PHYSIOLOGICAL REVIEWS   Vol. 78 No. 2 April 1998, pp. 429-466
Copyright ©1998 by the American Physiological Society

Transduction Mechanisms in Vertebrate Olfactory Receptor Cells

DETLEV SCHILD AND DIEGO RESTREPO

Physiologisches Institut, Universität Göttingen, Göttingen, Germany; and Department of Cellular and Structural Biology, School of Medicine, University of Colorado Health Sciences Center, Denver, Colorado

I. INTRODUCTION
II. GEOMETRY AND PASSIVE MEMBRANE PROPERTIES OF OLFACTORY RECEPTOR NEURONS
    A. Geometry
    B. Capacitance and Resting Membrane Resistance
    C. Resting Membrane Potential
    D. Membrane Time Constant
    E. Sensitivity and Single Odorant Molecule Detection
    F. Summary
III. CONDUCTANCES INVOLVED IN ACTION POTENTIALS
    A. Voltage-Gated Sodium Currents
    B. Voltage-Gated Calcium Currents
    C. Potassium Conductances
    D. Summary
IV. ODORANT RESPONSES
    A. Recording Methods
    B. Odorant-Induced "Cation" Current
    C. Optical Recordings of Odorant Responses
    D. Diversity of Transduction Mechanisms
    E. Responses of Olfactory Neurons in Culture
    F. Responses of Vomeronasal Neurons
    G. Summary
V. ADENOSINE 3',5'-CYCLIC MONOPHOSPHATE-MEDIATED SIGNALING IN OLFACTORY CELLS
    A. Olfactory Receptors, G_olf, and cAMP Formation
    B. Olfactory Cyclic Nucleotide-Gated Conductance
    C. Calcium-Activated Chloride Current
    D. Summary
VI. INOSITOL TRISPHOSPHATE-MEDIATED SIGNALING IN OLFACTORY CELLS
    A. InsP₃ Formation and Receptors
    B. Inositol Polyphosphate-Regulated Conductance(s)
    C. Electrophysiological Studies of InsP₃-Mediated Odor Responses
    D. Summary
VII. CALCIUM AS A THIRD MESSENGER IN OLFACTORY TRANSDUCTION
VIII. GASEOUS MESSENGERS IN OLFACTORY NEURONS (CARBON MONOXIDE AND NITRIC OXIDE)
IX. IONIC CONCENTRATIONS IN MUCUS AND ION HOMEOSTASIS
X. MULTIPLE PATHWAYS: IMPLICATIONS FOR OLFACTORY CODING
XI. SUMMARY AND PERSPECTIVES
REFERENCES

	ABSTRACT

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Schild, Detlev, and Diego Restrepo. Transduction Mechanisms in Vertebrate Olfactory Receptor Cells. Physiol. Rev. 78: 429-466, 1998.  Considerable progress has been made in the understanding of transduction mechanisms in olfactory receptor neurons (ORNs) over the last decade. Odorants pass through a mucus interface before binding to odorant receptors (ORs). The molecular structure of many ORs is now known. They belong to the large class of G protein-coupled receptors with seven transmembrane domains. Binding of an odorant to an OR triggers the activation of second messenger cascades. One second messenger pathway in particular has been extensively studied; the receptor activates, via the G protein G_olf , an adenylyl cyclase, resulting in an increase in adenosine 3',5'-cyclic monophosphate (cAMP), which elicits opening of cation channels directly gated by cAMP. Under physiological conditions, Ca²⁺ has the highest permeability through this channel, and the increase in intracellular Ca²⁺ concentration activates a Cl current which, owing to an elevated reversal potential for Cl, depolarizes the olfactory neuron. The receptor potential finally leads to the generation of action potentials conveying the chemosensory information to the olfactory bulb. Although much less studied, other transduction pathways appear to exist, some of which seem to involve the odorant-induced formation of inositol polyphosphates as well as Ca²⁺ and/or inositol polyphosphate-activated cation channels. In addition, there is evidence for odorant-modulated K⁺ and Cl conductances. Finally, in some species, ORNs can be inhibited by certain odorants. This paper presents a comprehensive review of the biophysical and electrophysiological evidence regarding the transduction processes as well as subsequent signal processing and spike generation in ORNs.

	I. INTRODUCTION

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The morphology of olfactory receptor neurons (ORNs) was first described by Schultze (314, 315; reviewed in Ref. 366) 140 years ago. How ORNs function, however, has remained a riddle that still is not completely solved. In 1985 and 1986, research on olfactory transduction took a decisive turn. Two reviews published by Getchell and Lancet described the functional properties of vertebrate ORNs (106) and the processes in olfactory perception ranging from biochemistry to systems theory (188). This wealth of knowledge had been accumulated using classical morphological and biochemical methods and classical electrophysiological tools, above all extracellular recordings. Anderson and Ache's study of 1985 (in the lobster) (7) and Trotier's study of 1986 (in the salamander) (343) were the first to report the use of the patch-clamp technique to provide a description of voltage-gated membrane currents, single-channel recordings, and odorant-induced currents in ORNs. The various patch-clamp recording configurations (115), other biophysical methods, and molecular biological techniques allowed a more detailed study of signal transduction in the ORN. Since then, a number of reviews and minireviews have appeared, each dealing with a specific topic in olfactory perception, e.g., perireceptor events (107, 270, 271), the structure and function of receptor proteins (52, 189, 283), ultrastructural studies (232, 233), second messenger signaling (2, 9, 10, 40, 43, 47, 74, 295, 298, 372), cyclic nucleotide-gated channels (151, 357, 369, 372), Ca²⁺-dependent Cl channels in ORNs (185), and adaptation and concentration coding (340, 344) as well as aspects of coding and information conveyance between olfactory epithelium and olfactory bulb (52, 149, 189, 247, 252, 304, 316, 322). However, a comprehensive review of cellular studies of olfactory transduction focused on electrophysiological and biophysical aspects is lacking.

Here we review the biophysical, electrophysiological, and some of the biochemical work done in ORNs over the past 10 years. Although we focus on the functional aspects of vertebrate olfactory transduction, we point out interesting parallels to chemosensory transduction mechanisms in invertebrate ORNs (2, 3, 137, 139, 140, 329), eukaryotic microorganisms (346), and vomeronasal receptor neurons (197). Molecular biological studies are mentioned where appropriate but are not covered comprehensively. The reader is referred to recent reviews covering biochemical and molecular biological aspects of olfactory transduction (2, 47, 52, 189, 247, 295, 298, 322). This review of electrophysiological and biophysical studies documents significant advances in our understanding of olfactory transduction in the last 10 years. More importantly, the comprehensive review of the literature makes a compelling argument that olfactory receptors are more complicated than heretofore imagined and reveals the limitations in our knowledge of the complexity of the mechanisms underlying olfactory transduction.

	II. GEOMETRY AND PASSIVE MEMBRANE PROPERTIES OF OLFACTORY RECEPTOR NEURONS

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Olfactory receptor cells in vertebrates are bipolar neurons with a small soma, a single dendrite, cilia or microvilli attached to the olfactory knob or vesicle formed at the dendritic end, and an axon projecting to the olfactory bulb (Fig. 1). The soma has a round or ellipsoidal shape with the short and long axes ranging from 4 to 15 µm and from 7 to 21 µm, respectively. Among the species studied using electrophysiological and biophysical techniques, the largest ORNs are found in newts (182) and salamanders [Ambystoma tigrinum (92) and Necturus maculosus (76)], whereas the smallest ORNs are found in zebrafish (Danio rerio) (65). The unbranched axon is ~200 nm in diameter, and its length, when assessed in isolated cell preparations, varies considerably. The dendrite is ~1-3 µm thick and between 5 and 120 µm long (253, 284). In a preparation of isolated receptor cells, the dendrites may shorten either due to the preparation procedure (307) or due to an excessive Ca²⁺ load (310). The dendrite ends in a dendritic knob (diameter 2-3 µm) from which several cilia issue. The number of cilia appears to vary approximately between 5 and 40 (76, 147, 148, 236, 253, 284). The diameter of a cilium is 100-250 nm (depending on the species), i.e., at or below the resolution limit of light microscopes, and its length is in the range between 5 and 250 µm (124, 166, 279, 284, 371). A cilium is thicker at its base and tapers down its length somewhat (231-233, 318). The typical ciliary 9 + 2 structure changes over the length of a cilium, because the number of microtubules is reduced. In frog, short and medium-sized (up to 90 µm) cilia move irregularly, whereas the long cilia (up to 250 µm) do not move (124, 166). In mammals, the cilia are immotile (231).

View larger version (79K): [in this window] [in a new window]   FIG. 1.   Top: scanning electron micrographs of human olfactory receptor cells. Left: olfactory mucosa viewed from side. Cell with long, slightly prominent dendrite is a typical olfactory neuron. Scale bar, 10 µm. Right: view of surface of olfactory mucosa shows an olfactory knob with 8 cilia. Scale bar, 1 µm. [From Morrison and Costanzo (253).] Bottom: schematic diagram of an olfactory neuron showing transduction compartments (cilia and dendritic knob), dendrite, soma, and beginning of axon as well as some of the characteristic properties of compartments. InsP3 , inositol trisphosphate; NO, nitric oxide.

A comparison of morphological data (124, 284) with electrophysiological reports clearly indicates a tendency for isolated cells to exhibit shorter cilia, although cilia as long as 130 µm have been found and recorded from (166). Occasionally, isolated ORNs lose all the cilia during the dissociation procedure. Deciliated olfactory receptor cells of the newt do not respond to odors (182). For a survey of the electrical properties of olfactory cilia in frog, see References 159, 164, 166.

Often during recordings in the whole cell mode of the patch-clamp technique, the seal with the patch pipette is established on the soma, which is the part of the cell most distant from the cilia. Because of this, space-clamp effects due to the fact that the resistance of the plasma membrane is finite can come into play, particularly in large ORNs with long cilia during odorant stimulation. The effects of space clamp in ORNs have been described by various investigators (166, 213, 276).

The receptor cells in vomeronasal organs (6, 20, 171, 318, 319) and a number of receptor cells in the olfactory epithelium of fish (78, 128, 250, 254, 338, 358, 367, 368), reptiles (112, 352), and amphibia (171) bear microvilli instead of cilia. Microvillous cells have also been reported in the olfactory epithelium of mammals, in particular, in humans (242, 251, 253, 300). There are no reports on their function, and there is controversy on whether they project axons to the olfactory bulb. Thus one study found that when horseradish peroxidase (HRP) is injected into the olfactory bulb, HRP reaction products can be found with a certain delay in microvillous cells of the olfactory epithelium (300), whereas another study failed to find axonal processes stemming from putative microvillar cells in rat (54). In the vomeronasal organ of mouse and rat, two different subclasses of receptor neurons have been described that express two different G proteins and project differentially to the accessory olfacory bulb (131). In the goldfish, there is a correlation between the reappearance of microvillous cells after axotomy and the behavioral responses to pheromones (368). Interestingly, ORNs and axons from the different kinds of chemosensory neuroepithelia, i.e., olfactory proper versus accessory olfactory or vomeronasal mucosa, can be labeled differentially by lectin agglutinins (94, 95, 239).

The resting electrical properties including the capacitance (C), the resting membrane resistance (R_m), and the resting potential (u_r) of ORNs have been measured in various vertebrate and invertebrate species. The measured capacitance of ORNs (Table 1) is in the range from 0.7 to 35 pF. The smallest and the largest values have been found in zebrafish (65) and in newt (F. Kawai, personal communication), respectively. With the assumption of a standard value of 1 µF/cm² (corresponding to 1 pF/100 µm²), the membrane surface of ORNs is thus in the range between 70 and 3,500 µm².

In patch-clamp recordings from ORNs, the seal resistance and the resting membrane resistance are often in the same range (between 1 and 40 G). This can lead to problems in the interpretation of data, and it is therefore particularly important to make a clear distinction between the resting input resistance (R_i) and R_m . The R_i value is usually measured in the whole cell configuration using voltage clamp. Starting at a holding voltage of about 80 mV, a small voltage pulse u of typically 10 mV is applied, and the resulting asymptotic current shift I is measured. The u/I determined at u_r gives the R_i , which is the combined resistance of R_m and the seal resistance (R_seal)

(1)

whereby the series (or access) resistance (R_s) of the pipette is neglected here because it should be at least two orders of magnitude smaller than R_m or R_seal . From Equation 1 it follows that the R_m can be obtained from R_i and R_seal (as measured in the cell-attached mode)

(2)

Seal resistances as reported in recordings from ORNs are in the range between 1 G (182) and 53 G (219). Clearly, the approximation R_m ~ R_i is better the smaller the R_i/R_seal .

The resting membrane potential u_r in ORNs, measured as the zero-current potential in the whole cell configuration of the patch-clamp technique, has been reported in the remarkably large range between 90 mV (98) and 30 mV (305). These widely differing values might be explained by species differences. However, Firestein and Werblin (92) explained the deviation of u_r (in their study 54 mV) from the K⁺ equilibrium potential (E_K; in their study 98 mV) by the membrane being permeable for ions other than K⁺. A deviation of u_r from K⁺ potential (u_K) could be brought about by three other conductances.

First, a Cl conductance (76, 165) with Cl potential (u_Cl) > E_K as reported in ORNs of the mudpuppy (76), newt (184), and the clawed frog (361) would raise u_r . In the frog, this explanation might, however, be incorrect because the Ca²⁺-activated Cl conductance (g_Cl) is half-activated at intracellular Ca²⁺ concentrations of ~5 µM (165), which is almost two orders of magnitude higher than the resting Ca²⁺ concentrations in ORNs as obtained in imaging studies in frog (309-311). Hence, the Ca²⁺-dependent Cl conductance in frog is presumably not activated at rest.

A second conductance that necessarily affects the measurements of u_r is the seal between plasma membrane and patch pipette. With g_seal (=1/R_seal) and g_Ko (=1/R_Ko) being the seal conductance and the resting K⁺ conductance, and with the assumption that other conductances are negligible at rest, the resting membrane potential is given by the equation (113) where f_seal = g_seal/(g_seal + g_Ko), f_Ko = g_Ko/(g_seal + g_Ko) are the fractional conductances with u_seal and u_K being the corresponding equilibrium potentials. Because u_seal ~0 mV, u_r can be approximated by

(3)

This shows that a "good seal resistance" is a relative concept and that u_r is reliably determined only if R_seal >> R_Ko , a requirement difficult to meet in small neurons with a high resting resistance.

Frings and Lindemann (99), for example, have reported u_r approximately equal to 85 mV with R_i ~5 G and R_seal ~40 G. Indeed, with u_K = 91 mV, Equations 2 and 3 give R_Ko ~7.5 G and u_r = 79 mV, which is in the range of the measured value. Whenever R_i and R_seal happen to be of the same order of magnitude, the seal short circuits the membrane potential. For example, with R_i = 4 G, R_seal = 10 G, and u_K = 97 mV, Equations 2 and 3 yield u_r = 57 mV (90). The short-circuiting effect of the seal shunt has been modeled and analyzed in detail by Pongracz et al. (276).

An additional problem arises if the seal dependence of a measured resting membrane potential affects further steps of data evaluation. For example, to investigate the Cl equilibrium potential in mudpuppy ORNs, Dubin and Dionne (76) recorded the Cl channel blocker-dependent shift u of current-voltage (I-V) curves recorded in the cell-attached mode. The resulting value u added to the resting membrane potential as subsequently measured in the whole cell mode (in this case u_r was 40 mV) gave u_Cl = 45 mV. However, the seal-dependent inaccuracy in the measurement of u_r affects the resulting value for u_Cl so that u_Cl is more negative than 45 mV, the exact value depending on R_seal .

A third conductance that can influence the measurement of R_m and u_r is an inward rectifying cation conductance (g_h), which is permeable for K⁺ and Na⁺ (P_K/P_Na = 5) and activates at hyperpolarized potentials. This conductance has been reported in ORNs of catfish (246), frog (343, 344), mouse (227), and rat (222). In other systems, a similar conductance is crucial for the generation of membrane potential oscillations (230). Whether or not this conductance underlies the membrane potential oscillations observed in ORNs (98, 205) is not known. Interestingly, however, Tokimasa and Akasu (339) have shown in sympathetic neurons that a conductance of the same type can be activated by increased levels of cAMP.

The product of the resistance R_m and the cell capacitance C determines the membrane time constant (_m). Average values of R_m × C are ~60 ms. The membrane can be regarded as an RC element or first-order low-pass filter with corner frequency (f_c) of ~16 Hz that dampens odor-induced current components for frequencies f > f_c . It thus rejects "high"-frequency noise (214). The time constants of ORNs were measured in various species using current injections into ORNs through a patch pipette. The values obtained with gigaohm seals are at least one order of magnitude larger than those obtained with sharp microelectrodes [e.g., 4.2 ms in the tiger salamander (226)]. Owing to the high resting resistance and, at least in some species, a relatively high capacitance, the time constant _m can be as long as ~100 ms [e.g., in squid (217), or salamander (92)]. This implies that, because of the high R_m , ORNs can be excited by very small currents but that the depolarization is relatively slow because of the long time constants. This was nicely demonstrated by Lynch and Barry (219) as well as by Maue and Dionne (227), who observed that the current supplied by the opening of a single ion channel was able to trigger an action potential. The underlying membrane biophysics have been analyzed in detail by Lynch and Barry (219). The delay of the action potential after channel opening depended on the current amplitude. For the smallest current that elicited an action potential, it was in the range of the membrane time constant.

The above-mentioned evidence that the current through a single channel can excite an olfactory neuron poses the question of whether a single odorant molecule binding to an odorant receptor can induce an action potential. Menini et al. (238) suggested this. These authors applied an odorant at concentrations well below the K_1/2 for the dependence of odor-induced steady-state current on odor concentration. The resulting current fluctuations were stereotyped, "quantal-like" in amplitude ranging from 0 pA (failure) to 9.5 pA, with an average response of ~1 pA (holding potential, 50 mV). Based on these data, Menini et al. (238) concluded that odors induce quantal-like current fluctuations presumably triggered by the binding of a single odorant molecule to its receptor. These data do, however, not prove that binding of one odorant molecule induces a quantal current. To show this, it would be necessary to demonstrate that the frequency of occurrence of the current fluctuations is equal to the frequency of binding of odor molecules to the receptor (23, 317). Moreover, the detection of a single molecule by a receptor cell would require that a quantal current could induce an action potential. Although this is not excluded, particularly in the case when amplification of channel activity occurs through opening of Ca²⁺-activated conductances (see sects. V and VI), the effect of quantal currents on membrane potential was not analyzed by Menini et al. (238).

On the other hand, Lowe and Gold (215) have shown that low concentrations of odorants elicit random current fluctuations in olfactory receptor cells from rat. These authors argue convincingly that these current fluctuations are not because of single molecular events. They show that the power spectrum for odor-induced currents is identical to the power spectrum of the current induced by increases in the concentration of the second messenger cAMP in the absence of odorants. They conclude that the current fluctuations induced by odorants are due to noise intrinsic to the transduction mechanism (intrinsic noise) and suggest that quantal detection is not a common property of olfactory receptor cells. Under basal conditions, the intrinsic noise may not be apparent because the steep dependence of current on cAMP concentration (Hill coefficient of 2-4) acts like a threshold. Addition of odorant would then elevate the level of activity above the threshold, thereby unmasking the intrinsic noise signals.

However, because of the differences in the experimental designs of the experiments carried out by Menini et al. (238) on the one hand, and Lowe and Gold (215) on the other, the data provided by Lowe and Gold do not rule out the possibility that the current fluctuations in the Menini study may be due to single molecular events (see also Ref. 295).

	III. CONDUCTANCES INVOLVED IN ACTION POTENTIALS

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Vertebrate olfactory receptor cells are neurons. They have an axon along which they send encoded information to the central nervous system (CNS). In this section we review some biophysical properties of the conductances that are involved in the generation of action potentials in ORNs.

As outlined in section II, the resting membrane potential appears to be determined by a very low resting K⁺ conductance, which consists of more than one type of channel: 1) an inward rectifiying conductance g_h (222, 227, 246, 343) and 2) a slowly inactivating delayed rectifier type of K⁺ conductance that has been observed in squid (217), fish (65, 246, 260), Chilean toad (69), frog (305, 343), salamander (92), rat (220, 342), and human (292). At the single-channel level, a slowly activating K⁺ current has been described (220) that could underlie the resting delayed rectifier conductance.

The reports of the voltage-gated Na⁺ conductances (g_Na) in ORNs for various species differ in some respects. 1) In the tiger salamander (92) and rat (280), it has been reported to be relatively insensitive to tetrodotoxin (TTX), whereas it is not in other species (65, 69, 246, 305). 2) It activates at about 60 mV in Xenopus laevis (305) and frog (277), 50 mV in catfish (246) and Chilean toad (69), 30 mV in squid (217) and salamander (92), and about 45 mV in rat (280, 342), human (292), salmon (260), zebrafish (65), and mudpuppy (76). This variability could be species dependent, but it could also be due to a voltage drop either along axonal compartments or across the pipette resistance. In the last case, the same voltage drop, i.e., the same right shift of I-V curves, is also seen for other conductances that are not localized to the axon.

The amplitudes of the Na⁺ currents show a considerable variability and, in some cells (292, 343), Na⁺ current could not be measured at all. The reason for this variability and occasional lack of Na⁺ currents in ORNs could be that the channels are primarily located in the axonal membrane, which is partially lost during preparation. This is consistent with the finding of Maue and Dionne (227) who were unable to observe single Na⁺ channels on the somata of mouse ORNs. Frings and Lindemann (99) found no effect of TTX when applied to the mucosal surface, whereas it blocked spike generation when applied to the interstitial surface of the mucosa. However, in the mudpuppy, Na⁺ channels were almost equally distributed on somata and dendrites (76).

Interestingly, cytosolic GTP (100 µM) shifts the inactivation curve of g_Na to the right (277). This effect is presumably not brought about by GTP itself, because the nonhydrolyzable analog guanosine 5'-O-(3-thiotriphosphate) (GTPS) exerts the same effect (277). Under perforated patch, the steady-state inactivation curve for g_Na overlaps with the inactivation curve measured in the whole cell configuration with GTPS in the pipette. These results indicate that a constitutively activated G protein maintains low half-inactivation voltages for g_Na in ORNs. These results open the possibility that g_Na might be modulated by G protein-coupled receptors.

High-voltage-ativated (HVA) Ca²⁺ currents in ORNs have been described in various species (65, 69, 92, 260, 305, 342, 343). The amplitude of this current seems to be relatively small, and in the whole cell configuration of the patch-clamp technique, the maximum current decreases with a time constant of ~3 min (305). This phenomenon, often called "washout," has been observed in many preparations and is presumably because of the diffusion between cytosol and patch pipette (14, 278). The HVA Ca²⁺ currents activate between 40 and 30 mV and have maximum amplitudes at ~0 mV; in some reports, the I-V relationship appears to be shifted to higher voltages, which may depend on high access resistances (13) or an elevated extracellular Ca²⁺ concentration (92, 305, 343). The HVA channels are thus primarily activated during the generation of action potentials, and the resulting Ca²⁺ influx presumably contributes to the repolarization by activating K⁺ channels (see sect. IIIC). Using quantitative ratiometric Ca²⁺ imaging with fluo 3 and FuraRed, Schild et al. (309) have shown that the HVA Ca²⁺ channels in Xenopus ORNs are primarily situated on the soma and the proximal dendrite. The resolution of the method does, however, not exclude that a small portion of Ca²⁺ influx could occur in other cellular compartments of ORNs.

In some species, a low-voltage-activated (LVA) Ca²⁺ current may also be involved in action potential formation. In the newt, Kawai et al. (153) have reported an LVA current being half-activated at 44 mV. Although this current is absent in some species [e.g., Xenopus (305), Chilean toad (69), and catfish (246)], its description is consistent with Trotier's finding of a TTX-insensitive inward current that could be blocked with Co²⁺ in salamander ORNs (343). This current lowers the threshold for action potential firing. It speeds up action potential generation by decreasing the inactivation of the voltage-gated g_Na during receptor potentials. The LVA Ca²⁺ current might therefore be particularly useful in relatively large ORNs that have high capacitances and long membrane time constants.

Repolarization of the action potential is achieved by the activation of various K⁺ channels: first, and above all, by a transient, 4-aminopyridine (4-AP)-sensitive K⁺ current described in squid (217), salamander (92), zebrafish (65), catfish (246), salmon (260), clawed frog (305), and rat (221). This current activates and inactivates rapidly; it activates at potentials more positive than the activation threshold of Na⁺ channels (221, 305), which precludes the traditional role of A currents in spike frequency modulation (64, 120). It rather contributes strongly to the repolarization of action potentials. However, in cultured ORNs of rat, which have a large delayed-rectifier conductance, the fast 4-AP-sensitive current as well as a Ca²⁺-dependent K⁺ current were absent (342).

Second, Ca²⁺-activated K⁺ conductances [g_K(Ca)] measured in various species (65, 69, 92, 227, 260, 305, 343) seem to contribute to the repolarization and an increase of the cell's impedance, whereby in the mouse, 130-pS channels with voltage-dependent kinetics and 80-pS channels with voltage-insensitive kinetics have been observed (227), presumably corresponding to BK and SK K⁺ channels (190, 196). In agreement with these findings, a fraction of the outward current in Xenopus ORNs was blockable by apamin (305), which blocks Ca²⁺-dependent K⁺ channels of the SK type (55). Among the various Ca²⁺ sources that could potentially activate g_K(Ca) , Ca²⁺ influx through HVA Ca²⁺ channels appears to play the major role because blocking the HVA current (69, 305, 343) or reducing the Ca²⁺ flux through HVA channels (92, 260) reduced or abolished the Ca²⁺-dependent conductances. Calcium influx-induced Ca²⁺ gradients are very steep, and the diffusion length of free cytosolic Ca²⁺ is on the order of 1 µm (143, 259). The Ca²⁺- activated conductances, or at least a major fraction of them, can therefore be supposed to be colocalized with HVA Ca²⁺ channels. This is also supported by the fact that the voltage-gated Ca²⁺-activated K⁺ current in ORNs, when activated by voltage steps under voltage clamp, activates simultaneously with the voltage steps (69, 92, 246, 305, 343).

Because most of the HVA Ca²⁺ channels seem to be situated on the soma and the proximal dendrite (309), we can conclude that g_K(Ca) is colocalized with HVA channels at the soma and the proximal dendrite. This is consistent with single-channel measurements in excised patches from olfactory neuronal soma membrane (227); it does, however, not preclude additional Ca²⁺-dependent K⁺ channels being localized elsewhere. Bacigalupo and co-workers (248, 249) have reported evidence for a Ca²⁺-dependent K⁺ conductance in the transduction compartments of the toad that can be activated by odorants (see sect. VII). Differentiation between conductances involved in action potential generation and others involved in receptor potential generation might turn out to be inappropriate because the same g_Na , g_H, and g_K(Ca) may be involved in both.

In conclusion, an action potential, initiated by a receptor potential and voltage-gated Na⁺ channels, goes through the voltage range positive to 30 mV, where HVA channels on the soma and the proximal dendrite activate. The resulting Ca²⁺ influx activates Ca²⁺-dependent K⁺ channels on soma and dendrite which, in concert with fast g_K , then contribute to the repolarization of the action potential. Maue and Dionne (227) describe two different types of g_K(Ca) , which opens the possibility that a g_K(Ca) , in addition to participating in the repolarization, may play an additional role in tuning the refractory period. The third conductance, which may play a role in both repolarization and tuning of the resting and poststimulus impedance, is the slow delayed rectifier-like K⁺ conductance (69, 92, 217, 246, 260, 292, 305, 343).

This particular combination of K⁺ conductances taking part in the repolarization seems to be very similar in all studied species, although an exact quantitative comparison is presently impossible. The general idea, however, appears to be that only very few K⁺ channels are open at rest, resulting in an extremely high resting resistance and a correspondingly high sensitivity. The K⁺ conductances that are open at rest, presumably the inward rectifier and the delayed rectifier, would not be able to repolarize the membrane back to the resting potential because the inward rectifier is closed at depolarized voltages and the delayed rectifier has too small a conductance. Repolarization seems to be done by transiently switching on the fast K⁺ conductance and the fast Ca²⁺-dependent K⁺ conductance.

	IV. ODORANT RESPONSES

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The classical recordings from single ORNs were made using either platinum-black electrodes or sharp pipettes (4, 105, 106). Although the sharp electrode recordings suffered from the leak between glass and membrane resulting in short recording durations, extracellular recordings proved useful to determine the differential responsiveness of ORNs to many stimuli (142, 325). The patch-clamp technique with gigaohm seal resistances provided several new methods of recording odorant responses: 1) tight-seal whole cell patch-clamp recordings under voltage-clamp or current-clamp conditions, 2) cell-attached patch-clamp recordings, 3) loose patch recording with a suction electrode (212), 4) perforated patch recordings using pore-forming molecules such as nystatin (76, 217) or gramicidin (361), and 5) sensory cilia attached in loose-clamp (99), tight-seal, or excised configuration (159, 164, 165). The tight-seal whole cell configuration allows voltage clamping of the membrane potential so that activation and inactivation of voltage-gated and odorant-induced conductances could be measured. However, the diffusion of molecules from and into the pipette changes the cell's milieu, resulting in loss of important factors in the cytoplasm. Washout of messengers and modulators can prevent physiological responses, especially when recording Ca²⁺ currents or odorant responses. The cell-attached mode allows the measurement of mainly capacitative currents across the patch that are associated with action potentials (219). The perforated-patch configuration prevents the diffusion of large molecules between pipette and cytosol, but the access resistance to the cell is higher than in the whole cell mode. This causes a current-dependent voltage drop across the pipette tip, and, in most patch-clamp amplifiers, the voltage-clamp control circuit is slowed considerably.

In addition to electronic amplifiers, charge-coupled device (CCD) and laser scanning microscope imaging techniques (296, 306) became available and were used to measure odorant responses, whereby the increase of intracellular Ca²⁺ was monitored as an indirect measure of excitation.

The first manuscripts to present studies of ORNs using the patch-clamp technique were published in 1985 (7) and 1986 (343). Trotier (343) described voltage-gated currents, single channels including the inward rectifier and currents induced by addition of odorants (7 µM isoamylacetate and 10 µM butanol).¹ Although the description of the odorant-induced current was short, it contained some important points that were confirmed later: the reversal potential of the odorant-induced current was ~0 mV, the I-V relationship was almost linear, and there was a considerable delay in the response (in this study ~2 s). Trotier (343) cautiously suggested as a working hypothesis that the odor-activated channels were nonselective cation channels modulated by a diffusible cytosolic metabolite. The equilibrium potential for both Cl and cations was ~0 mV in his study.

The history of the discovery of odor-activated currents in vertebrates took three main routes: 1) independently the laboratories of Firestein, Kurahashi, and Gold followed Trotier's working hypothesis and consequently carried out detailed analyses of the odor-activated currents. Kleene and Gesteland (165) discovered that what was believed to be an odor-gated "cation" current was actually a mixture of a Ca²⁺-permeable cation current and a Ca²⁺-activated Cl current. These experiments led to the present view of the cAMP-mediated second messenger cascade, which is described in section V. 2) Evidence regarding odorant-induced Ca²⁺ and cation currents that were different from the cAMP-gated current did not fit in the model of a cAMP-mediated second messenger cascade. Work from the laboratories of Restrepo, Ache, and Schild as well as from studies in insects (327, 328, 363, 373) indicated that there might be at least one more second messenger pathway mediated by inositol phosphates, Ca²⁺, or guanosine 3',5'-cyclic monophosphate (cGMP). This work is described in sections VI and VII. 3) Several studies reporting odorant-mediated currents did not primarily aim at analyzing elements of the cAMP or inositol trisphosphate (InsP₃) pathway. They rather pointed at the kinetic and biophysical characterization of odorant responses, which are reviewed in the rest of this section.

After Trotier's 1986 paper (343), one study² reported the block of the odorant-induced current by amiloride in frog ORNs (98), another reported an odorant-stimulated depolarization that reversed at 2.7 mV (8), and studies from two laboratories presented a detailed characterization of odor-activated currents in ORNs (91, 175, 179, 182). Kurahashi (175) characterized an odorant-gated cation conductance in newt ORNs using n-amyl acetate (10 mM) as odorant stimulus (Fig. 2). This current had the following properties: 1) it reversed at ~0 mV, the reversal potential shifting by ~57 mV upon reducing Na⁺ concentration by a factor of 10; 2) the current was maximum when the odorant was applied at the apical dendrite; 3) the I-V curve was nonlinear, showing a block of inward current at potentials more negative than 30 mV; 4) its permeabilities for alkali ions were P_Li/P_Na/P_K/P_Rb/P_Cs = 1.25:1:0.98:0.84:0.8; 5) removal of Ca²⁺ increased the odorant-gated current and prolonged its duration, but did not cause a change in reversal potential; and 6) removal of Cl did not change the reversal potential. The size of the odor-induced current was maximal when the odor plume was directed to the apical side of the cell. The pipette solutions in these studies contained 5 mM ethylene glycolbis(-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA) so that [Ca²⁺]_i was heavily buffered. In addition, in a separate manuscript, Kurahashi and Shibuya (183) suggested that the inactivation of the odorant-induced current was mediated by Ca²⁺ influx through the odor-regulated conductance.

View larger version (12K): [in this window] [in a new window]   FIG. 2.   A: odorant-induced currents recorded at various holding voltages (Vh; indicated to left of each current trace) from a newt (Cynops pyrrhogaster) olfactory receptor neuron. Current traces were arbitrarily shifted. Bottom trace indicates timing of n-amyl acetate (10 mM) application from a glass pipette positioned <20 µm away from apical dendrite. B: voltage dependence of odorant-induced current; replotted from A and additional records from same cell. [Modified from Kurahashi (175).]

Firestein and Werblin (91) stimulated terrestrial phase salamander ORNs using a mixture of acetophenone, n-amyl acetate, cineole, phenylethylamine, and triethylamine (each at a concentration between 0.1 and 1 mM). The reversal potential for cations (u_cat) and u_Cl were both at ~0 mV in this study. These authors confirmed and extended Trotier's finding of a purported cation current with respect to the following points: 1) again the I-V curve of the odor-activated currents was linear with a reversal potential of ~0 mV, 2) the delay of the odorant response was assessed more accurately (140-570 ms), 3) a dose-response curve extending from ~10⁵ to 10⁴ M for a mixture of stimuli was given, and 4) the odor response was shown to be cooperative with a Hill coefficient of 2.7. Any of the steps between binding of the odorant and flow of ions through odorant-activated channels could thus be cooperative, leading to a sigmoidal input-output (i.e., concentration-current) relationship (see also Ref. 87). In a later study, Firestein et al. (90) confirmed that the odorant sensitivity is restricted to the cilia and possibly the distal dendrite. They also found that the latency of the response was relatively independent of the stimulus concentration (10⁵ and 10⁶ M).

The spatial distribution of odorant sensitivity and of the odorant-induced current was carefully examined in a series of experiments in salamander ORNs by Lowe and Gold (212). Using an odorant mixture containing 2-hexylpyridine, isoamyl acetate, acetophenone, and cineole, these investigators determined that the odorant response increased approximately linearly as a function of the fraction of the ciliary bundle that was exposed to the odorant. No further increase in responsivity was found upon stimulation of the entire ciliary bundle and a fraction of the dendrite. This indicated that olfactory receptor sites are uniformly distributed along the length of the cilium and are excluded from the dendrite. In addition, the odorant-induced current was found to be localized to the cilia, and the resting K⁺ conductance was found to be much lower in the ciliary membrane than in the dendritic membrane. The pipette contained a pseudointracellular solution with 0.1 mM EGTA, pH 7.2, providing little buffering of intracellular Ca²⁺.

Obviously, in a sensory system that can possibly detect single molecules (215, 238), concentrations in the range between 10⁵ and 10⁶ M are high. However, the physical chemistry of lipophilic odorants in water is by no means trivial, and it has to be kept in mind that in the experiments described, the odorants were applied in the absence of mucus or odor-binding proteins, i.e., the perireceptor microenvironment (107, 139, 224, 235, 271) was far from physiological. The experiments by Frings and Lindemann (99), who recorded from single cilia of ORNs in an at least partly intact epithelium, are interesting in this respect because they recorded an increase in action potential rate upon application of cineole (dissolved in ethanol), whereby 9, 5, and 3 of 22 responding cells responded to odorant concentrations of 1, 10, and 100 pM, respectively (99).

Optical recordings of odorant-induced changes in [Ca²⁺]_i with CCD cameras or laser scanning microscopes have the advantage of spatial resolution, and they are minimally invasive, although the endogenous Ca²⁺ buffering capacity of the cells (158) is increased by adding the reporter dye (118). Optical recording of [Ca²⁺]_i was first used by Restrepo and co-workers (288, 290) to monitor odorant responsiveness of catfish ORNs. These and subsequent studies determined that odorants elicit increases in intracellular Ca²⁺ (16, 145, 192, 257, 288, 291, 292, 302, 337) and that removal of extracellular Ca²⁺ abolished the odorant-induced increases in [Ca²⁺]_i in ORNs, suggesting that the increase was due to influx of Ca²⁺ through the plasma membrane (16, 288, 291, 292, 302, 337). However, some olfactory neurons from bullfrog (Rana catesbeiana) were found to respond even in the absence of extracellular Ca²⁺ (302). Imaging experiments determined that shortly after stimulation, and in some ORNs even after prolonged stimulation, the increases in [Ca²⁺]_i took place at the apical end of the cell, suggesting that the source of Ca²⁺ was at the apical compartments of the cell (16, 257, 291, 302, 337). Interestingly, recent measurements with confocal microscopy in salamander (A. tigrinum) ORNs has demonstrated odor-induced changes in [Ca²⁺]_i in individual olfactory cilia (192). The increase in [Ca²⁺]_i in the cilia is rapid and transient and contrasts with a much slower and prolonged increase in [Ca²⁺]_i in the cell body.

Imaging of [Ca²⁺]_i has also been used to study channels and transporters involved in [Ca²⁺]_i regulation in ORNs and the regulation of these channels and transporters by second messengers. Thus it has been possible to localize the HVA Ca²⁺ current in ORNs (309), the Na⁺/Ca²⁺ antiport (136), and InsP₃-activated, Ca²⁺-permeable channels of the plasma membrane of ORNs (145, 311).

Studies of odorant responsiveness utilizing [Ca²⁺]_i imaging techniques have provided information on the odor specificity of neighboring ORNs. Takebayashi and co-workers (122, 303) imaged a partly dissociated preparation of mouse ORNs in which the cells retained their original spatial relationships. With the use of three different odorants in one study (122) and a series of aliphatic odorants in another study (303), the responsiveness of ORNs was estimated from the odor-induced increase in [Ca²⁺]_i . These authors were able to demonstrate that cells with a similar odor responsivity were located next to cells with a different odor responsivity. The distribution patterns were, however, not consistent with a completely random distribution.

Finally, optical recording with voltage-sensitive dyes (VSDs) has been used to determine neuronal activity in the olfactory system (60, 149). This technique has the advantage that the average activity in a volume of tissue can be determined upon stimulation with many odorants (150). Staining of the olfactory epithelium with VSDs could be used to study signal transduction in ORNs, which would be particularly useful in resolving questions about signaling between neighboring ORNs. However, the use of VSDs in the olfactory epithelium at high magnification has not been reported in refereed publications. Voltage-sensitive dyes have been used to record distributed activity within the olfactory epithelium and the olfactory bulb at low magnification (cf. Refs. 59, 61, 62, 155, 156, 356). These experiments have provided an important contribution of our understanding of patterns of responsivity to odors in the olfactory epithelium and form a basis for understanding olfactory quality coding. Studies of distributed patterns of responsivity with VSDs are not reviewed in detail here, and the reader is referred to recent reviews (60, 149).

4. Are ORNs modulated by the autonomic nervous system?

A further interesting point within the context of diversity in transduction mechanisms is the possible influence of the autonomic nervous system on olfactory transduction pathways. Some neurotransmitter receptor antagonists such as propranolol, a -adrenergic antagonist and a serotonin (5-HT₁) agonist, and the muscarinic antagonist atropine can block odorant responses in isolated ORNs via an unknown mechanism (

It has proven difficult to culture functionally responsive ORNs. The only report of odorant-induced changes in electrical activity is from cultured ORNs of rat bearing multiple long processes that showed odorant-induced depolarizations in conjunction with an increase in conductance. Interestingly, these cells also displayed a second type of odorant-activated current that resembled very much synaptic currents (275). Others have reported odorant-induced changes in second messenger formation and/or odorant-induced changes in [Ca²⁺]_i in cultured olfactory neuroblasts. Long-term cell cultures of neuroblasts from human fetal olfactory epithelium and olfactory neuronal cell cultures from rat embryo responded to stimulation by odorants with biochemically measured increases in cAMP or InsP₃ formation (297, 347), and human olfactory neuroblastoma cells (110) and neuronal cells cultured from adult human olfactory epithelium (354) have been shown to respond to odors with increases in intracellular Ca²⁺.

Cell cultures of olfactory neuroblasts have also been used in the study of olfactory mechanisms in invertebrates. Cultured ORNs of lobster responded to odorants even before they had developed processes (83), suggesting that these cells insert all transduction components in the soma membrane during their development. Cultured ORNs of the silkmoth Manduca sexta responded to a pheromone (synthetic bombycal) with a sequence of inward currents. The early component of the response to the pheromone was a Ca²⁺ current that has been suggested to be regulated by InsP₃ (328). This current was inhibited by Ni²⁺ and did not appear in the absence of Ca²⁺ or Ba²⁺ in the bath. It was followed by a slower inwardly rectifying current component with a reversal potential near 0 mV. The nonspecific current had the characteristics of a Ca²⁺-dependent nonspecific cation current described in insect ORNs (327). Finally, a third sustained component reversed near 0 mV and was proposed to be a protein kinase C-dependent cation current (327, 328).

Responses of single vomeronasal ORNs to odorants or pheromones have as yet not been reported. However, Trotier et al. (345) and Liman and Corey (200) have measured voltage-gated currents in microvillous vomeronasal neurons using the patch-clamp technique. Interestingly, Liman and Corey (200) report a lack of a response upon dialysis into the cytosol of vomeronasal receptor neurons with cAMP (200), whereas Taniguchi et al. (335, 336) report current responses upon dialysis of vomeronasal receptor cells of the turtle with cAMP, cGMP, or InsP₃ . Furthermore, Okamoto et al. (265) have shown a forskolin-induced increase of adenylyl cyclase in turtle vomeronasal receptor cells, but odorants that increase the activity of this enzyme in ORNs of other species had no effect. Hence, there appear to be some dissimilarities between the receptor cells in the main olfactory epithelium and those in the vomeronasal organ (197). This has been confirmed by Dulac and Axel's (77) finding that a family of putative receptor genes of the rat vomeronasal organ is unrelated to the receptors expressed in the rat main olfactory epithelium, and by studies from Berghard et al. (24) indicating that, unlike ORNs, vomeronasal neurons do not express mRNA for the G protein G_olf , for subunit 1 of the olfactory cAMP-gated channel (g_cn1) and for adenylyl cyclase III.³ Vomeronasal neurons do express mRNA encoding for the second subunit of the olfactory cAMP-gated channel (g_cn2) (24). In the absence of g_cn1 , g_cn2 does not open in response to increases in the concentration of cyclic nucleotides, but does open upon exposure to nitric oxide (NO) (44). A channel with characteristics consistent with those of g_cn2 opens in response to NO in patches excised from rat vomeronasal neurons (44) (see Ref. 197 for review).

	V. ADENOSINE 3',5'-CYCLIC MONOPHOSPHATE-MEDIATED SIGNALING IN OLFACTORY CELLS

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It is now generally believed that most odorants bind to seven transmembrane (TM) receptors that couple to G proteins resulting in second messenger formation (40, 46, 51, 52, 181, 189, 232, 262, 268, 274, 281, 283, 322, 324) (Fig. 3). A G protein called G_olf , which was first cloned on the basis of enriched expression in the olfactory system, is localized to the olfactory cilia (132, 234). Virtually identical G proteins have also been found in the striatum (119) and tissues outside the CNS such as the pancreas (365). Although no functional data are available directly implicating G_olf in olfactory transduction, it is most likely that G_olf couples certain olfactory receptors to cAMP formation by mediating stimulation of type III adenylyl cyclase (17). Accordingly, biochemical studies of adenylyl cyclase activity and measurements of cAMP formation in isolated olfactory cilia implicated a G protein-mediated increase in cAMP in olfactory transduction (267, 326).⁴ However, some potent odor stimuli failed to activate adenylyl cyclase (see sect. VI) (49, 326).

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