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Dynamic development of the first synapse impinging on adult-born neurons in the olfactory bulb circuit
© Katagiri et al; licensee BioMed Central Ltd. 2011
Received: 9 September 2010
Accepted: 1 November 2010
Published: 1 February 2011
The olfactory bulb (OB) receives and integrates newborn interneurons throughout life. This process is important for the proper functioning of the OB circuit and consequently, for the sense of smell. Although we know how these new interneurons are produced, the way in which they integrate into the pre-existing ongoing circuits remains poorly documented. Bearing in mind that glutamatergic inputs onto local OB interneurons are crucial for adjusting the level of bulbar inhibition, it is important to characterize when and how these inputs from excitatory synapses develop on newborn OB interneurons. We studied early synaptic events that lead to the formation and maturation of the first glutamatergic synapses on adult-born granule cells (GCs), the most abundant subtype of OB interneuron. Patch-clamp recordings and electron microscopy (EM) analysis were performed on adult-born interneurons shortly after their arrival in the adult OB circuits. We found that both the ratio of N-methyl-D-aspartate receptor (NMDAR) to α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor (AMPAR), and the number of functional release sites at proximal inputs reached a maximum during the critical period for the sensory-dependent survival of newborn cells, well before the completion of dendritic arborization. EM analysis showed an accompanying change in postsynaptic density shape during the same period of time. Interestingly, the latter morphological changes disappeared in more mature newly-formed neurons, when the NMDAR to AMPAR ratio had decreased and functional presynaptic terminals expressed only single release sites. Together, these findings show that the first glutamatergic inputs to adult-generated OB interneurons undergo a unique sequence of maturation stages.
Adult neurogenesis, a process encompassing the generation, maturation and synaptic integration of new neurons in the adult brain, represents a striking form of structural adult neural plasticity [1–3]. In the adult olfactory bulb (OB), nearly all newly recruited neurons become local interneurons . About 95% of the new cells differentiate into granule cells (GCs) and less than 4% become periglomerular cells [5, 6], with glutamatergic juxtaglomerular neurons accounting for the rest . The newly-formed interneurons project dendrites that establish synaptic contacts with pre-existing partners, becoming indistinguishable from other mature interneurons within few weeks [8–13].
In this study, we focused on the excitatory proximal inputs to newborn GCs. Pioneering studies on embryonal and postnatal development have shown that the first contacts to be formed are MC-GC dendrodendritic synapses at embryonic day (E)17. The formation of these first synaptic contacts is followed by the appearance of MC-GC axodendritic synapses at E18 [27, 28]. Thus, key synapses important for the OB functioning are already present at birth, when olfaction is crucial for the survival of rodent pups. In contrast to embryonic synaptogenesis, we found that axodendritic proximal synapses were the first synaptic contacts formed on developing new GCs, shortly after the newcomer migrates radially into the bulb. Using electrophysiological and morphological approaches, we demonstrated that proximal synaptic inputs on newborn GCs are functionally developed and profoundly evolved over time much before the appearance of their output synapses. The overall developmental pattern of the proximal inputs on newborn GCs may be essential to shape the functional impact of the adult neurogenesis and, as a consequence, for proper circuit function.
Structural reorganization of the glutamatergic synapses during GC maturation
Previous morphological studies have shown that adult-born GCs can be categorized into five distinct classes . To characterize the early steps of synaptic integration, we focused on class 3 to class 5 neurons. We thus excluded migrating neurons (class 1 and 2) from the present analysis, as these cells do not exhibit direct synaptic activity (data not shown). We injected a green fluorescent protein (GFP)-encoding lentiviral vector into the rostral migratory stream (RMS) to label migrating neuroblasts [12, 30]. At 3 days post-injection (dpi) of viruses, most GFP-positive cells were still navigating in the RMS, but a small proportion had already reached the bulb class 3 neurons (Figure 1A; see ). Their apical dendrites first reached the external plexiform layer a few days later (class 4). This step was followed by the formation of lateral branches with spines (class 5) (Figure 1A).
To examine early glutamatergic inputs contacting newborn GCs, we used pre-embedding immunolabeling for GFP combined with electron microscopy (EM). This ultrastructural analysis was performed in the deep portion of the GC layer, thus excluding TC axons located in the internal plexiform layer. At 7 dpi, EM analysis showed GFP-positive cells in the GC layer receiving asymmetric contacts from axon terminals. Serial sectioning revealed that some synapses at 7 dpi had a simple shape with a compact and homogeneous post-synaptic density (PSD) (7 over 13 sections) (Figure 1B, single cell; Figure 1C), whereas other synaptic junctions exhibited discontinuities of the PSD (6 over 13 sections; not shown). Synapses impinging onto more advanced maturing cells (21 dpi) and pre-existing GFP-negative cells exhibited more complex postsynaptic profiles, most notably, a doughnut-like shape with the PSD forming an almost complete ring around the central 'hole' (Figure 1B) (21 dpi cells: 5 over 13 sections; GFP-negative cells: 6 over 16 sections). Despite changes in the shape of the synaptic junctions, we did not observe any obvious differences in the ultrastructure of the presynaptic boutons contacting GC spines throughout the distinct maturation stages. All in all, these observations suggest that early proximal excitatory inputs on adult-generated GCs are formed within a few days of these cells arriving to the GC layer and that they then undergo a reorganization of PSDs (Figure 1D) during subsequent maturation stages.
Functional characterization of early proximal glutamatergic inputs on new GCs
To investigate the functional properties of developing synapses in the GC layer, we monitored electrophysiological events using whole-cell patch-clamp recordings (Figure 2A). We first analyzed the profiles for both inward Na+ currents and input membrane resistance (Rin). Previous studies showed that inward Na+ currents appeared mostly on class 5 newborn GCs; that is, one week after viral injection . The presence of the apical dendrite was visualized both by GFP labeling and by loading cells with biocytin during patch-clamp recordings (Figure 1A). As expected, total dendritic length increased together with the increase in amplitude of the Na+ current (see Additional file 1A). By contrast, an inverse correlation was found between the Na+ current amplitude and Rin (see Additional file 1B), known to decrease during neuronal maturation . Therefore, the amplitude of the voltage-gated Na+ current could be used as a reliable proxy of maturation stage (see Additional file 1C). Rather than categorizing recorded cells by the number of dpi (see Additional file 1D), we classified recorded individual GFP-positive GCs according to the combination of three parameters: 1) maximum Na+ current amplitude, 2) dendrite length, and 3) Rin values. Class 3 neurons, recorded at about 4 dpi, expressed a Na+ current of <150 pA, dendrite length <100 μm and Rin values >2.7 GΩ (see Additional file 1). Class 4 neurons, recorded at about 6 dpi, had a Na+ current amplitude ranging from 200 to 800 pA, dendrite length of up to 280 μm and Rin values of between 2.3 and 5 GΩ. Class 5 neurons, recorded at about 21 dpi, had a Na+ current amplitude >800 pA, dendrite length of up to 310 μm and Rin values <2.5 GΩ.
We then mapped connectivity for the recorded GCs using a minimal stimulation protocol delivered at proximal domains of GC dendrites. Previous studies suggested that excitatory glutamatergic inputs to the proximal dendrites of GCs originate from local collaterals of MC axons and centrifugal feedback projections from cortical regions . However, these two inputs differ substantially in their location in the GC layer; centrifugal inputs remain deep in the GC layer [16, 17]. Using this criterion, we focused our analysis on inputs showing the hallmarks of centrifugal feedback projections [17, 26], bearing in mind that patch recordings are performed from the cell body whereas the synapses on the EM analysis originate from the dendritic spines some distance away on proximal dendrites. Starting with a low-intensity stimulation that did not evoke any excitatory postsynaptic currents (EPSCs), we then increased the stimulus strength gradually until a fast EPSC appeared. Only when a stimulating electrode was placed into the GC layer (Figure 2A) near the soma of the class 3 neurons did we observe EPSCs in an "all or nothing" manner (Figure 2B and Additional file 2). In our previous report , using a similar position for the stimulating electrode we observed EPSCs similar to those evoked by the axodendritic inputs in the proximal domain of the apical dendrite (Figure 2A). Stimulation near this recruitment threshold often failed to induce an EPSC. Slightly increasing the stimulation intensity beyond that threshold prevented the occurrence of failures, without affecting the amplitude of successful responses (see Additional file 2). In all cases, further small increments in stimulus intensity did not affect EPSC amplitude.
We characterized the development of synaptic inputs for the three classes of newly-formed neurons by using the voltage-clamp technique to measure isolated AMPAR-mediated EPSCs at a holding potential of -70 mV. After EPSCs were recorded at +40 mV with the same stimulus strength, we applied the AMPAR antagonist 2,3-dioxo-6-nitro-1,2,3,4-tetrahydrobenzo[f]quinoxaline-7-sulfonamide (NBQX) at 10 μmol/l. Finally, we confirmed, using the NMDAR antagonist D,L-2-amino-5-phosphonopentanoic acid (D,L-APV) at 100 μmol/l, that EPSCs were mediated by NMDAR. Gabazine (SR-95531) at 10 μmol/l was used to block GABAA receptor-mediated events and was present throughout all experiments. Minimal stimulation evoked both AMPAR-mediated and NMDAR-mediated EPSCs in newborn GCs in a success or failure manner (Figure 2B, C). After we subtracted mean failure responses from all responses (that is, success and failure) to prevent the obscuration by the stimulus artifact, we compared AMPAR-mediated and NMDAR-mediated EPSC amplitude among classes. NMDAR-mediated current amplitude gradually increased with more advanced maturation (class 3: 1.8 ± 0.6 pA; class 4: 5.0 ± 1.6 pA; class 5: 8.8 ± 1.4 pA) (Figure 2D). By contrast, AMPAR-mediated EPSC amplitude did not differ between class 3 and class 4 GCs (class 3: 4.6 ± 1.0 pA; class 4: 4.4 ± 0.9 pA), but was significantly increased in class 5 neurons (19.3 ± 7.6 pA; p < 0.05) (Figure 2E). These findings show that stimulation of OB slices, when delivered at minimal intensity, is a useful approach to study the maturation of a single presynaptic terminal contacting a maturing adult-generated GC.
To exclude the possibility that the abrupt increase in AMPAR-mediated EPSC amplitude at class 5 might reflect the transition from regimens in which Na+ current is adequately clamped (class 3 and 4) to a regimen in which potentially it could not be clamped so well (class 5), we compared the amplitude of AMPAR-mediated outward EPSCs at the depolarized membrane potential. When these currents were derived from the subtraction of NMDAR-mediated components from EPSCs without NBQX, we obtained similar results (see Additional file 3). Taken together, these results suggest that NMDAR and AMPAR might have distinct roles during the maturation processes of adult-born GCs.
Functional maturation of NMDAR- and AMPAR-mediated EPSCs
The conductance of ionotropic glutamate receptors depends on subunit composition . We therefore used non-stationary fluctuation analysis to estimate single-channel conductance [35, 36]. The relationship between the variance and mean amplitude of NMDAR-mediated current was found to be skewed (Figure 3C). We fitted the initial slope to estimate the mean single-channel current underlying NMDAR-mediated EPSCs, and then calculated the single channel conductance. Results showed a gradual increase in NMDAR single-channel conductance over time (class 3: 19.7 ± 3.5 pS; class 4: 33.9 ± 8.2 pS; class 5: 52.0 ± 5.4 pS) (Figure 3D).
Mechanism of transmitter release at newly-formed synapses
Properties of proximal synapses on mature GCs
Although the amplitudes of AMPAR-mediated and NMDAR-mediated EPSCs recorded from mature GCs were significantly higher than those from class 3 GCs (AMPAR-mediated events 26.7 ± 8.2 pA; NMDAR-mediated events 11.8 ± 5.1 pA), the NMDAR/AMPAR ratios for mature and class 3 GCs were indistinguishable (Figure 7D). This ratio was maximal for class 4 and class 5 neurons (class 3: 0.47 ± 0.11; class 4: 1.30 ± 0.17; class 5: 0.88 ± 0.28; mature: 0.57 ± 0.28). These findings suggest that AMPARs are functionally dominant at proximal synapses in both class 3 neurons and mature GCs.
Formation of the first glutamatergic contacts
Class 1 and 2 cells are migratory neurons , which express AMPAR before reaching their final position in the bulb [8, 45]. AMPAR-mediated current recorded from these migrating cells showed an almost linear current-voltage relationship, indicative of Ca2+-impermeable AMPARs . By contrast, AMPAR-mediated currents at the proximal synapses of class 3 cells showed inward rectification (see Additional file 4) indicative of the presence of Ca2+-permeable AMPAR channels, as it has been recently demonstrated using flash photolysis of caged glutamate . Supporting this assumption is the large single channel conductance for AMPAR estimated from our non-stationary noise analysis (Figure 4D), as previously reported . Intracellular Ca2+ is thought to play an important role in regulating the migration and maturation of newborn neurons [45, 48]. Further experiments will determine whether this Ca2+ permeability mediates a stop signal that halts the radial migration of new GCs, as demonstrated for migrating neurons of the embryonic cortex .
Class 3 GCs that have just reached the OB show limited dendritic arborization (< 100 μm) and a virtual absence of spines; however, we detected early functional glutamatergic inputs at this time point (Figure 2). Meanwhile, newborn neurons in the hippocampus receive their first glutamatergic synapses about two weeks after developing in the GC layer . Further studies should investigate whether the difference in the onset of glutamatergic inputs might support differences in new neuron turnover between the OB and hippocampus, as previously reported .
We found that a single axon terminal formed synapses onto the dendrites of both GFP-positive and GFP-negative GCs (see Additional file 5). This suggests that immature and mature synapses share the same presynaptic input, assuming that GFP-negative cells represent mature pre-existing GCs. Notably, the distance between synapses in immature and pre-existing GC dendrites can be as small as < 1 μm (see Additional file 5). Such proximity suggests that new GCs may integrate into the mature circuit using pre-existing terminals, as previously described for new neurons of the adult dentate gyrus . Upon arrival of a new GC, glutamate release from the pre-existing terminals may trigger the extension of dendritic filopodia, resulting in the formation of postsynaptic specializations. At the same time, directed recruitment of mobile vesicles to nascent active zones in pre-existing terminals  may lead to the formation of additional presynaptic release sites (Figure 8B, bottom). One potential benefit of such a coordinated mechanism would be the ability to rapidly initiate the formation of new synapses.
Maturation of glutamatergic synapses at the postsynaptic site
When a presynaptic fiber contacts the proximal dendrites of class 3 neurons, mobile transport packets, containing presynaptic and postsynaptic proteins, are immediately recruited for synaptic maturation . We observed that the amplitude of EPSCs mediated only by NMDAR was higher in class 3 than in class 4 cells. Furthermore, NMDARs may also play an active role in keeping immature synapses free from AMPARs until an appropriate trigger signals the recruitment of this receptor, as described in developing circuits . Thus, the incorporation of NMDARs may precede the insertion of AMPARs during the early maturation stages in adult newborn GCs.
We also found a shorter duration of NMDAR-mediated currents in class 4 neurons than in class 3 neurons. Increasing evidence suggests an acceleration of NMDAR-mediated events during neuronal maturation, reflecting a switch in subunit composition . NMDARs consist of two obligate NR1 subunits and two NR2A-D or NR3A-B subunits, the recruitment of which depends on spatial and developmental signals . Our findings indicate that NMDAR channel conductance increased from low to high conductance during the maturation process, possibly because the NMDAR-mediated currents recorded in class 3 cells were presumably mediated by NR2D-containing receptors, known to support low conductance states . Reconstructed systems have shown that NR2D-containing receptors mediate currents with a long decay-time constant , consistent with the prolonged decay-time constant observed in class 3 cells. Additionally, morphological analyses have shown that the NR2 D subunit is mainly located at extrasynaptic sites . The slow rise time of NMDAR-mediated currents recorded from the centrifugal fibers of class 3 cells could result from the activation of extrasynaptic NR2D-containing receptors. Then, during maturation of newborn cells from class 3 to class 4, high-conductance NMDARs may be recruited from extrasynaptic to synaptic sites. This scenario would be consistent with the shortened rise time of NMDAR-mediated currents observed at the later maturation stages; however, pharmacological experiments should be conducted to directly test this possibility.
During development, glutamatergic synaptic currents undergo a characteristic pattern of maturation involving changes in the kinetics of NMDAR-mediated currents. The formation of 'silent' synapses, which display only NMDAR-mediated currents and which are later made functional through recruitment of AMPARs, is also a possibility. These changes in glutamatergic synapses during maturation have implications for the early development of neural networks  and for the mechanisms underlying the LTP of synaptic efficacy . In our study, we detected the highest NMDAR/AMPAR ratio in class 4 cells, even though axodendritic synapses are not 'silent' at the stage of class 3. This high NMDAR/AMPAR ratio declined with neuronal age as a result of AMPAR recruitment. This observation suggests that the development of axodendritic synapses after the class 4 stage (that is, from 6 dpi) somewhat recapitulates the mechanism of maturation described during brain development.
Changes in synapse ultrastructure on developing GCs
Our study found changes in functional release sites during the maturation process. Minimal stimulation experiments revealed the presence of a single functional release site in class 3 GCs and multiple functional release sites in both class 4 and class 5 GCs. Previous studies have shown that multiple vesicles might be released at each synaptic contact for each action potential fired [62, 63]. These changes could reflect a maturation of individual synapses, involving developmental switching from single to multiple release sites (Figure 8B, top) [62, 63]. Indeed, previous ultrastructural analyses have demonstrated that developing (class 3) neurons establish immature synapses characterized by small cluster of vesicles (that is, a group of >4 vesicles) and faint membrane specializations , whereas mature synapses contain large cluster of vesicles and substantial asymmetric membrane thickening  (and this study).
Each neurotransmitter release site in the mature brain functions in a binary mode, releasing one or zero quantum of neurotransmitter for each action potential [38, 64]. Therefore, many excitatory synapses in the mammalian brain can release a maximum of the contents of one vesicle . Alternatively, the changes described here may reflect changes in the number of synaptic appositions contacting a given GC (Figure 8B, bottom). A single synapse may first develop into a more complex contact (involving the presence of perforations) and then split into two synapses, still connected to the same single axon. Interestingly, LTP has been reported to initiate the perforation of spines, leading to the appearance of multiple spine boutons . Because LTP has been observed at the proximal synapses of GCs [25, 26], it is possible that the changes in the number of synaptic contacts we found might reflect some form of synaptic plasticity.
At later stages of maturation, we found that GCs revert to a single functional release site. The paired-pulse ratio was smaller after supraminimal stimulation than after minimal stimulation in mature, but not in class 5 GCs () (Figure 7B). This could arise through presynaptic events, such as successful release at most release sites triggered by the first supraminimal pulse, or through postsynaptic events, such as saturation of receptors after multivesicular release . Two possible models may thus be considered for the axodendritic synapse of mature GCs: individual boutons displaying multiple dense release sites (Figure 8B, top) or synaptic connections with multiple release sites in close proximity (Figure 8B, bottom). In both cases, multivesicular events occur in synchrony at the axodendritic synapse of more mature GCs.
In this study we found that, within a few days after having reached the OB, the first glutamatergic synapses underwent a dynamic developmental process. This proximal excitatory input is subjected to both physiological and morphological modifications before reciprocal dendrodendritic synapses start to operate at distal sites. Our results provide a conceptual framework for understanding mechanisms underlying the precise control of neural wiring through presynaptic and postsynaptic events during adult neurogenesis. It is remarkable that both basic physiological properties (this study) and the synaptic plasticity  were found to be continuously changing over time, even when the ultrastructure of newly-formed synapses became similar to that of mature counterparts. This finding contrasts with the observation that GCs in the adult dentate gyrus continue to change physiologically and morphologically after the formation of their first glutamatergic synapses [50, 68, 69]. We indicate here that the influence of adult OB neurogenesis may be gradually tuned by early glutamatergic synaptic transmission, depending on the activity of cortical structures downstream from the OB. Elucidating the origin of the glutamatergic centrifugal fibers impinging so early onto newborn GCs, and the molecular mechanisms that shape the integration of new neurons, has important implications both for understanding how OB circuits are refined by experience and for the successful use of stem cell-based replacement therapies in brain repair.
C57BL/6J male mice about eight weeks old (Janvier, France) were used for all experiments. All procedures were carried out in accordance with the EU Charter of Fundamental Rights (2000/C 364/01) and the European Communities Council Directive of 24 November 1986 (86/609/EEC), and were reviewed and approved by our institutional animal welfare committee.
For cytosolic GFP labeling, a custom-built lentivirus containing the GFP gene under the control of the PGK promoter was used to transduce adult-born neurons as previously described . Viruses (2.2 × 1010 transducing units (TU)/ml) were stored at -80°C. Immediately before injection, the lentiviral vectors were diluted in phosphate-buffered saline (PBS) to a final concentration of 15 ng of p24 protein per microliter.
For stereotaxic injections of lentiviral vectors, adult mice about six weeks old were anesthetized with a mixture of ketamine (Imalgene®, Merial, Lyon, France) (1.5% in PBS) and xylazine (Rompun®, Bayer Health Care, Puteaux France) (0.05%; 250 μl per mouse). Mice were mounted in a Kopf stereotaxic apparatus, and small craniotomies were created above the injection sites. Virus was injected into the rostral migratory stream (RMS) at the following co-ordinates in each hemisphere: anteroposterior +3.3 mm from bregma; mediolateral ± 0.82 mm from bregma; and dorsoventral -2.85 mm from pial surface. Injected mice were housed individually until used for electrophysiological experiments. Injections of the viral vector into the RMS specifically labeled adult-born migrating neuroblasts, with OB slices showing GFP expression in newborn interneurons only. We did not detect any diffusion from the injection site resulting in transduction of the resident OB cell population during the course of these experiments. This approach has been successfully used previously to characterize the morphological maturation of adult-born periglomerular cells .
Mice were deeply anesthetized with isoflurane (Mundipharma, Issy-les-Moulineaux, France) and then rapidly decapitated. Horizontal slices from the OB and frontal cortices were obtained after decapitation and brain removal. After cutting, slices (300 μm) were placed in oxygenated artificial cerebrospinal fluid (ACSF) at 35°C for 30 min. Slices were then kept in bubbled ACSF at room temperature. ACSF contained 124 mmol/l NaCl, 3 mmol/l KCl, 1.3 mmol/l MgSO4, 26 mmol/l NaHCO3, 1.25 mmol/l NaHPO4, 20 mmol/l glucose and 2 mmol/l CaCl (all chemicals from Sigma-Aldrich, Saint-Quentin, France).
Whole-cell patch-clamp recordings
Individual slices were placed in a submerged recording chamber and were continuously perfused with ACSF (1.5 ml/min) at room temperature. Whole-cell voltage-clamp recordings from GCs in the GC layer were obtained using a ×40 water-immersion objective, a halogen light source, differential interference contrast filters (all Olympus, Rungis, France), a charge-coupled device (CCD) camera (C7500; Hamamatsu, Japan) and an amplifier (EPC9/2; Heka Instrument, Port Washington, USA). Patch electrodes (6-10 MΩ) were filled with an internal solution (126 mmol/l Cs gluconate, 6 mmol/l CsCl, 2 mmol/l NaCl, 10 mmol/l Na-HEPES, 10 mmol/l D-glucose, 0.2 mmol/l Cs-EGTA, 0.3 mmol/l GTP, 2 mmol/l Mg-ATP, 0.2 mmol/l cAMP and 0.15% biocytin at pH 7.3 and 290-300 mOsm). Liquid junction potentials (9 mV) were compensated. Maximum Na+ current amplitude, measured by increasing the magnitude of depolarizing pulse by 10 mV increments, was mainly used to classify newborn GCs into three classes. Individual cell class was established from recording GFP-positive cells within the following range of days after virus injection: class 3 a 3 to 6 dpi; class 4 at 4 to 10 dpi and class 5 at 6 to 44 dpi (see Additional file 1D).
The stimulating electrode was placed near a recorded GC (Figure 2A) . Response amplitude was adjusted as required for a minimal stimulation protocol to stimulate only one axon that was directly presynaptic to the recorded GC. Stimulus intensity was gradually increased from a low level until an EPSC suddenly appeared in an all or nothing manner (see Additional file 2). Single axons were stimulated at 0.1 Hz, after checking that the mean EPSC amplitude for successful responses was not affected by small changes in stimulus intensity. Recordings were filtered at 10 kHz (Filter 1) and 2.9 kHz (Filter 2), digitized, and sampled at intervals of 20 to 450 μs (2.2 - 50 kHz) according to the requirements of individual protocols . Series resistance (< 30 MΩ) was monitored for stability throughout the recordings and if it had a change of >20%, the data were discarded. All experiments were performed in the presence of SR-95531 (gabazine) to block GABAA receptor-mediated currents.
Where i is the current carried by a single open channel, and N is the number of open channels. NMDAR-mediated EPSCs showed a skewed variance versus mean relationship. The single-channel conductance for NMDAR-mediated currents was estimated by fitting the initial slope of the relationship (Figure 3C). For AMPAR-mediated EPSCs, the variance versus mean relationship was parabolic. Single-channel conductance for AMPAR-mediated currents was therefore estimated using i, and the difference between holding potential and reversal potential (Figure 4C).
One or two slices were prepared from each virus-injected mouse and individual data were obtained from individual slices. Results are reported as mean ± SEM. A paired t-test was used for statistical analysis (Figure 6C; Figure 7C; Additional file 4). Kruskal-Wallis test followed by the Dunn multiple comparison test were also used to evaluate the data.
Tissue slices containing biocytin-loaded cells were fixed in 4% paraformaldehyde at 4°C overnight. Slices were then washed three times in 0.1 mol/l phosphate buffer pH 7.4, without resectioning, and incubated with PBS containing Alexa 546 conjugated-Streptavidin (Molecular Probes, http://www.probes.invitrogen.com) and 0.25% Triton-X for 2 hours at room temperature. After washing three times in phosphate buffer, slices were observed under a confocal microscope (TCS SP5; Leica Wetzlar, Germany).
Pre-embedding EM immunocytochemistry
Mice (7 dpi and 21 dpi) were anesthetized with pentobarbital and perfused with 2% PFA and 0.1% glutaraldehyde in sodium acetate buffer pH 6 for 2 minutes followed by 1 hour perfusion with 2% PFA and 0.1% glutaraldehyde in 0.1 mol/l borate buffer pH 9. Brains were post-fixed for 4 h with OBs cut into 70 μm coronal sections on a vibrating blade microtome (VT1200; Leica). The sections were cryoprotected with 30% sucrose and freeze-thawed three times to enhance antibody penetration. Sections were then processed for immunoperoxidase using primary antibodies against GFP (1:20,000; Chemicon International, http://www.chemicon.com). The peroxidase reaction product was silver-intensified and gold-toned as described previously . Serial thin sections were collected on copper slot grids and examined under a transmission electron microscope (JEM-1010; Jeol, Tokyo Japan) equipped with a side-mounted CCD camera (Mega View III, Olympus Soft Imaging Systems, Brandenburg Germany) at a magnification of 30,000. Synaptic contacts were analyzed in images taken from at least five consecutive sections. Glutamatergic (type 1) synapses were recognized by the presence of vesicles in the presynaptic terminal and by a prominent postsynaptic density (asymmetric junctions). Three-dimensional reconstructions were generated with the software Reconstruct (J.C. Fiala, Biology Department, Boston University, Boston, USA), using digital images acquired from each serial section.
This work was supported by the Fondation pour la Recherche Médicale "Equipe FRM", the Groupe "Novalis-Taitbout" and the Ecole des Neurosciences de Paris (ENP), Compagnia di San Paolo and Regione Piemonte (Ricerca Sanitaria Finalizzata 2006 and 2008). HK was supported by the Association Pasteur-Japon fellowship. MP is the recipient of a doctoral fellowship from 'Università Italo-Francese' (Progetto Vinci) and Servier. The laboratory is also supported by the Agence Nationale de la Recherche "ANR-09-NEUR-004" in the frame of "ERA-NET NEURON" of FP7 program by the European Commission. Dr. Lledo's visit to the Department of Molecular and Cellular Biology, Harvard University (Murthy's laboratory), was funded by the Philippe Foundation.
- Abrous DN, Koehl M, Le Moal M: Adult neurogenesis: from precursors to network and physiology. Physiol Rev. 2005, 85: 523-569. 10.1152/physrev.00055.2003.View ArticlePubMedGoogle Scholar
- Ming GL, Song H: Adult neurogenesis in the mammalian central nervous system. Annu Rev Neurosci. 2005, 28: 223-250. 10.1146/annurev.neuro.28.051804.101459.View ArticlePubMedGoogle Scholar
- Duan X, Kang E, Liu CY, Ming GL, Song H: Development of neural stem cell in the adult brain. Curr Opi Neurobiol. 2008, 18: 108-115. 10.1016/j.conb.2008.04.001.View ArticleGoogle Scholar
- Lledo PM, Merkle FT, Alvarez-Buylla A: Origin and function of olfactory bulb interneuron diversity. Trends Neurosci. 2008, 31: 392-400. 10.1016/j.tins.2008.05.006.PubMed CentralView ArticlePubMedGoogle Scholar
- Luskin MB: Restricted proliferation and migration of postnatally generated neurons derived from the forebrain subventricular zone. Neuron. 1993, 11: 173-189. 10.1016/0896-6273(93)90281-U.View ArticlePubMedGoogle Scholar
- Lois C, Alvarez-Buylla A: Long-distance neuronal migration in the adult mammalian brain. Science. 1994, 264: 1145-1148. 10.1126/science.8178174.View ArticlePubMedGoogle Scholar
- Brill MS, Ninkovic J, Winpenny E, Hodge RD, Ozen I, Yang R, Lepier A, Gascón S, Erdelyi F, Szabo G, Parras C, Guillemot F, Frotscher M, Berninger B, Hevner RF, Raineteau O, Götz M: Adult generation of glutamatergic olfactory bulb interneurons. Nat Neurosci. 2009, 12: 1524-1533. 10.1038/nn.2416.PubMed CentralView ArticlePubMedGoogle Scholar
- Carleton A, Petreanu LT, Lansford R, Alvarez-Buylla A, Lledo PM: Becoming a new neuron in the adult olfactory bulb. Nat Neurosci. 2003, 6: 507-518.PubMedGoogle Scholar
- Belluzzi O, Benedusi M, Ackman J, LoTurco JJ: Electrophysiological differentiation of new neurons in the olfactory bulb. J Neurosci. 2003, 23: 10411-10418.PubMedGoogle Scholar
- Mizrahi A: Dendritic development and plasticity of adult-born neurons in the mouse olfactory bulb. Nat Neurosci. 2007, 10: 444-452.PubMedGoogle Scholar
- Whitman MC, Greer CA: Synaptic integration of adult-generated olfactory bulb granule cells: basal axodendritic centrifugal input precedes apical dendrodendritic local circuits. J Neurosci. 2007, 27: 9951-9961. 10.1523/JNEUROSCI.1633-07.2007.View ArticlePubMedGoogle Scholar
- Grubb MS, Nissant A, Murry K, Lledo PM: Functional maturation of the first synapse in olfaction: development and adult neurogenesis. J Neurosci. 2008, 28: 2919-2932. 10.1523/JNEUROSCI.5550-07.2008.View ArticlePubMedGoogle Scholar
- Kelsch W, Lin CW, Lois C: Sequential development of synapses in dendritic domains during adult neurogenesis. Proc Natl Acad Sci. 2008, 105: 16803-16808. 10.1073/pnas.0807970105.PubMed CentralView ArticlePubMedGoogle Scholar
- Chen WR, Greer CA: Olfactory bulb. The synaptic Organization of the Brain. Edited by: Shepherd GM. 2004, New York: Oxford University Press, 165-216. 5Google Scholar
- Lledo PM, Alonso M, Grubb MS: Adult neurogenesis and functional plasticity in neuronal circuits. Nat Rev Neurosci. 2006, 7: 179-193. 10.1038/nrn1867.View ArticlePubMedGoogle Scholar
- Laaris N, Puche A, Ennis M: Complementary postsynaptic activity patterns elicited in olfactory bulb by stimulation of mitral/tufted and centrifugal fiber inputs to granule cells. J Neurophysiol. 2007, 97: 296-306. 10.1152/jn.00823.2006.PubMed CentralView ArticlePubMedGoogle Scholar
- Balu R, Pressler RT, Strowbridge BW: Multiple modes of synaptic excitation of olfactory bulb granule cells. J Neurosci. 2007, 27: 5621-5632. 10.1523/JNEUROSCI.4630-06.2007.View ArticlePubMedGoogle Scholar
- Rall W, Shepherd GM, Reese TS, Brightman MW: Dendrodendritic synaptic pathway for inhibition in the olfactory bulb. Exp Neurol. 1966, 14: 44-56. 10.1016/0014-4886(66)90023-9.View ArticlePubMedGoogle Scholar
- Price JL, Powell TP: The synaptology of the granule cells of the olfactory bulb. J Cell Sci. 1970, 7: 125-155.PubMedGoogle Scholar
- Price JL, Powell TP: An electron-microscopic study of the termination of the afferent fibres to the olfactory bulb from the cerebral hemisphere. J Cell Sci. 1970, 7: 157-187.PubMedGoogle Scholar
- Orona E, Rainer EC, Scott JW: Dendritic and axonal organization of mitral and tufted cells in the rat olfactory bulb. J Comp Neurol. 1984, 226: 346-356. 10.1002/cne.902260305.View ArticlePubMedGoogle Scholar
- de Olmos J, Hardy H, Heimer L: The afferent connections of the main and the accessory olfactory bulb formations in the rat: an experimental HRP-study. J Comp Neurol. 1978, 181: 213-244. 10.1002/cne.901810202.View ArticlePubMedGoogle Scholar
- Haberly LB, Price JL: Association and commissural fiber systems of the olfactory cortex of the rat. J Comp Neurol. 1978, 178: 711-740. 10.1002/cne.901780408.View ArticlePubMedGoogle Scholar
- Shipley MT, Adamek GD: The connections of the mouse olfactory bulb: a study using orthograde and retrograde transport of wheat germ agglutinin conjugated to horseradish peroxidase. Brain Res Bull. 1984, 12: 669-688. 10.1016/0361-9230(84)90148-5.View ArticlePubMedGoogle Scholar
- Gao Y, Strowbridge BW: Long-term plasticity of excitatory inputs to granule cells in the rat olfactory bulb. Nat Neurosci. 2009, 12: 731-733. 10.1038/nn.2319.PubMed CentralView ArticlePubMedGoogle Scholar
- Nissant A, Bardy C, Katagiri H, Murray K, Lledo PM: Adult neurogenesis promotes synaptic plasticity in the olfactory bulb. Nat Neurosci. 2009, 12: 728-730. 10.1038/nn.2298.View ArticlePubMedGoogle Scholar
- Hinds JW, Hinds PL: Synapse formation in the mouse olfactory bulb. I. Quantitative studies. J Comp Neurol. 1976, 69: 15-40. 10.1002/cne.901690103.View ArticleGoogle Scholar
- Hinds JW, Hinds PL: Synapse formation in the mouse olfactory bulb. II. Morphogenesis. J comp Neurol. 1976, 69: 41-61. 10.1002/cne.901690104.View ArticleGoogle Scholar
- Petreanu L, Alvarez-Buylla A: Maturation and death of adult-born olfactory bulb granule neurons: role of olfaction. J Neurosci. 2002, 22: 6106-6113.PubMedGoogle Scholar
- Panzanelli P, Bardy C, Nissant A, Pallotto M, Sassoè-Pognetto M, Lledo PM, Fritschy JM: Early synapse formation in developing interneurons of the adult olfactory bulb. J Neurosci. 2009, 29: 15039-15052. 10.1523/JNEUROSCI.3034-09.2009.View ArticlePubMedGoogle Scholar
- Takahashi T: Postsynaptic receptor mechanisms underlying developmental speeding of synaptic transmission. Neurosci Res. 2005, 53: 229-240. 10.1016/j.neures.2005.07.005.View ArticlePubMedGoogle Scholar
- Feldman DE, Knudsen EI: Experience-dependent plasticity and the maturation of glutamatergic synapses. Neuron. 1998, 20: 1067-1071. 10.1016/S0896-6273(00)80488-2.View ArticlePubMedGoogle Scholar
- Yashiro K, Philpot BD: Regulation of NMDA receptor subunit expression and its implication for LTD, LTP and metaplasticity. Neuropharmacology. 2008, 55: 1081-1094. 10.1016/j.neuropharm.2008.07.046.PubMed CentralView ArticlePubMedGoogle Scholar
- Barrera NP, Edwardson JM: The subunit arrangement and assembly of ionotropic receptors. Trends Neurosci. 2008, 31: 569-576. 10.1016/j.tins.2008.08.001.View ArticlePubMedGoogle Scholar
- Traynelis SF, Silver RA, Cull-Candy SG: Estimated conductance of glutamate receptor channels activated during EPSCs at the cerebellar mossy fiber-granule cell synapses. Neuron. 1993, 11: 279-289. 10.1016/0896-6273(93)90184-S.View ArticlePubMedGoogle Scholar
- Katagiri H, Fagiolini M, Hensch TK: Optimization of somatic inhibition at critical period onset in mouse visual cortex. Neuron. 2007, 53: 805-812. 10.1016/j.neuron.2007.02.026.View ArticlePubMedGoogle Scholar
- Zucker RS, Regehr WG: Short-term synaptic plasticity. Annu Rev Physiol. 2002, 64: 355-405. 10.1146/annurev.physiol.64.092501.114547.View ArticlePubMedGoogle Scholar
- Stevens CF, Wang Y: Facilitation and depression at single central synapses. Neuron. 1995, 14: 795-802. 10.1016/0896-6273(95)90223-6.View ArticlePubMedGoogle Scholar
- Isaac JTR, Lüthi A, Palmer MJ, Anderson WW, Benke TA, Collingridge GL: An investigation of the expression mechanism of LTP of AMPA receptor-mediated synaptic transmission at hippocampal CA1 synapses using failures analysis and dendritic recordings. Neuropharmacology. 1998, 37: 1399-1410. 10.1016/S0028-3908(98)00140-3.View ArticlePubMedGoogle Scholar
- Palmer MJ, Isaac JTR, Collingridge GL: Multiple developmentally regulated expression mechanism of long-term potentiation at CA1 synapses. J Neurosci. 2004, 24: 4903-4911. 10.1523/JNEUROSCI.0170-04.2004.View ArticlePubMedGoogle Scholar
- Davis BJ, Macrides F: The organization of centrifugal projections from the anterior olfactory nucleus, ventral hippocampal rudiment, and piriform cortex to the main olfactory bulb in the hamster: an autoradiographic study. J Comp Neurol. 1981, 203: 475-493. 10.1002/cne.902030310.View ArticlePubMedGoogle Scholar
- Luskin MB, Price JL: The topographic organization of associational fibers of the olfactory bulb system in the rat, including centrifugal fibers to the olfactory bulb. J Comp Neurol. 1983, 216: 264-291. 10.1002/cne.902160305.View ArticlePubMedGoogle Scholar
- Yamaguchi M, Mori K: Critical period for sensory experience-dependent survival of newly generated granule cells in the adult mouse olfactory bulb. Proc Natl Acad Sci USA. 2005, 102: 9697-9702. 10.1073/pnas.0406082102.PubMed CentralView ArticlePubMedGoogle Scholar
- Lin CW, Sim S, Ainsworth A, Okada M, Kelsch W, Lois C: Genetically increased cell-intrinsic excitability enhances neuronal integration into adult brain circuits. Neuron. 2010, 65: 32-39. 10.1016/j.neuron.2009.12.001.PubMed CentralView ArticlePubMedGoogle Scholar
- Platel JC, Dave KA, Bordey A: Control of neuroblast production and migration by converging GABA and glutamate signals in the postnatal forebrain. J Physiol. 2008, 586: 3739-3743. 10.1113/jphysiol.2008.155325.PubMed CentralView ArticlePubMedGoogle Scholar
- Darcy DP, Isaacson JS: Calcium-permeable AMPA receptors mediate glutamatergic signaling in neural precursor cells of postnatal olfactory bulb. J Neurophysiol. 2010, 103: 1431-1437. 10.1152/jn.00821.2009.View ArticlePubMedGoogle Scholar
- Koh DS, Geiger JR, Jonas P, Sakmann B: Ca(2+)-permeable AMPA and NMDA receptors in basket cells of rat hippocampal dentate gyrus. J Physiol. 1995, 485: 383-402.PubMed CentralView ArticlePubMedGoogle Scholar
- Komuro H, Kumada T: Ca2+ transients control CNS neuronal migration. Cell Calcium. 2005, 37: 387-393. 10.1016/j.ceca.2005.01.006.View ArticlePubMedGoogle Scholar
- Métin C, Denizot JP, Ropert N: Intermediate zone cells express calcium-permeable AMPA receptors and establish close contact with growing axons. J Neurosci. 2000, 20: 696-708.PubMedGoogle Scholar
- Li Y, Mu Y, Gage FH: Development of neural circuits in the adult hippocampus. Curr Top Dev Biol. 2009, 87: 149-174. full_text.View ArticlePubMedGoogle Scholar
- Imayoshi I, Sakamoto M, Ohtsuka T, Takao K, Miyakawa T, Yamaguchi M, Mori K, Ikeda T, Itohara S, Kageyama R: Roles of continuous neurogenesis in the structural and functional integrity of the adult forebrain. Nat Neurosci. 2008, 11: 1153-1161. 10.1038/nn.2185.View ArticlePubMedGoogle Scholar
- Toni N, Teng EM, Bushong EA, Aimone JB, Zhao C, Consiglio A, van Praag H, Martone ME, Ellisman MH, Gage FH: Synapse formation on neurons born in the adult hippocampus. Nat Neurosci. 2007, 10: 727-734. 10.1038/nn1908.View ArticlePubMedGoogle Scholar
- McAllister AK: Dynamic aspects of CNS synapse formation. Annu Rev Neurosci. 2007, 30: 425-450. 10.1146/annurev.neuro.29.051605.112830.PubMed CentralView ArticlePubMedGoogle Scholar
- Adesnik H, Li G, During MJ, Pleasure SJ, Nicoll RA: NMDA receptors inhibit synapse unsilencing during brain development. Proc Natl Acad Sci USA. 2008, 105: 5597-5602. 10.1073/pnas.0800946105.PubMed CentralView ArticlePubMedGoogle Scholar
- Flint AC, Maisch US, Weishaupt JH, Kriegstein AR, Monyer H: NR2A subunit expression shortens NMDA receptor synaptic currents in developing neocortex. J Neurosci. 1997, 17: 2469-2476.PubMedGoogle Scholar
- Cull-Candy SG, Leszkiewicz DN: Role of distinct NMDA receptor subtypes at central synapses. Sci STKE. 2004, 2004: re16-10.1126/stke.2552004re16.PubMedGoogle Scholar
- Piña-Crespo JC, Gibb AJ: Subtypes of NMDA receptors in new-born rat hippocampal granule cells. J Physiol. 2002, 541: 4-64.View ArticleGoogle Scholar
- Vicini S, Wang JF, Li JH, Zhu WJ, Wang YH, Luo JH, Wolfe BB, Grayson DR: Functional and pharmacological differences between recombinant N-methyl-D-aspartate receptors. J Neurophysiol. 1998, 79: 555-566.PubMedGoogle Scholar
- Momiyama A: Distinct synaptic and extrasynaptic NMDA receptors identified in dorsal form neurones of adult rat spinal cord. J Physiol. 2000, 523: 621-628. 10.1111/j.1469-7793.2000.t01-1-00621.x.PubMed CentralView ArticlePubMedGoogle Scholar
- Cline HT, Wu GY, Malinow R: In vivo development of neuronal structure and function. Cold Spring Harbor Symp Quant Biol. 1996, 61: 95-104.View ArticlePubMedGoogle Scholar
- Kerchner GA, Nicoll RA: Silent synapses and the emergence of a postsynaptic mechanism for LTP. Nat Rev Neurosci. 2008, 9: 813-825. 10.1038/nrn2501.PubMed CentralView ArticlePubMedGoogle Scholar
- Wadiche JI, Jahr CE: Multivesicular release at climbing fiber-Purkinje cell synapses. Neuron. 2001, 32: 301-313. 10.1016/S0896-6273(01)00488-3.View ArticlePubMedGoogle Scholar
- Oertner TG, Sabatini BL, Nimchinsky EA, Svoboda K: Facilitation at single synapses probed with optical quantal analysis. Nat Neurosci. 2002, 5: 657-664.PubMedGoogle Scholar
- Redman S: Quantal analysis of synaptic potentials in neurons of the central nervous system. Physiol Rev. 1990, 70: 165-198.PubMedGoogle Scholar
- Silver RA, Lubke J, Sakmann B, Feldmeyer D: High-probability uniquantal transmission at excitatory synapses in barrel cortex. Science. 2003, 302: 1981-1984. 10.1126/science.1087160.View ArticlePubMedGoogle Scholar
- Toni N, Buchs PA, Nikonenko I, Bron CR, Muller D: LTP promotes formation of multiple spine synapses between a single axon terminal and a dendrite. Nature. 1999, 402: 421-425. 10.1038/46574.View ArticlePubMedGoogle Scholar
- Xu-Friedman MA, Regehr WG: Structural contributions to short-term plasticity. Physiol Rev. 2004, 84: 69-85. 10.1152/physrev.00016.2003.View ArticlePubMedGoogle Scholar
- Zhao C, Teng EM, Summers RGJr, Ming GL, Gage FH: Distinct morphological stages of dentate granule neuron maturation in the adult mouse hippocampus. J Neurosci. 2006, 26: 3-11. 10.1523/JNEUROSCI.3648-05.2006.View ArticlePubMedGoogle Scholar
- Toni N, Laplagne DA, Zhao C, Lombardi G, Ribak CE, Gage FH, Schinder AF: Neurons born in the adult dentate gyrus form functional synapses with target cells. Nat Neurosci. 2008, 11: 901-907. 10.1038/nn.2156.PubMed CentralView ArticlePubMedGoogle Scholar
- Sassoè-Pognetto M, Wässle H, Grünert U: Glycinergic synapses in the rod pathway of the rat retina: cone bipolar cells express the alpha 1 subunit of the glycine receptor. J Neurosci. 1994, 14: 5131-5146.PubMedGoogle Scholar
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