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Department of Neurobiology, School of Medicine, Yale University, New Haven, Connecticut
Submitted 3 January 2006; accepted in final form 12 July 2006
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ABSTRACT |
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INTRODUCTION |
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).
The structure and function of the network formed by mitral secondary dendrites and granule cell dendrites are relatively clear (Andres 1965; Chen et al. 2000; Hirata 1964; Isaacson and Strowbridge 1998; Jahr and Nicoll 1982; Price and Powell 1970; Rall et al. 1966; Schoppa et al. 1998; Yokoi et al. 1995). In contrast, the glomerular local network is little understood. Olfactory glomeruli are probably the most clearly demarcated module for multineuronal information processing in the brain (Chen and Shepherd 2006). Most receptor neurons expressing a given olfactory receptor send their axons to converge onto two glomeruli (Mombaerts et al. 1996; Ressler et al. 1994; Vassar et al. 1994). Functional studies show that a given odor elicits a specific map of glomerular activation reflecting its binding to several different receptors (Johnson and Leon 2000; Mori et al. 1999; Rubin and Katz 1999; Stewart et al. 1979; Xu et al. 2003). A glomerulus thus forms a structural and functional unit in which the olfactory signals are initially processed as the basis for olfactory perception.
Morphological studies have provided a basic picture of the synaptic organization of the glomeruli (Pinching and Powell 1971a,b). Each glomerulus is surrounded by a shell of small neurons and glia. These neurons have been termed juxtaglomerular (JG) cells, consisting of three types: periglomerular (PG) cells, short axon (SA) cells, and external tufted (ET) cells (Pinching and Powell 1971c; Shipley and Ennis 1996). Olfactory axons terminate within the glomeruli and make synapses with the dendritic tufts of mitral/tufted cells and some intrinsic PG cells. Mitral/tufted and PG cells also make numerous dendrodendritic connections (reviewed in Shepherd et al. 2004).
Understanding the intrinsic membrane properties of JG cells is clearly a necessary step toward understanding the neural basis of olfactory signal processing within the glomeruli. Of particular interest are the cells in the glomerular and external plexiform layer region that show intensive spike bursts to olfactory nerve (ON) input (Shepherd 1963). Recent studies have provided intracellular recordings of a long-lasting depolarizing potential that underlies the bursts of spikes (McQuiston and Katz 2001; Wellis and Scott 1990). Both persistent sodium current (Hayar et al. 2004b) and low-threshold calcium current (McQuiston and Katz 2001) have been proposed to contribute to the long-lasting depolarizing potential. The persistent sodium current–driven bursts have been shown to coordinate the glomerular activity (Hayar et al. 2004a). The long duration of calcium spikes was critical for activating inhibitory responses in the glomerulus (Murphy et al. 2005). In view of this new evidence, a deeper understanding is needed of the critical role of the dendritic properties of PG cells for processing of odor input at the glomerular level. We thus used patch-clamp recordings and calcium imaging techniques in rat olfactory bulb slices to investigate the calcium currents involved in generating the long-lasting plateau potential and analyzed their functional roles in the glomerular network. Parts of this work were previously published in abstract form (Zhou et al. 2002, 2003).
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METHODS |
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Slices of olfactory bulbs from Sprague–Dawley (SD) rats (16–23 days) of either sex were obtained as previously described (Chen and Shepherd 1997). Briefly, rats were decapitated after urethane (1.5 mg/g) anesthesia. After removal of the skull, the head was quickly immersed in 4°C normal artificial cerebral spinal fluid (ACSF) oxygenated with 95% O2-5% CO2. Two bulbs were carefully dissected out and then sliced into 300 to 400 µm slices using a vibrating slicer (Campden Instruments, Loughborough, UK). Slices were incubated in 37°C ACSF for 30–60 min and then maintained at room temperature. The normal ACSF contained (in mM, pH 7.4): NaCl 124, KCl 3, MgSO4 1.3, CaCl2 2, NaHCO3 26, NaH2PO4 1.25, and glucose 10.
Experiments were performed mostly at room temperature (about 25°C) or otherwise specified at about 35°C with a TC-344B temperature controller (Warner Instruments, New Haven, CT). Under infrared differential interference contrast (IR-DIC) video microscopy, whole cell patch-clamp recordings were performed using an Axoclamp-2A amplifier (Axon Instruments, Union City, CA). Data were acquired with Clampex 8.0 through an AD/DA converter Digidata 1320A (Axon Instruments). Micropipettes were fabricated with a P97 Puller (Sutter Instruments, San Rafael, CA). The resistance ranged from 2 to 6 M
. The micropipette intracellular solution contained (in mM): potassium gluconate 130, Mg-ATP 4, Na2-GTP 0.3, HEPES 20, and Oregon-green 488 BAPTA-1 (Molecular Probes, Eugene, OR) 0.2. Olfactory nerve stimulation was delivered by a SECO SC-100 stimulator through a bipolar electrode (MCE-100; Rhodes Medical Instruments, Woodland Hills, CA).
Ionic channel blocker drugs were applied by bath perfusion or local puff. A glass pipette (about 1 µm) filled with a high concentration of 50 mM Ni2+ and 500 µM tetrodotoxin (TTX) (and food dye) was put near a recorded JG cell soma or its dendrites in the glomerulus. Weak pressure was applied by a Picospritzer II valve (General Valve, Fairfield, NJ) to puff the TTX and Ni2+ on the soma or dendrites.
Two-photon calcium imaging
Calcium-sensitive dye Oregon-green was loaded into the cells by whole cell recording pipettes. For calcium imaging we used a custom-built two-photon microscopy setup. For excitation the Tsunami Ti:sapphire lasers (Spectra Physics, Mountain View, CA) provided an approximately 100 fs, 810 to 830 nm pulse at 80 MHz, pumped by a 10 W Millennia Xs laser (Spectra Physics). Image scanning and acquisition were controlled by an Olympus Fluoview 300 confocal system mounted on an upright microscope (BX50WI, Olympus) equipped with a water immersion objective (x40, NA 0.8). The calcium signal was usually recorded in line-scan mode. The sampling interval was 2.02 ms. Images of cell morphology were usually taken in a stack of images in the z-axis. Recordings of fluorescence and electrophysiological signals were synchronized by an A-65 Timer (Winston Electronics, St. Louis, MO). Fluorescence signals were outputted from Fluoview 300 in EXCEL format files and represented as 
F/F0 = (F – F0)/F0, where F0 is the average of baseline fluorescence before stimulation. Data were reported as means ± SE, and the Student's t-test was applied to compare group data.
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RESULTS |
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Olfactory glomeruli in the rat appear surrounded by a shell of small neurons (Pinching and Powell 1971c). Under IR-DIC, these so-called juxtaglomerular (JG) cells, which include periglomerular (PG) cells, external tuft (ET) neurons, and short axon (SA) neurons, could be visually identified. PG cells usually have round and smaller cell bodies compared with ET and SA neurons. ET cells have relatively larger somata and are often found to have one thick trunk entering a glomerulus. The final identification of these cell types was made after cell morphology reconstruction with two-photon images according to previous criteria (Shepherd et al. 2004; Shipley and Ennis 1996). ET cells receive monosynaptic inputs from the ON; PG cells receive poly- or no synaptic inputs from the ON. ET cell axons were seen directed toward the EPL and to other areas, whereas PG cell axons remain in the glomerular area; some PG cells lack axons.
Of 177 recorded JG neurons with images reconstructed by two-photon microscopy, there were 86 PG, 87 ET, and four SA neurons; 24 PG (27.9%), 65 ET (74.7%), and no SA neurons were found to have plateau potentials (PPs). As shown in Fig. 1, the plateau potentials were evoked by ON stimulation (Aa) and DC injection on soma (Ab). The plateaus always outlasted the brief stimuli, with durations of 106.0–649.3 ms, averaging 234.8 ± 12.8 ms (n = 89). The plateau amplitudes were 41.1 ± 1.2 mV, ranging from 25.1 to 57.5 mV. The plateau potential usually, but not always, followed spike discharges. In most cells (61 of 89) there was an afterhyperpolarizing potential (AHP) after the plateau potential (6.3 ± 0.4 mV). Figure 1Ac is a typical voltage–current (V–I) response of an ET cell showing plateau potentials. Hyperpolarizing current could evoke rebound depolarization. The rebound depolarization produced a PP (single arrowhead) when it reached the threshold of about –40 mV, comparable to that of the calcium spike previously reported (Hayer et al. 2004b). A sag (horizontal arrow) in the sustained response to hyperpolarizing current suggested a hyperpolarization-activated nonselective cation current (H-current, Ih).
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; see also Bufler et al. 1992). As shown in Fig. 2, A and Ca, ET cells had a clearly imaged axon extending to the EPL, but no secondary dendrites. Ionic basis of plateau potential
Plateau potentials were evoked by both ON stimulation and soma current injection. The PG cells with PPs gave evidence of both mono- and polysynaptic inputs from the ON. ET cells received monosynaptic ON inputs. Soma stimulation-induced PPs displayed an all-or-nothing property (Fig. 3A). The production of PPs was all-or-nothing from certain membrane potentials, although the plateau duration was variably affected by other factors such as membrane potential and recording temperature. As reported previously the plateau duration was shorter at body temperature than at room temperature (McQuiston and Katz 2001). Figure 3B shows that depolarized membrane potentials partially inactivated the PPs. In any case, the duration of the PPs always outlasted the brief stimulation (usually 20 ms for current injection at soma). This, together with the all-or-nothing property, suggested a regenerative mechanism underlying the PPs.
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) that low-threshold calcium channels are responsible for the generation of the PP, which they therefore designated as a low-threshold spike (LTS). Our results showed that high-threshold Ca2+ channels also contribute to maintaining the PPs. In another study (Murphy et al. 2005), a novel type of low-threshold calcium channel was reported to be responsible for calcium spike generation in JG cells; however, they are not blocked by nickel but by cadmium.
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Dendritic calcium conductance and plateau potentials
The differential location of Ca2+ conductances in soma and dendrites is critical to their functional roles. This is especially relevant for JG cells because of the involvement of their dendrites in dendrodendritic synaptic interactions within the glomerulus. Experimental analysis of this problem is difficult because JG cells are small neurons and their dendrites are too small for patch recordings. We therefore applied the two-photon microscopic imaging technique to study Ca2+ signals in the JG cell dendrites.
Single Na+ spikes did not evoke obvious changes of Ca2+ intracellular concentration ([Ca2+]i) in JG cell dendrites (Fig. 5Ab). Dendritic [Ca2+]i increases were mostly correlated with evoked (Fig. 5Aa) and spontaneous PPs recorded in soma (Fig. 5B) (n = 15). The onset of [Ca2+]i increases (see Fig. 5) indicated that they were not induced by Na+ action potentials preceding each plateau potential. This is more obviously shown in Fig. 5Bb, where a spontaneous PP appeared with 50 µM Ni2+ in bath. The amplitude was decreased but the PP was not abolished, and the onset phase of the [Ca2+]i increase was delayed after the first three Na+ action potentials, which enabled us to see the close correlation between PP (not Na+ spikes) and dendritic [Ca2+]i increase. As shown in Fig. 5Aa, the time course of the [Ca2+]i increase (blue trace) matched well with the integral of the membrane potential (green trace), indicating a close correlation between the plateau potential and dendritic [Ca2+]i increase. This is similar to an in vivo study in cortex showing that dendritic [Ca2+]i increase always correlated with dendritic complex spikes (Helmchen et al. 1999). However, as shown in Fig. 5C, in this cell, six of 26 trials of robust [Ca2+]i increase were observed in the dendrites without a PP in the soma. Comparing the time course of the [Ca2+]i increase in the dendrites and membrane potential in the soma, it suggests that the [Ca2+]i increase in the dendrites was not ascribed to backpropagation of the somatic Na+ action potential or the PP. The most likely explanation is that soma stimulation triggered the Ca2+ channels in the dendrites to generate a PP, which then propagated to the soma during which it was variably inhibited en route. This would be similar to in vivo experiments (Helmchen et al. 1999) in cortical cells in which inhibitory inputs prevented propagation of dendritic complex spikes to the cell soma, which facilitated the recording of [Ca2+]i increases in dendrites without observing the usual complex spikes in the soma.
The dendritic contribution of Ca2+ channels to the PPs was further studied by local puffs of Ca2+ and Na+ channel blockers. When Ni2+ and TTX were puffed on the soma, Na+ action potentials were blocked but plateau potentials remained unaffected (Fig. 6A, red trace) (n = 6). The blockade of Na+ action potentials showed that the blockers were effectively applied and restricted to the soma. Somatic application of Ni2+ did not block the plateau potential. Ni2+ and TTX puffed on the dendrites blocked the plateau potential but not the Na+ spikes (Fig. 6A, green trace). As shown in Fig. 6B, perfusion of 250 µM Ni2+ completely blocked the PP but not the Na+ action potentials. These results further supported the generation of PPs by dendritic Ca2+ conductance.
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We finally explored how the plateau potential modulates neuronal input–output transformation from the glomerular level. We first found that PPs modulated the incoming excitatory postsynaptic potentials (EPSPs) at different time points. If the synaptic input appeared during a PP, its EPSP decreased dramatically to 25–1% of its control (Fig. 7A, n = 7). A plateau potential thus could effectively modulate the amplitudes of EPSPs, particularly near the crest of the potential. The PPs had average amplitudes of 41.4 ± 1.2 mV and reached average membrane potential levels of –23.5 ± 1.7 mV. The EPSP amplitude modulation (AM) thus could reflect shunting of the EPSPs and reduced driving potentials. These values were recorded in the soma; presumably the PP amplitudes were higher and the membrane potentials were more depolarized at the site of generation in the dendrites as a result of the electrotonic decay between the dendrites and soma.
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With regard to generation of the plateau potential, the example in Fig. 7 shows that, compared with the control recording (Fig. 7Ba, gray trace), the second and third EPSPs were 39 and 74% larger than their counterparts (see also Fig. 7Bb). The second and third EPSPs were even larger (4 and 7%, respectively) than the first one. This boosting effect must be largely attributed to the dendritic Ca2+ conductance because blocking Na+ channels had little effect on producing PPs (Fig. 4B above). We conclude that at least some JG cells tune repetitive high-frequency inputs into low-frequency outputs in the form of PPs.
Plateau potentials fired in a rhythmic way (Fig. 8) at low frequencies. Figure 8A shows the nearly 2-Hz PPs induced by a hyperpolarizing current. We studied the frequency of the rhythmic PPs by varying the frequency of injected current pulses in the soma (n = 16). As shown in Fig. 8, when a PP was elicited there was a nonresponsive period for the next PP generation (287.6 ± 78 ms, n = 16), which limited the PPs to firing at a low frequency of 2.6 ± 0.8 Hz (n = 16). This property enables the JG cells with PPs to act as a low-pass filter to generate low-frequency outputs. The JG cells thus uncouple the incoming firing rate of inputs from various sources (Hayer et al. 2005) and set the output rate by the intrinsic rhythmic PPs or bursts. JG cells with PPs may thus serve as pacemakers in an oscillatory glomerular network.
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DISCUSSION |
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; Reyes 2001). The present study extends previous work and indicates that PPs in juxtaglomerular (JG) cells of the olfactory bulb constitute an attractive model for analyzing not only the properties of PPs but also their roles in information processing.
We have shown that a subtype of JG cell reliably generates PPs that can last as long as 650 ms (average 234.8 ms) with an average amplitude of 41 mV. As with PPs in other neurons, the PP in JG cells has been reported to be a Ca2+ potential because it could be blocked by the low-threshold Ca2+ channel blockers Ni2+ and alpha-methyl-alpha-phenylsuccinimide (McQuiston and Katz 2001). The present study has confirmed their result with Ni2+ blockade. We also found that, although Cd2+—a high-threshold Ca2+ channel blocker—could not block the generation of the PPs it could reduce the potential in amplitude and duration (Fig. 4). A high-threshold Ca2+ conductance could therefore also play a role in maintaining the plateau. In a recent study (Murphy et al. 2005), PG cell calcium spikes were also reported to be carried by calcium channels activated at low voltage; however, this novel type of low-threshold calcium channels was not blocked by Ni2+, but by Cd2+ and dihydropyridine. The discrepancy may be explained by the difficulties in clearly classifying the calcium channels, the selectivity of the channel blockers, and the heterogeneity of the cells in the periglomerular area (Kosaka et al. 1998).
Our Ca2+ imaging studies showed that a large dendritic [Ca2+]i increase was always produced together with the PPs, further confirming the Ca2+ mechanism underlying the plateau potential. JG cell PPs were Ca2+ potentials. Some JG cells showed no delayed-rectifier K+ current (Puopolo and Belluzzi 1998), which could further prolong the Ca2+ potential into a long-lasting PP. Most cells showing PPs have an afterhyperpolarizing potential (AHP), which could be activated by intracellular Ca2+ accumulation (Egan et al. 1993; Sah 1996) and be involved in terminating the PPs.
Our Ca2+ imaging and pharmacological studies showed that the JG cell PPs were generated by Ca2+ channels in the dendrites. The dendritic [Ca2+]i increase always correlated with the dendritic but not somatic PPs (Fig. 5). The PPs were blocked by puffs of Ni2+ on the dendrites but not soma (Fig. 6). Our evidence thus indicates that JG cell PPs are dendritic Ca2+ potentials, generated in the dendrites by dendritic Ca2+ conductance and then propagated to the soma during which they undergo variable inhibition en route.
Dendrites are well known as sites for synaptic inputs and synaptic processing. It is also now well established that they are active rather than passive neuronal structures with a diversity of ion channels (Häusser et al. 2000; Migliore and Shepherd 2002; Reyes 2001). Different channels play differential roles in the integration of synaptic inputs (Reyes 2001). The JG cells appear to be attractive models for in depth analyses of these roles. This is partly on the basis of their intimate relation to the olfactory glomeruli, perhaps the clearest example of an anatomical, molecular, and functional unit in the nervous system. It is also on the basis that the JG cell dendrites both receive a well-defined input, from the olfactory receptor cell axons, and engage in dendrodendritic interactions with mitral and tufted cell dendrites within the glomerulus. Through the JG cell dendritic outputs, the dendritic Ca2+ PPs thus can play significant roles in the networks that control glomerular input–output functions.
Among the properties that are likely to be important in these roles, we have shown (see Fig. 7B) that the dendritic calcium conductance first amplifies the incoming EPSPs to produce an all-or-nothing PP. Once produced, the PP then modulates subsequent EPSPs (Fig. 7, A and B), by shunting the EPSP conductance and by moving closer to the reversal potential of 
-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) and NMDA receptors to reduce the EPSP amplitudes. The JG cells can thus uncouple most of the specific temporal information of presynaptic inputs, arriving from disparate parts of the olfactory epithelium, from its postsynaptic output, thereby acting as a filter to convert the high-frequency olfactory axonal inputs into low-frequency olfactory glomerular outputs in the form of stereotyped PPs.
The firing frequencies of JG cell PPs appear to be mainly set by their intrinsic membrane properties. In addition to a Ca2+ conductance, a hyperpolarization-activated cationic current (H-current, Ih) appears to be present (see Figs. 1 and 8). Immunocytochemistry has shown strong hyperpolarization-activated cyclic nucleotide-sensitive nonselective cation (HCN) channel expression in neurons of the glomerular region (Holderith et al. 2003; Santoro et al. 2000). Electrophysiological studies have also shown that both PG and ET cells exhibit an Ih current (Cadetti and Belluzzi 2001). As a pacemaker current (Luthi and McCormick 1998), Ih may work with Ca2+ and Ca2+-activated K+ currents to drive rhythmic PPs. In this mechanism, the Ca2+ PP activates a Ca2+-activated K+ current and inhibitory GABAergic current (Murphy et al. 2005; Smith and Jahr 2002), which hyperpolarizes the membrane. Membrane hyperpolarization alone and/or activated Ih depolarizes the membrane to activate a low-threshold Ca2+ current to produce the next PP. These mechanisms cycle to produce rhythmic PPs or bursts (Fig. 8A). Persistent sodium current was reported to be critical for burst firings (Hayar et al. 2004b). Here we have shown that calcium current could drive the rhythmic burst (Fig. 8A).
The rhythmic property of the PPs was investigated by stimulating the cells with different frequencies. The PP had a nonresponsive period during which no further PPs could be induced (Fig. 8B). These nonresponsive periods determined the firing frequency of PPs at around 2.6 Hz. This is not in conflict with a previous report (Hayar et al. 2004b) that ET cells could be entrained by ON inputs 
10 Hz. As shown in Fig. 8B, sodium spikes could still be produced in the nonresponsive period to PP production. Earlier in vivo extracellular recordings in the rabbit showed that ON volleys produced in PG cells a facilitation period within 40 ms and a later suppression period lasting 200 ms (Shepherd 1971). Later work in the cat showed a similar phenomenon referred to as "glomerular transmission attenuation" (Freeman 1974a,b). A study using optical imaging showed a paired-pulse depression (PPD) phenomenon acting on the glomerular signal within a 500 ms period (Senseman 1996). The mechanism underlying the PPD phenomenon is still not clear. Earlier work suggested a synaptic inhibitory mechanism (Shepherd 1971). Both presynaptic (Ennis et al. 2001; Keller et al. 1998) and postsynaptic inhibition could participate in glomerular PPD; however, PPD could still be produced without presynaptic signal depression of the ON input (Senseman 1996). The nonresponsive period of the plateau potential reported in this study suggests that the glomerular PPD could be occurring at the cellular level.
In conclusion, some juxtaglomerular cells of olfactory bulb produce long-lasting plateau potentials that are dependent on low-threshold calcium channels. The calcium channels are mainly located in dendrites, where they integrate the inputs and produce low-frequency rhythmic output near the theta range, which is close to that of the olfactory activities linked to the breathing rhythm. We suggest that these juxtaglomerular cells may serve as pacemaker cells to synchronize the low-frequency glomerular oscillation.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: Z. Zhou, Department of Neurobiology, School of Medicine, Yale University, 333 Cedar Street, New Haven, CT 06510
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REFERENCES |
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Andres K. Der Feinbau des bulbus olfactorius der ratte unter besonderer berÜksichtgung der synaptischen verbindungen. Z Zellforsch Mikrosk Anat 65: 530–561, 1965.[CrossRef][Web of Science][Medline]
Bufler J, Zufall F, Franke C, and Hatt H. Patch-clamp recordings of spiking and nonspiking interneurons from rabbit olfactory bulb slices: membrane properties and ionic currents. J Comp Physiol A Sens Neural Behav Physiol 170: 145–152, 1992.[Medline]
Cadetti L and Belluzzi O. Hyperpolarisation-activated current in glomerular cells of the rat olfactory bulb. Neuroreport 12: 3117–3120, 2001.[CrossRef][Web of Science][Medline]
Chen WR and Shepherd GM. Membrane and synaptic properties of mitral cells in slices of rat olfactory bulb. Brain Res 745: 189–196, 1997.[CrossRef][Web of Science][Medline]
Chen WR and Shepherd GM. The olfactory glomerulus: a cortical module with specific functions. J Neurocytol 34: 353–360, 2006.[CrossRef]
Chen WR, Xiong W, and Shepherd GM. Analysis of relations between NMDA receptors and GABA release at olfactory bulb reciprocal synapses. Neuron 25: 625–633, 2000.[CrossRef][Web of Science][Medline]
Egan TM, Dagan D, and Levitan IB. Properties and modulation of a calcium-activated potassium channel in rat olfactory bulb neurons. J Neurophysiol 69: 1433–1442, 1993.
Ennis M, Zhou FM, Ciombor KJ, Aroniadou-Anderjaska V, Hayar A, Borrelli E, Zimmer LA, Margolis F, and Shipley MT. Dopamine D2 receptor-mediated presynaptic inhibition of olfactory nerve terminals. J Neurophysiol 86: 2986–2997, 2001.
Freeman WJ. Attenuation of transmission through glomeruli of olfactory bulb on paired shock stimulation. Brain Res 65: 77–90, 1974a.[CrossRef][Web of Science][Medline]
Freeman WJ. Relation of glomerular neuronal activity to glomerular transmission attenuation. Brain Res 65: 91–107, 1974b.[CrossRef][Web of Science][Medline]
Häusser M, Spruston N, and Stuart GJ. Diversity and dynamics of dendritic signaling. Science 290: 739–744, 2000.
Hayar A, Karnup S, Ennis M, and Shipley MT. External tufted cells: a major excitatory element that coordinates glomerular activity. J Neurosci 24: 6676–6785, 2004a.
Hayar A, Karnup S, Shipley MT, and Ennis M. Olfactory bulb glomeruli: external tufted cells intrinsically burst at theta frequency and are entrained by patterned olfactory input. J Neurosci 24: 1190–1199, 2004b.
Hayer A, Shipley MT, and Ennis M. Olfactory bulb external cells are synchronized by multiple intraglomerular mechanisms. J Neurosci 25: 8197–8208, 2005.
Helmchen F, Svoboda K, Denk W, and Tank DW. In vivo dendritic calcium dynamics in deep-layer cortical pyramidal neurons. Nat Neurosci 2: 989–996, 1999.[CrossRef][Web of Science][Medline]
Hirata Y. Some observations on the fine structure of synapses in the olfactory bulb of mouse, with particular reference to the atypical synaptic configurations. Arch Histol Jpn 24: 303–317, 1964.[Medline]
Holderith NB, Shigemoto R, and Nusser Z. Cell type-dependent expression of HCN1 in the main olfactory bulb. Eur J Neurosci 18: 344–354, 2003.[CrossRef][Web of Science][Medline]
Isaacson JS and Strowbridge BW. Olfactory reciprocal synapses: dendritic signaling in the CNS. Neuron 20: 749–761, 1998.[CrossRef][Web of Science][Medline]
Jahr CE and Nicoll RA. An intracellular analysis of dendrodendritic inhibition in the turtle in vitro olfactory bulb. J Physiol 326: 213–234, 1982.
Johnson BA and Leon M. Modular representations of odorants in the glomerular layer of the rat olfactory bulb and the effects of stimulus concentration. J Comp Neurol 422: 496–509, 2000.[CrossRef][Web of Science][Medline]
Keller A, Yagodin S, Aroniadou-Anderjaska V, Zimmer LA, Ennis M, Sheppard NF Jr, and Shipley MT. Functional organization of rat olfactory bulb glomeruli revealed by optical imaging. J Neurosci 18: 2602–2612, 1998.
Kosaka K, Toida K, Aika Y, and Kosaka T. How simple is the organization of the olfactory glomerulus? The heterogeneity of so-called periglomerular cells. Neurosci Res 30: 101–110, 1998.[CrossRef][Web of Science][Medline]
Luthi A and McCormick DA. H-current: properties of a neuronal and network pacemaker. Neuron 21: 9–12, 1998.[CrossRef][Web of Science][Medline]
McQuiston AR and Katz LC. Electrophysiology of interneurons in the glomerular layer of the rat olfactory bulb. J Neurophysiol 86: 1899–1907, 2001.
Migliore M and Shepherd GM. Emerging rules for the distributions of active dendritic conductances. Nat Rev Neurosci 3: 362–370, 2002.[CrossRef][Web of Science][Medline]
Mombaerts P, Wang F, Dulac C, Chao SK, Nemes A, Mendelsohn M, Edmondson J, and Axel R. Visualizing an olfactory sensory map. Cell 87: 675–686, 1996.[CrossRef][Web of Science][Medline]
Mori K, Nagao H, and Yoshihara Y. The olfactory bulb: coding and processing of odor molecule information. Science 286: 711–715, 1999.
Murphy GJ, Darcy DP, and Isaacson JS. Intraglomerular inhibition: signaling mechanisms of an olfactory microcircuit. Nat Neurosci 8: 354–364, 2005.[CrossRef][Web of Science][Medline]
Pinching AJ and Powell TP. The neuropil of the periglomerular region of the olfactory bulb. J Cell Sci 9: 379–409, 1971a.
Pinching AJ and Powell TP. The neuropil of the glomeruli of the olfactory bulb. J Cell Sci 9: 347–377, 1971b.
Pinching AJ and Powell TP. The neuron types of the glomerular layer of the olfactory bulb. J Cell Sci 9: 305–345, 1971c.
Price JL and Powell TP. The synaptology of the granule cells of the olfactory bulb. J Cell Sci 7: 125–155, 1970.
Puopolo M and Belluzzi O. Functional heterogeneity of periglomerular cells in the rat olfactory bulb. Eur J Neurosci 10: 1073–1083, 1998.[CrossRef][Web of Science][Medline]
Rall W, Shepherd GM, Reese TS, and Brightman MW. Dendrodendritic synaptic pathway for inhibition in the olfactory bulb. Exp Neurol 14: 44–56, 1966.[CrossRef][Web of Science][Medline]
Ressler KJ, Sullivan SL, and Buck LB. Information coding in the olfactory system: evidence for a stereotyped and highly organized epitope map in the olfactory bulb. Cell 79: 1245–1255, 1994.[CrossRef][Web of Science][Medline]
Reyes A. Influence of dendritic conductances on the input–output properties of neurons. Annu Rev Neurosci 24: 653–675, 2001.[CrossRef][Web of Science][Medline]
Rubin BD and Katz LC. Optical imaging of odorant representations in the mammalian olfactory bulb. Neuron 23: 499–511, 1999.[CrossRef][Web of Science][Medline]
Sah P. Ca2+-activated K+ currents in neurones: types, physiological roles and modulation. Trends Neurosci 19: 150–154, 1996.[CrossRef][Web of Science][Medline]
Santoro B, Chen S, Luthi A, Pavlidis P, Shumyatsky GP, Tibbs GR, and Siegelbaum SA. Molecular and functional heterogeneity of hyperpolarization-activated pacemaker channels in the mouse CNS. J Neurosci 20: 5264–5275, 2000.
Schoppa NE, Kinzie JM, Sahara Y, Segerson TP, and Westbrook GL. Dendrodendritic inhibition in the olfactory bulb is driven by NMDA receptors. J Neurosci 18: 6790–6802, 1998.
Senseman DM. High-speed optical imaging of afferent flow through rat olfactory bulb slices: voltage-sensitive dye signals reveal periglomerular cell activity. J Neurosci 16: 313–324, 1996.
Shepherd GM. Neuronal systems controlling mitral cell excitability. J Physiol 168: 101–117, 1963.
Shepherd GM. Physiological evidence for dendrodendritic synaptic interactions in the rabbit's olfactory glomerulus. Brain Res 32: 212–217, 1971.[CrossRef][Web of Science][Medline]
Shepherd GM, Chen WR, and Greer CA. Olfactory bulb. In: The Synaptic Organization of the Brain, edited by Shepherd GM. New York: Oxford Univ. Press, 2004.
Shipley MT and Ennis M. Functional organization of olfactory system. J Neurobiol 30: 123–176, 1996.[CrossRef][Web of Science][Medline]
Smith TC and Jahr CE. Self-inhibition of olfactory bulb neurons. Nat Neurosci 5: 760–766, 2002.[Web of Science][Medline]
Stewart WB, Kauer JS, and Shepherd GM. Functional organization of rat olfactory bulb analysed by the 2-deoxyglucose method. J Comp Neurol 185: 715–734, 1979.[CrossRef][Web of Science][Medline]
Vassar R, Chao SK, Sitcheran R, Nunez JM, Vosshall LB, and Axel R. Topographic organization of sensory projections to the olfactory bulb. Cell 79: 981–991, 1994.[CrossRef][Web of Science][Medline]
Wellis DP and Scott JW. Intracellular responses of identified rat olfactory bulb interneurons to electrical and odor stimulation. J Neurophysiol 64: 932–947, 1990.
Xu F, Liu N, Kida I, Rothman DL, Hyder F, and Shepherd GM. Odor maps of aldehydes and esters revealed by functional MRI in the glomerular layer of the mouse olfactory bulb. Proc Natl Acad Sci USA 100: 11029–11034, 2003.
Yokio M, Mori M, and Nakanishi S. Refinement of odor molecule tuning by dendrodendritic synaptic inhibition in the olfactory bulb. Proc Natl Acad Sci USA 92: 3371–3375, 1995.
Zhou ZS, Xiong WH, Chen WR, Hines ML, and Shepherd GM. Dendritic origin of a plateau potential in juxtaglomerular cells in the rat olfactory bulb. Soc Neurosci Abstr 28: 561.12, 2002.
Zhou ZS, Xiong WH, Chen WR, Hines ML, and Shepherd GM. Dynamic interaction between synaptic inputs and plateau potentials in olfactory bulb juxtaglomerular (JG) cells. Soc Neurosci Abstr 29: 821.16, 2003.
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Vdt). Stimuli were normalized with the plateau potential threshold. B: dependency of plateau potential on resting membrane potential. Ba: membrane potential responses to a series of current-pulse injections evoked from 2 different potential levels in the presence of 1 mM tetrodotoxin (TTX). Holding current was +102 and –286 pA, respectively, for –45 and –80 mV. Bb: plot of plateau potential amplitude against different levels of step depolarization. Plateau potential amplitude (
60 ms after the somatic potential. Arrowhead indicates a suppressed plateau potential at soma. D: cell image showing the recording site (arrowhead) on the dendrite. Somatic potentials in Aa, Ab, and C were induced by 20 ms intracellular current injection of 0.11, 0.10, and 0.21 nA, respectively. Resting membrane potential was –58 mV. Scale bars in C indicate the 20 mV for membrane potential, 50% for fluorescence change, and 200 ms for timescale.
