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REPORT
1Department of Anatomy, Howard University College of Medicine, Washington, District of Columbia; 2Departments of Cellular Biology and Anatomy and Pharmacology, Toxicology and Neuroscience, Louisiana State University Health Sciences Center, Shreveport, Louisiana; and 3Department of Anatomy and Neurobiology, University of Tennessee Health Science Center, Memphis, Tennessee
Submitted 14 November 2006; accepted in final form 25 December 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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). The cell bodies of GCs are densely packed within the GC layer (GCL) and they also occur more superficially, interspersed with mitral cell somata within the mitral cell layer (MCL). The population of superficial GCs (sGCs) is substantial; the rat MCL contains nearly 100,000 of these cells (Frazier and Brunjes 1988), but only roughly 40,000 mitral cells (Meisami 1989). The apical dendrites of both GC subtypes extend radially into the external plexiform layer (EPL), where they form dendrodendritic synapses with mitral and tufted cells. The mitral/tufted cell (M/TC)–GC dendrodendritic synapses have been well studied. Glutamate released by M/TC lateral dendrites activates 
-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) and N-methyl-D-aspartate (NMDA) receptors on GCs that, in turn, drive GABAergic feedback/feedforward inhibition of M/TCs (Aroniadou-Anderjaska et al. 1999; Chen et al. 2000; Isaacson and Strowbridge 1988; Schoppa et al. 1998).
Little is known about the physiological properties and functions of the two GC subtypes. In this regard, it is noteworthy that deep (d)GCs have been reported to express the Group I metabotropic glutamate receptor (mGluR) subtype mGluR5 and that the GCL and several other nonolfactory regions express the highest mGluR5 levels in the brain (Romano et al. 1995; Sahara et al. 2001). Immunohistochemical staining in the EPL indicates that mGluR5 is present on GC apical dendrites that are apposed to presynaptic glutamatergic synapses from M/TC lateral dendrites (van den Pol 1995). This localization of mGluR5 suggests that it plays a role in mediating dGC dendritic responses to glutamatergic inputs from M/TCs. In agreement with these observations, we recently reported that activation of mGluR5 directly excites dGCs (Heinbockel and Ennis 2003; Heinbockel et al. 2007). By contrast, mGluR5 staining of MCs and of the entire MCL is weak or absent (van den Pol 1995), suggesting that sGCs do not express mGluR5. MCs and the MCL densely express the other Group I mGluR subtype, mGluR1 (Fotuhi et al. 1993; Heinbockel et al. 2004; Sahara et al. 2001; Shigemoto et al. 1992; van den Pol 1995). However, it is not known whether sGCs express mGluR1. In the present study, we used patch-clamp electrophysiology and intracellular staining in olfactory bulb slices from wildtype mice and mice with targeted deletions of mGluR subtypes to determine whether excitability of sGCs is regulated by mGluR1, rather than by mGluR5. Parts of this study were previously published in abstract form (Heinbockel et al. 2005).
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Wildtype mice (C57BL/6J, Jackson Laboratory, Bar Harbor, ME) and mGluR1–/– and mGluR5–/– mutant mice (C57BL/6J background) from our colony were used in agreement with Institutional Animal Care and Use Committee and National Institutes of Health guidelines. The generation of mGluR1 and mGluR5 mutant mice was previously described (Chiamulera et al. 2001; Conquet et al. 1994). Mutant mice were maintained by heterozygous mating and were genotyped by polymerase chain reaction (PCR) of DNA from tail tip digests, as described previously (Heinbockel et al. 2004). Homozygous mGluR1–/– mice were also identified phenotypically by their mobility deficit at several weeks of age (Conquet et al. 1994).
Juvenile (21- to 31-day-old) mice were decapitated and the MOBs were dissected out and immersed in artificial cerebrospinal fluid (ACSF; see following text) at 4°C, as previously described (Heinbockel et al. 2004). Horizontal slices (400 µm thick) were cut parallel to the long axis using a vibratome (Vibratome Series 1000, Ted Pella, Redding, CA). After 30 min at 30°C, slices were incubated in a holding bath at room temperature (22°C) until use. For recording, a brain slice was placed in a recording chamber mounted on a microscope stage and maintained at 30 ± 0.5°C by superfusion with oxygenated ACSF flowing at 2.5–3 ml/min.
Electrophysiology
Visually guided recordings were obtained from cells in the MCL and GCL with near-infrared differential interference contrast optics and a BX50WI microscope (Olympus Optical, Tokyo, Japan) equipped with a charge-coupled detector (CCD) camera (CCD 100, Dage, Stamford, CT) and zooming coupler (Optem, Fairport, NY). Images were displayed on a monochrome monitor (Dage HR120, Dage-MTI, Michigan City, IN). Recording pipettes (5–8 M
) were pulled on a Flaming-Brown P-97 puller (Sutter Instrument, Novato, CA) from 1.5-mm OD borosilicate glass with filament. Seal resistance was routinely >1 G and liquid junction potential was 9–10 mV; reported measurements were not corrected for this potential. Data were obtained using an Axopatch 200B or Multiclamp 700A amplifier (Axon Instruments/Molecular Devices, Sunnyvale, CA). Signals were low-pass Bessel filtered at 2 kHz and digitized on videotape (AR Vetter, Rebersburg, PA) and computer disc (Axoscope/Clampex, Axon Instruments). Data were also collected through a Digidata 1200A (Axopatch) or Digidata 1322A (Multiclamp) Interface (Axon Instruments) and digitized at 10 kHz. Holding currents were generated under manual control by the recording amplifier. Membrane resistance was calculated from the amount of steady-state current required to hyperpolarize the cell by 10 mV, typically from –70 to –80 mV, or, in voltage clamp, from currents elicited by negative voltage pulses (10 mV, 1 s, 0.05 Hz). Current–voltage (I–V) plots were obtained from –130 to –40 mV using a voltage-step protocol (10-mV step intervals, 500 ms/step, Vhold = –70 mV) in the presence of blockers of fast synaptic transmission (see following text) and tetrodotoxin (TTX). To compare the agonist-induced current in sGCs and dGCs, the mean current at –70 mV in the absence of agonist was subtracted from that in the presence of agonist (Fig. 3). Numerical data are expressed as means ± SE. Tests for statistical significance (P < 0.05) were performed using paired Student's t-test.
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). The standard pipette-filling solution consisted of (in mM) K gluconate 125, MgCl2 2, HEPES 10, Mg2ATP 2, Na3GTP 0.2, NaCl 1, and EGTA 0.2. The following drugs were bath applied: L-2-amino-5-phosphonopentanoic acid (AP5, APV), 6-cyano-7-nitroquinoxaline-2-3-dione (CNQX), (2S,2'R,3)-2-(2',3'-dicarboxycyclopropyl)glycine (DCG-IV), (RS)-3,5-dihydroxyphenylglycine (DHPG), (2-(3-carboxypropyl)-3-amino-6-(4-methoxyphenyl)-pyridazinium bromide (gabazine, SR-95531), and TTX. Chemicals and drugs were supplied by Sigma–Aldrich (St. Louis, MO) and Tocris Cookson (Ellisville, MO). Histology
Biocytin (0.05–0.1%, Molecular Probes/Invitrogen, Carlsbad, CA) was added to the pipette solution to allow histological confirmation of recorded cells. After recording, slices were fixed in 4% paraformaldehyde in 0.1 M Na+ phosphate buffer or in 2.5% paraformaldehyde in phosphate buffer containing 15% saturated picric acid + 1.5% glutaraldehyde. The slices were embedded in 10% gelatin, sectioned at 60- to 100-µm thickness using a vibratome, and processed using immunoperoxidase methods (ABC Elite and DAB kits, Vector Labs, Burlingame, CA) to permanently stain the recorded cells. Photomicrographs were obtained using a microscope equipped with a digital camera (Optronics MicroFire, Goleta, CA). The brightness, contrast, color balance, and sharpness of the images were altered to optimize visibility of fine dendrites using Adobe Photoshop CS (San Jose, CA).
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RESULTS |
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, n = 128 sGCs and dGCs from wildtype and knockout mice). Additionally, the lack of membrane bistability distinguished GCs from MCs (Heinbockel et al. 2004; Heyward et al. 2001). Identification of six sGCs and 14 dGCs was subsequently verified by histological examination of the soma location within the MCL (Fig. 1A) or GCL (Fig. 1B), respectively. Group I mGluR agonist depolarizes superficial granule cells and increases firing rate
Intrinsic properties did not differ significantly between sGCs and dGCs from wildtype mice. sGCs were characterized as follows: membrane potential, Vm = –68.4 ± 1.5 mV; input resistance: 892.0 ± 78.0 M; ability to generate spikes: 94.7% (n = 35). dGCs showed the following characteristics: Vm = –66.2 ± 1.3 mV (P = 0.18 vs. sGCs); input resistance: 947.5 ± 106.2 M (P = 0.2 vs. sGCs); ability to generate spikes: 92.6% (n = 46). As shown in Fig. 1C, bath application of the selective Group I mGluR agonist DHPG (50 µM) depolarized and increased the spontaneous discharge of sGCs (
mV = 16.2 ± 1.7 mV, n = 14) recorded in slices from wildtype mice. The effects of DHPG were similar to those observed in dGCs in wildtype mice (Fig. 1D), as previously described (mV in dGCs mice = 15.3 ± 3.2 mV, n = 10; Heinbockel and Ennis 2003; Heinbockel et al. 2007).
To determine whether the effects of DHPG on sGCs were mediated by direct activation of mGluRs, DHPG was applied in the presence of blockers of ionotropic glutamate and 
-aminobutyric acid (GABA) receptors (synaptic blockers: CNQX, 10 µM; APV, 50 µM; gabazine, 5 µM). Under these conditions, application of DHPG resulted in substantially smaller, but nevertheless consistent, membrane depolarization (mV in synaptic blockers: 6.9 ± 0.7 mV, n = 14; Fig. 1C, bottom trace), similar to the effect observed in dGCs (mV in synaptic blockers: 8.0 ± 1.1 mV, n = 11; Heinbockel et al. 2007; Fig. 1D, bottom trace). We previously showed that DHPG directly depolarizes MCs and increases their firing rate by activation of mGluR1 (Heinbockel et al. 2004). We therefore interpret the greater effect of DHPG on sGCs and dGCs in the absence of synaptic blockers to result from 1) direct depolarization of GCs by the agonist and 2) indirect excitation as a consequence of increased glutamate release from MCs. The latter effect is eliminated when DHPG is applied in the presence of synaptic blockers, resulting in reduced depolarization.
DHPG excitation of sGCs is present in mGluR5–/– mice and absent in mGluR1–/– mice
We next took advantage of mice with targeted deletions of the mGluR5 or mGluR1 receptor genes (Chiamulera et al. 2001; Conquet et al. 1994) to investigate the role of these two Group I receptor subtypes in the DHPG-evoked depolarization of sGCs and dGCs. We examined the current–voltage (I–V) relationship (see METHODS) of DHPG-evoked responses of GCs recorded in slices from wildtype, mGluR1 mutant (mGluR1–/–), and mGluR5 mutant (mGluR5–/–) mice. In the presence of synaptic blockers and TTX, DHPG (50 µM) consistently evoked an inward current in sGCs in slices from the mGluR5–/– mice (n = 5 cells from five animals; Fig. 2Aa). By contrast, no DHPG-evoked currents were observed in sGCs in slices from mGluR1–/– mice at any membrane potential tested (n = 4 cells from four animals; Fig. 2Ab). Opposite results were observed for dGCs (Fig. 3). No effect of DHPG was observed on dGCs in slices from mGluR5–/– mice (n = 6 cells; Fig. 2Ba), but inward currents were consistently observed in slices from mGluR1–/– mice (n = 6 cells; Fig. 2Bb).
DHPG-evoked currents in GCs recorded in slices from wildtype mice were maximal at voltages near the GC resting membrane potential (–90 to –70 mV). To compare the magnitude of DHPG-evoked currents in wildtype and knockout mice, we quantified the amplitude of DHPG-induced inward current at –70 mV (Fig. 3). Amplitudes of DHPG-evoked currents were similar in sGCs (–7.9 ± 1.9 pA) and dGCs (–6.3 ± 3.0 pA) from wildtype mice and in sGCs from mGluR5–/– mice (7.2 ± 0.9 pA; Fig. 3). No significant (P > 0.05) currents were observed in dGCs from mGluR5–/– mice or in sGCs from mGluR1–/– mice. The DHPG-evoked current in dGCs from mGluR1–/– mice was substantially larger than that of all other groups (–19.6 ± 4.0 pA, P = 0.02 compared with dGCs from wildtype mice). The reason for this is unclear but it may reflect a compensatory change in the number or affinity of mGluR5 in mGluR1–/– mice.
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DISCUSSION |
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). Our findings that mouse sGCs express mGluR1, whereas mouse dGCs express mGluR5, therefore suggest that these two GC subtypes are functionally as well as anatomically distinct. These results are consistent with previous reports of anatomical and physiological differences between these two populations of GCs.
At resting membrane potential, DHPG elicited an inward current in sGCs and in dGCs. For dGCs, the DHPG current reversed in polarity at about –100 mV and appeared to decrease at potentials positive to –50 mV. This suggests that the current might involve multiple components, similar to DHPG currents in several brain areas (Anwyl 1999). We recently showed that a major component of the DHPG-mediated inward current in dGCs is mediated by closure of K+ channels associated with a K+ leak current (Heinbockel et al. 2007). Activation of Group I mGluRs is known to reduce leak K+ currents in several systems (Glaum and Miller 1992; Guerineau et al. 1994; McCormick and von Krosigk 1992; Takeshita et al. 1996). Other components of the current may involve nonselective cation conductances (Congar et al. 1997; Crepel et al. 1994; Guerineau et al. 1995; Raggenbass et al. 1997) or electrogenic Na+/Ca2+ exchangers (Keele et al. 1997; Lee and Boden 1997; Staub et al. 1992).
Activation of mGluR1 induced an inward current in sGCs that appears to be similar to the DHPG-evoked, mGluR1-mediated current in rat and mouse MCs (Heinbockel et al. 2004). In both MCs and sGCs, DHPG produces a parallel inward shift of the I–V relationship at all membrane potentials tested. The current in MCs involves multiple components and is eliminated by a combination of K+ and Ca2+ channel blockers and intracellular calcium chelation. Additional experiments are necessary to delineate the properties of mGluR1-mediated current in sGCs.
Our finding that dGCs express mGluR5 but not mGluR1 is consistent with observations that GCL somata exhibit immunocytochemical staining for mGluR5, that this staining is strongest within the deep GCL (Sahara et al. 2001), and that the GCL stains weakly for mGluR1 (Martin et al. 1992; Sahara et al. 2001; van den Pol 1995), although faint mGluR1 expression was previously observed in this layer with in situ hybridization (Shigemoto et al. 1992). Our finding that sGCs express mGluR1 is also consistent with immunohistochemical staining and in situ hybridization studies showing high levels of mGluR1 expression in the MCL (Fotuhi et al. 1993; Martin et al. 1992; Masu et al. 1991; Sahara et al. 2001; Shigemoto et al. 1992; van den Pol 1995). However, our results show that mGluR1 expression in the mouse MCL is not restricted to MCs, but also includes sGCs. In the rat, M/TC dendrites, but not GC dendrites, stain strongly for mGluR1 (van den Pol 1995). Based on the present results, the dendrites of mouse sGCs should also stain for mGluR1. These apparent discrepancies may be explained by 1) species differences in subcellular localization of mGluR1, 2) differential GC expression of mGluR1 splice variants in mice versus rats, or 3) the region of the EPL that was sampled in the van den Pol (1995) study of mGluR1 staining in the rat (see following text).
Previous anatomical studies showed that two or three GC subtypes occur in rodents and rabbits, which exhibit different soma locations and/or distributions of apical dendrites and dendritic spines within the EPL. Apical dendrites of rat sGCs typically extend into the superficial EPL, where they exhibit their highest spine densities, whereas apical dendrites of rat dGCs are largely confined to the deep EPL, where they exhibit their highest spine densities (Orona et al. 1983). These anatomical differences are correlated with differential staining of the superficial versus deep rat EPL for a variety of markers, including glutamic acid decarboxylase (Mugnaini et al. 1984) and certain GABAA (Panzanelli et al. 2005) and AMPA receptor subunits (Montague and Greer 1999). In the rabbit, however, apical dendrites of some dGCs reach the superficial EPL, where they exhibit their highest spine densities, apical dendrites of some sGCs are restricted to the deep EPL, and apical dendrites of a third GC subtype (intermediate GCs) exhibit high spine densities in both the superficial and deep EPL (Mori et al. 1983). Normal heterozygous littermates of Purkinje Cell Degeneration mice also exhibit three GC subtypes, which resemble the three rabbit GC subtypes (Greer 1987). Our results indicate that apical dendrites of some dGCs of C57BL/6J mice reach the most superficial portions of the EPL (Fig. 1D), like apical dendrites of dGCs seen in this mouse strain and in the rabbit, but unlike apical dendrites of most rat dGCs. We did not examine responses of the third GC subtype (intermediate GCs) in this study, so it is not known whether it expresses mGluR1, mGluR5, or both.
The apical dendrites of superficial and deep GCs are thought to interact with lateral dendrites of different M/TC subtypes at different depths within the EPL. The lateral dendrites of most TCs ramify within the superficial and middle (intermediate) EPL, so they presumably interact with apical dendrites of sGCs (Macrides et al. 1985; Mori 1987; Mori et al. 1983; Orona et al. 1984). By contrast, the lateral dendrites of MCs ramify within the middle and deep EPL, so they presumably interact with the apical dendrites of dGCs. Our results suggest that mGluR1 might therefore play a role in responses of sGCs to TC input, whereas mGluR5 might play a role in responses of dGCs to MC input. However, this TC–sGC and MC–dGC relationship should not be considered an absolute segregation because three different TC subtypes occur, which extend lateral dendrites within the superficial, middle, or even deep portions of the EPL (Scott and Harrison 1987). Additionally, two MC subtypes occur that extend lateral dendrites preferentially within the middle or deep EPL zones (Kiyoshi et al. 1982; Orona et al. 1984).
Little is known about the roles of different subpopulations of GCs in olfactory processing. Earlier in vivo recordings in rats provided evidence that sGCs are functionally distinct from dGCs (Wellis and Scott 1990). Odor stimulation induced spikes in most (six of eight) GCs located within about 150 µm of the MCL, but only in one of four deeper GCs, even though spiking could be induced in the dGCs by lateral olfactory tract stimulation. Anatomical findings suggest that dGCs may be more heavily targeted by excitatory projections from higher-order olfactory structures than sGCs. Inputs to MOB from olfactory cortical structures (such as piriform cortex, periamygdaloid nucleus, nucleus of the lateral olfactory tract, anterior olfactory nucleus) terminate preferentially in the GCL (Laaris et al. 2007; Luskin and Price 1983). Recent studies suggest that the physiological influence of these cortical inputs is focused on dGCs in the GCL (Laaris et al. 2007). Thus dGCs may play a more important role than sGCs in feedback regulation of the MOB by olfactory cortical structures.
In the mature olfactory bulb, GCs are continuously generated from progenitor cells originating from the rostral migratory stream (Lois and Alvarez-Buylla 1994; Luskin 1993; Smith and Lushkin 1998; Wichterle et al. 2001). Adult-born GCs distribute to the MCL and GCL, where many survive and become functionally integrated into the MOB circuitry (Belluzzi et al. 2003; Carleton et al. 2003) depending on olfactory input (Corotto et al. 1994; Fiske and Brunjes 2001; Frazier-Cierpial and Brunjes 1989; Najbauer and Leon 1995; Rochefort et al. 2002). There is evidence that different subsets of adult-born GCs migrate into the MCL and superficial GCL. Adult-born GCs that express the ETS transcription factor Er81 distribute to the superficial GCL and MCL (Stenman et al. 2003). Similarly, adult-born GCs that distribute specifically to the MCL express the leucine-rich repeat membrane protein 5T4 (Imamura et al. 2006). Based on the present findings, it is reasonable to speculate that sGCs that express Er81 or 5T4 may also express mGluR1.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: T. Heinbockel, Dept. of Anatomy, Howard University College of Medicine, 520 W Street, NW, Washington, DC 20059 (E-mail: theinbockel{at}howard.edu)
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REFERENCES |
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