min after adding RP to the dish (Figure 2–1). Half maximal recovery is detected at 1.67 µl/ml RP. Alone, 10-5 M PPADS, a known inhibitor of purinoceptors, does not significantly affect vibration dependent discharge of nematocysts in animals having healthy hair bundles. After 1 hr exposure in calcium-free seawater, including PPADS in the bath with the RP gives normal recovery of vibration dependent discharge at 3.3 µl/ml RP, but essentially no recovery at lower RP levels (Figure 2–1). In the presence of PPADS, the half-maximal dose of RP is increased to approximately 2.88 µl/ml. Inasmuch as purinoceptors can function as ligand gated channels (i.e., P2X receptors) or as typical G protein linked receptors (i.e., P2Y receptors), inhibitors of the calcium second messenger pathway were tested. In the presence of 3.3 µl/ml RP, the calcium calmodulin inhibitor, W7, inhibits re-


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Figure 2–1. Effects of PPADS on recovery of vibration dependent discharge of nematocysts. Anemones were exposed to calcium-free seawater for 1 hr to disrupt structural integrity and function of hair bundles. Upon returning to calcium-containing seawater, discharge of nematocysts into test probes vibrating at 55 Hz was tested 10 min after co-adding 10-5 M PPADS, an inhibitor of purinoceptors, (or not, controls) and repair protein concentrate at the dose indicated. Data points indicate the mean number (± SEM) of microbasic p-mastigophore nematocysts counted per field of view for a total of 4 test probes, each touched to a separate anemone. Data for controls receiving RP only (triangles) are plotted alongside data for experimental animals exposed to PPADS and RP (circles).

covery at 10-9 M W7 (Figure 2–2A). In animals having healthy hair bundles, W7 does not affect vibration dependent discharge. In the presence of 3.3 µl/ml RP, 10-9 M nifedipine, a dihydropyridine inhibitor of L-type voltage gated calcium channels, inhibits recovery of vibration dependent discharge of nematocysts (Figure 2–2B). Alone, nifedipine does not affect vibration dependent discharge of nematocysts.

Calcium Imaging of Excised Tentacles Dynamics of intracellular calcium ions were monitored in excised tentacles using the calcium fluorochrome, fluo 3. Upon returning tentacles to calciumcontaining seawater after 1 hr exposure to calcium-free seawater, levels of intracellular calcium increase as indicated by a brightening of fluorescence intensity, in this case, to a mean of 1.28 times the initial intensity (Figure 2–3). Adding 3.3 µl/ml RP further increases levels of intracellular calcium as indicated by brightening to 1.47 times the initial intensity, but with a decrease in intracellular calcium occurring by 7 min, a time when vibration dependent discharge normally recovers. After 7 min, calcium levels again

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Figure 2–2. Effects of W7 or nifedipine on recovery of vibration dependent discharge. Anemones were exposed to calcium-free seawater for 1 hr to disrupt structural integrity and function of hair bundles. Upon returning to calcium-containing seawater, specimens were loaded with W7, an inhibitor of calcium/calmodulin, or with nifedipine, an inhibitor of L-type calcium channels, then transferred to fresh seawater. 3.3 µl/ml RP was added and discharge into test probes vibrating at 55 Hz was tested 10 min later. Mean discharge (± SEM, n = 4 for each data point) is plotted for specimens loaded with (A) W7 or (B) nifedipine at the dose indicated.

increase and remain elevated through 30 min (Figure 2–3). Recent data indicate substantial rearrangements of the hair bundle continue after the first 7 min in RP, despite the full recovery of vibration sensitivity at 7 min. It is striking how much movement is observed in the hair bundle during repair. Because exposure to calcium-free seawater might alone alter resting levels of intracellular calcium, excised tentacles from animals having healthy hair bundles were exposed to ATP to determine whether activated purinoceptors increase intracellular calcium. Untreated control tentacles having healthy hair bundles exhibit relatively constant calcium levels over


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Figure 2–3. Relative fluorescence intensity of fluo labeled tentacles during repair of hair bundles. Excised tentacles were mounted onto slides and then labeled with 5 µM fluo-3 AM to monitor intracellular calcium. Tentacles were exposed to calciumfree seawater for 1 hr to disrupt hair bundles and then returned to calcium-containing seawater fortified with 3.3 µl/ml RP. Images were collected before and after exposure to calcium-free seawater and at intervals after exposure to RP. Data points indicate relative fluorescence intensity with the intensity of the initial image adjusted to 1.00. Each point plotted represents mean relative fluorescence (± SEM, n = 4).

time as indicated by relatively stable intensity of fluo fluorescence. Mean intensity over 10 min is 1.02 times the initial fluorescence intensity (Figure 2–4A). Upon perfusing 10-6 M ATP, levels of intracellular calcium increase and remain elevated through 10 min with mean fluorescence intensity of 1.31 times the initial intensity (Figure 2–4B, 2–5). Calcium oscillations are evident in each of the individual experiments (e.g., Figure 2–5). However, because the oscillations are not synchonized between trials, they are obscured in mean data based on four trials (Figure 2–4). At 10-12 M ATP, calcium levels increase less dramatically as indicated by a mean fluorescence intensity of 1.18 times the initial intensity (Figure 2–4C). Commercial antibodies to P2X purinoceptors inhibit the increase in calcium normally induced by 10-6 M ATP (Figure 2–6A). In the combined presence of anti-P2X antibodies and 106 M ATP, mean fluorescence intenstity is 0.99 that of initial intensity. Likewise, in the presence of 10-5 M PPADS, the increase in intracellular calcium normally induced by 10-6 M ATP is inhibited (Figure 2–6B). In the combined presence of 10-6 M ATP and 10-5 M PPADS, mean fluorescence intensity is 0.95 that of initial intensity. Washing out the PPADS or P2X antibodies permits a modest increase in intracellular calcium (Figure 2–6).

Figure 2–4. Relative fluorescence intensity of fluo labeled tentacles following exposure to extracellular ATP. Excised tentacles were mounted onto slides and then labeled with 5 µM fluo-3 AM to monitor intracellular calcium. Tentacles either received no further treatment (A) controls, or were exposed to (B) 10-6 M ATP, or (C) 10-12 M ATP. Images were collected both before the perfusion of ATP and at intervals afterwards. Data points indicate relative fluorescence intensity with the intensity of the initial image (before perfusion) adjusted to 1.00. Each point plotted represents mean relative fluorescence (± SEM, n = 4).

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Figure 2–5. Spatio-temporal effects of µM ATP on fluo fluorescence. Excised tentacles were mounted and loaded with fluo 3AM as described above. Difference images are shown, each obtained by subtracting the before perfusion image from a specific after perfusion image obtained X minutes after the perfusion was initiated. The value for X is shown in the upper lefthand corner of each panel. A positive difference (gray to white areas) indicates a local increase in intracellular calcium. Contrast was adjusted comparably for the entire series so that comparisons between images of the panel faithfully represent changes in levels of intracellular calcium. Scale bar = 7 µm.

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Figure 2–6. Relative fluorescence intensity of fluo labeled tentacles following exposure to extracellular ATP in the presence of anti-P2X antibodies or PPADS. Excised tentacles were mounted onto slides and then labeled with 5 µM fluo-3 AM to monitor intracellular calcium. Tentacles were first incubated in (A) anti-P2X antibodies or (B) 10-4 M PPADS and then perfused with the same solution fortified with 10-6 M ATP. After 5 min, both the ATP and the putative blocker were washed out and fluorescence monitored for an additional 11 min. Data points indicate relative fluorescence intensity with the intensity of the initial image (before ATP perfusion) adjusted to 1.00. Each point plotted represents mean relative fluorescence (± SEM, n = 4).

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Effects of ATP on Vibration Dependent Discharge of Nematocysts A 5 min exposure to 10-6 M ATP is sufficient to abolish vibration dependent discharge (Figure 2–7A) with recovery requiring approximately 6 hr (Figure 2–7B). The detrimental effects of ATP on vibration dependent discharge are prevented by PPADS. In the combined presence of ATP and 10-4 M PPADS, vibration dependent discharge is inhibited at ATP concentrations of 10-9 M ATP or higher. However, vibration dependent discharge is immediately restored when the ATP and PPADS are washed out (Figure 2–8). A comparable effect is observed in animals preloaded with 10-8 M W7 and then exposed to ATP, where vibration dependent discharge is inhibited at ATP concentrations of 10-7 M or higher (Figure 2–9). Vibration sensitivity is fully restored upon washing out the ATP. Likewise, vibration dependent discharge is protected from µM ATP by 10-7 M nifedipine (Figure 2–10). At all concentrations of nifedipine, washing out the ATP fully restores vibration dependent discharge at all concentrations of nifedipine (Figure 2–10).

DISCUSSION It appears that purinoceptors are activated during the repair process because PPADS, a known inhibitor of purinoceptors, decreases the efficacy of repair proteins to restore vibration dependent discharge. On the other hand, activating purinoceptors in animals having healthy hair bundles abolishes vibration dependent discharge of nematocysts with recovery requiring approximately 6 hr. In healthy animals, vibration dependent discharge is protected from ATP by PPADS, indicating an involvement of purinoceptors in this loss of vibration sensitivity. Activated purinoceptors may activate calcium second messenger pathways since W7, a known inhibitor of calmodulin, inhibits the repair process in experimental animals exposed to RP. Likewise, W7 protects vibration dependent discharge from ATP in animals having healthy hair bundles. Nifedipine, a known inhibitor of L-type calcium channels, inhibits the repair process in experimental animals exposed to RP. In addition, nifedipine protects vibration dependent discharge from ATP. Thus, the activated purinoceptors may activate calcium channels to induce calcium transients in the tentacle epidermis. Calcium imaging indicates calcium transients follow exposure to ATP except in the presence of purinoceptor inhibitors or P2X antibodies. Elevated intracellular calcium may activate calcium calmodulin (CAM) to facilitate the repair process in experimental animals damaged by exposure to calcium-free seawater. In the absence of repair proteins, CAM appears to have a destructive activity. Thus, it would be essential to the repair process that RP arrive at the damaged hair bundle before or simultaneously to extracellular ATP. Apparent-

Figure 2–7. Effects of ATP on vibration dependent discharge of nematocysts. (A) Animals with healthy hair bundles received no additional treatment (control) or were exposed 5 min to ATP at the dose indicated on the X axis. After removing the ATP, discharge of nematocysts was tested at 55 Hz, a preferred frequency. (B) Recovery of vibration dependent discharge after 5 min in 10-6 M ATP. Discharge was tested at 30 min intervals after removing the ATP. Data points indicate the mean number (± SEM) of microbasic p-mastigophore nematocysts counted per field of view for a total of 4 test probes, each touched to a separate anemone.

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Figure 2–8. Effects of PPADS and ATP on vibration dependent discharge. Specimens with healthy hair bundles were placed in the combined presence of 10-4 M PPADS and ATP at the concentration indicated on the X axis. Vibration dependent discharge was assayed at 55 Hz (A) after 5 min in the combined presence of PPADS and ATP or (B) 1 min after these reagents were washed out. Data points indicate the mean number (± SEM) of microbasic p-mastigophore nematocysts counted per field of view for a total of 4 test probes, each touched to a separate anemone.

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Figure 2–9. Effects of W7 and ATP on vibration dependent discharge. Specimens with healthy hair bundles were preloaded with 10-8 M W7 for 20 min, then transferred to fresh seawater. Nematocyst discharge into test probes vibrating at 55 Hz was tested (A) 5 min after adding ATP to the final concentration indicated on the X axis, or (B) 1 min after the ATP was washed out. Data points indicate the mean number (± SEM) of microbasic p-mastigophore nematocysts counted per field of view for a total of 4 test probes, each touched to a separate anemone.

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Figure 2–10. Effects of nifedipine and ATP on vibration dependent discharge. Specimens with healthy hair bundles were preloaded with nifedipine at the dose indicated on the X axis for 20 min, then transferred to fresh seawater. Nematocyst discharge into test probes vibrating at 55 Hz was tested (A) 5 min after adding 10-6 M ATP to the dish, or (B) 1 min after the ATP was washed out. Data points indicate the mean number (± SEM) of microbasic p-mastigophore nematocysts counted per field of view for a total of 4 test probes, each touched to a separate anemone.

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ly, RP triggers ATP secretion by the sensory neuron located at the center of the hair bundle (Watson, Venable, Hudson, and Repass, 1999).The reversible inhibition of vibration dependent discharge observed at relatively high concentrations of ATP in animals preloaded with W7 or nifedipine is interesting in that it suggests that ATP may be affecting vibration sensitivity by at least two mechanisms, only one of which depends on CAM. Whereas the CAM dependent inhibition requires a 6 hr recovery, the CAM independent inhibition is rapidly reversible. Taken together, it seems reasonable to speculate that components of the repair protein mixture regulate purinoceptors to enhance the repair process while also preventing the destructive potential of activated purinoceptors. The apparent involvement of purinoceptors in repair places at least two classes of molecules at the extracellular surface of damaged hair bundles: repair proteins, several different protein complexes each having an estimated mass of 2000 kD; and ATP. The ATP may serve both as an energy source for ATPase domains residing in at least some of the RP protein complexes (Watson et al., 1999), and also as a ligand for purinoceptors which initiate calcium transients associated with the repair process. Preliminary results from ongoing experiments suggest that elevated cytoplasmic calcium activates calcium calmodulin (CAM) to reorganize the actin cytoskeleton of stereocilia.

Acknowledgments: I appreciate the technical expertise of Ms. Stacy Venable-Thibodeaux and the financial support from NIH R01-GM52334.

REFERENCES Assad, J. A., Shepherd, G. M. G., & Corey, D.P., (1991) Tip link integrity and mechanotransduction in vertebrate hair cells. Neuron, 7, 985–994. Crawford, A. C., Evans, M. G., & Fettiplace, R., (1991) The actions of calcium on the mechano-electrical transducer current of turtle hair cells. Journal of Physiology, 434, 369–398. Holstein, T., & Tardent, P., (1984) An ultrahigh-speed analysis of exocytosis: nematocyst discharge. Science, 223, 830–833. Housley, G. D., Raybould, N. P., & Thorne, P. R. (1998) Fluorescence imaging of the Na+ influx via P2X receptors in cochlear hair cells. Hearing Research, 119, 1–13. Kroese, A. B. A., Das, A., and Hudspeth, A. J. (1989) Blockage of the transduction channels of hair cells in the bullfrog’s sacculus by aminoglycoside antibiotics. Hearing Research, 37, 203–218. Mariscal, R. N., (1984) Cnidaria: Cnidae. In “Biology of the Integument, vol. I, Invertebrates.” (Bereiter-Hahn, J., Maltoltsy, A. G., Richards,, K. S., eds.), pp.57–67, Springer-Verlag, Berlin/New York. Mire, P., & Watson, G. M., (1997) Mechanotransduction of hair bundles arising from multicellular complexes in anemones. Hearing Research, 113, 244–234.


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Osborne, M. P., & Comis, S. D. (1990) Action of elastase, collagenase and other enzymes upon linkages between stereocilia in the guinea-pig cochlea. Acta Otolaryngology, 110, 37–45. Preyer, S., Hemmert, W., Zenner, H. P., & Gummer, A. W., (1995) Abolition of the receptor potential response of isolated mammalian outer hair cells of the guinea pig cochlea by hair bundle treatment with elastase: a test of the tip link hypothesis. Hearing Research, 89, 187–193. Skaer, R. J., & Picken, L. E. R., (1965) The structure of the nematocyst thread and the geometry of discharge in Corynactis viridis (Allman). Philosophical Transactions of the Royal Society of London, Series:B, 250, 131–164. Watson, G. M., & Hudson, R. R., (1994) Frequency and amplitude tuning of nematocyst discharge by proline. Journal of Experimental Zoology, 268, 177–185. Watson, G. M., & Mire, P., (1999) A comparison of hair bundle mechanoreceptors in sea anemones and vertebrate systems. Current Topics in Developmental Biology, 43, 51–84. Watson, G. M., Mire, P., & Hudson, R. R., (1997) Hair bundles of sea anemones as a model system for vertebrate hair bundles. Hearing Research, 107, 53–66. Watson, G. M., Mire, P., and Hudson, R. R., (1998a) Frequency specificity of vibration dependent discharge of nematocysts in sea anemones. Journal of Experimental Zoology, 281, 582–593. Watson, G. M., Mire, P., & Hudson, R. R., (1998b) Repair of hair bundles in sea anemones by secreted proteins. Hearing Research, 115, 119–128. Watson, G. M., and Roberts, J., (1995) Chemoreceptor-mediated polymerization and depolymerization of actin in hair bundles of sea anemones. Cell Motility and the Cytoskeleton, 30, 208–220. Watson, G. M., Venable, S., Hudson, R. R. & Repass, J. J. (1999) ATP enhances repair of hair bundles in sea anemones. Hearing Research, 136, 1–12. Zhao, Y., Yamoah, E. N., & Gillespie, P. G., (1996) Regeneration of broken tip links and restoration of mechanical transduction in hair cells. Proceedings of the National Academy of Sciences, USA, 94, 15469–15474.

3 Fast Transducer Adaptation, Physiological Implications and Underlying Mechanisms Anthony J. Ricci, Ph.D.

MECHANO-ELECTRIC TRANSDUCTION Hair cells are mechano-receptors responsible for detecting motions over a wide frequency range. A common mechanism is thought to be responsible for establishing the broad frequency range over which these cells operate. This same mechanism is thought to maintain a high sensitivity over a wide range of stimulus magnitudes (Hudspeth, 1989). An array of actin filled stereocilia, embedded in the cuticular plate, project from the apical end of the hair cell in increasing heights toward, in some cases, a true kinocilium (Tilney, DeRosier, and Melroy, 1980; Slepecky and Chamberlain, 1982; Tilney, Egelman, DeRosier, and Saunder, 1983). Mechano-electric transducer (MET) channels are located near the tops of the stereocilia (Hudspeth, 1982; Jaramillo and Hudspeth, 1991; Lumpkin and Hudspeth, 1995). It is hypothesized that the channels are at either or both ends of a thin filamentous element, the tip-link, that joins stereocilia in adjacent rows and are aligned along the hair bundle’s axis of sensitivity (Pickles et al., 1989). Present theory suggests that deflection of the hair bundle increases (toward the kinocilium) or decreases (away from the kinocilium) tension in the tip-link which in turn is thought to directly translate this tension to a mechanically-gated channel is tethered to either or both ends of the link (Howard and Hudspeth, 1988; Denk, Holt, Shepherd, and Corey, 1995; Gillespie and Corey, 1997). An adaptation process has been described which is thought to restore the tip-links to a constant tension regardless of the hair bundle’s steady-state position, in order to extend the dynamic range of the hair bundle while maintaining sensitivity (Eatock, 45

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Corey, and Hudspeth, 1987; Crawford, Evans, and Fettiplace, 1989; Hacohen, Assad, Smith, and Corey, 1989; Assad and Corey, 1992). Recently, it has been shown that adaptation can be extremely fast, and it has been postulated that adaptation may impart a mechanical tuning to hair bundles (Ricci and Fettiplace, 1997; Ricci, Wu, and Fettiplace, 1998). Furthermore, evidence suggests that there may be multiple mechanisms involved in adaptation (Ricci et al., 1998; Wu, Ricci, and Fettiplace, 1999). It may be that the function of the fast adaptation process is to provide a tuning mechanism to the hair bundle, while the slower adaptation process maintains the operating range of the hair bundle. The purpose of this review is first to synopsize existing data regarding fast adaptation, emphasizing evidence that implicates adaptation as a mechanical tuning mechanism. The second part of the chapter will present evidence supporting the contention that there may be multiple mechanisms of adaptation.

METHODS The turtle auditory papilla has been a productive preparation for studying tuning mechanisms in auditory hair cells for over twenty years (Crawford and Fettiplace, 1978; Crawford and Fettiplace, 1980; Crawford and Fettiplace, 1981; Art, Crawford, Fettiplace, and Fuchs, 1982; Crawford and Fettiplace, 1985; Art and Fettiplace, 1987; Crawford et al., 1989; Art, Wu, and Fettiplace, 1995; Tucker and Fettiplace, 1996). Turtle hair cells have been employed to investigate afferent nerve tuning, electrical resonance as a tuning mechanism, efferent regulation of tuning, activation and adaptation of MET channels and mechanical properties of hair bundles. The recent modification of the turtle preparation, in which patch-clamp recordings from the hair cell in the intact papilla could be made, has allowed for more quantitative investigations of transducer adaptation (Ricci and Fettiplace, 1997). Transducer currents can be recorded at a much more routine rate, the magnitudes of the currents are much larger and adaptation is more robust than has been previously reported.

ADAPTATION IS FAST Adaptation manifests itself as a decrease in current amplitude during a constant stimulus. The rate and extent of adaptation vary with stimulus magnitude; however, as seen in Figure 3–1, the rate of adaptation remains fast for stimuli which activate up to about 50% of the maximal current (Hacohen et al., 1989; Ricci and Fettiplace, 1997). For small displacement steps adaptation could be almost complete. In addition, fast adaptation was found to be linear, being equally as fast in response to positive and negative stimuli (Wu et al., 1999). Initially, the rates of adaptation were

Figure 3–1: Mechano-electric transducer currents elicited from an auditory hair cell recorded from the intact papilla. Stimuli are shown above the current responses. Hair cells were voltage clamped at 70 mV. The right hand panels are an expanded view of a 50 nm response showing fast adaptation. A single exponential decay is fit (heavy line) to the current giving a time constant of 0.9 ms.

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measured by fitting a single exponential to the decay in transducer current elicited by a small deflection of the hair bundle (Figure 3–1). The response chosen was one eliciting less than 50% of the maximal current in order not to obscure the measurement due to saturation or perhaps the onset of a second component. Rates of adaptation were found to be less than 1 ms. To date, the fastest adaptation rate measured has been ~100µs, comparable to the stimulus rise time. Wu et. al., (1999), demonstrated that the stimulus rise time is critical in setting the rate of adaptation. It is possible then that the adaptation rate is being limited by the speed of the stimulus, such that faster stimuli may lead to faster rates of adaptation. This result may also explain the variability in measured adaptation rates between investigators, where other methods of hair bundle stimuli would be considerably slower.

TONOTOPIC VARIATIONS IN TRANSDUCTION Plotting the time constant of adaptation against hair cell location along the papilla showed an exponential decrease in adaptation rate (Ricci and Fettiplace, 1997). The time constant of adaptation varied in a manner similar to the tonotopic frequency map of the auditory papilla determined from afferent nerve fiber recordings (Figure 3–2, left) (Fettiplace and Fuchs, 1999). The solid line in Figure 3–2 (left) is the tonotopic map of the papilla derived from afferent nerve recordings (Crawford and Fettiplace, 1980). The correlation between adaptative rate and tonotopic map was the first clue that the rate of fast adaptation might be a tuning mechanism or at least a high pass filter for hair cells. A high pass filter had been reported previously in afferent nerve recordings, but the etiology was unknown (Crawford and Fettiplace, 1980). A tonotopic increase in the magnitude of the transducer current has also been found (Figure 3–2, right). Since the rate of adaptation is directly proportional to calcium entry, it has been postulated that an increase in channels per stereocilia would increase the calcium load per stereocilia thus speeding adaptation (Ricci and Fettiplace, 1998). In an attempt to determine if the number of channels per stereocilia increased tonotopically, an estimate of channel density was made. The number of stereocilia per bundle increases tonotopically, so it would be predicted that the magnitude of the transducer current would increase regardless of a change in density (Hackney, Fettiplace, and Furness, 1993). The dashed lines of Figure 3–2 illustrate this phenomenon where either 1,3 or 5 channels is multiplied by the single channel current and then by the number of stereocilia for that position to give a theoretical maximal current (Crawford et al., 1991; Hackney et al., 1993). The solid line represents a graded change in the number of channels per stereocilia, varying linearly from 2 to 5. This line most closely matches the measured current responses. Thus, a tonotopic increase in the number of channels per stereocilia may in part underlie the variation in adaptation rates measured.

TRANSDUCER ADAPTATION, PHYSIOLOGICAL IMPLICATIONS & MECHANISMS

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Figure 3–2: The left-hand panel plots adaptation time constant against location. The solid line represents the tonotopic map of the papilla measured from primary afferent neurons (Crawford and Fettiplace, 1980). Cells were binned in 0.1 increments of position and the maximal transducer currents averaged and plotted in the right-hand panel (±SEM). Numbers of channels per stereocilia were estimated, as described in the text. The dashed lines represent predicted transducer channels for 1, 3 or 5 channels per stereocilia, while the solid line represents a tonotopic increase in channels per stereocilia, from 2 to 5.

CALCIUM SENSITIVITY OF ADAPTATION Calcium Buffering It has been known for some time that adaptation is a Ca2+-dependent process (Eatock et al., 1987; Corey and Hudspeth, 1983; Crawford et al., 1989; Hacohen et al., 1989; Crawford, Evans, and Fettiplace, 1991). Lowering external Ca2+ levels can slow and even abolish adaptation (Crawford et. al., 1991; Ricci and Fettiplace, 1997) (Figure 3–3 bottom). This has led to the question of whether adaptation can be maintained under normal endolymph environment. Endolymph is a high potassium, low calcium solution bathing the hair bundle. Ca2+ has been measured near 60M in turtle papilla (Crawford et al., 1991) and closer to 30M in mammalian cochlea (Bosher and Warren, 1978). The sensitivity to external Ca2+ is to some degree determined by the properties and concentrations of intraciliary Ca2+ buffer (Crawford et al., 1991; Ricci and Fettiplace, 1997; Ricci et al., 1998). By lowering the concentration of the intraciliary Ca2+ chelator, it is possible to maintain adaptation

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Figure 3–3: Example of the response of two cells (upper and lower panels) from a similar region of the papilla (~0.53 from apex) to lowering external calcium from 2.8 mM to 50M (left and right, respectively). The cell at the top had 0.1 mM BAPTA and maintains adaptation while the bottom cell had 3 mM of the calcium chelator and adaptation is compromised. With BAPTA there is a similar effect on open probability and adaptation.

at physiologic concentrations of Ca2+ (Ricci and Fettiplace, 1997; Ricci et al., 1998). The sensitivity of adaptation to intraciliary buffer concentration has allowed for the estimate of the endogenous Ca2+ buffering concentration(Ricci et al., 1998). Perforated patch recordings were compared to whole-cell recordings using different BAPTA concentrations for these estimates. It was found that a gradient existed in the endogenous buffer concentration with the levels increasing with frequency. The range of concentrations for endogenous buffer ranged from ~0.1 mM to greater than 0.5 mM (Ricci et al., 1998). Similar concentrations gradients have been estimated in mammalian cochlea. At these concentrations of Ca2+ buffer, adaptation can be maintained at low external Ca2+ concentrations (Oberholtzer, Buettger, Summers, and Matschonsky, 1988). As an aside, the sensitivity of a particular cell to either the intraciliary level of Ca2+ buffer or of external Ca2+ was also dependent on the magni-


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tude of the transducer current (Ricci et al., 1998). The larger the transducer current the less sensitive to either variable. This implies that there is a summation of Ca2+ within the stereocilia in cells with large currents, supporting the hypothesis that there are increasing numbers of channels per stereocilia.

Calcium Permeability of Transducer Channel Coupling Ca2+ imaging experiments with electrical recordings has allowed for the demonstration of a direct relationship between Ca2+ entry and the rate of adaptation (Ricci and Fettiplace, 1998). That is, the rate of adaptation was directly proportional to the Ca2+ entering the stereocilia. This relationship might explain in part why the rate of adaptation is sensitive to the stimulus rise-time. The faster the rise-time, the greater the peak Ca2+ achieved in the stereocilia. Ca2+ can both block and permeate the transducer channel so that lowering external Ca2+ increased the total magnitude of the transducer current while decreasing the proportion of the current carried by Ca2+ (Crawford et al., 1991; Ricci and Fettiplace, 1998). It was further demonstrated that the monovalent ion could alter Ca2+ permeability, presumably through an anomalous mole fraction effect, so that more Ca2+ entered the stereocilia with K+ as the monovalent ion than with Na+. These surprising results might explain the necessity for the unusual composition of endolymph. Having a solution of high K+ and low Ca2+ will maximize the total current through the transducer channels while maintaining a high Ca2+ permeability for regulating adaptation (Ricci and Fettiplace, 1998). In addition, the positive endocochlear potential will increase the driving force for Ca2+ through the channel, thus speeding adaptation even further (Ricci and Fettiplace, 1998). Interestingly, similar types of ionic interactions have been described in saccule hair cells using a macroscopic measuring technique (Lumpkin, Marquis, and Hudspeth, 1997). However, quantitatively the reported Ca2+ permeabilities were quite different between end organs. These data might suggest variations in transducer channel permeability properties between hair cells operating over different frequency ranges. This type of mechanism is frequently used in other sensory systems such as the visual and olfactory as a means of regulating adaptation. A variation in transducer channel Ca2+ permeability might be a mechanism underlying the tonotopic changes in adaptation rate.

Ca2+ Feedback Predicted by Ca2+ Gradients How then do Ca2+ buffers alter adaptation? Increasing the concentration of the Ca2+ chelator BAPTA decreases the rate of adaptation and shifts the channel activation curve leftward, suggesting that less free Ca2+ is available for binding at the intraciliary site that triggers adaptation. Since Ca2+ is lower at this site, adaptation responds by opening more channels in order to restore Ca2+ to some set level. In this way, adaptation acts as a feedback loop regulating the intraciliary Ca2+ concentration. For this to occur,

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the adaptation trigger site must be at a distance away from the source of Ca2+ where the buffer can be effective. If this idea of Ca2+ feedback is accurate, then there is a position within the stereocilia where at steady state the Ca2+ remains constant regardless of the buffer concentration or the external Ca2+ concentration. Naraghi and Neher (1997) have demonstrated that a length constant can be defined for a given Ca2+ buffer. The length constant represents the distance from a source at which a buffer can be effective. The length constant is determined by several factors including, the forward rate constant (how fast it binds), the affinity (Kd) (how tightly it binds) and by its concentration (Naraghi and Neher, 1997). BAPTA is a fast high affinity buffer. Increasing BAPTA concentration will effectively shorten its length constant, making it more effective at altering adaptation. In a nonfeedback situation, a plot of Ca2+ concentration against distance from the source would have steeper profiles as the buffer concentration is elevated, so steady state levels would be achieved closer to the source. However, in a perfect feedback system, it would be predicted that the plots would intersect near the Ca2+ binding site position (see Figure 3–4). That is, the feedback system would operate to maintain Ca2+ at a constant level at the Ca2+ sensor, despite the difference in steepness of the gradient. A three-dimensional model of Ca2+ diffusion within the stereocilia was developed in order to test the feedback model and to predict the location of the Ca2+ binding site (Ricci et al., 1998). Having measured the shift in transducer activation curves caused by either changing external Ca2+ or by altering internal Ca2+ buffering, and having directly determined Ca2+ entry under different external Ca2+ conditions, allowed us to estimate the Ca2+ gradient down the stereocilia. Measured values for the dimensions of the stereocilia as well as for Ca-ATPases levels were used in the model (Sneary, 1988; Hackney et al., 1993; Tucker and Fettiplace, 1995; Yamoah et al., 1998). The model was capable of predicting the gradients in intraciliary Ca2+, demonstrating an intersection between gradients of different buffer concentrations at points between 20 to 40nm from the channel (see Figure 3–4). This supports the hypothesis that adaptation can be described as a feedback system, and also serves to define a region very close to the transducer channel for the Ca2+ binding domain.

ADAPTATION AS A MECHANICAL TUNING MECHANISM While investigating the effects of external Ca2+ and internal Ca2+ buffering on adaptation it was found that a proportion of cells demonstrated oscillations in the transducer currents that were Ca2+-dependent (Ricci et al., 1998) (Figure 3–5). The frequency of the oscillations varied with the initial fast time constant of adaptation and with characteristic frequency of the hair cell. This tonotopic variation in transducer oscillations suggests that


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Figure 3–4: Estimation of Ca2+ gradient down a single stereocilia for different buffer concentrations. The intersection site predicts the location of the Ca2+ sensor. That the profiles intersect supports the idea of a Ca2+-dependent feedback system regulating fast adaptation. This Figure is modified from Ricci et. al. (1998).

adaptation may provide an as yet uncharacterized tuning mechanism for hair cells (Ricci et al., 1998). The oscillations are also predicted by a Ca2+ feedback system regulating fast adaptation (Wu et al., 1999). Thus far we have demonstrated that the rate of fast adaptation is dictated by several factors including the size of the transducer current, the stimulus rise time, the intraciliary Ca2+ buffer, the extracellular Ca2+ concentration and the position along the papilla from which the cell was recorded. The tonotopic variations in adaptation rate led to the suggestion that adaptation may be a mechanical tuning mechanism for hair cells. This was supported by the observation of Ca2+-dependent oscillations in the transducer current. Manipulations of Ca2+ homeostasis has also allowed for the estimate of the endogenous Ca2+ buffer concentration as well as a prediction for the location of the Ca2+ binding site, between 20 to 40nm from

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Figure 3–5: Example of mechanical oscillations of a transducer current elicited by application of 50M external Ca2+. 1 M EGTA was the internal Ca2+ buffer. No oscillation was observed in higher concentrations of external Ca2+. These oscillations may underlie a mechanical tuning mechanism in hair cells. Imax for this cell was 1.3 nA.

the channel. Adaptation responses to alterations in Ca2+ homeostasis are consistent with the hypothesis that fast adaptation can be depicted as a Ca2+-dependent feedback system that is tightly coupled to channel opening.

MULTIPLE MECHANISMS OF ADAPTATION Present theories regarding the mechanism of transducer adaptation involve active motors, thought to be myosin isozymes, being attached to the transducer channels (Hacohen et al., 1989; Assad and Corey, 1992; Gillespie and Hudspeth, 1993; Gillespie et al., 1993; Hudspeth and Gillespie, 1994; Metcalf, Chelliah, and Hudspeth, 1994; Hasson, Heintzelman, Santos-Sacchi, Corey, and Mooseker, 1995; Gillespie, 1996; Gillespie, Hasson,


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Garcia, and Corey, 1996; Burlacu, Tap, Lumpkin, and Hudspeth, 1997; Gillespie, 1997; Gillespie and Corey, 1997; Steyger, Gillespie, and Baird, 1998). The motors move the channel up and down the stereocilia in a Ca2+dependent manner, increasing or decreasing tension on the channel and thus resetting the operating range of the transducer channels. Data in support of this model come from a variety of sources; however, direct evidence is limited largely due to the technical difficulties of the experiment (see Chapter 1 by Dr. Corey in this book). One basic problem with this model is that the kinetics of fast adaptation are too fast to be dictated by a myosin cycle. Previous measures of adaptation rates on which the myosin model is based are 1 to 2 orders of magnitude slower than those reported here for fast adaptation. This discrepancy has led us to investigate whether multiple adaptation processes might be involved. To date, there are several lines of data suggesting there might be multiple mechanisms of adaptation. None of the present data is direct or conclusive, nor does this body of data directly refute the myosin hypothesis. However, the growing body of data suggest the mechanisms involved in adaptation are more complex than previously thought.

Kinetics The decay of the transducer current during a longer test pulse is best fit by a double exponential, one having a time constant near 1 ms and the second with a time constant in the tens of milliseconds (Wu et al., 1999) (Figure 3–6). The relative proportion of slow to fast components increases as the magnitude of the stimulus grows, perhaps suggesting different operating ranges. This result is appealing because the second time constant is similar to that reported in other preparations and could be driven by a motor system. A single Ca2+ binding site model cannot predict this decay.

Ca2+ Buffers Another line of evidence supporting multiple mechanisms for adaptation is that particular Ca2+ buffers can differentially alter the rate of adaptation as compared to the steady-state open probability of the channel. As mentioned earlier, Naraghi and Neher (1997) have demonstrated that a length constant can be assigned to a given mobile buffer based upon the concentration, the forward rate constant (how fast it binds) and the affinity (how tightly it binds). For example, NitroBAPTA is a low affinity buffer with a forward rate constant similar to BAPTA. It would be predicted to be an effective buffer close to the Ca2+ source but ineffective farther away from the source. Comparing the effectiveness of this buffer with BAPTA showed that NitroBAPTA could alter the rate of adaptation but was ineffective at altering the activation curve for the channels, thus uncoupling these two

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Figure 3–6: Adaptation to larger stimuli induces a double exponential decay in the transducer current. The thicker solid line indicates the fit. Ca2+ buffer was 0.1 mM BAPTA. Imax was 1.5 nA. The second component increases as larger deflections of the hair bundle are employed (Wu et al., 1999).

processes (Ricci et al., 1998). This means perhaps that there are two sites regulating adaptation with the close site governing the rate of fast adaptation and the distant site dictating the open probability. Another test is to use a high affinity buffer with a slower rate constant. This buffer will be effective further from the channel but would be ineffective close to the channel. EGTA is such a buffer. Comparison of EGTA with BAPTA showed that EGTA was ineffective at altering the adaptation rate but was as effective as BAPTA at shifting the transducer activation curve (Ricci et al., 1998). This result is consistent with the notion that there are at least two processes involved in regulating adaptation, one close to the channel dictating the rate of adaptation and the other located farther away from the channel and regulating the activation curve. Alone the data are not conclusive. One drawback is that the two measurements are inherently different. The rate of adaptation is influenced by the dynamic change in Ca2+, while the position of the activation curve is a steady-state measurement. This difference


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may in part be responsible for the differential buffer sensitivity, where a difference in the Kd, a steady-state buffer property, alters the activation curve, not the rate of adaptation and the forward rate constant, a dynamic variable, alters the rate of adaptation and not the activation curve. Either interpretation is valid and important for understanding mechanisms involved in regulating adaptation.

PHARMACOLOGICAL EVIDENCE Phosphate Analogs Alter Slow but Not Fast Adaptation Several pieces of pharmacological data support the hypothesis of multiple adaptation mechanisms. Phosphate analogs interfere with the myosin cycle preventing the power stroke (Yamoah and Gillespie, 1996). Phosphate analogs have been used to block adaptation, in fact shifting the transducer activation curve to the right (Yamoah and Gillespie, 1996). In an effort to separate two components of adaptation, vanadate, a phosphate analog, and butyldiene monoxamine (BDM), a myosin antagonist, were used to try to block adaptation. Both compounds shifted the activation curve to the right; however, neither altered the fast component of adaptation (Wu et al., 1999). In addition, a single exponential curve better fit the current decay as compared to a double exponential, supporting the idea that multiple mechanisms of adaptation exist. These data suggest that myosin is not involved in fast adaptation but may be involved in a second slower component of adaptation.

Cyclic Nucleotides Cyclic nucleotides can alter the position of the transducer activation curve (Ricci and Fettiplace, 1997). Cyclic AMP shifts the activation curve to the right, requiring larger stimuli to activate the transducer channels. No effect was found on the rate of adaptation. The actions of cAMP were also found to be independent of Ca2+ (Ricci and Fettiplace, 1997). This suggests the cyclic nucleotide effect might be on the second component of adaptation.

Calmodulin And finally, calmodulin antagonists block the rate of adaptation but do not shift the transducer activation curve (Walker et al., 1993; Walker and Hudspeth, 1996). Together these data suggest that multiple mechanisms may be involved in adaptation. One mechanism appears to be tightly coupled to the transducer channel, located very close to the channel, while a second mechanism appears to be more distant.

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TWO-SITE FEEDBACK MODEL OF ADAPTATION Recently, we have developed a model of transducer adaptation that is meant to serve as a template in which to further investigate possible mechanisms involved in adaptation (Wu et al., 1999). The model is a two-site Ca2+ feedback model that uses the three-dimensional Ca2+ diffusion model as the starting point (Ricci et al., 1998). No assumptions regarding the mechanical properties or mechanisms underlying adaptation are included; rather, two simple generic Ca2+ regulated processes are described. Three simple reactions underlie the basic model. K1 Ca2+ + Bs ======= Ca2+Bs K-1 Here B is the binding site regulating adaptation, the subscript (s) referring to either the first or second site. Ca2+ entry through the transducer channels binds at the adaptation trigger sites (Bs). The concentration of Ca2+B regulates the kinetics of the conformational change of a modulating protein (M): (Ca2+ B)ka M ======= M* kb The position of the activation curve is dictated by the concentration of M* and scaled linearly as: Xa = I * M*I + I I where: and are constants and the M* concentration is integrated over each specified binding site region. Each of the two binding sites is set up similarly. The sum of the two sites controls the set point of Xa. Differences between the first and second site include: 1. The forward rate constant, ka, of the conformational change is set to match the difference in adaptation rates so that the first site is two orders of magnitude faster than the second site. 2. The Kd of the Ca2+ binding, k-1/k1, is dictated by the gradient of Ca2+ down the stereocilia and the location of the binding site within this gradient (Ricci et. al 1998). 3. The first site is located 20 to 40nm from the Ca2+ source, as indicated by the intersection of the Ca2+ gradients described earlier.


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The second site is located 150 to 200nm from the source and was set somewhat arbitrarily to give the best match to the data. 4. The operating range for each site overlapped but was set based on the relative proportion of slow vs. fast components, determined from the double exponential fit to the current decay for a series of different magnitude displacements. This relatively simple feedback model was capable of reproducing many of the characteristics of transducer adaptation. Figure 3–7 is an example of current responses produced by the model in response to increasing stimulus magnitude where 0.1 mM BAPTA was the buffer. These traces show a single exponential decay to small steps, with a slower second component becoming predominant when larger magnitude stimuli were used. The model was capable of reproducing a variety of properties previously measured for transducer adaptation including: 1. Double exponential decay of the current. 2. Sensitivity to Ca2+ buffer concentration, with increasing buffer concentrations slowing the rate of adaptation and shifting the activation curve leftward.

Figure 3–7: Two site Ca2+ feedback model current responses. Using a three dimensional profile of Ca2+ diffusion down the stereocilia predicts many of the measured properties of transducer adaptation. These traces illustrate that the model can reproduce the set of current responses to increasing displacement steps in a very similar manner to experimental data. Figure 7 is modified from Wu et. al. (1999).

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3. Sensitivity to external Ca2+, again lowering external Ca2+ slowed the rate of adaptation and shifted the plot of transducer activation leftward. 4. Sensitivity to the stimulus rise-time, so that slowing the risetime of stimulus slowed the rate of adaptation and shifted the activation curve. 5. A single site would not reproduce the measured responses. Interestingly, the model produced current traces remarkably similar to the vanadate experiments when the second site was removed. 6. Capable of generating oscillations in the transducer current that were Ca2+-dependent. Altering the kinetics of the fast component varied the frequency of the oscillations.

Implications Regarding a Mammalian Cochlea The ability of adaptation to work on the microsecond time scale suggests that it is capable of working at frequencies used in mammalian systems. That the stimulus rise time alters the rate of adaptation suggests that adaptation can be a tunable filter. The hair bundle and the transducer channels are not limited by the membrane time constant and so can provide high frequency input to hair cells. Is there a mechanical correlate to fast adaptation? Mechanical oscillations have been reported in hair cells (Crawford and Fettiplace, 1985; Benser et al., 1996). Also, movements correlating to adaptation have been described (Howard and Hudspeth, 1987; Howard and Hudspeth, 1988; Assad and Corey, 1992). Hair bundles would be required to move much smaller distances in order to alter transducer properties than would the cell body. These properties make fast adaptation a possible candidate to be the active process in mammalian cochlea. It does not rule out the importance of outer hair cell motility, but it does provide an alternative or additional hypothesis that does not have either the mechanical or electrical limitations of outer hair cell motility. One possibility, though completely speculative, is that the outer hair cell motility provides a bias for the hair bundle either pressing the bundle into the tectorial membrane or moving it away from the membrane, thereby providing a tonic offset to the hair bundle position. This role being slower would bypass the constraints outlined above.

FUTURE DIRECTIONS As our knowledge regarding transduction and adaptation increases, so do the complexities of the system. The realization that so little is really under-


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stood about the mechanisms regulating adaptation and transduction is both exciting and humbling. What are the mechanical correlates of fast adaptation? Is fast adaptation intrinsic to the channel, a conformational change in the channel or a separate molecule tightly coupled to the channel? Are there synergistic interactions between transducer channels? Might adaptation underlie some component of the active process in the mammalian ear? The molecular nature of the channel is still unknown. How the channel is tethered to the cytoskeleton or the membrane is unknown. Is there a biochemical regulation of transduction or adaptation? Cyclic nucleotide experiments suggest there is some modulation, but what is the mechanism of this response and where is the site of action? The list of hair cell properties that vary tonotopically is continually growing. Hair bundles vary in number and height of stereocilia. The size of the transducer current increases tonotopically, presumably by increasing the number of channels per stereocilia. The adaptation rate decreases tonotopically, presumably at least in part due to the change in number of channels per stereocilia. The concentration of endogenous Ca2+ buffer increases tonotopically both in the stereocilia and in the soma. The number of Ca2+ channels and Ca2+-activated potassium channels increases tonotopically. The number of Ca2+ hot-spots, presumably representing the number of synaptic release sites increases tonotopically. The distribution of subunits of the Bk-channel varies tonotopically (Jones et al., 1999). The presence of inward rectifying channels as well as delayed rectifying channels varies tonotopically as well (Art and Goodman, 1996; Goodman and Art, 1996b; Goodman and Art, 1996a). What are the signals dictating these numerous tonotopic variations found in so many aspects of signal processing in the ear? Are similar variations found in mammalian systems? At what point in development is the tonotopic template established? What are the signaling mechanisms? Can it be restored with new hair cell formation? Recent investigations regarding adaptation of the mechano-electric transducer channels have revealed a variety of new and exciting findings. Most importantly, adaptation may provide a mechanical tuning mechanism for hair cells. Secondly, it appears that multiple mechanisms may be involved in generating adaptation. Uncovering these mechanisms and understanding how these properties are varied tonotopically are issues for future research.

Acknowledgments: The data presented in this review were obtained in collaboration with Robert Fettiplace. Yu-Cherng Wu programmed the modeling experiments both for the Ca2+ gradients as well as for the two-site model. This work was supported by NIH grants DC-01362 to RF and DC-03896 to AJR as well as a Deafness Research Award to AJR.

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REFERENCES: Art, J. J., Crawford, A. C., Fettiplace R., & Fuchs P. A. (1982). Efferent regulation of hair cells in the turtle cochlea. Proceedings of the Royal Society of London Series: B Biological Sciences, 216, 377–384. Art, J. J., & Fettiplace, R. (1987). Variation of membrane properties in hair cells isolated from the turtle cochlea. Journal of Physiology (London), 385, 207–242. Art, J. J., & Goodman, M. B. (1996). Ionic conductances and hair cell tuning in the turtle cochlea. Annals of the New York Academy of Sciences, 781, 103–122. Art, J. J., Wu, Y. C., & Fettiplace, R. (1995). The calcium-activated potassium channels of turtle hair cells. Journal of General Physiology, 105, 49–72. Assad, J. A., & Corey, D. P. (1992). An active motor model for adaptation by vertebrate hair cells. Journal of Neuroscience, 12, 3291–3309. Benser, M. E., Marquis, R. E., & Hudspeth, A. J. (1996). Rapid, active hair bundle movements in hair cells from the bullfrog’s sacculus. Journal of Neuroscience, 16, 5629–5643. Bosher, S. K., & Warren, R. L. (1978). Very low calcium content of cochlear endolymph, an extracellular fluid. Nature, 273, 377–378. Burlacu, S., Tap, W. D., Lumpkin, E. A., & Hudspeth, A. J. (1997). ATPase activity of myosin in hair bundles of the bullfrog’s sacculus. Biophysical Journal, 72, 263–271. Corey, D. P., & Hudspeth, A. J. (1983). Kinetics of the receptor current in bullfrog saccular hair cells. Journal of Neuroscience, 3, 962–976. Crawford, A. C., Evans, M. G., & Fettiplace, R. (1989). Activation and adaptation of transducer currents in turtle hair cells. Journal of Physiology (London). 419, 405–434. Crawford, A. C., Evans, M. G., and Fettiplace, R. (1991). The actions of calcium on then mechano-electrical transducer current of turtle hair cells. Journal of Physiology (London), 434, 369–398. Crawford, A. C., & Fettiplace, R. (1978). Ringing responses in cochlear hair cells of the turtle (proceedings). Journal of Physiology (London), 284, 120P–122P. Crawford, A. C., & Fettiplace, R. (1980). The frequency selectivity of auditory nerve fibres and hair cells in the cochlea of the turtle. Journal of Physiology (London), 306, 79–125. Crawford, A. C., & Fettiplace, R. (1981). An electrical tuning mechanism in turtle cochlear hair cells. Journal of Physiology (London), 312, 377–412. Crawford, A. C., & Fettiplace, R. (1985). The mechanical properties of ciliary bundles of turtle cochlear hair cells. Journal of Physiology (London), 364, 359–379. Denk W., Holt J. R., Shepherd, G. M., & Corey, D. P. (1995). Calcium imaging of single stereocilia in hair cells: localization of transduction channels at both ends of tip links. Neuron, 15, 1311–1321. Eatock, R. A., Corey, D. P., & Hudspeth, A. J. (1987). Adaptation of mechanoelectrical transduction in hair cells of the bullfrog’s sacculus. Journal of Neuroscience, 7, 2821–2836. Fettiplace, R., & Fuchs, P. A. (1999). Mechanisms of hair cell tuning. Annual Review of Physiology, 61, 809–834. Gillespie, P. G. (1996). Deaf and dizzy mice with mutated myosin motors. Nature Medicine, 2, 27–29.


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Gillespie, P. G. (1997). Multiple myosin motors and mechanoelectrical transduction by hair cells. Biological Bulletin. 192, 186–190. Gillespie, P. G., & Corey, D. P. (1997). Myosin and adaptation by hair cells. Neuron, 19, 955–958. Gillespie, P. G., Hasson, T., Garcia, J. A., & Corey, D. P. (1996). Multiple myosin isozymes and hair-cell function. Cold Spring Harbor Symposium on Quantitative Biology, 61, 309–318. Gillespie, P. G., & Hudspeth, A. J. (1993). Adenine nucleoside diphosphates block adaptation of mechanoelectrical transduction in hair cells. Proceedings of the National Academy of Science, USA, 90, 2710–2714. Gillespie, P. G., Wagner, M. C., & Hudspeth, A. J. (1993). Identification of a 120 kd hair-bundle myosin located near stereociliary tips. Neuron, 11, 581–594. Goodman, M. B., & Art, J. J. (1996a). Positive feedback by a potassium-selective inward rectifier enhances tuning in vertebrate hair cells. Biophysical Journal, 71, 430–442. Goodman, M. B., & Art, J. J. (1996b). Variations in the ensemble of potassium currents underlying resonance in turtle hair cells. Journal Physiology (London), 497, 395–412. Hackney, C. M., Fettiplace, R., & Furness, D. N. (1993). The functional morphology of stereociliary bundles on turtle cochlear hair cells. Hearing Research, 69, 163–175. Hacohen, N., Assad, J. A., Smith, W. J., & Corey, D. P. (1989). Regulation of tension on hair-cell transduction channels: displacement and calcium dependence. Journal of Neuroscience, 9, 3988–3997. Hasson, T., Heintzelman, M. B., Santos-Sacchi, J., Corey, D. P., & Mooseker, M. S. (1995). Expression in cochlea and retina of myosin VIIa, the gene product defective in Usher syndrome type 1B. Proceedings of the National Academy of Sciences, USA, 92, 9815–9819. Howard, J., & Hudspeth, A. J. (1987). Mechanical relaxation of the hair bundle mediates adaptation in mechanoelectrical transduction by the bullfrog’s saccular hair cell. Proceedings of the National Academy of Sciences, USA, 84, 3064–3068. Howard, J., & Hudspeth, A. J. (1988). Compliance of the hair bundle associated with gating of mechanoelectrical transduction channels in the bullfrog’s saccular hair cell. Neuron, 1, 189–199. Hudspeth, A. J. (1982). Extracellular current flow and the site of transduction by vertebrate hair cells. Journal of Neuroscience, 2, 1–10. Hudspeth, A. J. (1989). How the ear’s works work. Nature 341, 397–404. Hudspeth, A. J., & Gillespie, P. G. (1994). Pulling springs to tune transduction: adaptation by hair cells. Neuron, 12, 1–9. Jaramillo, F., & Hudspeth, A. J. (1991). Localization of the hair cell’s transduction channels at the hair bundle’s top by iontophoretic application of a channel blocker. Neuron, 7, 409–420. Jones, E. M. C., Gray-Keller, M., and Fettiplace, R. (1999). The role of Ca2+ activates K+ channel splice variants in the tonotopic organization of turtle cochlea. Journal Physiology, 518, 653–665. Lumpkin, E. A., & Hudspeth, A. J. (1995). Detection of Ca2+ entry through mechanosensitive channels localizes the site of mechanoelectrical transduction in hair cells. Proceedings of the National Academy of Sciences, USA, 92, 10297–10301.

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Lumpkin, E. A., Marquis, R. .E., & Hudspeth, A. J. (1997). The selectivity of the hair cell’s mechanoelectrical-transduction channel promotes Ca2+ flux at low Ca2+ concentrations. Proceedings of the National Academy of Sciences, USA, 94, 10997–11002. Metcalf, A. B., Chelliah, Y., & Hudspeth, A. J. (1994). Molecular cloning of a myosin I beta isozyme that may mediate adaptation by hair cells of the bullfrog’s internal ear. Proceedings of the National Academy of Sciences, USA, 91, 11821–11825. Naraghi, M., & Neher, E. (1997). Linearized buffered Ca2+ diffusion in microdomains and its implications for calculation of Ca2+ at the mouth of a calcium channel. Journal of Neuroscience, 17, 6961–6973. Oberholtzer, J. C., Buettger, C., Summers, M. C., & Matschinsky, F. M. (1988). The 28k-Da calbindin-D is a major calcium-binding protein in the basilar papilla of the chick. Proceedings of the National Academy of Sciences, USA, 85, 3387–3390. Pickles, J. O., Brix, J., Comis, S. D., Gleich, O., Koppl, C., Manley, G. A., & Osborne, M. P. (1989). The organization of tip links and stereocilia on hair cells of bird and lizard basilar papillae. Hearing Research, 41, 31–41. Ricci, A. J., & Fettiplace, R. (1997). The effects of calcium buffering and cyclic AMP on mechanoelectrical transduction in turtle auditory hair cells. Journal of Physiology (London), 501, 111–124. Ricci, A. J., & Fettiplace, R. (1998). Calcium permeation of the turtle hair cell mechanotransducer channel and its relation to the composition of endolymph (published erratum appears in Journal of Physiology (London) 1998, Mar. 15, 507 (Pt 3): 939), 506, 159–173. Ricci, A. J., Wu, Y. C., & Fettiplace, R. (1998). The endogenous calcium buffer and the time course of transducer adaptation in auditory hair cells. Journal of Neuroscience, 18, 8261–8277. Slepecky, N., & Chamberlain, S. C. (1982). Actin in cochlear hair cells-implications for stereocilia movement. Archives of Oto-Rhino-Laryngology, 234, 131–134. Sneary, M. G. (1988). Auditory receptor of the red-eared turtle: I. General ultrastructure. Journal of Comparative Neurology, 276, 573–587. Steyger, P. S., Gillespie, P. G., & Baird, R. A. (1998). Myosin is located at tip link anchors in vestibular hair bundles. Journal of Neuroscience, 18, 4603–4615. Tilney, L. G., DeRosier, D. J., & Mulroy, M. J. (1980). The organization of actin filaments in the stereocilia of cochlear hair cells. Journal of Cell Biology, 86, 244–259. Tilney, L. G., Egelman, E. H., DeRosier, D. J., & Saunder, J. C. (1983). Actin filaments, stereocilia, and hair cells of the bird cochlea. II. Packing of actin filaments in the stereocilia and in the cuticular plate and what happens to the organization when the stereocilia are bent. Journal of Cell Biology 96, 822–834. Tucker, T., & Fettiplace, R. (1995). Confocal imaging of calcium micordomains and calcium extrusion in turtle hair cells. Neuron, 15, 1323–1335. Tucker, T., & Fettiplace, R. (1996). Monitoring calcium in turtle hair cells with a calcium-activated potassium channel. Journal of Physiology (London), 494, 613–626. Walker, R. G., & Hudspeth, A. J. (1996). Calmodulin controls adaptation of mechanoelectrical transduction by hair cells of the bullfrog’s sacculus. Proceedings of the National Academy of Sciences, USA, 93, 2203–2207. Walker, R. G., Hudspeth, A. J. & Gillespie, P. G. (1993). Calmodulin and calmodulin-binding proteins in hair bundles. Proceedings of the National Academy of Sciences, USA, 90, 2807–2811.


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Wu, Y. C., Ricci, A. J., & Fettiplace, R. (1999). Two components of transducer adaptation in auditory hair cells. Journal of Neurophysiology, 82, 2171–2181. Yamoah, E. N., & Gillespie, P. G. (1996). Phosphate analogs block adaptation in hair cells by inhibiting adaptation-motor force production. Neuron, 17, 523–533. Yamoah, E. N., Lumpkin, E. A., Dumont, R. A., Smith, P. J., Hudspeth, A. J., & Gillespie, P. G. (1998). Plasma membrane Ca2+ -ATPase extrudes Ca2+ from hair cell stereocilia. Journal of Neuroscience, 18, 610–624.

4 “Hearing” in a Primitive Animal Patricia Mire, Ph.D

Hair bundles on tentacles of sea anemones, primitive marine invertebrates, are similar in many respects to hair bundles in the acousticolateralis system of vertebrates. In both systems, hair bundles are composed of actin-based stereocilia interconnected distally by tip links and proximally by other linkages. The hair bundles are mechanoelectric transducers with an asymmetric sigmoidal response to graded stimuli. Transduction currents adapt during sustained hair bundle deflection and are sensitive to aminoglycoside antibiotics and calcium-free buffers. Anemone mechanotransduction also differs in some respects from that in vertebrates. Hair bundles in anemones arise from a multicellular complex and are radially symmetrical. Mechanoelectric responses are less sensitive to small deflections and completely adapt during sustained deflection. Hair bundles in anemones are morphodynamically modulated by chemoreceptors and self repair following damage. In this chapter, a model of signal transduction for anemone hair bundles is discussed. The model is a modification of the tip link model of transduction for vertebrate hair bundles that incorporates the atypical features of anemone hair bundles. Unique features of anemone hair bundles are presented as adaptations for functioning in prey detection in a noisy environment.

INTRODUCTION Sea anemones are primitive marine invertebrates that use hair bundles to detect potential prey (reviewed in Watson and Mire-Thibodeaux, 1994; Watson and Mire, 1999). At first glance, sea anemones appear to be unlikely subjects for research into hearing (Figure 4–1A). The tubular body col67

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umn has a mouth surrounded by one or more whorls of tentacles at one end and an adhesive foot attached to the substrate at the opposite end. The tentacles are used for capturing prey that is then ingested whole (Shick, 1991). Upon closer inspection, one finds numerous hair bundles located on the surface of the tentacles (Figure 4–1B). Protruding into the seawater, the hair bundles are poised to receive vibrations traveling through the water column. Of obvious significance to the anemone’s survival, is the detection of vibrations produced by nearby swimming prey. Anemone hair bundles are intriguing mechanoreceptors both in form and function. On the one hand, they are similar in many respects to hair bundles of the acousticolateralis system of vertebrates (Watson, Mire, and Hudson, 1997). On the other hand, they possess unique attributes that set them apart from typical vertebrate hair bundles (Mire and Watson, 1997). This chapter attempts to reconcile these similarities and differences into a working model of signal transduction in anemone hair bundles.

SIMILARITIES BETWEEN ANEMONE AND VERTEBRATE HAIR BUNDLES Actin-based Stereocilia Hair bundles in anemones are composed of actin-based stereocilia as are vertebrate hair bundles (Mire-Thibodeaux and Watson, 1994a; Watson and Roberts, 1995). Within stereocilia, most of the actin is in the filamentous form (F-actin), as demonstrated by confocal microscopy of tentacles incubated in fluorescently-tagged phalloidin, a compound that binds to the filamentous form of actin (Figure 4–2A). Within the apical cytoplasm of cells beneath hair bundles are pools of globular or monomeric actin (G-actin) as visualized in confocal micrographs of tentacles incubated with DNAse 1, which binds to the globular form of actin (Figure 4–2B). Filaments of approximately 7 nm diameter, consistent with F-actin, are routinely observed in TEM micrographs of anemone hair bundle stereocilia (Figure 4–2C). Treatment with cytochalasin, a compound that destabilizes F-actin (Cooper, 1987), obliterates hair bundles on anemone tentacles and renders the animals insensitive to vibrations (Watson and Hessinger, 1992). The finding that hair bundles of anemones and those of their relatives within the phylum Cnidaria are composed of an actin cytoskeleton is surprising for two reasons. Cnidarians are considered to be the simplest animals to possess a nervous system and are therefore phylogenetically far removed from vertebrates (Barnes, Calow, and Olive, 1993). Moreover, hair bundles in other invertebrate phyla are composed of microtubule-based kinocilia (Budelmann, 1988). Although the evolutionary significance of actin-based versus microtubule-based hair bundles is unclear, the fact that anemone

Figure 4–1: Close-up photograph and FESEM micrograph of an anemone. (A) Living sea anemone, Haliplanella lineata, in side view. (B) FESEM micrograph of the tentacle surface featuring numerous hair bundles. Scale bars = 2 mm in A and 10 µm in B.

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Figure 4–2: Micrographs showing actin in the tentacle epidermis. (A) Confocal image of phalloidin labeled hair bundles protruding from the tentacle surface. (B) Confocal optical section taken beneath the hair bundles of DNAse labeled tentacle epidermis. (C) TEM micrograph of hair bundle stereocilia in oblique section. Scale bar = 10 µm for A, B and 0.1 µm for C.

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hair bundles are actin-based makes anemone hair bundles all the more interesting for comparison to vertebrate hair bundles.

Extracellular Linkages Within hair bundles of anemones, stereocilia are interconnected by a variety of extracellular linkages similar to those seen interconnecting stereocilia of vertebrate hair bundles (discussed in detail in Watson et al., 1997). The linkages include a basal network of filaments, distal long lateral links and short cross links, and tip links (Figure 4–3). The tip links interconnect the tops of shorter stereocilia to the sides of adjacent longer stereocilia. Anemone tip links are of the approximate dimensions of vertebrate tip

Figure 4–3: High resolution FESEM micrograph of hair bundle in side view. Black arrowheads indicate extracellular linkages interconnecting stereocilia. Scale bar = 1 µm.

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links, but are bifurcated at the insertion to the shorter stereocilium instead of to the longer stereocilium as is the typical vertebrate case. Both in anemones and in vertebrates, distal linkages are better preserved with ruthenium red in the fixative, indicating that they are likely composed of glycosaminoglycans, at least in part (Slepecky and Chamberlain, 1985; Watson et al., 1997). The precise composition of linkages is not known for either anemones or vertebrates.

Mechanotransduction Hair cells in anemone tentacles are mechanoelectric transducers as are hair cells in vertebrate hearing organs (Mire and Watson, 1997). When anemone hair bundles are deflected with puffs of seawater delivered from a nearby pipette, transients in membrane currents are detected during loose-patch recording from hair cells. The current transients are graded with stimulus strength and either positively or negatively directed (Figure 4–4A). The positive currents are relatively small in amplitude, ranging from approximately 5 to 30 pA, and saturate abruptly to large deflections. The negative currents are larger in amplitude, ranging from approximately 10 to 150 pA, and require larger deflections to saturate. Thus, anemone mechanoelectric responses are asymmetrically sigmoidal in relation to stimulus strength as are other known mechanotransducers (Figure 4–4B). Anemone mechanotransducer currents also adapt to prolonged step deflection of the hair bundles as do vertebrate mechanotransducer currents. However, adaptation in anemones differs in some respects from adaptation in vertebrates and will be discussed in more detail below.

Pharmacology Pharmacological experiments indicate that mechanotransduction in anemones and in vertebrates exhibits similar sensitivities to several agents (Mire and Watson, 1997: Watson et al., 1997). Among the inhibitory agents are the aminoglycoside antibiotics, streptomycin and gentamicin, which reversibly abolish mechanotransduction at 10-4 M antibiotic (Figure 4–5A). Investigations in vertebrate hair cells suggest that these drugs inhibit mechanotransduction by plugging the transduction channels (reviewed in Hudspeth, 1992). Other treatments, such as brief exposure to elastase or to calcium-free buffers, also cause a loss of signal transduction both in anemones (Figure 4–5B) and vertebrates (Osborne and Comis, 1990; Assad, Shepherd, and Corey, 1991; Crawford, Evans, and Fettiplace, 1991; Preyer, Hemmert, Zenner, and Gummer, 1995; Watson, Mire, and Hudson, 1998); however, these treatments are thought to act on the linkages between stereocilia, specifically by severing tip links.

Figure 4–4: Mechanoelectrical response of hair cells to stimuli of graded strength. (A) Family of membrane currents obtained from loose-patch, in situ recording of hair cells during step deflections of the hair bundles. (B) Stimulus/response plot based on data from n=5–25 hair cells. Data points indicate mean peak current ± SEM.

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Figure 4–5: Loose-patch, in situ recordings of membrane currents from hair cells subjected to deflection of their hair bundles. Continuous traces are stacked with the sequence indicated by the number at the left margin. Seawater puff stimuli to deflect hair bundles are indicated by vertical bars superimposed onto the traces. (A) Effects of streptomycin on membrane currents. Puff stimuli containing streptomycin are indicated with an “S” placed beneath the bar. (B) Effects of calcium-free seawater on membrane currents. Calcium-free seawater perfusion begins at the arrow.

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DIFFERENCES BETWEEN ANEMONE AND VERTEBRATE HAIR BUNDLES Multicellular Complex Perhaps the most striking difference between anemone and vertebrate hair bundles is that each anemone hair bundle arises from a multicellular complex instead of arising from a single hair cell (Peteya, 1975). In the anemone case, a central sensory neuron is surrounded by 2 to 4 supporting cells (Figure 4–6A). The sensory neuron contributes 4 to 6 large diameter stereocilia and a single kinocilium to the center of the hair bundle (Figure 4–6B). Each supporting cell contributes approximately 50 to 100 small diameter stereocilia to the hair bundle periphery (Watson and Roberts, 1995). Although the sensory neuron makes afferent contact with the underlying nerve net (Figure 4–6C), the supporting cells are not thought to be in direct communication with the nerve net (Mire-Thibodeaux and Watson, 1994a).

Radial Symmetry Unlike vertebrate hair bundles which are bilaterally symmetrical, anemone hair bundles are radially symmetric (Figure 4–7). Stereocilia of each supporting cell are arranged in a staircase array with the shortest ones at the bundle periphery and the tallest ones near the center of the hair bundle. The tallest stereocilia of the supporting cells converge onto the large diameter stereocilia of the sensory neuron. Hence, supporting cells on opposite sides of the sensory neuron are mirror images of each other. The situation is complicated further by a twist superimposed onto the radial symmetry. Electrophysiological evidence suggests that the so-called “supporting cells” are the transducers of the primary receptor current (Mire and Watson, 1997). Indeed, it is from these “supporting cells” that the mechanically-induced, streptomycin-sensitive membrane currents described above are derived. Hence, although the name “supporting cell” has historical precedence, these cells appear to function in an analogous fashion to hair cells in vertebrate hearing organs. Like vertebrate hair cells, anemone “supporting cells” are non-neuronal in nature. Thus far, the sensory neurons of anemone hair bundles have eluded electrophysiological investigations. The location of the sensory neuron in the center of the hair bundle precludes loose-patch recording of membrane currents from these cells without simultaneously interfering with hair bundle displacement. Intracellular recording has not yet been successful due to the fragility and small size of the sensory neurons.

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Figure 4–6: Micrographs of multicellular complexes from which hair bundles arise. (A) TEM micrograph of complex in longitudinal section showing the sensory neuron (sn) surrounded by supporting cells (sc). (B) TEM micrograph of hair bundle in oblique section showing kinocilium (k) and large diameter stereocilia (ls) from the sensory neuron and small diameter stereocilia (ss) from supporting cells. (C) Stacked confocal optical sections of DiI labeled tentacle showing a sensory neuron (sn) with hair bundle apically and long processes basally extending to other neurons. Scale bar = 0.6 µm for A, B and 10 µm for C.

Sensitivity and Adaptation Electrophysiological data indicate that anemone mechanotransduction differs in at least two respects from vertebrate mechanotransduction. One difference is in sensitivity of the response. In anemone hair bundles step deflections of less than 1µm give no measurable responses (Mire and Watson, 1997). The mechanosensitive range of anemone hair bundles is from 1µm to 4 to 7 µm depending on whether the response is positive or negative (refer to Figure 4–4B). In vertebrate hair bundles, the entire mechanosensitive


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Figure 4–7: FESEM micrograph of hair bundle viewed from above. Scale bar = 1 µm.

range is less than a micrometer (reviewed in Hudspeth, 1989). The second difference is in adaptation of the response. Both in vertebrates and in anemones, transduction currents begin to return to baseline levels even while the hair bundle is deflected during prolonged stimulation (Figure 4–8A). However, in anemones, adaptation is complete such that the current reaches baseline (Figure 4–8B) and actually overshoots baseline (Figure 4–8C) when the hair bundle is deflected for 1 sec or more. In the vertebrate case, transduction currents incompletely adapt to prolonged hair bundle displacement such that the current does not fully return to baseline until cessation of the stimulus (reviewed in Corey and Assad, 1992; Hudspeth and Gillespie, 1994) .

Cell-cell Communication Within the Complex A dilemma presented by the multicellular nature of anemone hair bundles is how signaling by the cells is coordinated so that the complex functions

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Figure 4–8: Loose patch, in situ recordings of membrane currents from a hair cell during hair bundle deflections of equal strength but increasing duration. Durations of stimuli indicated by the black bars beneath the corresponding current traces.

as a mechanoreceptive unit during vibration detection. Part of the answer may be that the cells of the complex communicate through gap junctions. Preliminary evidence using a modification of the FRAP (Fluorescence Recovery After Photobleaching) approach indicates that epidermal cells directly below hair bundles are dye coupled (Figure 4–9). Moreover, dye coupling appears to be enhanced when the overlying hair bundle is deflected with a puff of seawater from a nearby pipette. For these experiments, whole tentacles are loaded with Calcein, a gap junction permeable fluorescent dye, threaded onto fine human hair strands, and then viewed microscopically.


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Figure 4–9: Photomicrographs of the tentacle epidermis during a FRAP experiment monitoring Calcein fluorescence. (A) Transmitted light and (B–F) epifluorescence images after photobleaching. (A1) Contour plot of initial fluorescence before photobleaching. (B1–F1) Corresponding contour plots after photobleaching showing the upper 20 % of gray levels indicated in white. Arrowheads indicate the position of the base of a hair bundle that is deflected immediately before image E in the series. Scale bar = 15 µm.

Using transmitted light, a hair bundle is centered in the field of view of a cooled CCD camera. Using epifluorescence, a first image is taken of the epidermal area before photobleaching. A second image is taken after the area has been photobleached for approximately 5 min, at which point gray levels have been reduced by approximately 50 %. A third, fourth, and fifth image are taken at 1 min intervals to monitor passive recovery of fluorescence. The hair bundle protruding from the epidermis is then deflected with a puff of seawater from a nearby micropipette. A sixth image is taken immediately after the deflection, then a seventh and an eighth image at 1 min intervals. Preliminary cytochemical evidence suggests that the gap junctions are composed of a connexin 43-like protein. Using a commercially available antibody to a mouse connexin 43 protein and a fluorescently-labeled

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secondary antibody, immunofluorescence is detected in cells giving rise to hair bundles (Figure 4–10). Although gap junctions are known to occur in vertebrate hearing and balance organs, including junctions between supporting cells and hair cells, the precise function of the gap junctions in these organs is not yet understood (Dunn and Morest, 1975; Oesterle, Cunningham, and Rubel, 1992; Kikuchi, Kimar, Paul, and Adams, 1995; Masuda, Yamazaki, Kanzaki, and Hosoda, 1995). However, gap junctions are obviously crucial to hearing in humans as certain forms of nonsyndromic deafness have been linked to mutations in connexin genes (reviewed in Simon and Goodenough, 1998; Simon, 1999).

Chemoreceptor-induced Morphodynamics A remarkable feature of anemone hair bundles is that they are morphodynamically tunable (Watson and Hessinger, 1989, 1992, 1994; Mire-Thibodeaux and Watson, 1994a). Morphological adjustments to the hair bundles are regulated by the activity of two classes of chemoreceptor located on the apical surface of supporting cells. When receptors for N-acetylated sugars are activated, the hair bundles elongate from 1 to 2 µm and vibration sensitivity shifts to lower frequencies and smaller amplitudes. Subsequent activation of receptors for certain amino acids results in shortening of the hair bundles to control lengths and a shifting of vibration sensitivity to higher frequencies and larger amplitudes (Watson and Hudson, 1994). Sugar-induced elongation of the hair bundles involves activation of a cAMP second messenger pathway resulting in polymerization of actin (Watson and Hessinger, 1992; Mire-Thibodeaux and Watson, 1994b; Watson and Roberts, 1995). Recent evidence suggests that chemoreceptor activation may induce more widespread adjustments to the hair bundle in addition to the known effects on actin dynamics. Loose-patch recordings show an absence of mechanically induced membrane current transients from a previously responsive cell within 5 min perfusion of 10-7 M N-acetylneuraminic acid (NANA) to the bath (Figure 4–11). The hair cell is still unresponsive at 15 min in NANA alone. However, if repair proteins (RP, see the following paragraph) are added to the bath, spontaneous transients in membrane current are detected at 5 min, even though no mechanically-induced responses occur at this time point. At 25 min after the addition of RP, mechanically-induced membrane current transients are restored, but, surprisingly, the transients have reversed polarity. Upon washout of NANA and RP, the hair cell once again becomes mechanically unresponsive, although small spontaneous transients occur both before and after addition of RP. At 20 min after addition of RP, mechanically-induced transients are restored but have returned to their original polarity. Line profiles of gray values taken across live hair bundles imaged with video-enhanced light

Figure 4–10: Confocal micrographs of connexin 43 immunofluorescence in the tentacle epidermis. (A) Optical section showing hair bundles (arrowheads) in profile. (B) Apical optical section of epidermal cells beneath hair bundles. Double arrowhead indicates one of many examples of punctate fluorescence. (C) Deeper optical section showing patch of labeled cells at the region of nerve net. Double arrowhead indicates a string of fluorescent dots which appear to lie at the periphery of the soma. Scale bar = 5 µm in A, B and 7 µm in C.

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Figure 4–11: Loose patch, in situ recordings of membrane currents from a hair cell during hair bundle deflections of equal strength and duration. Continuous traces are stacked within the bold horizontal lines with the sequence indicated at the left margin. Treatments (indicated at the right margin) included perfusion of seawater alone (sw controls), after the perfusion of N-acetylneuraminic acid (NANA) alone at times indicated, after the perfusion of N-acetylneuraminic acid with added repair proteins (NANA RP) at times indicated, washout with seawater (sw washout), and after perfusion of seawater with added repair proteins (sw RP) at the times indicated.

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microscopy indicate that sugar stimulation induces reorientation of the hair bundles in addition to inducing elongation. The NANA induced reorientation appears to be especially pronounced at the middle and bases of the hair bundles (Figure 4–12). We recently began using FESEM to look more closely at the morphological changes to the hair bundles associated with activation of the NANA receptor. Our preliminary FESEM evidence

Figure 4–12: Line profiles of morphological changes in a live hair bundle with chemoreceptor activation. Plots show gray levels across the diameter (in pixels) of a hair bundle viewed from the side at the tip (top), middle (center), and base (bottom) in seawater alone (left) and 5 min after addition of N-acetylneuraminic acid (NANA) (right).

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supports our observations with light microscopy on live hair bundles that substantial reorientation of the hair bundle occurs with NANA stimulation. In particular, there appear to be changes in the positions of small diameter stereocilia relative to the long axis of the hair bundle (Figure 4–13). Obviously, chemoreceptor modification of hair bundle structure and function is more complicated than previously appreciated and will be the focus of further investigation.

Protein-mediated Repair Following Trauma Structural integrity of hair bundles can be disrupted by a variety of treatments including exposure to calcium-free buffers in both anemones and vertebrate hearing systems (Watson et al., 1997, 1998). In chickens, some function is restored to damaged hair cells within 12 hours following a mild trauma, i.e., 10 min exposure to calcium-free buffer (Zhao, Yamoah, and Gillespie, 1996). In this case, the hair bundles are thought to be repaired by utilizing pre-existing linkages because the process does not require the synthesis of new proteins. In anemones that have been exposed to calcium-free seawater for 1 hr to cause extensive damage to hair bundles, normal hair bundle morphology and vibration sensitivity recover after 4 hr of returning the animals to calcium-containing seawater (Watson et al., 1998). In the anemone case, repair of hair bundles requires the synthesis of new proteins (refer to Chapter 2 by G.M. Watson for details and an update of the repair process in anemone hair bundles).

THE WORKING MODEL Modification of the “Tip Link Model” to Anemone Hair Bundles One model for how vertebrate hair bundles transduce mechanical signals into electrical responses is the “tip link model” (Pickles, Comis, and Osborne, 1984; Howard, Roberts, and Hudspeth, 1988). According to this model, fine linkages connecting the tips of shorter stereocilia to the sides of adjacent, longer stereocilia called tip links are directly connected at one or both ends to transduction channels embedded in the membrane of one or both stereocilia. When hair bundles are deflected in the positive direction, that is, toward the taller stereocilia, tip links are strained and pull open the transduction channels. The transduction channels allow a cation influx that leads to depolarization of the hair cell. When hair bundles are deflected in the negative direction, toward the shorter stereocilia, tip links are slack allowing transduction channels to close. Closure of all transduction channels results in a small hyperpolarization of the hair cell. The hair cell hyperpolarizes because there is a slight amount of tension exerted on tip links even when the bundle is at rest, such that approximately 15 % of the

Figure 4–13: FESEM micrographs of hair bundles viewed from the side. (A) A hair bundle from an anemone fixed after exposure to seawater alone. (B) A hair bundle from an anemone fixed after 5 min exposure to N-acetylneuraminic acid. Arrows indicate the general orientation of distal regions of small diameter stereocilia. Scale bar = 1 µm.

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transduction channels are open resulting in a slight leak current. Lateral deflections of the hair bundle, that is, perpendicular to a line bisecting the bundle, have no measurable effect on transduction current. We have developed a model for mechanotransduction of anemone hair bundles that borrows from the tip link model while also incorporating both the similarities and differences to vertebrate hair bundles (Figure 4–14). In our model, “supporting cells” on opposite sides of the complex are oppositely polarized hair cells (Mire and Watson, 1997). When the hair bundle is deflected, tip links on one side of the bundle are stretched pulling open transduction channels while tip links on the opposite side of the bundle are slack allowing transduction channels to close. Thus, on one side of the complex, the hair cell depolarizes due to cation influx while on the opposite side of the complex the hair cell hyperpolarizes due to cessation of the leak current. Since the hair bundles are free standing, they resonate to vibrations of particular frequencies, which is determined in part by their lengths. When

Figure 4–14: Cartoon diagram of the model for signal transduction in the multicellular complex of a hair bundle. When the hair bundle is at rest (middle), the cells are at resting membrane potential (white). As the hair bundle oscillates, hair cells on opposite sides of the complex alternately depolarize (black) and hyperpolarize (gray). During the depolarization phase, cations (small black dots) pass through gap junctions from the hair cells to the sensory neuron.


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a hair bundle oscillates to vibrations at stimulatory frequencies, the hair cells of the complex alternately depolarize and hyperpolarize. The depolarization component is larger than the hyperpolarization component resulting in a net depolarization of the hair cells. Cations pass from the hair cells to the sensory neuron through gap junctions. The sensory neuron integrates the signal and eventually fires an action potential to send the message through the nerve net to effector cells downstream.

Special Mechanisms for Dealing With a Noisy Environment and Dynamic Nature In contrast to the relatively quiet and protected environment of cochlear or saccular hair bundles, anemone hair bundles function in an extremely noisy and volatile environment. The seawater into which the hair bundles protrude contains many living and nonliving sources of vibrations. The anemones typically live in the intertidal zone and therefore are exposed to substantial wave action. The water column contains vibrations from a variety of moving, living creatures some of which are significant to the anemone, such as potential prey. The anemones themselves generate “noise”. The tentacle epidermis contains a diverse array of cells in addition to the multicellular complexes giving rise to hair bundles. Interspersed with hair bundles are motile cilia that arise from certain epidermal cells. These cilia frequently contact hair bundles in whip-like fashion with sufficient force to cause small deflections. Many of the ways in which hair bundle mechanotransduction in anemones differs from that in vertebrates may be adaptations to their noisy environment. Reduced sensitivity of the hair bundle response is obviously beneficial if it is to function in the presence of background vibrations. For example, reduced sensitivity would be advantageous when a hair bundle is being “whipped” by nearby beating cilia. The deflections produced from contact with the motile cilia are periodic, but relatively small, and thus could go undetected by the hair bundle. In theory, multicellularity may allow the hair bundle to filter out random, inconsistent signals. In this case, primary signals coming from alternately depolarizing hair cells of the complex would have to be of sufficient periodicity and duration to depolarize the sensory neuron before signals would be propagated through the nerve net. However, the anemones have an acute detection of specific frequencies. Previous studies indicate that if vibrations occur at stimulatory frequencies, very few cycles of vibration are necessary to elicit an organismal response (Watson, Venable, and Mire, 2000). Adaptation of the transduction current would be beneficial to reset the hair cells when hair bundles are deflected for prolonged durations by stimuli that ultimately may be ignored. For anemones, such stimulation could come in the form of objects pressing against the hair bundles, such

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as pieces of rocks or shells carried by the water currents. Complete adaptation of the transduction current would not only diminish the signal from prolonged deflections, as is the case with incomplete adaptation, but actually negate the signal. Although the mechanism behind complete adaptation is unknown at this time, it may be due, at least in part, to the relatively high calcium concentration in seawater (on the order of 12 mM in seawater compared to approximately 50 µM in endolymph of the cochlea). In vertebrate hair cells, adaptation is sensitive to extracellular calcium concentration. For example, increasing the calcium concentration 0.1 to 1 mM increases the rate and extent of adaptation in turtle hair cells (reviewed in Fettiplace, 1992). However, more recent studies indicate that the mechanism behind adaptation is likely to be more complicated than a simple, direct calcium regulation (see Chapters 1 and 3 contributed by D.P. Corey and A. Ricci respectively, for recent progress on adaptation mechanisms). Some of the differences between anemone hair bundles and vertebrate hair bundles are probably due to differences in the physiological function of the hair bundles. Anemone hair bundles detect vibrations produced by swimming prey. Radial symmetry likely bestows multidirectional sensitivity to the hair bundle. This ability would be especially crucial in the detection of vibrations produced by swimming prey, which could approach from any direction in the water column. Hair bundles possessing radial symmetry and arising from a multicellular complex could have the added benefit of detecting direction while at the same time being multidirectionally sensitive. When the hair bundle oscillates, the positions of the signaling hair cells could supply information to the complex about the location of the vibrational source. The multicellular complex of anemone hair bundles must work at a frequency relevant to detecting meaningful vibrations. Gap junctional communication between the hair cells and the sensory neuron affords an efficient and rapid method of signal transmission. Critical to the detection and capture of prey is an elegant modulation of the hair bundles by chemoreceptors. The sugars that tune hair bundles to lower frequencies are found coating the surface of prey. Swimming prey produce vibrations corresponding to the lower frequencies to which the hair bundles tune upon activation of the sugar receptor. Hence, as prey approaches the anemone, sugars that diffuse from the prey surface activate chemoreceptors that in turn cause the hair bundles to elongate. The longer hair bundles oscillate to vibrations produced by swimming prey. Signaling by the complexes is communicated to effector cells downstream to predispose nematocysts to discharge if the prey contacts the tentacle. Nematocysts are large (approximately 15 by 5 µm) secretory products consisting of a capsule and an eversible tubule. During discharge the tubule rapidly everts (Holstein and Tardent, 1984) penetrating the prey to inject potent toxins that cause paralysis. In addition to their effects on prey, discharging nematocysts may also damage nearby hair bundles (P. Mire, personal ob-


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servation). Thus, during tuning of hair bundles and nematocyst discharge the hair bundles are likely in peril of structural damage. The degree of twisting of the hair bundles may be a mechanism to minimize destruction of linkages during elongation. In addition, the ability to self-repair may rescue damaged hair bundles from what would otherwise be fatal, structural damage both during the detection and capture of prey.

Acknowledgments: I thank Jason Nasse for assistance with FESEM and Glen Watson for assistance with TEM. This work is supported by The National Science Foundation, Grant number IBN9807782.

REFERENCES Assad, J. A., Shepherd, G. M. G., & Corey, D. P. (1991). Tip link integrity and mechanotransduction in vertebrate hair cells. Neuron, 7, 985–994. Barnes R. S. K., Calow P., & Olive P. J. W. (1993). The Invertebrates: A New Synthesis. (p. 488). Cambridge, MA; Blackwell Science. Budelmann, B. U. (1988). In J. Atema, R. R. Fay, A. N. Popper, & W. N. Tavolga (Eds.), Sensory Biology of Aquatic Animals (pp. 757–782). New York: Springer. Cooper, J. A. (1987). Effects of cytochalasin and phalloidin on actin. Journal of Cell Biology, 105, 1473–1478. Corey, D. P., & Assad, J. A. (1992). In D. P. Corey & S. D. Roper (Eds.), Sensory Transduction, (pp. 325–342). New York: Rockefeller University Press. Crawford, A. C., Evans, M. G., & Fettiplace, R. (1991). The actions of calcium on the mechano-electrical transducer current of turtle hair cells. Journal of Physiology, 434, 369–398. Dunn, R. A., & Morest, D. K. (1975). Receptor synapses without synaptic ribbons in the cochlea of the cat. Proceedings of the National Academy of Sciences, USA, 72, 3599–3603. Fettiplace, R. (1992). In D. P. Corey, & S. D. Roper (Eds.), Sensory Transduction (pp. 341–356). New York: Rockefeller University Press. Holstein, T., & Tardent, P. (1984). An ultrahigh-speed analysis of exocytosis: nematocyst discharge. Science, 223, 830–833. Howard, J., Roberts, W. M., & Hudspeth A. J. (1988). Mechanoelectrical transduction by hair cells. Annual Review of Biophysics and Biophysical Chemistry. 17, 99–124. Hudspeth, A. J. (1989). How the Ears Work. Nature, 341, 397–404. Hudspeth, A. J. (1992). In D. P. Corey & S. D. Roper (Eds.), Sensory Transduction (pp 357–370). New York: Rockefeller University Press. Hudspeth, A. J., & Gillespie, P. G. (1994) Pulling strings to tune transduction: adaptation by hair cells. Neuron 12, 1–9. Kikuchi, T., Kimar, R. S., Paul, D. L., & Adams, J. C. (1995). Gap junctions in the rat cochlea: immunohistochemical and ultrastructural analysis. Anatomical Embryology, 191, 101–118.

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Masuda, M., Yamazaki, K., Kanzaki, J., & Hosoda, Y. (1995). Ultrastructural evidence of cell communication between epithelial dark cells and melanocytes in vestibular organs of the human inner ear. Anatomical Record, 242, 267–277. Mire, P., & Watson, G. M. (1997). Mechanotransduction of hair bundles arising from multicellular complexes in anemones. Hearing Research, 113, 224–234. Mire-Thibodeaux, P., & Watson, G. M. (1994a). Morphodynamic hair bundles arising from sensory cell/ supporting cell complexes frequency-tune nematocyst discharge in sea anemones. Journal of Experimental Zoology, 268, 282–292. Mire-Thibodeaux, P., & Watson, G. M. (1994b). Cyclical morphodynamics of hair bundles in sea anemones: second messenger pathways. Journal of Experimental Zoology, 270, 517–526. Oesterle, E. C., Cunningham, D. E., & Rubel, E. W. (1992). Ultrastructure of hyaline, border, and vacuole cells in chick inner ear. Journal of Comparative Neurology, 318, 64–82. Osborne, M. P., & Comis, S. D. (1990). Action of elastase, collagenase and other enzymes upon linkages between stereocilia in the guinea pig cochlea. Acta Otolaryngology, 110, 37–45. Peteya, D. J. (1975). The ciliary-cone sensory cell of anemones and cerianthids. Tissue and Cell, 7, 243–252. Pickles, J. O., Comis, S. D., & Osborne, M. P. (1984). Cross-links between stereocilia in the guinea pig organ of Corti, and their possible relation to sensory transduction. Hearing Research, 15, 103–112. Preyer, S., Hemmert, W., Zenner, H. P., & Gummer, A. W. (1995). Abolition of the receptor potential response of isolated mammalian outer hair cells by hairbundle treatment with elastase: a test of the tip-link hypothesis. Hearing Research, 89, 187–193. Shick, J. M. (1991). A Functional Biology of Sea Anemones. New York: Chapman and Hall. Simon, A. M. (1999). Gap junctions: more roles and new structural data. Trends in Cell Biology, 9, 169. Simon, A. M., & Goodenough, D. A. (1998). Diverse functions of vertebrate gap junctions. Trends in Cell Biology, 8, 477–483. Slepecky, N., & Chamberlain, S. C. (1985). The cell coat of inner ear sensory and supporting cells as demonstrated by ruthenium red. Hearing Research, 17, 281–288. Watson, G. M., & Hessinger, D. A. (1992). Receptors for N-acetylated sugars may stimulate adenylate cyclase to sensitize and tune mechanoreceptors involved in triggering nematocyst discharge. Experimental Cell Research, 198, 8–16. Watson, G. M., & Hessinger, D. A. (1989). Cnidocyte mechanoreceptors are tuned to the movements of swimming prey by chemoreceptors. Science, 243, 1589–1591. Watson, G. M., & Hessinger, D. A. (1994). Antagonistic frequency tuning of hair bundles by different chemoreceptors regulates nematocyst discharge. Journal of Experimental Biology, 187, 57–73. Watson, G. M., & Hudson, R. R. (1994). Frequency and amplitude tuning of nematocyst discharge by proline. Journal of Experimental Zoology, 268, 177–185.


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Watson, G. M., & Mire, P. (1999). A comparison of hair bundle mechanoreceptors in sea anemones and vertebrate systems. Current Topics in Developmental Biology, 43, 51–84. Watson, G. M., & Mire-Thibodeaux, P. (1994). The cell biology of nematocysts. International. Review of Cytology, 156, 275–300. Watson, G. M., Mire, P., & Hudson, R. R. (1997). Hair bundles of sea anemones as a model system for vertebrate hair bundles. Hearing Research, 107, 53–66. Watson, G. M., Mire, P., & Hudson, R. R. (1998). Repair of hair bundles in sea anemones by secreted proteins. Hearing Research, 115, 119–128. Watson, G. M., & Roberts, J. (1995). Chemoreceptor-mediated polymerization and depolymerization of actin in hair bundles of sea anemones. Cell Motility and Cytoskeleton, 30, 208–220. Watson, G. M., Venable, S., & Mire, P. (2000). Rhythmic sensitization of nematocyst discharge in response to vibrational stimuli. Journal of Experimental Zoology, 286 (in press). Zhao, Y., Yamoah, E. N. & Gillespie, P. G. (1996). Regeneration of broken tip links and restoration of mechanical transduction in hair cells. Proceedings of the National Academy of Science, USA, 94, 15469–15474.

5 Brn-3.1 is Required for Development and Survival of the Hair Cell Allen F. Ryan, Ph.D.

TRANSCRIPTION FACTORS AND DEVELOPMENTAL CONTROL The development of individual cell types is determined by selective expression of different combinations of genes. Critical to this selective control are transcription factors (TFs), which regulate gene expression by binding to specific DNA sequences present in the regulatory elements of each gene, in a combinatory code. Different combinations of TFs lead to differences in gene expression. Of particular interest in developmental biology are TFs encoded by so-called “master regulatory genes.” These genes control major aspects of the development of different cells, tissues or organs. Many such master regulatory elements have been identified. They are exemplified by the factors MyoD and Myf5, that play major roles in the development of muscle cells (Rudnicki et al., 1993). The POU-domain TFs contain several family members that have been identified as master regulatory factors for different cell types (Ryan and Rosenfeld, 1997). This TF family is identified by a homeodomain (HD) similar to that encoded by other homeobox genes, paired with a second homologous domain, the POU-specific domain, that is unique to this family. The POU-HD is necessary for high-affinity DNA binding, while the POU-specific domain appears to required for the sequence specificity of DNA binding. A member of the POU-domain TF family, Pit- 1, is associated with the development of a small number of cell types in the anterior pituitary, and mutations in this gene are responsible for several forms of pituitary dwarfism (Li et al., 1990). 93

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UNC-86: ANCESTOR OF A HAIR CELL REGULATORY FACTOR? The POU-domain factor Unc-86 serves as a master regulator for the development of a number of neuronal phenotypes in the nematode C. elegans. These include the mechanosensory neurons responsible for the sense of touch (Gu, Caldwell, and Chalfie, 1996). This is of particular relevance for the development of the inner ear. Via an intermediary LIM-HD factor, Mec-3, Unc-86 controls the expression of two proteins that are thought to be elements of the mechanosensory receptor channel of these neurons (Tavernarakis and Driscoll, 1997). Moreover, mechanoreceptor neurons similar to those of C. elegans have been proposed as possible evolutionary ancestors of hair cells (Jorgensen, 1989). It therefore seemed possible that related TFs are involved in the development of the hair cell. Three orthologues of Unc-86 have been identified in mammals: Brn-3.0, Brn-3.1 and Brn.3.2 (also known as Brn-3a, Brn-3c and Brn-3b, respectively). Initial studies of Brn-3.0 found that it was expressed in developing inner ear ganglion cells, but not hair cells (Ryan, Simmons, and Crenshaw, 1991). When the additional orthologues Brn-3.1 and Brn-3.2 were cloned, hybridization histochemistry revealed that while Brn-3.2 was also expressed in inner ear ganglion cells, Brn-3.1 expression in the inner ear was limited to hair cells (Ryan, Luo, McEvilly, Erkman, and Rosenfeld, 1996).

BRN-3.1 EXPRESSION IN HAIR CELLS: BIRTH TO DEATH Developmentally, Brn-3.1 expression was first noted in vestibular hair cell precursors of the mouse on embryonic day 12.5 (e 12.5) and in the rat on embryonic day 16.5. In the cochlea, expression was first seen in the mouse basal turn on e 14.5 (Ryan, 1997), and in the rat on embryonic day 17.5 (Erkman et al. 1996). In e 14.5 mouse and e 17.5 rat cochlea, hair cell precursors cannot be distinguished from adjacent cells based on morphological or positional characteristic. However, at the earliest stages of expression, Brn-3.1 mRNA appeared to be present only in hair cell precursors, since the inner and outer hair cell precursors were clearly separate, and other nearby cells fated to become supporting cells were not labeled with Brn-3.1 probes. However, at this earliest stage, weaker label sometimes appeared between the hair cell precursor cell body, in the upper organ of Corti, and the basilar membrane (Figure 5–1). This observation may mean that expression of Brn-3.1 is initiated prior to separation of the hair cell precursor from the basement membrane, although brief expression in an underlying supporting cell cannot be ruled out.

BRN-3.1 IS REQUIRED FOR DEVELOPMENT AND SURVIVAL OF THE HAIR CELL

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Figure 5–1. In situ hybridization showing expression of Brn-3.1 mRNA in the organ of Corti of the e18.5 rat. In the basal turn (Base), expression is seen in both inner and outer hair cell precursors. In the lower middle turn (Mid) only one precursor is labeled, while none is labeled in the apex (Apex). In the middle turn, weaker expression extends in a band from the labeled hair cell precursor to the basilar membrane (arrow). This suggests that the cell has not yet released from the epithelial basement membrane, or that expression occurs in an underlying cell.

Unlike the temporal expression pattern of many developmental genes, strong expression of Brn-3.1 continued in auditory and vestibular hair cells into adulthood (Figure 5–2). While Brn-3.1 was expressed in a few other neurons during embryonic development, hair cells were the only postnatal cells that expressed this factor. The timing of hair cell expression, and the specific and continued production in hair cells throughout life, suggested that Brn-3.1 might represent a master regulatory gene for this cell type.

DELETION OF BRN-3.1 PREVENTS HAIR CELL DEVELOPMENT A targeted deletion of Brn-3.1 was performed by Erkman et al. (1996) and Xiang et al. (1997, 1998) using homologous recombination. When bred to homozygosity, animals null for Brn-3.1 were characterized by failure of hair cell development. Hair cell precursors never exhibited cuticular plates, stereocilia, or other morphological characteristics of this cell type. By birth, the precursor cells had largely lost their position in the upper half of the organ of Corti, and were virtually indistinguishable from other cells

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Figure 5–2. Expression of Brn-3.1 mRNA in hair cells of the utricular (Ut) and sacular (Sa) maculae of the adult mouse.

in the sensory epithelium. At postnatal day 7, the organ was highly disorganized, and the supporting cells of the organ of Corti had failed to develop their characteristic morphologies. It was not possible to identify hair cell or supporting cell types in the organ of Corti of Brn-3.1 null mice (Figure 5–3). In the vestibular sensory epithelia, failure of hair cell differentiation was similarly absent. However, accessory structures such as the cupulae, otolithic membrane and otoconia appeared normal. The inner ear defect was the only abnormality noted in Brn-3.1 null mice. Spiral ganglion neurons were normal in Brn-3.1 null mice at birth. However, they exhibited rapid degeneration, especially in the middle


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Figure 5–3. Disorganization in the organ of Corti (oC) of a mouse null for Brn-3.1 at postnatal day 14. While the tectorial membrane (TM), basilar membrane (BM), spiral limbus (SLim) and spiral ligament (SLig) are normal, none of the cell types normally present in the sensory epithelium is recognizable.

cochlear turn, until most spiral ganglion neurons were lost by 14 days after birth. Similarly, vestibular sensory neurons were initially present but were lost in the perinatal period.

THE HUMAN HOMOLOGUE OF BRN-3.1 IS A DEAFNESS GENE When the deletion of Brn-3.1 was performed, the genomic locus of the Brn-3.1 gene in mice suggested that it might be a candidate for the human nonsyndromic hereditary deafness DFNA 1, at 5q3I (Erkman et al., 1996). However, subsequent fine mapping indicated that the Brn-3.1 (POU4F3) locus in humans was about 2 cm away from DFNA 1, which was subsequently identified as a mutation in the diaphanous gene. However, this location allowed Vahava et al. (1998) to identify another form of nonsyndromic deafness (DFNA 15), that also mapped to 5q3I but did not involve diaphanous, as a mutation in the human Brn-3.1 gene locus (POU4F3). This dominant mutation, which produces a late onset, progressive deafness, results in a truncated molecule lacking the POU-HD. Since the POU-HD is required for high-affinity DNA binding of POU-domain TFs, and since unlike many other TFs POU-domain factors

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do not appear to form dimers, it seemed unlikely that the mutation in DFNA 15 would result in a molecule with dominant negative properties. It seemed more likely that the mutation produced a non-functional molecule, and that the phenotype in DFNA 15 patients was the result of haplo-insufficiency. To examine this hypothesis, the hearing of mice with only a single copy of the Brn-3.1 gene (Brn-3.1 +/-) was evaluated at several ages. No difference was found in the hearing of wild-type (Brn-3.1 +/+) and heterozygous (Brn-3.1 +/- up to 24 months of age, inconsistent with haplo-insufficiency. This suggests that either the short life span of mice does not allow haplo-insufficiency to manifest, or that the DFNA 15 mutation results in a dominant negative mutation, perhaps due to competitive protein binding with another nuclear factor (Keithley, Erkman, Bennett, Luo, and Ryan, 1999).

CONCLUSIONS The data reviewed above indicate that the POU-domain TF Brn-3.1 is required for the differentiation of hair cells from their precursors in the sensory epithelia of the inner ear. Because Brn-3.1 appears to occur only in cells that are destined to become hair cells, it is unlikely to be responsible for fate determination of this cell type. However, the initial expression of Brn-3.1 occurs within one day of the final division of cells destined to become hair cells (Ruben, 1967). Brn-3.1 expression also appears to begin before hair cell precursors lose contact with the basement membrane of the sensory epithelium. Taken together, these observations suggest that commitment to the hair cell fate and expression of Brn-3.1 are tightly linked. It is even possible that the same signals that mediate commitment to the hair cell fate initiate expression of Brn-3.1. The upstream regulation of POU-domain genes is not well understood. However, factors that control Brn-3.1 expression may be involved in the determination of hair cell fate. Brn-3.1 expression continues in hair cell throughout life, and a mutation in the human Brn-3.1 gene produces late onset, progressive deafness. These observations suggest that Brn-3.1 also serves as a maintenance factor for hair cells. Unc-86 is expressed in C. elegans mechanoreceptor precursors and neurons, while Brn-3.1 is expressed in murine hair cell precursors and hair cells. In addition, mutations in both genes have severe effects upon the development of the corresponding cell type (Erkman et al., 1996; Tavernarakis and Driscoll, 1997). These similarities support the hypothesis that an evolutionary relationship may exist between primitive mechanoreceptor neurons and vertebrate hair cells. The downstream genes that are targets of Brn-3.1 in hair cells are unknown at the present time. However, given the lack of hair cell differentia-


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tion seen in Brn-3.1 null mice, this TF appears to control, either directly or indirectly, the expression of many of the genes that are required for the normal structure and function of these cells. Supporting a more direct role for Brn-3.1 is the observation that, in other systems, Brn-3 family TFs have been shown to interact with the promoters of a wide variety of neuronal genes, including those encoding synaptic proteins (Lakin et al., 1995; Morris et al., 1996)), neurotransmitters (Miltone, Bessis, Changeux, and Latchman, 1995, 1996) and neurofilaments (Smith et al., 1997). In addition, this TF family contributes to the regulation of other genes whose expression might be expected to vary during development, including Bcl-2 (Smith et al., 1998), genes containing hormone response elements (Budhram-Mahadeo, Parker, and Latchman, 1998), and iNOS (Gay, Dawson, Murphy, Russell, and Latchman, 1998). Brn-3 family members have also been demonstrated to regulate the promoters of genes in viruses that show preferences for neurons (Brownlees et al., 1999). Brn-3.1 appears to function as a master regulatory factor during hair cell differentiation. Other factors have recently been shown to make major contributions to hair cell development, incuding Notch/Delta signaling proteins (Lanford et al., 1999) and the TF Math I (Bermingham et al., 1999). As the relationships between these factors, Brn-3. 1, and other developmental regulators are elucidated, a more complete picture of the molecular control of hair cell development will doubtless emerge.

Acknowledgments: Supported by NIH/NIDCD Grant DC00139, by the Research Service of the Veterans Administration, and by the Duaei Hearing Research Fund.

REFERENCES Bermingham, N.A., Hassan, B.A., Price, S.D., Vollrath, M.A., Ben-Arie, N., Eatock, R.A., Bellen, H.J., Lysakowski, A., & Zoghbi, H.Y. (1999). Math I: an essential gene for the generation of inner ear hair cells. Science, 284, 1837–1841. Brownlees, J., Gough, G., Thomas, S., Watts, P., Cohen, J., Coffin, R., & Latchman, D.S. (1999). Distinct responses of the herpes simplex virus and varicella zoster virus immediate early promoters to the cellular transcription factors Brn-3a and Brn-3b. International Journal of Biochemistry and Cell Biology, 31, 451–461. Budhram-Mahadeo, V., Parker, M., & Latchman, D.S. (1998). POU transcription factors Brn-3a and Brn-3b interact with the estrogen receptor and differentially regulate transcriptional activity via an estrogen response element. Molecular Cell Biology, 18, 1029–1041. Erkman, L., McEvilly, R.J., Luo, L., Ryan, A.E., Hoosmand, F., O’Connell, S.M., Keithley, E.M., Rappaport, D.H., Ryan, A.F., & Rosenfeld, M.G. (1996). Requirement for Brn-3.1 and Brn-3.2 in auditory and visual system development. Nature, 381, 603–606.

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Gay, R.D., Dawson, S.J., Murphy, W.J., Russell, S.W., & Latchman, D.S. (1998). Activation of the iNOS gene promoter by Brn-3 POU family transcription factors is dependent upon the octamer motif in the promoter. Biochimica et Biophysica Acta, 1443, 315–322. Gu, G., Caldwell, GA, Chalfie, M. (1996). Genetic interactions affecting touch sensitivity in Caenorhabditis elegans. Proceedings of the National Academy of Sciences, USA, 93, 6577–6582. Jorgensen, J.M. (1989). Evolution of octavolateralis sensory cells. In S. Coombs, P. Goerner, H. Muenz (Eds) The Mechanosensory Lateral Line: Neurobiology and Evolution. (115–145) New York: Springer Verlag. Keithley, E.M., Erkman, L., Bennett, T., Luo, L., & Ryan, A.F. (1999). Effects of a hair cell transcription factor, Brn-3.1, deletion on homozygous and heterozygous mouse cochleas in adulthood and aging. Hearing Research, 134, 71–76, Lakin, N.D., Morris, P.J., Theil, T., Sato, T.N., Mèorèoy, T., Wilson, M.C., & Latchman, D.S. (1995). Regulation of neurite outgrowth and SNAP-25 gene expression by the Brn-3a transcription factor. Journal of Biological Chemistry, 270, 15858–15863. Lanford, P.J., Lan, Y., Jiang, R., Lindsell, C., Weinmaster, G., Gridley, T., & Kelley, M.W. (1999). Notch signalling pathway mediates hair cell development in mammalian cochlea. Nature Genetics, 21, 289–292. Li, S., Crenshaw, E.B., Rawson, E.J., Simmons, D.M., Swanson, L.W. & Rosenfeld, M.G. (1990). Dwarf locus mutants lacking three pituitary cell types result from mutations in the POU-domain gene Pit-1. Nature, 347, 528–533. Milton, N.G., Bessis, A., Changeux, J.P., & Latchman, D.S. (1995). The neuronal nicotinic acetylcholine receptor alpha 2 subunit gene promoter is activated by the Brn-3b POU family transcription factor and not by Brn-3a or Brn-3c. Journal of Biological Chemistry, 270, 15143–15147. Milton, N.G., Bessis, A., Changeux, J.P., & Latchman, D.S. (1996). Differential regulation of neuronal nicotinic acetylcholine receptor subunit gene promoters by Brn-3 POU family transcription factors. Biochemical Journal, 317, 419–423. Morris, P.J., Lakin, N.D., Dawson, S.J., Ryabinin, A.E., Kilimann, M.W., Wilson, M.C., & Latchman, D.S. (1996). Differential regulation of genes encoding synaptic proteins by members of the Brn-3 subfamily of POU transcription factors. Molecular Brain Research, 43, 279–285. Ruben, R.J. (1967). Development of the inner ear of the mouse: a radioautographic study of terminal mitoses. Acta Otolaryngology, Supplement, 220, 1–44. Rudnicki, M.A., Schnegelsberg, P.N., Stead, R.H., Braun, T., Arnold, H.H., & Jaenisch, R. (1993). MyoD or Myf-5 is required for the formation of skeletal muscle. Cell, 75, 1351–1359. Ryan, A.F. (1997). Transcription factors and the control of inner ear development. Seminar in Cell and Developmental Biology, 8, 249–256. Ryan, A.F., Luo, L., McEvilly, R., Erkman, L., & Rosenfeld, M.G. (1996). A transcription factor specific for mammalian hair cells is expressed throughout life. Abstract Association for Research in Otolaryngology, 19, 575. Ryan, A.K., & Rosenfeld, M.G.. (1997). POU domain family values: flexibility, partnerships and developmental codes. Genes and Development, 11, 1207–1225. Ryan, A.F., Simmons, D.M., & Crenshaw, E.B. (1991). Gene expression in normal and abnormal inner ears. Annals of the New York Academy of Sciences, 630, 129–132.


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Smith, M.D., Ensor, E.A., Coffin, R.S., Boxer, L.M., & Latchman, D.S. (1998). Bcl-2 transcription from the proximal P2 promoter is activated in neuronal cells by the Brn-3a POU family transcription factor. Journal of Biological Chemistry, 273, 16715–16722. Smith M.D., Morris, P.J., Dawson, S.J., Schwartz, M.L., Schlaepfer, W.W., & Latchman, D.S.(1997). Coordinate induction of the three neurofilament genes by the Brn-3a transcription factor. Journal of Biological Chemistry, 272, 21325–21333. Tavernarakis, N., & Driscoll, M. (1997). Molecular modeling of mechanotransduction in the nematode Caenorhabditis elegans. Annual Review of Physiology, 59, 659–689. Vahava, 0., Morell, R., Lynch, E.D., Weiss, S., Kagan, M.E., Ahituv, N., Morrow, J.E., Lee, M.K., Skvorak, A.B., Morton, C.C., Blumenfeld, A., Frydman, M., Friedman, T.B., King, M.C., & Avraham, K.B. (1998). Mutation in transcription factor POU4F3 associated with inherited progressive hearing loss in humans. Science, 279, 1950–1954 Xiang, M., Gan, L., Li, D., Chen, Z.Y., Zhou, L., O’Malley, BW., Klein, W., & Nathans, J. (1997). Essential role of POU-domain factor Bm-3c in auditory and vestibular hair cell development. Proceedings of the National Academy of Sciences, USA, 94, 9445–9450. Xiang, M., Gao, W.Q., Hasson, T., & Shin, J.J. (1998). Requirement for Bm-3c in maturation and survival, but not in fate determination of inner ear hair cells. Development, 125, 3935–3946.

6 Enhancing Signal Discrimination by Means of a Metabotropic Glutamate Receptor Adam W. Hendricson Grace Athas, Ph.D. Paul S. Guth, Ph.D.

Vestibular afferent fibers produce action potentials at rest. The generation of these action potentials requires their activation by the hair cell transmitter, for the afferents are silent in the absence of transmitter release from the hair cells (Guth, Aubert, Ricci, Norris, 1991). The release of transmitter from resting hair cells is made possible by a steady, depolarizing K+ current which flows into the stereocilia from the endolymph through transduction channels. This depolarizing current causes the opening of voltagegated Ca2+ channels, which in turn leads to transmitter release. The tonic release of transmitter from hair cells, and its activation of afferents, means that afferent synapses in the vestibular periphery are “on” at rest. Thus, motion-induced signals need only modulate the ongoing activity, without having to turn the network of synapses “on” or “off.” This motif—graded transmitter release—is also found at the photoreceptor-bipolar cell synapse in the retina (Tessier-Lavigne, 1991). Such a system is more closely attuned to minute changes in transmitter release than an ordinary synapse, which can only be modulated in a positive direction (i.e., above “rest”, the absence of transmitter release). Assuming that baseline discharge is rest, the hair cell-afferent synapse can be modulated above and below its set point, making the conveyance into the brain of information about the position and velocity of the head a far more spatially sensitive operation. This seemingly efficient arrange103

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ment comes with a price, however. The price is that the ongoing activity presents a background against which mechanically-evoked signals must be discriminated. We propose that the solution selected by vertebrate evolution to this problem of signal discrimination may take the shape of a presynaptic metabotropic receptor for glutamate (mGluR). The hypothesis under consideration is that mGluRs on vestibular hair cells (VHCs) are recruited under conditions of mechanically-evoked activity, but not at rest. When recruited, the mGluRs provide positive feedback enhancement of hair cell transmitter release and thus enhanced afferent firing, but in the evoked mode only.

BACKGROUND The amino acid glutamate is arguably the most important excitatory neurotransmitter of the vertebrate nervous system, implicated in functions ranging from the long term potentiation which precedes learning (Balschun, Manahan-Vaughn, Wagner, Behnisch, Reymann, & Wetzel, 1999; Morris, Knevett, Lerner, & Bindman, 1999) to the excitotoxic neuronal damage which succeeds stroke (Pizzi, Fallacara, Arrighi, Memo, and Spano, 1993; Chen, Surmeier, & Reiner, 1999; Pizzi, 1996). Its receptors are both ionotropic (iGluRs; ion-channel linked) and metabotropic (G-protein linked) (Conn & Pin, 1997). The metabotropic family of glutamate receptors is divided into three groups (I, II, and III) whose various agonists and antagonists have been extensively described (Pin, DeColle, Bessis, & Acher, 1999). It is generally accepted that group I mGluRs enhance the activity of phospholipase C, and that Group II and III mGluRs decrease the activity of adenylyl cyclase (Conn et al., 1997). First characterized in the mid-1980s (Sladeczek, Pin, Recasens, Bockaert, & Weiss, 1985; Sugiyama, Ito, & Hirono, 1987), these neuromodulatory receptors are ubiquitous in distribution, varied in function, and complex in pharmacology. Their distribution in the CNS includes, but is not necessarily limited to, the hippocampus (Nicolle, Columbo, Gallagher, & McKinney, 1999; Schools, & Kimelber, 1999), hypothalamus (Schrader & Tasker, 1997), cerebellum (Pizzi et al., 1993; Vetter, Garthwaite, & Batchelor, 1998; Pizzi, 1996), cerebral cortex (Beaver, Ji, & Daw, 1999; Strasser, Lobner, Behrens, Canzoniero, & Choi, 1998), striatum (Sladeczek et al., 1985; Mao & Wang, 1999), olfactory bulb (Schoppa & Westbrook, 1999; Wada, Shigemoto, Kinoshita, Ohishi, & Mizuno, 1998), neural retina (Gafka, Vogel, & Linn, 1999; Linn & Gafka, 1999), and auditory (Kleinlogel, Oestreicher, Arnold, Ehrenberger, & Felix, 1999) and vestibular hair cells (Guth et al., 1998). Metabotropic glutamate receptors have also been reported to exist in the mammalian myocardium (Gill, Pulido, Mueller, & McGuire, 1999) and may exist at the Drosophila neuromuscular junction (Peterson, Fetter, Noordermeer, Goodman, & DiAntonio, 1999). The best characterized neuronal phenomena mediated

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by mGluRs are excitotoxicity (Colwell & Levine, 1999) and long-term potentiation (Morris et al., 1999; Balschun et al., 1999). The role of mGluRs in the vestibular system came under scrutiny as the result of a series of observations concerning the relationship between resting and mechanically- or electrically-evoked discharge at VHC-afferent nerve synapses (Guth et al., 1991). Under a variety of conditions, the release of neurotransmitter from unstimulated hair cells (the so-called “tonic” release made possible by an inward K+ current at the apical end of the cells) displays different kinetics than does transmitter release evoked by mechanical, chemical, or electrical stimuli. This led to the concept of a presynaptic (i.e., hair cell) autoreceptor which is activated solely by evoked discharge of neurotransmitter from hair cells, and thus modulates evoked phenomena while having no effect on tonic release of transmitter. Contemporaneous observations of the responsiveness of VHCs to drugs, such as quisqualic acid, known to activate mGluRs led to the concept that this hair cell autoreceptor might be glutamatergic (Prigioni, Russo, Valli, & Masetto, 1990; Valli, Prigioni, Zucca, Botta, & Guth, 1985). The evidence that mGluRs exist on VHCs, and that they may mediate a differential release of glutamate in resting versus mechanically stimulated mode is derived from pharmacology, molecular biology, and immunocytochemistry (ICC). Administered alone, the mGluR antagonists (RS)-1aminoindan-1,5-dicarboxylic acid (AIDA) and (S)-4-carboxyphenylglycine (4CPG) do not alter ampullar afferent firing rates (Guth et al., 1998). Antagonists of the mGluRs are effective at reducing transmitter release evoked by mGluR agonists, which have been shown pharmacologically to act at presynaptic receptors (Guth et al., 1998; Prigioni et al., 1990; Valli et al., 1985). Messenger RNA expression profiling studies from our lab have shown mGluR1 to be localized to VHCs from the SCC (Guth et al., 1998), results which have recently been confirmed and expanded by ICC experiments showing both mGluR1a and 5 to exist on SCC hair cells. The above data are the foundation for our hypothesis that presynaptically located mGluRs may mediate positive feedback loops in afferent neurotransmission from the vestibular apparatus to the brain. By this model, mechanically-evoked release of glutamate from hair cells activates presynaptic mGluRs, which increase transmitter release via a mechanism which remains speculative (Guth, Perin, Norris, & Valli, 1998), but which based on evidence from other systems may involve modulation of K+ channels (Sharon, Vorobiov, & Dascal, 1997), Ca2+ channels (Guatteo, Mercuri, Bernardi, & Knopfel, 1999), intracellular Ca2+ metabolism (Bianchi, Young, & Wong, 1999), PKC activation (Hori, Takai, & Takahashi, 1999), or, speculatively, the transmitter release apparatus itself. In this way, a mechanical signal transduced by the vestibular neuroepithelium is elevated above the tonic release of transmitter from resting VHCs. Positive feedback of glutamate on its own release via presynaptic group I mGluRs has been documented in 105

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cerebral cortex (Herrero, Miras-Portugal, & Sanchez-Prieto, 1992) and hypothalamus (Schrader et al., 1997). In brain, activation of group I mGluRs can lead to Ca2+ influx through voltage-dependent channels which is readily discernible via electrophysiological (patch or voltage clamping of neurons) or photochemical (imaging the fluorescence of calcium-sensitive dyes such as fura-2) recording techniques (Guatteo et al., 1999).

METHODS Whole Labyrinth Preparation (Guth, Norris, Guth, Quine, & Williams, 1986) Leopard frogs (Rana pipiens) are anesthetized by chilling to 40° F, sacrificed by decapitation, and caudally pithed. The roof of the mouth is cut into two halves, and the tissues overlying the bone and cartilage of the otic capsule are cut away. The ventral portion of the otic capsule and the melanophorecovered membranes which ensconce the labyrinth itself are removed, exposing the ampullar nerve. The nerve is severed near the point at which it delves between the auditory papillae. The remainder of the soft tissue and bone surrounding the otic capsule are cut away and the labyrinth is transplanted into a 20 ml tissue bath superfused with frog Ringer’s solution (104.6 mM NaCl, 2.5 mM KCL, 13 mM NaHCO3, 1.7 mM NaH2PO4·H20, 1.8 mM CaCl2, 1mM MgCl2·H20, 5mM glucose in ddH20 ((double-distilled water)), adjusted to pH 7.18-7.22) at a continuous rate of 5 ml/min. The nerve stump is pulled via gentle suction into a glass pipette filled with Ringer’s solution and connected to a Ag/AgCl electrode. Action potentials recorded from the severed nerve are amplified 100–fold by a differential amplifier (WPI), converted into an analog measure of frequency by a slope/height window discriminator, and collected in 1sec bins by a rate/interval monitor (Frederick Haer and Co). The rate/interval analyzer discharges an analog measure of frequency once per second into an analog-to-digital-converter board (Axon Instruments), which in turn discharges into a microcomputer (Gateway Computing) running the Axoscope 7.0 electrophysiological recording suite (Axon Instruments), where a linear representation of action potential frequency is displayed and simultaneously written onto hard media for offline analysis. Drugs dissolved in Ringer’s are applied either by introduction into the reservoir of Ringer’s (bath exchange) or by close microinjection onto the neuroepithelium at a rate of 50 µl/min via a single-port pipette fed by four separate lines. Control injections of Ringer’s alone are given to ascertain that they do not alter the discharge rate, and Ringer’s is run through the common injection port for 30 sec after every drug injection to flush the remnants of the drug out of the port in preparation for the next injection. (See Figure 6–1)


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Amp FRC L

PC

S U AC

HC

Figure 6–1. Schematic of isolated vestibular organ preparation. Nerve in suction elecrode innervates posterior semi-circular canal. Amp: AC amplifier. FRC: frequency to rate converter. AC: Anterior canal. HC: Horizontal canal. PC: Posterior canal. S: Saccule. L: Lagena

In order to evoke increases in transmitter release via mechanical movement of stereocilia bundles, a bimetallic strip coupled to a glass pushrod is exposed via voltage amplifier (ThorLabs) to a set voltage which reverses itself in phase with a sine wave from a function generator (Tektronix). The resulting inward and outward flexion of the bimetallic strip causes the tip of the pushrod to gently indent the duct of the inferior SCC. This perturbation of the canal wall causes a concomitant movement of the gelatinous cupula, which overlays the hair-cell studded crista, both of which reside inside the ampulla of the canal. Movement of the cupula causes deflection of the hair cell stereocilia bundles which project into it, leading to changes in hair cell membrane voltages and thus, transmitter release. These upward and downward oscillations in transmitter release can be discerned on a physiograph configured (as described above) to reflect frequency of action potentials in the afferent nerve (see Figure 6–9 in Results section).

Isolation of Vestibular Hair Cells (Housley, Norris, & Guth, 1989; Hudspeth & Lewis, 1988; Lewis & Hudspeth, 1983). Leopard frogs (Rana pipiens) are chilled, pithed, and sacrificed by decapitation. The superior portion of the head, sectioned sagitally, is placed in external medium (119mM NaCl, 3 mM KCL, 8mM Na2HPO4· 2 H20, 2mM NaH2PO4· 2H20, 2mM CaCl2· 2H20, 1mM MgCl2·H2O, and 3mM glucose,

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adjusted to pH 7.25). After exposing the inner ear by removing the ventral portions of the otic capsule, the whole labrynth is removed and placed in dissociation medium (122mM NaCl, 3mM KCl, 8mM Na2HPO4, 2 mM NaH2PO4 • H20, 0.02 mM CaCl2 • 2H2O, and 3mM glucose, pH adjusted to 7.25). The cristae of the SCCs are dissected from the remainder of the labyrinth and trimmed of excess tissue. The tissue is next moved via fine forceps (A. Dumont Instruments) to a glass well plate (Pyrex) filled with dissociation medium containing the proteolytic enzyme trypsin (0.05%; Sigma) for 30 minutes. Next, the cristae are transferred for 1 minute into a well containing FBS, then into a final well containing bovine serum albumin (500 µg/µl; Sigma) for 15 minutes. The cristae are removed and placed on a Sylgard platform designed to fit the stage of an inverted microscope (Nikon). Under the scope, gentle agitation with a fine glass wisp is used to separate the hair cells from their tissue substrate. After allowing the cells to settle to the bottom of the dish for 5 minutes, a 500µl/min perfusion of external medium is begun which will persist for the duration of the experiment. Patch clamping may now be performed. In the case of immunocytochemical (ICC) experiments, cells are isolated as described, except they are wisped from the cristae onto glass slides coated with concanavalin-A (Sigma) instead of onto a Sylgard platform.

Amplification of mRNA (Eberwine, Yeh, Miyasharo, Cao, Nair, & Finnel, 1999; Eberwine, Spencer, Miyasharo, Mackler, & Finnel, 1992; Mackler, Brooks, & Eberwine, 1992; Van Gelder, vonZastrow, Yool, Dement, Barchas, Eberwine, 1993). Under stringent RNAse precautions (latex gloves, molecular grade reagents, diethylpyrocarbonate-treated and autoclaved ddH20, autoclaved plasticware and baked glassware), the cytoplasm of SCC hair cells are aspirated into a patch pipette, then transferred into a 600 µL microcentrifuge tube containing 15.5 µL ddH20, 2 µL 10X electrode buffer, 2.5 mM each of 4 deoxynucleotide triphosphates (dNTPs), and 10U avian myeloma virus reverse transcriptase (Seikagaku). After 2 min of centrifugation, the pipette and its contents are incubated at 37° C for 60–90 min, then precipitated with ethanol and centrifuged for 15 min at 4°C. Upon aspiration of the ethanol, the pellet is air-dried and resuspended in 20µL ddH20. The cDNA/mRNA complex is heat-denatured at 95° C for 3 min, and cooled rapidly over ice. Upon transfer, 1µL ddH20, 4µL 10X 2nd strand buffer, 2.5 mM of 4 dNTPs, 1U/1µL T4 DNA polymerase, and 1U/1µl Klenow fragment are added to catalyze the formation of 2nd strand cDNA. After 5 hours of incubation at 14° C, 48 µL ddH20, 10 µL S1 buffer, and 1 U/µL S1 nuclease (Boehringer) are added and the solution is incubated again at 37° C for 5 hours to remove hairpin loops. The pellet is extracted using phenol-


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chlorophorm extraction followed by ethanol precipitation and centrifugation. Following air-aspiration of the ethanol, the pellet is resuspended in 20 µL ddH20. Blunt ends are obtained by the addition of 2.5 µL 10X KFI buffer, .5 µL of each of 4 dNTPs (from a 10µM stock), and .5 U/0.5 µL Klenow fragment. After incubation, phenol-chloroform extraction and transfer, ethanol precipitation, and centrifugation, the pellet is aspirated, air-dried, and resuspended in 20 µL ddH20. Drop dialysis is then employed to remove unincorporated nucleotides. Double-stranded cDNA has now been isolated, and can be used to make aRNA. Nine µl ddH20, 2 µL 10X amplification buffer, 1µL DTT (100mM), 2 µL each ATP, GTP, UTP (2.5 mM), 1µL CTP (1mM), 15 µCi alpha-32-P-CTP, .5 RNAsin (ribonuclease inhibitor), and 1µL T7 RNA polymerase are added to 2 µL of the dialyzed cDNA extract and allowed to incubate at 37° C for 4 hours. Phenol-chloroform extraction and ethanol precipitation follow, as described above. An aliquot of RNA product is run on a denaturing 1% agarose gel in 1X MOPS to ascertain the extent of RNA degradation as well as the amount of incorporated radiolabel. A 103 fold amplification of the mRNA in the starting material has now been obtained. RNA is next reamplified by repeating the above steps, starting with 1st strand cDNA synthesis. The 2nd round of amplification yields a 106 fold increase in mRNA, and the radiolabelled, amplified mRNA is ready to be used as a probe for expression profiling.

Expression Profiling (Mackler et al., 1992) Following treatment with restriction enzymes (37° C, 2 hours), the cDNA clones are linearized. Equimolar amounts of the linearized clones are denatured (95° C for 5 min), transferred to nitrocellulose paper, and UVcrosslinked in a slot blot apparatus (Bio-Dot ST, Bio Rad). The individual slot blots are placed in hybridization bottles with 50% formamide, 5X SSC, 5X Denhardt’s solution, and 100 µg/ml salmon sperm DNA, for 3 hours at 42° C. Next, the amplified aRNAs from VHCs are heat denatured (5 min at 95° C), cooled (ice), and added to the bottle containing the slot blot. After 48 hours at 40° C in a hybridization oven (Bellco), the slot blot filters are washed (RNAse-free), first under low stringency conditions (twice, 2xSSC and 0.1% SDS for 40 min at 42° C), and then under high stringency (0.1xSSC and 0.1 % SDS for 30 min at 42° C). After air-drying of the slot blots, autoradiography is performed with an exposure time of 16 to 96 hours. Intensity of the hybridization signal as compared to controls allows for the determination of relative levels of expression. Also, densitometry or phosphoroimaging analysis can be used to quantify expression profiles. Vector cDNA (pUC18) is used as a negative control and to provide a background level, while housekeeping genes such as GAPDH, Fra-1, or Fra-2 can be used as positive controls. (See Figure 6–2)

110 AA A5A' A

P atch Pi pette

AAAAA AA AA A 5'

5'

AAAAA

5'

Figure 6–2. Flow Diagram for Single-Cell mRNA Amplification (aRNA®)


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IMMUNOCYTOCHEMISTRY Immunocytochemistry was performed on SCC (semicircular canal) hair cells enzymatically isolated as described in Probe-clip press-seal incubation chambers (Sigma) coated with a 2mg/ml solution of Concanavalin-A (Sigma) in standard frog Ringer’s. The chambers are rinsed with PBS for 10 minutes after the cells are isolated. Next, a fixative consisting of 1.08 ml paraformaldehyde, 1.40 ml saturated picrid acid 2.0 ml concentrated PBS, and ddH 20 up to a 10 ml total volume is added to the wells and allowed to sit for 20 minutes. The wells are rinsed with PBS three times at the end of the fixative incubation period. After rinsing, 2 blocking solutions (A and B) are added sequentially for 30 minutes each. Blocking solution A consists of PBS + 0.02% Triton X-100 (Sigma) + 1% BSA + 0.01% NaN3. Blocking solution B consists of PBS + 0.4% Triton X-100 + 1% BSA + 1% goat serum + 0.01% FCS + 0.1% FCS + 0.01% NaN3. Primary antibodies (mGluR1a and mGluR5, rabbit polyclonal, Pharmingen Corp.) diluted 1:100 in PBS + 0.4% Triton X-100 + 1% BSA + 1% goat serum + 0.1% FCS + 0.01 % NaN3 are added to the chambers for 40 hours at 4°C in a humid atmosphere. After incubation of the hair cells with the primary antibody, the chambers are washed three times with a solution of ice-cold PBS + 0.2% Triton X-100, 5 minutes per wash. The secondary antibody (fluorescine isothiocyanatetagged, goat anti-rabbit) is diluted 1:165 in a solution consisting of PBS + 0.2% Triton X-100 + 1% BSA + 1% goat serum + 0.1% FCS + 0.01% NaN3. This solution is added to the incubation chambers for 60 minutes at room temperature in a humid atmosphere. The chambers are then rinsed 3 times with PBS, 5 minutes per wash. Finally, the incubation chambers are rinsed once with ddH20, mounted on glass slides using Biomedia Gelmount, and visualized under a fluorescence microscope (Nikon). Slides are stored in the dark at 4°C. Negative controls are performed by following the above procedure with the exception of the addition of the primary antibody, in order to ensure that the secondary antibody is not binding to the cells in a non-specific manner, leading to false positive results.

RESULTS The Effects of mGluR Agonists in the SCC ACPD is a non-selective mGluR agonist. Application of ACPD to the neuroepithelium of the posterior SCC produced increases in firing rates of ampullar afferents (see Figure 6–3A). Figure 6–3B depicts the relationship between concentrations of ACPD applied and its facilitatory effects on afferent firing. The EC50 is approximately 40 µM. Quisqualate, a poorly selective glutamatomimetic, is a group I mGluR agonist (Littman, Chase,

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Figure 6–3. A: Resting firing rate, multiunit recording from ampullar nerve, frog posterior SCC. Dosedependently, the mGluR agonist t-ACPD increased resting multi-unit afferent discharge. Arrows mark start of drug injections; response to 100µM t-ACPD lasted 7 1.5 min (n8). B: Dose-response curve for t-ACPD facilitation of afferent discharge. Spontaneous firing rate reversibly increased to 182 8.9% (n8) of control values upon 100µM t-ACPD administration (in the absence of agonist, baseline noise was 8 2.9%). EC50 for t-ACPD action was 35.4 8.5 µM; the dose-response saturated above 300µM (n4). Nfr: nerve firing rate. Con: artificial perilymph control injection.

Renzi, Garlin, Koerner, Johnson, 1995). It causes an increase in afferent firing in the SCC (Annoni, Cochran, & Precht, 1984; Prigioni et al., 1990). tADA, weakly selective for mGluR5 over mGluR1, produced modest facilitation of afferent firing at 1 to 3 mM (data not shown). The group III agonist L-AP4 had no effect on afferent firing.


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Site of action of ACPD In an attempt to determine the site of action of ACPD, the whole labyrinth was bathed in artificial perilymph (AP) containing 0.1 mM Ca 2+ and 10 mM Mg2+. This alteration in ionic composition of the bathing medium is designed to inhibit hair cell transmitter release (Annoni, Cochran, & Precht, 1984; Bobbin, Bledsoe, Winbery, & Jenison, 1985; Soto & Vega, 1988; Valli, Zucca, Prigioni, Botta, Casella, & Guth, 1985). Under these conditions, resting afferent activity ceased, underscoring the idea that the afferents themselves are silent unless stimulated by the hair cell transmitter (Guth et al., 1991). ACPD was now incapable of causing increases in ampullar afferent activity, indicating that its site of action was probably the hair cell, and not the afferent neurons per se. To demonstrate that the afferent neurons were still capable of responding, injections of AP with 1 mM Glu were made. These produced a rapid and dramatic rise in afferent firing rates (Figure 6–4). High potassium (15 mM) AP also produced increases in afferent firing under these conditions (data not shown). This concentration of potassium is capable of depolarizing the afferents past their threshold for action potential discharge.

125 Hz 2+

Low Ca , high Mg

5 min

2+

Con ACPD 100 µM ACPD 100 µM

Glu ACPD Glu 1mM 100 µM 1 mM

Figure 6–4. Resting discharge from the whole ampullar nerve in the presence of a high-Mg2, low-Ca2 Ringer’s solution, which has been shown to block afferent transmission by impeding transmitter release (Valli et al., 1985). While 1mM glutamate increased discharge rate, the response to 100µM t-ACPD was negligible compared to control. These findings indicate that the receptor for t-ACPD may be presynaptic.

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Testing the Possibility that ACPD Could be Acting at Non-mGluR Sites Acetylcholine receptors Acetylcholine (ACh), the primary efferent transmitter, has been shown to cause an increase in ampullar afferent firing rates by an action on hair cell ACh receptors (Bernard, Cochran, & Precht, 1985; Guth, Dunn, Kronomer, & Norris, 1994; Guth et al., 1986; Guth, Dunn, Kronomer, & Norris, 1994; Guth & Norris, 1996; Housley, Norris, & Guth, 1990; Norris, Housley, Williams, Guth, & Guth, 1988). Since ACPD has been shown to increase transmitter release (Schrader et al., 1997), and mimics ACh in the SCC in causing facilitation of afferent firing, it seemed reasonable to test whether ACPD was acting indirectly through ACh (i.e., by stimulating the release of ACh from efferent endings). In the SCC, it is the activation of muscarinic receptors that is responsible for the facilitation caused by ACh (Guth et al., 1998). In the SCC, atropine at low concentrations (e.g., 3 µM) antagonizes the facilitatory effect of applied ACh (Guth et al., 1998). Therefore, atropine (3 µM) was applied by bath substitution. The responses to ACh and ACPD on ampullar nerve afferent activity before and after atropine were compared (Figure 6–5). While this concentration of atropine blocked the effect of ACh, no interaction between atropine and ACPD was seen. This suggests that in the SCC, ACPD neither acts by releasing ACh from efferent endings, nor by acting on muscarinic receptors. Non-mGluR glutamate receptors Other so-called selective agonists for mGluRs, such as quisqualate and 2,3 dicarboxycyclopropylglycine, have been seen to have measurable activities

125 Hz 5 min

ACPD ACh 100 µM 300 µM

3 µM Atropine

ACh ACPD Con ACh ACPD 300 µM 100 µM 300 µM 100 µM

Figure 6–5. A 3µM concentration of atropine reduced the facilitation of afferent firing caused by 300µM ACh. Atropine had no effect on the response to 100µM tACPD, however. This indicates that the receptor activated by t-ACPD is pharmacologically distinct from the muscarinic receptor.


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at AMPA and NMDA receptors (Wilsch, Pidoplichko, Opitz, Shinozaki, & Reyman, 1994). Both AMPA and NMDA receptors appear to exist in the SCC (Cochran & Correia, 1995; Devau, Lehouelleur, & Sans, 1993; Guth et al., 1998; Prigioni et al., 1990; Prigioni, Russo, & Masetto, 1994; Soto, Flores, Erostegui, & Vega, 1994; Zucca, Akoev, Maracci, & Valli, 1993). To eliminate the possibility that ACPD, used as an mGluR test-probe, was acting on NMDA receptors, either directly or in concert with the transmitter (Conn et al., 1997) the NMDA selective antagonist D(-)-2-amino-5-phosphono pentanoic acid (AP-5) was employed. Figure 6–6 depicts the antagonism of NMDA-induced facilitation of SCC afferent firing by AP-5. Note that the ACPD-induced facilitation is unaffected. This suggests that at the concentrations used in these studies, ACPD is not acting on or influencing an NMDA-type receptor. The enhancement of the responses to ACh and ACPD after AP-5 was unexpected but is reproducible and as yet, unexplained.

Antagonism of ACPD by mGluR antagonists Two different mGluR antagonists, (S)-4-carboxyphenyl glycine (4CPG) and (RS)-1-aminoindan-1,5 dicarboxylic acid (AIDA), were pitted against the facilitatory effect of ACPD. Both are said to be group I-selective, competitive mGluR antagonists. 4CPG antagonized ACPD in a dose-dependent manner, and at 1 mM completely blocked the effect of 100 µM ACPD (Figure 6–7A and 6-7B). AIDA was partially effective at 1 mM and completely effective at 3 mM against 100 µM ACPD (data not shown). These results suggest, if the reported selectivity of these antagonists is to be trusted, that ACPD may be activating group I mGluRs. When applied alone, neither 4CPG (Figure 6–7A) nor AIDA (not shown) affected resting activity.

125 Hz 5 min

ACh 300 µM

1 mM AP-5

Con ACPD 100 µM

NMDA 100 µM

ACPD 300 µM

NMDA Con ACh 300 µM 100 µM

Figure 6–6. A continuous recording from a single SCC. The NMDA blocker AP-5, applied by bath substitution, reduced the response to NMDA, while having no effect on the responses to ACh and t-ACPD.

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Figure 6–7. A. The facilitation of SCC afferent discharge caused by 100µM t-ACPD is blocked entirely by co-injection of 100µM t-ACPD with 1mM 4CPG, a selective, competitive, group I mGluR antagonist. B. 4CPG dose-dependently reduces the effect of t-ACPD on afferent discharge. Note that 4CPG alone does not reduce firing rate, indicating that group I mGluRs may not be activated in the absence of evoked discharge (n8; IC50217.3 73.8 µM). Data normalized to the effect of 100µM t-ACPD (red circle).

Antagonism of Mechanically Evoked Activity by 4CPG As shown, 4CPG appears to block group I mGluRs at their site of interaction with ACPD. The definitive test of our hypothesis concerning the physiological role of mGluRs is to assess their role in modulating mechani-


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cally-evoked activity at the hair cell-afferent synapse. To this end, we employ a peizo-electric bimorph device which, coupled to a glass pushrod, induces a gentle, sinusoidal perturbation of the duct of the inferior SCC. This perturbation displaces hair cell stereocilia in the hair cell-studded cristae of the canal, causing repetitive increases and decreases in transmitter release, resulting in sinusoidal oscillation of afferent discharge (Figure 6–8). Preliminarily, we have observed a dampening of the increase in afferent activity evoked in this manner following the application of 4CPG (Figure 6–9). Findings such as these, if valid, strongly support our hypothesis that group I mGluRs on vestibular hair cells function to amplify mechanically evoked activity at the hair cell efferent synapse, while having little to no effect on resting discharge.

Expression profiling of individual vestibular hair cells Using expression profiling, we have investigated the phenotype of metabotropic glutamate receptors (mGluR1-4) from SCC, saccular, and utric-

L S

PC Bimorph device

U AC

HC

Voltage amplifier

Function generator

Figure 6–8. Schematic of the mechanically-evoked activity preparation.

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CPG 1mM

100Hz

5 min.

AP

CPG 1mM

(bimorph stimulus) Figure 6–9. Mechanically evoked activity is reduced by blockade of group I mGluRs by 4-CPG. AP: artificial perilymph control. Bimorph stimulus: sine wave corresponding to the inward and outward movement of the bimorph pushrod against the canal duct. All injections were 60 seconds in length.

ular hair cells. Figure 6–10 displays representative slot blots using radioactive amplified RNA (aRNA) from the three cell types as probes (SCC and saccule; n=3, utricle; n=1). Note the strong hybridization signal for mGluR1. Since the blots were subjected to a high stringency wash and the negative control plasmid (pUC18) showed no signal, we can be confident that the strong signal for mGluR1 is valid. No probe for mGluR5 was available at the time of this study. The mGluR subtypes 2 and 3 are members of the group II mGluRs and subtype 4 is a member of the Group III mGluRs. This assay failed to detect the mRNA for any of these receptors. Thus evidence is presented only for the presence of Group I mGluRs in type II vestibular hair cells (the only type of hair cells found in frog vestibular organs).

Immunocytochemistry ICC experiments using antibodies for mGluR1a and mGluR5 showed immunoreactivity to be concentrated at the basal poles of SCC hair cells (Figure 6–11). These findings suggest a concentration of these receptor proteins at the transmitter release site.

DISCUSSION The SCCs provide the brain with constant information about movement of the head, irrespective of which way the head turns, and which way the


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SCC

Utr

Sac

CaCC GluH1

Blank

fra1

fra2

mGluR1 mGluR2 mGluR3

P.Ctrl (1) pUC18 Blank(2,3) mGluR4

Blank

Figure 6–10. Slot blots containing cDNA hybridized to radioactively labeled aRNA from isolated SCC, utricular, and saccular VHCs. Strong hybridization signals are seen for the positive controls (fra-1 and fra-2) and for mGluR1. Faint to no hybridization signals are seen for the AMPA receptor (GluH1), mGluRs 1–4, and the negative control (pUC18).

hair cell stereocilia are bent. This is accomplished by having hair cells release their transmitter continuously at rest so that the afferents fire under non-stimulated conditions, thus allowing bi-directional modulation of the resting activity by the graded release (or inhibition of release) of hair cell transmitter in response to graded mechanical stimulation. However, a problem arises precisely from this continuous resting activity. When an important excitatory stimulus is to be transmitted to the afferents, discrimination of this signal may be compromised by the constant background activity. If, however, a mechanically-evoked signal of importance could cause a disproportionately large release of hair cell transmitter, then signal discrimination could be maintained or enhanced. This function could be served by a group I mGluR acting as a positive feedback autoreceptor, as it does in the central nervous system (Herrero et al., 1992; Schrader et al., 1997). Group I mGluRs are coupled negatively to K+ channels (Sharon et al., 1997), which would produce depolarization, activation of Ca2+ channels, and increased transmitter release. They are also coupled positively to L-

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A

B

Figure 6–11. Immunocytochemistry confirmed the presence of (A) mGluRs 1a and (B) 5 on VHCs. Column A, top: Representative SCC hair cell shows heavy mGluR1a immunoreactivity at apical and basal poles, respectively. Middle: phase-contrast micrograph of SCC hair cells used in negative control (no primary antibody; see Methods). Bottom: fluorescence micrograph of cells from middle panel. Negligible fluorescence confirms validity of top panel. Column B, top: mGluR5 immunoreactivity is concentrated primarily at basal pole. Middle and bottom panels: same as column A.

type Ca2+ currents in cerebellar granule cells (Chavis, Fagni, Bockaert, & Lansman, 1995), which could lead directly to increased transmitter release. In addition, there is an mGluR coupled to phosphoinositide hydrolysis whose activation leads to increased intracellular Ca2+, and increased glutamate release (Herrero et al., 1992). Lee and Boden (1997) adduced evidence that ACPD causes depolarization by activating a Ca2+-Na+ exchanger. Logic predicates that the recruitment of hair cell mGluRs in the evoked but not resting mode could be accomplished in two ways: either the mGluRs could be located extrasynaptically, as occurs in some brain


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glutamatergic synapses (Baude, Nusser, Molnar, McIlhinney, & Somogyi, 1995; Lujan, Nusser, Roberts, Shigemoto, & Somogyi, 1996), or they could have low affinities for glutamate. Extrasynaptic mGluRs would not respond to the lower concentrations of transmitter released in the resting mode, because at this slow rate of release, the diffusion distance between synapse and receptor would degrade the transmitter concentration to a level below the threshold concentration for significant mGluR activation. Similarly, mGluRs with low affinities would require a higher concentration of transmitter than that provided by the resting mode in order to undergo significant activation. Because the affinity of glutamate for the mGluRs is about an order of magnitude greater than for the AMPA receptor (the dominant glutamate receptor on vestibular afferents) (Guth et al., 1998), the former possibility seems most likely. The work of Lujan et al (1997) in hippocampus also favors the former possibility, in that group I mGluRs were localized to an annulus approximately 60 nm away from the synapse. These authors suggest that “the distinct patterns of mGluR distribution may reflect specific spatial requirements for different...effector mechanisms” (Author note - such as resting and evoked activity). Thereby, the distant mGluRs would not be stimulated by the limited transmitter release of the resting mode, but only when synaptic transmitter concentrations reached levels sufficient for diffusion to mGluRs spatially distinct (perhaps 60 nm) from the transmitter release site. This view is supported by the work of Scanziani et al (1997) in brain, who demonstrated activation of mGluRs only when the release of glutamate was enhanced by increased frequency of stimulation. In our preparation, the mGluRs are probably not involved in resting mode activity, as attested by the inability of the selective group I mGluR antagonists, 4CPG and AIDA, to affect resting firing rates. As shown, ACPD clearly increases afferent firing rates of the SCC in a dose-dependent manner. Its site of facilitatory action is the hair cell, as shown by its lack of effect when hair cell transmitter release is inhibited in low Ca2+-high Mg2+ solution. ACPD does not seem to work indirectly through ACh or NMDA receptors. Finally, ACPD’s effect is blocked by the group I mGluR antagonist 4CPG, a drug which appears capable of reducing mechanically-evoked activity in the isolated vestibular organ preparation. These findings strongly support the hypothesis of an mGluR on hair cells of the SCC which may be exclusively involved in the evoked-mode activation of the afferents. It bears mentioning that even though the mGluRs may be recruited physiologically only in the evoked mode, they are still available for activation by exogenous mGluR agonists in the resting mode, as in the present research. The question arises: are these results, obtained in an amphibian, generalizable to the mammalian inner ear? Transmitters and their functions seem to be conserved throughout vertebrate evolution. The underpinnings of this assertion come from the writings of Florey (1967, 1972) and Michel-

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son (1974). Both authors agree that, although transmitters themselves are unchanged throughout the animal kingdom, their functions may change from phylum to phylum but within any one phylum (e.g., vertebrate), their functions are the same. Therefore, it may be assumed that results concerning transmitters obtained in amphibians may be applied to all vertebrates. To reiterate, the hypothesis under consideration is that there are functional, positive-feedback autoreceptors on vestibular hair cells which are active when hair cells are mechanically stimulated, and inactive when they are not. The use of pharmacological agents alone rarely provides incisive testing of such hypotheses, largely because the incisiveness of drugs as hypothesis testers depends upon their selectivities (or lack thereof). Early on, pharmacologists learn the sometimes painful lesson that a substance’s selectivity is inversely proportional to the number of publications in which it appears—only rarely does a drug influence only one receptor or enzyme, and the selectivity is always dose-dependent. All of the above notwithstanding, drugs are often very useful guideposts and easily used hypothesis-testing tools. The results of pharmacological experiments encourage or dissuade the investigator from more incisive tests of the hypothesis. The present case in a classic example. With confidence in our hypothesis buoyed by pharmacological investigations, we further compiled molecular and immunocytochemical evidence which seems to paint a clear picture of the phenotype, disposition, and function of mGluRs on vestibular hair cells. Based on these results, further electrophysiological, immunohistochemical, and molecular biological tests are underway to provide a more detailed description of the distribution and functional significance of mGluRs in the vestibular periphery. Finally, it seems worth mentioning that neurotransmission at the hair cell-afferent synapse is unusual in that there is tonic transmitter release which is modulated up or down according to the extent of membrane depolarization (and other factors) (Guth et al., 1998). Thus, transmitter release in the hair cell is graded primarily according to the transductional signal, and not dependent on action potential arrival at the release site (in any case, hair cells generally do not generate action potentials). It may be that the positive feedback autoreceptor will turn out to be a general feature of such graded synapses.

Acknowledgements: This research was supported by NIH grant DC-00303 and the PhRMA Foundation. The authors wish to thank Samara Shipon and Nieka Harris for technical assistance.

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7 Additional Studies on the Role of ATP as a Neuromodulator in the Organ of Corti Richard P. Bobbin, Christopher S. LeBlanc, Manisha Mandhare, and Margarett S. Parker

The function of various chemicals in the cochlea are slowly being defined. For example, there is no doubt that acetylcholine (ACh) functions as the major neurotransmitter of the medial olivocochlear neurons at the outer hair cells (OHCs) and that glutamate functions as the primary neurotransmitter released by the inner hair cells (IHCs) onto the primary auditory nerve dendrites (Bledsoe, Bobbin, and Puel, 1988; Bobbin, Bledsoe, Winbery, and Jenison, 1985; Eybalin, 1993; Puel, 1995). On the other hand, the neurotransmitter role of adenosine triphosphate (ATP) in the cochlea remains to be elucidated. Receptors for ATP are present on a large number of cells in the organ of Corti of the cochlea (Bobbin et al., 2000). This is demonstrated with immunohistochemistry in Figure 7–1. An antibody was used that binds to an intracellular segment of the ATP receptor protein called P2X2 and a chemical that is converted to a blue precipitate at the binding site. As shown in Figure 7–1A, the blue precipitate (ATP receptor protein) is present in OHCs, Hensen’s cells, Deiters’ cells, inner and outer pillar cells, IHCs and inner sulcus cells. The darkest staining was observed in Hensen’s cells and in Deiters’ cells. Reisner’s membrane was also stained but it is not shown in the Figure. Figure 7–1A shows staining on the stereocilia indicating that receptors may be present on these structures as suggested by Housley, Greenwood, and Ashmore (1992) and Housley et al. (1999). However, staining of the stereocilia and the surface of the reticular lamina in the control (antigen mixed with primary antibody), although lighter than the staining with primary antibody alone, makes it difficult to definitively 129

Figure 7–1. Results of immunohistochemistry of guinea pig cochlea in toto to detect P2X2 protein. A: A P2X2 antibody ( Product # 1APR-003; Alomone labs, Jerusalem, Israel) to an intracellular portion of the ATP receptor subunit was used as a primary antibody that was subsequently detected using Vectastain ABC-AP kit that yields a blue precipitate at the receptor sites (Vector Labs, Burlingame, CA). The tissue was then dehydrated and imbedded for sectioning. Mid-modiolar sections through turn three of the cochlea demonstrate that P2X2 protein is present in OHCs, Deiters’ cells, Hensen’s cells, pillar cells, IHCs and inner sulcus cells. Staining of the stereocilia was also observed. Cells in Reisner’s membrane were also stained but are not shown. B: Cells of a control cochlea (P2X2 primary antibody mixed with antigen) from approximately the same turn as shown in A. Light non-selective staining was observed in the stereocilia and in some of the fat globules in Hensen’s cells. Non-selective staining was observed in the tectorial membrane (not shown) and this staining was also observed when the primary antibody was absent. Staining was absent from the IHCs, OHCs, Deiters’ cells, pillar cells, the cytoplasm of Hensen’s cells, inner sulcus cells and Reisner’s membrane.

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state whether or not the receptors are present or absent on the stereocilia. The tectorial membrane and the lipid globules in Hensen’s cells exhibited non-specific staining, that is, staining in the absence of primary antibody (i.e., P2X2 antibody) (Figure 7–1B). Overall, our results confirm and extend the immunohistochemistry results of Housley et al. (1999) in guinea pig and Xiang, Bo, and Burnstock (1999) in rat and the in situ hybridization results of Housley, Luo, and Ryan (1998) in rat and Parker, Larroque, Campbell, Bobbin, and Deininger (1998) in guinea pig. The role of ATP at these receptors in the physiological functioning of the cochlea is currently still very speculative (see reviews: Bobbin, 1996, 1997; Bobbin, Chen, Nenov, and Skellett, 1998; Bobbin et al., 2000; Housley, 1997). There are ATP receptors on cell surfaces exposed to endolymph and ATP receptors on cell surfaces exposed to perilymph. There is no doubt that the ATP receptors exposed to endolymph are involved in fluid and ion transport (Bobbin et al., 2000). Our laboratory has focused on ATP receptors exposed to perilymph and their possible roles in ion movement and as neuromodulators of cochlear mechanics (Bobbin et al., 1998; Chen, Skellett, Fallon, and Bobbin, 1998; Skellett, Chen, Fallon, Nenov, and Bobbin, 1997). This chapter discusses additional experiments aimed at exploring the roles of extracellular ATP and its receptors exposed to perilymph in the organ of Corti.

A WORKING HYPOTHESIS: ATP AND ATP RECEPTORS LOCATED ON DEITERS’ CELLS MODULATE COCHLEAR MECHANICS Research into the mechanics of individual cells in the organ of Corti has focused on the outer hair cells (OHCs) and their somatic motility in response to voltage alterations (Brownell, 1996; Bobbin, 1996; Xue, Mountain, and Hubbard, 1995). Little attention has been given to Deiters’ cells, the only additional cell type in the organ of Corti that has been reported to move in response to a stimulus. Dulon, Blanchet, and Laffon (1994) demonstrated that Deiters’ cells move and alter their tension in response to an increase in internal calcium levels. Since ATP induced activation of P2X2 ATP receptors results in an inward current of Na+ and Ca2+ in Deiters’ cells (Ashmore and Ohmori, 1990; Dulon, Moataz, and Mollard, 1993), ATP may also induce a stiffness change in Deiters’ cells. A model on how a change in the stiffness of Deiters’ cells may alter cochlear mechanics has been presented by Kolston and Ashmore (1996). By puffing ATP onto the Deiters’ cell and measuring the response, Dulon (1995) suggested that the P2X2 ionotropic ATP receptors were localized near where the Deiters’ cell “cups” the OHC at the base of the OHC and so the receptors are exposed to perilymph. Definitive evidence for a

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perilymph exposed location is shown in Figure 7–2. Figure 7–2 shows an ATP-evoked response in a patch of Deiters’ cell membrane removed from a location near the nucleus [and so exposed to perilymph]. The ATP induced a large inward current typical of the P2X2 receptors (Chen, Parker, Barnes, Deininger, and Bobbin, 2000; Chen and Bobbin, 1998; Chen, LeBlanc, and Bobbin, 1997; and Chen et al., 1998). Thus, there is no doubt that P2X2 ionotropic ATP receptors are localized on the perilymph surface of Deiters’ cells.

EXPERIMENTAL EVIDENCE FOR AND AGAINST THE HYPOTHESIS Agonists on Distortion Product Otoacoustic Emissions (DPOAEs) The DPOAEs reflect the mechanical motion of the cochlear partition (Kemp, 1998; Mills, 1998). When placed in perilymph, agonists that acti-

Figure 7–2. Current trace of an ATP induced response recorded from an outside-out patch of membrane removed from an area of Deiters’ cell located near the nucleus at a holding potential of -70 mV (Bobbin, R. P. and Ricci, A., unpublished data). The upward deflection at the onset and the downward deflection at the offset of the delivery of the ATP is an electrical artifact due to the switching of the ATP delivery system.


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vate ATP receptors decrease the magnitude of the cubic DPOAE and increase the magnitude of the quadratic DPOAE (Bobbin et al., 2000; Kujawa, Erostegui, Fallon, Crist, and Bobbin, 1994). In addition, these agents alter cochlear sensitivity as monitored by changes in the summating potential (SP), the compound action potential of the auditory nerve (CAP), and N1 latency (Bobbin and Thompson, 1978; Kujawa, Erostegui et al., 1994). Thus, the tentative conclusion was reached that these drugs when placed in perilymph acted on P2X2 ATP receptors to alter cochlear mechanics. Although an action of these drugs at other cell types in the cochlea [such as pillar cells or Hensen’s cells which posses ATP receptors] cannot be excluded (Chen et al., 1998), the data are consistent with the effects being due, at least in part, to activation of ATP receptors exposed to perilymph and located on Deiters’ cells (Bobbin et al., 1998). Current evidence suggests that the OHCs and their motility may be the structural and mechanical basis for the cochlear amplifier (active process) that provides the low level sensitivity to the function of the cochlea (Brownell, 1996). Deiters’ cells, through their anatomical support at the base and apex of the OHCs, provide a direct means for controlling the force against which the OHCs must move. In this manner, the Deiters’ cells may adjust the set point and the operating range of the OHCs (Bobbin et al., 2000).

Antagonists on the Quadratic DPOAE Both suramin and PPADS, ATP antagonists, reversibly affect the quadratic DPOAE when placed in the perilymph compartment (Chen et al., 1998; Kujawa, Fallon, and Bobbin, 1994; Skellett et al., 1997). Frank and Kossl (1996) suggested that the quadratic DPOAE is a sensitive indicator of the “set point” of the cochlear amplifier. Thus, the effects of these drugs on the quadratic DPOAE may be evidence that endogenous ATP alters the set point of the cochlear amplifier. After a period of silence, turning on the primaries results in a slow increase followed by a decrease in the magnitude of the quadratic DPOAE over time (Figure 7–3; Brown, 1988; Kirk and Johnstone, 1993; Whitehead, Lonsbury-Martin, and Martin, 1991; Kujawa, Fallon, and Bobbin, 1995; Kujawa, Fallon, Skellett, and Bobbin, 1996; Lowe and Robertson, 1995). As shown in Figure 7–3, this change in the magnitude of the quadratic over time is dramatically affected by PPADS (Chen et al., 1998). In the presence of PPADS immediately upon turning on the primaries, the magnitude of the quadratic DPOAE is reduced more than 10 dB. During continuous exposure to the primaries, the quadratic DPOAE remains suppressed (0.3 M PPADS) or it is suppressed and then slowly increases in magnitude returning to near control values over about a 10 min period (1 mM, PPADS; Chen et al., 1998; Bobbin et al., 2000). Our premise is that PPADS is acting by blocking the actions of endogenous extracelluar ATP, although the actions of PPADS may also be in

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Figure 7–3. Effect of PPADS on quadratic f2 -f1 DPOAE (f2 - f1 = 1.25 kHz) amplitude alterations during continuous primary stimulation (f1 =6.25 kHz, f2 = 7.5 kHz, L1 = L2 = 60 dB SPL). Response amplitude as recorded following the second control perfusion (AP2), following increasing concentrations of PPADS (a, 0.033 mM; b, 0.1 mM; c, 0.33 mM; d, 1.0 mM) and a post drug wash (e, W1). The AP2 trace is repeated in each frame for reference. Pooled errors are shown in the upper right hand corner of each frame. Each data point represents a 10-spectra average and required 5 sec to complete. The break in response amplitude trace (C-D) represents 1 min during which the primaries were turned off. Points A - F (a) were used to calculate magnitudes of component amplitude changes for statistical analysis. Symbols represent means of n = 6. Noise floor averaged about -12 dB. (From “Additional pharmacological evidence that endogenous ATP modulates cochlear mechanics,” by C. Chen, R. A. Skellett, M. Fallon, and R. P. Bobbin, 1998. Hearing Research, 118, 47–61. Copyright 1998 by Hearing Research. Reprinted with permission.)

part due to its ability to prevent the breakdown of ATP by inhibiting an ectoATPase (Bobbin et al., 1998). In any case, the effects of PPADS on the quadratic DPOAE are supportive of the hypothesis that the magnitude of the quadratic DPOAE is greatly affected by endogenous extracellular ATP possibly acting on Deiters’ cells (Bobbin et al., 1998; Chen et al., 1998; Skel-


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lett et al., 1997). In addition, the quadratic DPOAE may be affected by efferent activity. It is a well known fact that the efferent nerve fibers enervating the organ of Corti affect cochlear mechanics (Mountain, 1980; Siegel and Kim, 1982; Puel and Rebillard, 1990; Kujawa, Glattke, Fallon, and Bobbin, 1993; Kujawa, Glattke, Fallon, and Bobbin, 1994; Lieberman, Puria, and Guinan, 1996). Kirk and Johnstone (1993) suggested that the activity of the efferent nerve fibers totally accounts for the change in the magnitude of the quadratic DPOAE over time that is shown in Figure 7–3.

Do the Efferent Nerve Fibers Have a Role in the Changes in the Quadratic DPOAE Magnitude? There is no doubt that the efferent nerve fibers entering the cochlea are active during sound exposure to the ipsilateral or contralateral ear. Mountain (1980) and Siegel and Kim, (1982) demonstrated that electrical activation of the efferent nerve fibers affects the magnitude of both the quadratic and cubic DPOAE and the effects were blocked by curarae. Puel and Rebillard (1990) demonstrated the suppression of the cubic DPOAE with noise presented to the contralateral ear. Kujawa, Glattke et al. (1993 and 1994) confirmed these results and showed they were due to acetylcholine acting on a receptor similar to the nicotinic receptor studied at the single OHC by Erostegui, Norris, and Bobbin (1994). These effects are blocked by strychnine, an antagonist of the acetylcholine receptor on the OHCs (i.e., the alpha 9 containing nicotinic receptor; for review see Bobbin, 1996 and 1997; Bobbin and LeBlanc, 1999). The transmitter of the efferent nerve endings synapsing on the OHCs is well known to be acetylcholine (Bobbin, 1996 and 1997). Kirk and Johnstone (1993) demonstrated that contralateral sound suppressed the quadratic DPOAE and the effects were blocked by strychnine. While monitoring the cubic DPOAE in cat, turning on the primaries activated the ipsilateral efferent nerve fibers within one second producing approximately a 3 dB suppression of the cubic DPOAE (Liberman et al., 1996; Liberman and Kujawa, 1999). Contralateral sound stimulation activated additional efferent nerve fibers to induce an additional 2 dB of suppression for a total efferent suppression of the cubic DPOAE of about 5 dB (Liberman et al., 1996; Liberman and Kujawa, 1999). Conflicting data have been presented as to whether ipsilateral activation of efferent nerve fibers can alter the quadratic DPOAE. Kirk and Johnstone (1993) presented compelling evidence that the ipsilateral efferent nerves acting via the neurotransmitter GABA, and not acetylcholine, play a large role in the decrease in the magnitude of the quadratic DPOAE that was illustrated in Figure 7–3. They reported that strychnine had no effect whereas tetrodotoxin, a neuronal sodium channel blocker, and bicuculline, a GABA antagonist, abolished the decrease in the quadratic DPOAE. In contrast, both Lowe and Robertson (1995) and Kujawa et al. (1995) failed

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to detect an effect of sectioning the efferent nerve fibers in the brain stem on the decrease in the quadratic DPOAE. Kujawa et al. (1995) did report a significant lessening of the decrease in the quadratic DPOAE over time with curarae, an acetylcholine antagonist, and a non-significant lessening with bicuculline. Bicuculline has been shown to block the effects of acetylcholine at the alpha 9 containing nicotinic receptor on the OHCs (Kujawa, Glattke et al., 1993 and 1994; Erostegui et al., 1994). Since sectioning the efferent nerve fibers in the brain stem had no effect, Kujawa et al. (1995) attributed the lessening of the decrease to nonselective effects of the curarae and bicuculline and not due to block of the action of the ipsilateral activated efferent nerve fibers (Kujawa et al., 1995). We reasoned that if the lessening of the decrease in the quadratic DPOAE over time by curarae and bicuculline reported by Kujawa et al. (1995) was actually due to blockade of the efferent neurotransmitter acetylcholine, then strychnine, a more selective blocker of the neurotransmitter acetylcholine at the OHCs, should act just like curarae and bicuculline. Figure 7–4 illustrates preliminary results indicating that strychnine (10 M) does act like curarae and bicuculline in that it lessened the decrease occurring in the magnitude of the quadratic DPOAE over time. This time course and magnitude of the strychnine effect is very similar to the results that Kujawa et al. (1995) obtained with curarae and bicuculline. Both in the Kujawa et al. (1995) and in the Lowe and Robertson (1995) studies large differences were found between runs within the same animal. Because of these large variations, the data were normalized by assigning zero to the initial value for each run. To extract the strychnine sensitive component from the total quadratic DPOAE change over time, the values obtained after strychnine perfusion were subtracted from the average values obtained after artificial perilymph perfusions (before and after strychnine). This strychnine sensitive component in the magnitude of the quadratic DPOAE is given in Figure 7–5. The magnitude approaches 3 dB similar to the ipsilateral suppression of the cubic DPOAE magnitude reported by Lieberman et al. (1996). The change appears to be composed of two phases, an early phase and a late phase. Figure 7–5 illustrates that an early strychnine sensitive enhancement of the quadratic DPOAE (about 0.5 dB) is followed by a late phase of strychnine sensitive suppression that appears to start at about 3 min after primary onset and slowly increases in magnitude until it reaches a maximum at about 9 min. A similar late onset for ipsilateral suppression of the quadratic DPOAE is seen in a few of the figures shown by Kirk and Johnstone (1993). Whether the early and late phases observed in Figure 7–5 are related to the slow and fast components of efferent suppression described by Sridhar, Liberman, Brown, and Sewell (1995) remains to be determined. Based on the pharmacology with three drugs, curarae, strychnine and bicuculline, an efferent component in the time varying alterations in the magnitude of the quadratic DPOAE has been conclusively demonstrated.


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Figure 7–4. Effect of strychnine (stry) on quadratic f2 -f1 DPOAE (f2 - f1 = 1.25 kHz) amplitude alterations during continuous primary stimulation (f1 =6.25 kHz, f2 = 7.5 kHz, L1 = L2 = 70 dB SPL). Response amplitude as recorded following 15 min perfusions of a control (AP), strychnine (10 M free base) and a post drug wash (wash). The primaries were off during the perfusions and turned on approximately 2 min following termination of the perfusions. The break in response amplitude trace (9 – 10.5 min) represents 1.5 min during which the primaries were turned off. Noise floor averaged about -12 dB. For additional methodology see Chen, Skellett, et al., (1998).

Given that there is an effect of efferent nerve fiber activity on the magnitude of the quadratic DPOAE in response to ipsilateral sound stimulation, then efferent nerve fiber activation must play a role in the response of the quadratic over time following administration of PPADS (Chen et al., 1998; Bobbin et al., 2000). This is certainly possible given that PPADS has little influence on the afferent portion of the loop, as indicated by the slight effect on the CAP (Chen et al., 1998). Future experiments such as combining strychnine with PPADs will be carried out to help in determining this role.

Antagonists and Agonists Action on Cochlear Microphonics Cochlear microphonics (CM) reflect the current passing through the transduction channels in the stereocilia of the IHCs and OHCs. If the ATP re-

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Figure 7–5. Ipsilateral efferent induced change in the quadratic f2 -f1 DPOAE (f2 - f1 = 1.25 kHz) amplitude during continuous primary stimulation (f1 =6.25 kHz, f2 = 7.5 kHz, L1 = L2 = 70 dB SPL) as indicated by the strychnine sensitive component. Response amplitude was normalized by assigning zero to the first value recorded in each run. To generate the graph the strychnine values were subtracted from the average of the AP and wash values for each set of runs and then these subtracted strychnine values were averaged (n = 4 strychnine perfusions in 3 animals). The break in response amplitude trace (9–10.5 min) represents 1.5 min during which the primaries were turned off. Noise floor averaged about -12 dB.

ceptors on Deiters’ cells were activated and Deiters’ cell processes moved then this might be expected to alter the distances between the basilar membrane, reticular lamina and the tectorial membrane. These changes in distances should alter the orientation of the stereocilia at rest and possibly alter the transduction current passing through the stereocilia (i.e., CM). Based on this model, blockade of endogenous ATP might be expected to alter the current passing through the stereocilia or CM magnitude. To date, studies indicate that the ATP antagonists, PPADs and suramin, when placed in perilymph have no effect on the CM intensity function recorded single ended from the scala vestibuli in the basal turn of the cochlea (Kujawa, Fallon, et al., 1994; Skellett et al., 1997; Chen et al., 1998 ; LeBlanc and Bobbin, 1999). To counter the argument that these single ended recordings


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do not reflect near threshold values, the 1 V CM was recorded using an electrode on the round window or an electrode in scala vestibuli of the basal turn before and after administration of artificial perilymph and PPADs to the perilymph compartment. Subtracting the dB values necessary to evoke a 1 V CM after artificial perilymph from those dB values after PPADS administration demonstrates no effect (± 1 dB) of the drug (data not shown). Consistent with this lack of effect of antagonists, only high concentrations of agonists had an effect on CM and the effect was slight (Kujawa, Erostegui et al., 1994). The failure to detect a change in the CM induced by ATP antagonists placed in perilymph probably means that endogenous ATP acting on receptors exposed to perilymph has no effect on the transduction current. On the other hand, the results may be evidence for the hypothesis that ATP has an action on Deiters’ cells. For any possible effect of Deiters’ cells on stereocilia position will most likely be negated by adaptation and result in an unchanged CM.

Antagonists Action on the Tonotopic Distribution Along the Cochlear Partition An alteration in tension or length of Deiters’ cells may not affect transduction current but it may alter the mechanics of the cochlear partition. If this occurs then the place of maximal stimulation on the basilar membrane or the tonotopic organization will be shifted. To test this hypothesis, the technique previously described by Bobbin, Fallon, Li, and Berlin (1991) was utilized. Briefly, the dB level necessary to record a 1 V CM utilizing a wire on the round window was compared to the dB necessary to record a one microvolt CM utilizing a differential recording from wires inserted into holes made in the basal turn in the scala vestibuli and scala tympani in response to a range of frequencies of sound. The dB difference between the intracochlear electrodes and the round window electrode isoresponse functions was calculated and defined as sensitivity. Thus, a negative number indicates greater sensitivity (lower sound pressure level) at the intracochlear recording site. The high frequency slope of this CM difference function is used as an indication of the characteristic frequency of that place (18–20 kHz in Figure 7–6). As shown in Figure 7–6, PPADS 1 mM perfused through the perilymph compartment had no detectable influence on this function when compared to that obtained after artificial perilymph perfusion. Thus, it was concluded that if endogenous ATP affects the tonotopic distribution of frequency along the partition, the technique used was not sufficiently sensitive to detect the change. For example, lowering body temperature 10°C has been reported to induce only a 250 Hz basal shift of the excitation pattern along the cochlear partition (deBrey and Eggermont, 1978), which is well below our sensitivity. However, the negative results are consistent with a model suggesting that altering the tension or stiffness of Deiters’ cells will not alter the position of the tuning along the basilar

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Figure 7–6. Lack of change in the sensitivity of a location in the basal turn of the guinea pig cochlea (18–20 kHz) after perfusion of artificial perilymph (AP) and PPADS (1 mM) through the perilymph compartment of the cochlea for 15 min. The dB necessary to evoke a 1 V RMS CM in response to various frequencies of sound, isoresponse function, was recorded from the round window and an intracochlear set of electrodes (differential: scala tympani-scala vestibuli) placed about 1 mm from the round window niche. The dB difference between the intracochlear electrodes and the round window electrode isoresponse functions was calculated and defined as sensitivity. A negative number indicates greater sensitivity (lower sound pressure level at the intracochlear recording site).

membrane (Kolston and Ashmore, 1996). Likewise, Murugasu, and Russell (1996) demonstrated that efferent nerve transmitter action at the OHCs does not alter the stiffness of the cochlear partition or position of the tuning of the basilar membrane. Therefore, the lack of effect of PPADS on the tonotopic distribution may be consistent with our hypothesis regarding the role of extracellular ATP.

ATP Antagonists Action on N1 Latency PPADS applied to the perilymph compartment decreases CAP magnitude and increases N1 latency (Figure 7–7; Chen et al., 1998; LeBlanc and Bob-


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Figure 7–7. Effect of PPADS on CAP, N1 latency, negative SP and CM as a function of stimulus intensity. Shown are functions recorded after pre-drug artificial perilymph perfusion #2 (AP2; 15 min), after perfusion with increasing concentrations (0.033 - 1.0 mM; 15 min each) of PPADS, and after a second post-drug wash with artificial perilymph (W2; 15 min). Functions recorded after 0.033 and 0.1 mM PPADS and wash 1 are not shown for clarity. Data are represented as means ± S.E. across 5 animals. (From “Additional pharmacological evidence that endogenous ATP modulates cochlear mechanics,” by C. Chen, R. A. Skellett, M. Fallon, and R. P. Bobbin, 1998. Hearing Research, 118, 47–61. Copyright 1998 by Hearing Research. Reprinted with permission.)

bin, 1999). In terms of dB sound pressure equivalents (i.e., the dB change in the intensity of the sound necessary to achieve the predrug value), PPADS increases N1 latency about 30 dB (Chen et al., 1998) compared to a

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6 dB change in the peak-to-peak magnitude of the CAP (Chen et al., 1998; LeBlanc and Bobbin, 1999). This result contrasts with the effects observed with most drugs. For example, salicylate (Puel, Bledsoe, Bobbin, Ceasar, and Fallon, 1989; Puel, Bobbin, and Fallon, 1989), glutamate antagonists (Litmann, Bobbin, Fallon, and Puel, 1989) and nimodipine (Bobbin, Jastreboff, Fallon, and Littman, 1990) suppress CAP magnitude to a greater or equal dB equivalent than they increase N1 latency. It is well documented that a change in body or cochlear temperature (Brown, Ian Smith, and Nuttall, 1983) and occasionally intense sound exposure (Puel, Bobbin and Fallon, 1988a; Puel, Bobbin and Fallon, 1988b) will alter N1 latency more than CAP magnitude. As shown in Figure 7–8, lowering the body temperature of a guinea pig to 31°C from the standard 38°C resulted in about a 30 dB equivalent shift in N1 latency compared to about a 5 dB equivalent shift in CAP magnitude. Similarly after exposure to a 6 kHz 95 dB intense tone, a 6 kHz evoked CAP was affected less than N1 latency (Puel et al., 1988a). The mechanisms contributing to any change in N1 latency could be extensive (Brown et al., 1983). In general, the global effect of lowering body temperature is to slow all biological processes, for example, synaptic

38°C 31°C

Figure 7–8. Effects of altering body temperature on cochlear potentials evoked by 10 kHz tone pips. Shown are peak-to-peak values obtained by a single electrode in scala vestibuli of the basal turn. For methods of recording see Chen et al., 1998.


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transmission and nerve conduction. Lowering body temperature slows basilar membrane travel time (deBrey and Eggermont, 1978). In the intense tone study, Puel et al. (1988a) speculated that the 6 kHz traveling wave evoking the N1 latency was “slowed” as it passed through the damaged area at the 8 kHz place (1⁄ 2 octave shift of the 6 kHz intense tone) and once it reached the 6 kHz place it evoked a nearly normal CAP. Therefore, it is tempting to speculate that PPADS, which also affects N1 latency more than CAP magnitude, does so by slowing basilar membrane travel time.

ATP Antagonist Interaction with Intense Sound Exposure To further test the hypothesis that endogenous ATP affects the function of the cochlea, PPADS was administered to the perilymph compartment during an exposure to a moderately intense sound (95 dB SPL; 15 min; 6.7 kHz). This level of intense sound apparently induces only a temporary loss in cochlear mechanics (Puel et al., 1988a; Puel et al., 1995). Therefore, it was reasoned that if endogenous ATP has an action on Deiters’ cells to alter their stiffness then PPADS by blocking the action of ATP may alter the effects of this intense sound on cochlear mechanics. The methods used were very similar to those used previously (Puel et al., 1988a and 1988b; Puel et al., 1995). Animals were exposed to a moderately intense sound (95 dB SPL; 15 min; 6.7 kHz) 10 min after the start of a perfusion with artificial perilymph (intense sound group) or 1 mM PPADS (intense sound plus PPADS group) and the tone was terminated 10 min before the end of perfusion. A third group received PPADS alone (PPADS alone group). Cochlear potentials evoked by 10 kHz tone bursts were recorded from an electrode in basal turn scala vestibuli. As shown in recordings of the cochlear microphonics (Figure 7–9) and N1 latency (Figure 7–10), the effects of the moderately intense sound combined with PPADS were larger than the effects induced by intense soundalone or PPADS alone. This is especially interesting since the intense tone by itself induced no significant shift in CM or N1 latency. In addition, the enhancing effect was readily reversed by washing out the drug (Figures 79 and 7-10).

Why No Effect of the Intense Sound on CM? The lack of effect of the intense 6.7 kHz tone on CM requires discussion. There is no doubt that intense sound reduces the magnitude of the CM (e.g., Nakajima, Hubbard, and Mountain, 2000). The CM in the LeBlanc and Bobbin (1999) study (Figure 7–9) was recorded with an electrode in the perilymph scala at the same place as the CM was recorded in Figure 7–6. According to Figure 7–6, the place of maximal stimulation for that recording site was about 18–20 kHz. Thus, if the 6.7 kHz intense tone dam-

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Figure 7–9. Effects of treatment with (a) an intense sound alone (6.7 kHz, 95 dB SPL, 15 min), ( b ) a combination of the intense sound with 1 mM PPADs, and (c) 1 mM PPADS alone, on 56 dB SPL soundevoked CM. Shown are values obtained after pre-treatment artificial perilymph perfusion (AP), after treatment (treatment) and after washes with artificial perilymph (wash). Data are displayed as means ± S.E. across n = 5 animals per treatment. The * indicates a significant (P < 0.05) difference from AP and # indicates a significant (p

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