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. 2017 Dec 6;119(3):1005–1018. doi: 10.1152/jn.00306.2017

Altered synaptic transmission and maturation of hippocampal CA1 neurons in a mouse model of human chr16p11.2 microdeletion

Hung-Chi Lu 1,2,3, Alea A Mills 4, Di Tian 1,2,3,✉
PMCID: PMC7864230  PMID: 29212915

Abstract

The pathophysiology of neurodevelopmental disorders is often observed early in infancy and toddlerhood. Mouse models of syndromic disorders have provided insight regarding mechanisms of action, but most studies have focused on characterization in juveniles and adults. Insight into developmental trajectories, particularly those related to circuit and synaptic function, will likely yield important information regarding disorder pathogenesis that leads to symptom progression. Chromosome 16p11.2 microdeletion is one of the most common copy number variations associated with a spectrum of neurodevelopmental disorders. Yet, how haploinsufficiency of chr16p11.2 affects early synaptic maturation and function is unknown. To address this knowledge gap, the present study focused on three key components of circuit formation and function, basal synaptic transmission, local circuit function, and maturation of glutamatergic synapses, in developing hippocampal CA1 neurons in a chr16p11.2 microdeletion mouse model. The data demonstrate increased excitability, imbalance in excitation and inhibition, and accelerated maturation of glutamatergic synapses in heterozygous deletion mutant CA1 neurons. Given the critical role of early synaptic development in shaping neuronal connectivity and circuitry formation, these newly identified synaptic abnormalities in chr16p11.2 microdeletion mice may contribute to altered developmental trajectory and function of the developing brain.

NEW & NOTEWORTHY The synaptic pathophysiology underlying neurodevelopmental disorders often emerges during infancy and toddlerhood. Therefore, identifying initial changes in synaptic function is crucial for gaining a mechanistic understanding of the pathophysiology, which ultimately will facilitate the design of early interventions. Here, we investigated synaptic and local circuit properties of hippocampal CA1 neurons in a human chr16p11.2 microdeletion mouse model during early postnatal development (preweaning). The data demonstrate increased neuronal excitability, excitatory/inhibitory imbalance, and accelerated maturation of glutamatergic synapses. These perturbations in early hippocampal circuit function may underlie the early pathogenesis of the heterozygous chr16p11.2 microdeletion, which is often associated with epilepsy and intellectual disability.

Keywords: chr16p11.2 microdeletion, excitation/inhibition, glutamatergic synapse, hippocampus CA1, silent synapse

INTRODUCTION

Chromosome copy number variations (CNVs) are frequently associated with neurodevelopmental disorders, such as autism spectrum disorder (ASD), schizophrenia, and intellectual disability (ID) (Girirajan et al. 2013; Levy et al. 2011; Malhotra and Sebat 2012; Merikangas et al. 2015; Pinto et al. 2014). Microdeletion at human chromosome 16p11.2 is one of the most common CNVs in ASD and accounts for ~0.5–1.0% of ASD cases (Kumar et al. 2008; Marshall et al. 2008; Sebat et al. 2004; Weiss et al. 2008). Human chr16p11.2 microdeletion is clinically heterogeneous, and the most common neurological presentations are language deficits, intellectual impairment, ASD, anxiety, attention deficit hyperactivity disorders (ADHD), and epilepsy (Hanson et al. 2010; McCarthy et al. 2009; Shinawi et al. 2010; Zufferey et al. 2012). Despite the broad clinical heterogeneity, the high prevalence of neurological deficits among individuals carrying the chr16p11.2 microdeletion indicates that genes in this region, individually or collectively, play critical roles in neurodevelopment and brain function.

Several studies have revealed multiple abnormalities at behavioral, biochemical, transcriptional, and electrophysiological levels in mouse models of human chr16p11.2 microdeletion (Blumenthal et al. 2014; Horev et al. 2011; Kusenda et al. 2015; Portmann et al. 2014; Pucilowska et al. 2015; Tian et al. 2015; Yang et al. 2015). Two investigations in zebrafish have also demonstrated the significance of several candidate genes in the pathogenesis of chr16p11.2 microdeletion (Blaker-Lee et al. 2012; Golzio et al. 2012). However, how the chr16p11.2 microdeletion affects early synapse development and function have not been adequately studied.

Intellectual impairment is reported in a high percentage of patients with chr16p11.2 microdeletion (Hanson et al. 2010; Shinawi et al. 2010; Zufferey et al. 2012). The affected individuals typically present with learning disabilities and reduced intelligence quotient scores (Hanson et al. 2010; Shinawi et al. 2010; Zufferey et al. 2012). Given the importance of the hippocampus in learning, memory, and cognition (Squire and Wixted 2011; Squire et al. 2015), and its frequent involvement in patients with neurodevelopmental disorders, we hypothesize hippocampal synaptic dysfunction plays a significant pathogenic role in intellectual impairment associated with chr16p11.2 microdeletion (Boronat et al. 2015; Bostrom et al. 2016; Boucher et al. 2012). Consistent with this hypothesis, a mouse model for human chr16p11.2 microdeletion exhibits deficits in a hippocampus-associated learning task (Tian et al. 2015). In the same mouse model, electrophysiological studies of the Schaffer collateral (SC)-CA1 synapses from P30–P35 mice demonstrated normal α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor (AMPAR)-mediated synaptic transmission and normal N-methyl-d-aspartate receptor (NMDAR)-mediated plasticity, but showed altered metabotropic glutamate receptor (mGluR)-mediated synaptic plasticity (Tian et al. 2015). These initial observations have provided important insights into the pathophysiology of the hippocampus in juvenile chr16p11.2 microdeletion mice, but the influence of the microdeletion on developmental maturation of hippocampal synaptic transmission and function is still unknown. Recent reports show that the rate of maturation of synapses at the hippocampal formation can be influenced by genes implicated in neurodevelopmental disorders (Clement et al. 2012, 2013; Peng et al. 2016; Qiu et al. 2014). Experiments that examine the effect of chr16p11.2 microdeletion on synapse development will provide insights into developmental and adult circuit dysfunction, with relevance to human pathophysiology.

In this study, we performed patch-clamp recording on postnatal days 13–15 (P13–P15) and days 20–22 (P20–P22) hippocampal CA1 neurons in a mouse model of the human chr1 6p11.2 microdeletion. We specifically characterized basal synaptic transmission, local connectivity, and synaptic maturation, which are key parameters influencing the formation and functions of the local circuitry of the hippocampus. The combined electrophysiological data demonstrate increased excitability, imbalance in excitation and inhibition, and accelerated maturation of glutamatergic synapses in chr16p11.2 microdeletion mice.

MATERIALS AND METHODS

Animals.

Mice carrying a deletion of the syntenic region of human chr16p11.2 (Horev et al. 2011) have been previously analyzed (Blumenthal et al. 2014; Horev et al. 2011; Pucilowska et al. 2015; Tian et al. 2015). Mice used in this study have been backcrossed onto the congenic C57BL6/N (Charles River) background for more than 20 generations. Wild-type (WT) and heterozygous mutant (Mut) mice were group housed and kept on a 12:12-h light-dark cycle with unrestricted access to food and water. All experimental procedures were approved by the Institutional Animal Care and Use Committee at the Children’s Hospital Los Angeles and conformed to National Institutes of Health guidelines. The experimenter was blind to the genotypes in all experiments and during data analysis.

Electrophysiology.

Whole cell patch-clamp recordings were performed on dorsal hippocampal CA1 neurons from P20–P22 male mice in all experiments. In addition, P13–P15 male mice were used in experiments designed to investigate AMPA-silent synapses. Mice were euthanized by rapid decapitation and the brains were immediately submerged in ice-cold high sucrose dissection buffer (HSDB) for 1 min. Four-hundred-micrometer coronal sections were cut on a Leica VT1000 vibratome in ice-cold HSDB, incubated in artificial cerebrospinal fluid (aCSF) at 30°C for 15 min and then transferred to aCSF at 25°C. Slices were further recovered at room temperature for 1 h before recording. HSDB was composed of the following (in mM): 87 NaCl, 75 sucrose, 2.5 KCl, 1.2 NaH2PO4, 30 NaHCO3, 25 glucose, 20 HEPES, 5 Na-ascorbate, 3 Na-pyruvate, 2 thiourea, 10 MgSO4, and 0.5 CaCl2. aCSF was composed of the following (in mM): 119 NaCl, 2.5 KCl, 1 MgCl2, 2 CaCl2, 26 NaHCO3, 1.23 NaH2PO4, and 10 glucose. All buffers used in dissection, recovery, and recording were supplemented with a mix of 95% O2-5% CO2 to maintain the pH at 7.4. All recordings were performed in aCSF at room temperature with a Multiclamp 700B microelectrode amplifier (Molecular Device, Sunnyvale, CA). Signals were low-pass filtered at a frequency of 1 kHz and digitized at 10 kHz using Digidata 1440A amplifier and Clampex9 software (Molecular Device). Series resistance was monitored continuously during recording, and experiments were discarded if the measurement changed by >10%. All internal solutions (resistance: 4–7 MΩ; pH: 7.2–7.4; and osmolarity: 290–300 mosM) used in this study contained 6.7 mM biocytin and 80 µM Alexa-Fluor 594 hydrazide with the addition of other components as described below for individual experiments. In experiments in which stimulations were applied, the stimulation intensities were recorded and reported in Table 1. No differences were observed in the stimulation intensities for wild-type and mutant neurons across all experiments except for the silent synapse measurement on neurons of P20–P22. We consistently placed the stimulating electrodes in the middle of striatum radiatum and ~300 μm from the recorded neurons, so the positioning effects on the evoked responses were minimized. All pulses were 200 µs in duration and delivered at 15-s interpulse intervals unless otherwise specified. All data were analyzed using Minianalysis (version 6.0.3. Synaptosoft) or Clampfit software (version 10, Molecular Device).

Table 1.

Stimulation intensities used in the experiments where stimuli were applied to the Schaffer collaterals

WT, μA
 Mut, μA
Experiment Figure Average SE Average SE P Value
E/I ratio 3 11.56 0.469438 11.47 0.416724 0.889095
PPR (single cell) 4 13.10 0.289499 12.97 0.350057 0.77129
PPR (field) 4 31.13 1.193297 29.58 1.117752 0.368854
A/N ratio 4 13.17 0.35746 13.27 0.337592 0.840304
TBS mEPSC 5 12.91 0.285614 12.96 0.297639 0.718755
NR2A/NR2B ratio 6 12.46 0.209884 12.64 0.205267 0.54467
Silent synapse (P13–15) 7 9.55 0.381584 9.31 0.314558 0.624436
Silent synapse (P20–22) 7 10.21 0.270954 9.02 0.326096 0.008701**
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WT, wild type; Mut, mutant; E/I, excitatory/inhibitory; PPR, paired pulse ratio; A/N, AMPA/NMDA; TBS, theta-burst stimulation; mEPSC, miniature excitatory postsynaptic current; P, postnatal day.

**

P < 0.01.

Current-clamp recording was performed to measure the resting membrane potential (RMP) (Graves et al. 2012). Specifically, after a gigaseal was obtained, the membrane was carefully broken to avoid a leaking current >50 pA. The membrane potentials were recorded for 10 min, and the measurements during the last 5 min were used to calculate the RMP. The electrodes were filled with the following (in mM): 131 K-gluconate, 20 KCl, 8 NaCl, 10 HEPES, 2 EGTA, 2 NaATP, and 0.3 NaGTP.

To measure membrane resistance (Rm) and membrane capacitance (Cm), neurons were recorded for 15 min, and the measurements during the last 5 min were analyzed. Neurons were discarded if the values of these two measurements fluctuated more than 20% from the average values. The electrodes were filled with the following (in mM): 131 K-gluconate, 20 KCl, 8 NaCl, 10 HEPES, 2 EGTA, 2 NaATP, and 0.3 NaGTP.

To determine the current-voltage (I–V) curve, membrane current at 11 voltage steps (−100, −80, −60, −40, −20, 0, 20, 40, 60, 80, and 100 mV) were measured. Specifically, neurons were held at each step for 400 ms. The peak current for the entire step and the average current for the last 100 ms were analyzed. The voltage step cycle was repeated twice for each neuron, and the current measurements for each step were averaged and used for the data analysis. The electrodes were filled with the following (in mM): 131 K-gluconate, 20 KCl, 8 NaCl, 10 HEPES, 2 EGTA, 2 NaATP, and 0.3 NaGTP.

Spontaneous and miniature excitatory postsynaptic currents (sEPSCs and mEPSCs) were recorded sequentially from the same neurons. Specifically, neurons were held at −70 mV and recorded for 15 min in the presence of 50 µM picrotoxin (PTX) and 1 µM strychnine (Stry) to isolate sEPSCs. Immediately afterwards, 200 nM tetrodotoxin (TTX) were added to aCSF and recording was continued for another 15 min to record mEPSCs. The recording electrodes were filled with the following (in mM): 131 K-gluconate, 20 KCl, 8 NaCl, 10 HEPES, 2 EGTA, 2 NaATP, and 0.3 NaGTP.

Spontaneous and miniature inhibitory postsynaptic currents (sIPSCs and mIPSCs) were also recorded sequentially from the same neurons. Specifically, neurons were held at −70 mV and recorded for 15 min in the presence of kynurenic acid to isolate sIPSCs. Then, 200 nM TTX were added to the aCSF and recording was continued for another 15 min to record mIPSCs. The recording electrodes were filled with the following (in mM): 145 KCl, 10 NaCl, 1 MgCl2, 10 HEPES, 2 EGTA, 2 NaATP, and 0.3 NaGTP.

To investigate the excitability of CA1 neurons, action potential frequency and current relationship (f-I) was determined (Sun et al. 2016). Specifically, 11 current steps [0, 10, 20, 30, 40, 50, 60, 70, 80, 90, and 100 pA; 2-s duration; 10 s interstimulus interval (ISI)] were applied to neurons, and the action potential frequency for each current step was calculated. The electrodes were filled with the following (in mM): 131 K-gluconate, 20 KCl, 8 NaCl, 10 HEPES, 2 EGTA, 2 NaATP and, 0.3 NaGTP.

Excitation/inhibition (E/I) ratios were determined by measuring excitatory and inhibitory input onto individual CA1 neurons as described previously (Qiu et al. 2014). The SCs were stimulated with a series of pulses to elicit either EPSCs or IPSCs. EPSCs were measured by evoking glutamate receptor-mediated current (derived from both AMPAR and NMDAR) at a holding potential of −40 mV [the reversal potential for γ-aminobutyric acid receptor (GABAR) and glycine receptor (GlyR)-mediated current]. Each neuron received 20–30 stimulations at a 15-s ISI. Neurons were subsequently depolarized to 0 mV in the presence of 50 μM kynurenic acid to block GluR-mediated current. The resultant IPSCs were mediated by monosynaptic inhibition through GABAR and GlyR. Again, each neuron received 20–30 stimulations at a 15-s ISI. The E/I ratio onto the same neuron was calculated by dividing the peak excitatory current (AMPAR plus NMDAR) by peak inhibitory (GABAR plus GlyR) current. The recording electrodes were filled with the following (in mM): 120 CsMeSO4, 15 CsCl, 10 TEA-Cl, 8 NaCl, 10 HEPES, 2 EGTA, 5 QX-314, 4 MgATP, 0.3 NaGTP, and 10 phosphocreatine.

AMPAR-mediated current was measured by evoking SCs at a holding potential of −70 mV in the presence of 50 µM PTX and 1 µM Stry to occlude inhibitory current. Each neuron received 20–30 stimulations. Then, 50 µM DNQX were bath applied to block AMPARs. Once the AMPAR current disappeared, NMDAR current was isolated by depolarizing neurons at +40 mV. Each neuron received 20–30 stimulations. The AMPA/NMDA (A/N) current ratio onto a single neuron was calculated by dividing the peak AMPAR current by peak NMDAR current. The recording electrodes were filled with the following (in mM): 120 CsMeSO4, 15 CsCl, 10 TEA-Cl, 8 NaCl, 10 HEPES, 2 EGTA, 5 QX-314, 4 MgATP, 0.3 NaGTP, and 10 phosphocreatine.

The paired pulse ratio (PPR) was determined by recording either single cell EPSCs or field EPSCs triggered by two consecutive pulses (stimulus 1 and 2) at varying interstimulus intervals (ISIs) of 25, 50, 75, 100, 200, 300, 500, and 1,000 ms. Single-cell EPSCs were measured by evoking SCs at a holding potential of −70 mV, and field EPSCs were measured at 0 mV. PPR was calculated by dividing EPSC slope in response to stimulus 2 by that to stimulus 1. The recording electrodes for single-cell EPSC recording were filled with the following (in mM) 120 CsMeSO4, 15 CsCl, 10 TEA-Cl, 8 NaCl, 10 HEPES, 2 EGTA, 5 QX-314, 4 MgATP, 0.3 NaGTP, and 10 phosphocreatine. The recording electrodes for field EPSC recording (0.5–1.0 MΩ) were filled with the following (in mM) 119 NaCl, 2.5 KCl, 1 MgSO4, 2 CaCl2, 26 NaHCO3, 1.23 NaH2PO4, and 10 glucose.

To determine the contribution of extrasynaptic AMPARs to mEPSCs, we used a long-term potentiation (LTP) protocol to induce lateral diffusion of AMPARs (especially GluR1) from extrasynaptic to the synaptic sites and recorded mEPSCs before and after LTP induction (Oh et al. 2006; Rouach et al. 2005). Specifically, neurons were held at −70 mV and baseline mEPSCs were recorded for 10 min in the presence of 50 µM PTX, 1 µM Stry, and 200 nM TTX. Then, PTX, Stry, and TTX were washed out with aCSF for 5 min. Afterwards, stimulations of increasing intensities were applied to SCs to trigger evoked EPSCs; the stimulation intensities were determined as those that induced stable responses. The theta-burst stimulations (TBSs) consisted of two trains of stimuli delivered 20 s apart. Each train was composed of five stimulus epochs 200 ms apart, with each epoch consisting of five pulses of 10–15 µA at 100 Hz. After TBS, PTX, Stry, and TTX were added back to aCSF and post-TBS mEPSCs were recorded for 10 min after the reapplication of PTX, Stry, and TTX. The last 5 min of the recording were analyzed. The electrodes (4–7 MΩ) were filled with (in mM) 131 K-gluconate, 20 KCl, 8 NaCl, 10 HEPES, 2 EGTA, 2 NaATP, and 0.3 NaGTP.

For NR2A and NR2B quantification, the NMDAR-mediated current was first measured by stimulating the SCs at a holding potential of +40 mV in the presence of DNQX. Then, 3 µM ifenprodil (IFN) were added to the bath for 3 min to block NR2B-mediated current. Twenty stimulations of SCs at 10-s ISIs were applied to measure both the NMDAR-mediated and NR2A-mediated current. The NR2B current was calculated by subtracting NR2A from NMDAR current (peak or charge transfer). The NR2A/NR2B ratio was calculated by dividing the peak NR2A current (or charge transfer) by the peak NR2B current (or charge transfer). The recording electrodes were filled with the following (in mM) 120 CsMeSO4, 15 CsCl, 10 TEA-Cl, 8 NaCl, 10 HEPES, 2 EGTA, 5 QX-314, 4 MgATP, 0.3 NaGTP, and 10 phosphocreatine.

To examine AMPA-silent synapses, a minimum stimulation protocol was used to activate a small number of axons to estimate the proportion of AMPA-silent synapses (Bateup et al. 2013; Qiu et al. 2014). First, the AMPAR current was measured at −70 mV in the presence of 50 µM PTX and 1 µM Stry. Stimulus intensities were adjusted to produce successful AMPAR current interleaved with failures, such that the percentages of successful responses were between 40 and 60%. Each neuron received 20 stimulations. Afterwards, neurons were voltage clamped at +40 mV and 20 more compound currents were recorded under the same stimulation intensity as in AMPAR current measurement. The percentages of NMDAR-only silent synapses were calculated by subtracting the failure rate of AMPAR current from that of the compound current. The recording electrodes were filled with the following (in mM): 120 CsMeSO4, 15 CsCl, 10 TEA-Cl, 8 NaCl, 10 HEPES, 2 EGTA, 5 QX-314, 4 MgATP, 0.3 NaGTP, and 10 phosphocreatine.

Statistical analysis.

Statistical analyses were performed using GraphPad software (version 6; La Jolla, CA). Two-tailed unpaired Student’s t-test was used for data analysis between wild-type and mutant mice. All data are presented as the average ± SE values.

RESULTS

Intrinsic membrane properties and current-voltage relationship.

We first investigated the intrinsic membrane properties of CA1 neurons. Mutant CA1 neurons showed a mild elevation in RMP (Fig. 1A; WT: −66.59 ± 0.30 mV, 15 neurons from 5 mice; Mut: −64.80 ± 0.22 mV, 15 neurons from 5 mice; ****P < 0.0001). However, membrane resistance (Fig. 1B; WT: 92.95 ± 15.22 MΩ, 15 neurons from 5 mice; Mut: 75.36 ± 14.24 MΩ, 15 neurons from 5 mice; P > 0.05), and membrane capacitance (Fig. 1C; WT: 43.42 ± 4.27 pF, 15 neurons from 5 mice; Mut: 49.80 ± 4.48 pF, 15 neurons from 5 mice; P > 0.05) did not differ significantly between wild-type and mutant CA1 neurons. These results indicated that the intrinsic membrane properties were largely unaffected in mutant CA1 neurons.

Fig. 1.

Fig. 1.

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Intrinsic membrane properties and current-voltage (I–V) relationship. A: resting membrane potential (RMP) was mildly increased in mutant CA1 neurons [n = wild type (WT): 15 neurons from 5 mice; Mut: 15 neurons from 5 mice]. B and C: membrane resistance (Rm) and membrane capacitance (Cm) were indistinguishable between wild-type and mutant CA1 neurons (n = WT: 15 neurons from 5 mice; Mut: 15 neurons from 5 mice). D: representative image of a recorded CA1 neuron (backfilled with Alexa-Fluor 594). E: representative traces of current-voltage relationship in wild-type and mutant CA1 neurons. Both peak (denoted by arrows) and steady-state responses (the last 100 ms denoted by horizontal bars) were subjected to analysis. F: mild increase in peak current response in mutant CA1 neurons (n = WT: 22 neurons from 7 mice; Mut: 18 neurons from 6 mice). G: steady-state responses were indistinguishable between wild-type and mutant CA1 neurons (n = WT: 22 neurons from 7 mice; Mut: 18 neurons from 6 mice). In all figures described in this study, data are presented as means ± SE. Numbers in parentheses represent those of neurons (1st number) and mice (2nd number). Statistical significance was determined by two-tailed, unpaired Student’s t-test for A–G. *P < 0.05, ****P < 0.0001.

Next, we applied a step protocol to determine neuronal current-voltage (I–V) relationship (Fig. 1, E–G). Both the peak current, representing the responses of voltage-gated ion channels and ionotropic receptors with fast kinetics, and the steady-state current, representing ligand-gated channels and metabotropic receptors with slow kinetics, were quantified (Fig. 1E). Mutant CA1 neurons showed slightly increased peak current at each voltage step (Fig. 1F, *P < 0.05), suggesting the presence of small changes in the quantities and/or types of fast-responding ion channels and/or receptors. In contrast, the steady-state current was indistinguishable between wild-type and mutant CA1 neurons (Fig. 1G), suggesting that the slow-response ligand-gated channels and metabotropic receptors were unaltered in mutant CA1 neurons.

Taken together, the above studies revealed a small reduction in RMP and mildly increased peak current of the I–V curve in mutant CA1 neurons. Although these changes were small in magnitude, they could contribute to the increased excitability of the mutant CA1 neurons (see below).

Synaptic transmission and spontaneous neuronal firing.

Next, we investigated the excitatory and inhibitory synaptic transmission by recording EPSCs and IPSCs, respectively. Mutant CA1 neurons exhibited significantly increased sEPSC frequency (WT: 1.18 ± 0.13 Hz, 24 neurons from 6 mice; Mut: 1.90 ± 0.23 Hz, 22 neurons from 6 mice; **P < 0.01) and mildly increased mEPSC frequency (Fig. 2, A and B; WT: 0.63 ± 0.06 Hz, 24 neurons from 6 mice; Mut: 0.98 ± 0.14 Hz, 22 neurons from 6 mice; *P < 0.05) without any changes in sEPSC amplitude (WT: 34.03 ± 1.35 pA, 24 neurons from 6 mice; Mut: 34.91 ± 1.75 pA, 22 neurons from 6 mice; P > 0.05) or mEPSC amplitude (Fig. 2, A and C; WT: 30.36 ± 0.81 pA, 24 neurons from 6 mice; Mut: 31.16 ± 1.24 pA, 22 neurons from 6 mice; P > 0.05), indicating elevated excitatory synaptic transmission. In contrast, no difference was observed in the frequency of sIPSCs or mIPSCs between wild-type and mutant CA1 neurons (Fig. 2, D and E; sIPSC frequency: WT: 3.92 ± 0.10 Hz, 27 neurons from 7 mice; Mut: 3.90 ± 0.09 Hz, 19 neurons from 5 mice; P > 0.05; mIPSC frequency: WT: 2.67 ± 0.07 Hz, 27 neurons from 7 mice; Mut: 2.60 ± 0.09 Hz, 19 neurons from 5 mice; P > 0.05). There was a mild increase in sIPSC amplitude (Fig. 2, D and F; WT: 65.60 ± 1.79 pA, 27 neurons from 7 mice; Mut: 72.01 ± 1.78 pA, 19 neurons from 5 mice; *P < 0.05), whereas mIPSC amplitude was unchanged in mutant CA1 neurons (Fig. 2, D and F; WT: 45.35 ± 1.05 pA, 27 neurons from 7 mice; Mut: 46.97 ± 0.95 pA, 19 neurons from 5 mice; P > 0.05).

Fig. 2.

Fig. 2.

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Significant increases in excitatory synaptic activities in mutant CA1 neurons. A: representative traces of spontaneous and miniature excitatory postsynaptic currents (sEPSCs and mEPSCs) in wild-type and mutant mice. B: both sEPSC and mEPSC frequencies were increased in mutant mice (n = WT: 24 neurons from 6 mice; Mut: 22 neurons from 6 mice). C: sEPSC and mEPSC amplitudes were not changed in mutant mice (n = WT: 24 neurons from 6 mice; Mut: 22 neurons from 6 mice). D: representative traces of spontaneous and miniature inhibitory postsynaptic currents (sIPSCs and mIPSCs) in wild-type and mutant mice. E: sIPSC and mIPSC frequencies were indistinguishable between wild-type and mutant mice (n = WT: 27 neurons from 7 mice; Mut: 19 neurons from 5 mice). F: sIPSC amplitude was mildly increased in mutant CA1 neurons. mIPSC amplitude was not changed (n = WT: 27 neurons from 7 mice; Mut: 19 neurons from 5 mice). G: representative traces of action potential from wild-type and mutant CA1 neurons injected with 10-, 50-, and 100-pA currents. H: action potential frequencies were significantly increased in mutant CA1 neurons at all current steps (n = WT: 15 neurons from 5 mice; Mut: 15 neurons from 5 mice). *P < 0.05, **P < 0.01.

We further determined excitability of CA1 neurons by studying the relationship between firing frequency and injected current. The firing-current (f-I) curve demonstrated that the frequencies of action potentials (Fig. 2, G and H) were significantly increased in mutant CA1 neurons at all current steps (0–100 pA; the values for 3 representative current steps shown in Fig. 2G: WT: 10 pA, 0.64 ± 0.23 Hz, 15 neurons from 5 mice; Mut: 1.67 ± 0.28 Hz, 15 neurons from 5 mice; **P < 0.01; 50 pA: WT: 7.19 ± 0.28 Hz, 15 neurons from 5 mice; Mut: 8.41 ± 0.32 Hz, 15 neurons from 5 mice; **P < 0.01; 100 pA: WT: 12.63 ± 0.28 Hz, 15 neurons from 5 mice; Mut: 13.83 ± 0.32 Hz, 15 neurons from 5 mice; **P < 0.01), indicating increased excitability of the mutant CA1 neurons.

Taken together, the observations of significant elevation in EPSC frequency and increased action potential frequencies in the presence of mild changes in IPSCs indicated overall increased activities at the excitatory synapses in mutant CA1 neurons. They also suggested an imbalance of excitation and inhibition. This prompted us to investigate whether E/I ratios were altered in mutant CA1 neurons.

Altered balance of excitation and inhibition in mutant CA1 neurons.

To determine the E/I ratio, we measured excitation and inhibition onto the same CA1 neurons using a minimum stimulation intensity protocol (Bateup et al. 2013; Qiu et al. 2014). Stimulation of the Schaffer collaterals (SCs) evoked EPSCs and IPSCs at holding potentials of −40 mV and 0 mV, respectively (Fig. 3A). There was a significant increase in evoked EPSCs (WT: −269.41 ± 13.92 pA, 17 neurons from 5 mice; Mut: −338.42 ± 19.70 pA, 17 neurons from 5 mice; **P < 0.01) and no change in evoked IPSCs (WT: 354.30 ± 17.19 pA, 17 neurons from 5 mice; Mut: 358.60 ± 12.34 pA, 17 neurons from 5 mice; P > 0.05; Fig. 3, B and C) in mutant CA1 neurons. In addition, E/I ratios, determined by dividing EPSCs by IPSCs from the same CA1 neurons, were significantly elevated in mutant CA1 neurons (WT: 77.15 ± 3.42%, 17 neurons from 5 mice; Mut: 93.92 ± 4.24%, 17 neurons from 5 mice; **P < 0.01; Fig. 3D). These results demonstrated significantly increased activities at the excitatory synapses, resulting in an imbalance between excitation and inhibition in hippocampal CA1 neurons of chr16p11.2df/+ mice.

Fig. 3.

Fig. 3.

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Imbalance of excitation and inhibition in mutant CA1 neurons. A: experimental design to measure the evoked excitatory and inhibitory postsynaptic currents (eEPSCs/eIPSCs). B: Representative traces of eEPSCs and eIPSCs in wild-type and mutant mice. C: eEPSCs were significantly increased in mutant mice, whereas eIPSCs were indistinguishable between wild-type and mutant mice (n = WT: 17 neurons from 5 mice; Mut: 17 neurons from 5 mice). D: the ratio of evoked excitatory (E) to inhibitory (I) postsynaptic current was increased in mutant mice (n = WT: 17 neurons from 5 mice; Mut: 17 neurons from 5 mice). aCSF, artificial cerebrospinal fluid; Vh, holding potential; KA, kynurenic acid. **P < 0.01.

Altered presynaptic neurotransmission and AMPAR-mediated current in mutant CA1 neurons.

The elevated excitation and E/I ratios in mutant CA1 neurons could come from two sources. First, presynaptic transmission could be increased. Second, the AMPAR- and/or NMDAR-mediated response could be altered.

We first evaluated presynaptic transmission by measuring PPRs at the SC-CA1 synapses using both single-cell patch-clamp and field recording (Fig. 4, A–D). In single-cell recording, mutant CA1 neurons showed reduced PPRs at all ISIs (ISI of 25 ms, WT: 119.35 ± 8.41%, 15 neurons from 5 mice; Mut: 86.93 ± 3.01%, 15 neurons from 5 mice; **P < 0.01. ISI of 50 ms: WT: 128.93 ± 9.18%, 15 neurons from 5 mice; Mut: 98.63 ± 4.16%, 15 neurons from 5 mice; **P < 0.01; ISI of 75 ms: WT: 130.56 ± 10.88%, 15 neurons from 5 mice; Mut: 100.11 ± 2.49%, 15 neurons from 5 mice; *P < 0.05; Fig. 4, A and B). Similar results were seen in field recording (ISI of 25 ms: WT: 184.50 ± 5.82%, 16 neurons from 5 mice; Mut: 160.90 ± 5.93%, 12 neurons from 4 mice; **P < 0.01; ISI of 50 ms: WT: 197.39 ± 5.57%, 16 neurons from 5 mice; Mut: 172.79 ± 6.00%, 12 neurons from 4 mice; **P < 0.01; ISI of 75 ms: WT: 187.07 ± 5.11%, 16 neurons from 5 mice; Mut: 169.09 ± 5.39%, 12 neurons from 4 mice; *P < 0.05; Fig. 4, C and D). These results indicated that presynaptic neurotransmitter release was increased at the SC-CA1 synapses.

Fig. 4.

Fig. 4.

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Increased presynaptic transmission onto and postsynaptic α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor (AMPAR)-mediated response from mutant CA1 neurons. A: representative traces of paired pulse responses determined by patch-clamp recording. B: paired pulse ratios at different interstimulus intervals (ISIs) were reduced in mutant CA1 mice (n = WT: 15 neurons from 5 mice; Mut: 15 neurons from 5 mice). C: representative traces of paired pulse responses determined by field recording. D: paired pulse ratios at different ISIs were reduced in mutant CA1 mice (n = WT: 16 neurons from 5 mice; Mut: 12 neurons from 4 mice). E: experimental design to measure AMPAR-mediated and N-methyl-d-aspartate receptor (NMDAR)-mediated current (IAMPA and INMDA). F: representative traces of IAMPA and INMDA in wild-type and mutant CA1 neurons. G: mutant CA1 neurons exhibited significantly increased AMPAR-mediated current and no change in NMDAR-mediated current (n = WT: 15 neurons from 5 mice; Mut: 15 neurons from 5 mice). H: mutant CA1 neurons showed a significant increase in A/N ratios (n = WT: 15 neurons from 5 mice; Mut: 15 neurons from 5 mice). *P < 0.05, **P < 0.01, ****P < 0.0001.

We then measured AMPAR- and NMDAR-mediated currents at holding potentials of −70 and +40 mV in the presence AMPAR inhibitors at +40 mV, respectively (Fig. 4E). Mutant CA1 neurons exhibited increased AMPAR-mediated current (Fig. 4F; WT: −340.96 ± 19.61 pA, 15 neurons from 5 mice; Mut: −451.13 ± 26.66 pA, 15 neurons from 5 mice; **P < 0.01) and no change in NMDAR-mediated current (Fig. 4G; WT: 491.74 ± 30.51 pA, 15 neurons from 5 mice; Mut: 486.33 ± 24.97 pA, 15 neurons from 5 mice; P > 0.05). In addition, the ratios of AMPAR-mediated and NMDA-mediated current were significantly increased in mutant CA1 neurons (Fig. 4H; WT: 70.23 ± 2.59%, 15 neurons from 5 mice; Mut: 93.24 ± 3.47%, 15 neurons from 5 mice; ****P < 0.0001). These results demonstrated that AMPAR-mediated response was increased, whereas NMDAR-mediated response was unchanged, at the SC-CA1 synapses from chr16p11.2df/+ mice.

The increase in evoked AMPAR-mediated response could result from an increase in presynaptic neurotransmitter release, upregulated surface expression of AMPAR at the synaptic site, and/or an elevated level of dendritic AMPAR at the extrasynaptic site (which can be activated under evoked condition). The reductions in PPRs in mutant CA1 neurons (Fig. 4, A–D) supported a presynaptic mechanism. The unaltered mEPSC amplitude in mutant CA1 neurons (Fig. 2C) suggested that the surface expression of AMPAR at the synaptic sites was unchanged.

To determine if mutant CA1 neurons contained a higher level of extrasynaptic AMPAR than the wild type, we applied TBS to induce lateral diffusion and measured mEPSCs before and after TBS (Fig. 5A). The rationale is that the degree of lateral diffusion is proportional to the size of the extrasynaptic AMPAR pool along the dendrites (Makino and Malinow 2009; Oh et al. 2006; Rouach et al. 2005). The amplitude of lateral diffusion can be calculated as the change in mEPSCs before and after TBS (∆mEPSCpost-pre). As shown in Fig. 5C, the pre-TBS mEPSC amplitudes were similar between wild-type and mutant CA1 neurons (WT: 28.13 ± 0.63 pA, 21 neurons from 7 mice; Mut: 29.69 ± 0.46 pA, 21 neurons from 7 mice; P > 0.05), consistent with the results described in Fig. 2. Application of TBS resulted in a significant difference in mEPSC amplitude between the two genotypes (Fig. 5C) (WT: 31.17 ± 0.51 pA, 21 neurons from 7 mice; Mut: 34.82 ± 0.72 pA, 21 neurons from 7 mice; ***P < 0.001). More importantly, ∆mEPSCpost-pre for mutant neurons was significantly higher than that for the wild-type (Fig. 5D) (WT: 3.04 ± 0.37 pA, 21 neurons from 7 mice; Mut: 5.45 ± 0.60 pA, 21 neurons from 7 mice; **P < 0.01), indicating that more AMPARs were available at the postsynaptic site as the result of TBS. In addition, mutant CA1 neurons showed higher mEPSC frequencies both before and after TBS (Fig. 5E) (Pre-TBS, WT: 0.65 ± 0.04 Hz, 21 neurons from 7 mice; Mut: 0.93 ± 0.05 Hz, 21 neurons from 7 mice; ***P < 0.001; Post-TBS, WT: 0.87 ± 0.04 Hz, 21 neurons from 7 mice; Mut: 1.19 ± 0.08 Hz, 21 neurons from 7 mice; **P < 0.01), indicating a higher rate of neurotransmister release in the mutant SC-CA1 synapses. To make sure that the differential effect of TBS on the amplitudes of ∆mEPSCpost-pre was not due to differences in stimulation intensity applied to wild-type and mutant neurons during TBS, we recorded the stimulation intensities and found no difference (Table 1).

Fig. 5.

Fig. 5.

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Increased extrasynaptic AMPAR in mutant CA1 neurons. A: experimental design to measure mEPSCs before and after theta-burst stimulation (TBS). B: representative traces of mEPSCs before and after TBS in wild-type and mutant CA1 neurons. C: mEPSC amplitude was indistinguishable between wild-type and mutant CA1 neurons before TBS. However, mutant CA1 neurons showed increased mEPSC amplitude compared with wild-type CA1 neurons after TBS (n = WT: 21 neurons from 7 mice; Mut: 21 neurons from 7 mice). D: ∆mEPSCpost-pre was higher in mutant CA1 neurons compared with wild-type CA1 neurons (n = WT: 21 neurons from 7 mice; Mut: 21 neurons from 7 mice). E: mEPSC frequency was elevated in mutant CA1 neurons before and after TBS compared with wild-type CA1 neurons (n = WT: 21 neurons from 7 mice; Mut: 21 neurons from 7 mice). F: schematic of AMPAR distribution at the synaptic and extrasynaptic sites in wild-type and mutant CA1 neurons before and after TBS. Stry, strychnine; TTX, tetrodotoxin. **P < 0.01, ***P < 0.001.

The results described in Fig. 5 were consistent with the hypothesis that mutant CA1 neurons contained a higher level of dendritic AMPAR at the extrasynaptic site (Fig. 5F), and this, along with an increase in presynaptic neurotransmitter release (Fig. 4, A–D), contributed to the elevation of evoked AMPAR-mediated responses in mutant CA1 neurons.

Altered development of the SC-CA1 synapses.

Development and maturation of the SC-CA1 synapses are characterized by increases in AMPAR expression (hence increased A/N ratio), switching of the NMDAR subunit from NR2B to NR2A (hence increased NR2A/NR2B current ratio), and reduction in AMPA-silent synapses (Clement et al. 2012; Hanse et al. 2013; Qiu et al. 2014; Rauner and Köhr 2011; Sans et al. 2000; Shipton and Paulsen 2013; Tsien 2000). Increased A/N ratios in mutant CA1 neurons (Fig. 4) suggest that maturation of the SC-CA1 synapses might be altered. To test this hypothesis, we measured NR2A, NR2B, and NR2A/NR2B ratios and quantified AMPA-silent synapses in wild-type and mutant CA1 neurons.

To measure NR2A and NR2B current, we first determined NMDAR-mediated current as aforementioned (Fig. 4) and calculated both peak current and charge transfer (Fig. 6, A and B). No differences were observed in either parameter between wild-type and mutant CA1 neurons (peak current: WT: 420.85 ± 23.63 pA, 24 neurons from 8 mice; Mut: 375.42 ± 15.48 pA, 24 neurons from 8 mice; P > 0.05; charge transfer: WT: 34.08 ± 1.64 pC, 24 neurons from 8 mice; Mut: 29.30 ± 1.89 pC, 24 neurons from 8 mice; P > 0.05; Fig. 6, C and D), consistent with data presented in Fig. 4. NR2A current, isolated by the application of the NR2B receptor blocker IFN, did not demonstrate differences in peak current or charge transfer between two genotypes (peak current: WT: 223.41 ± 10.32 pA, 24 neurons from 8 mice; Mut: 229.87 ± 8.70 pA, 24 neurons from 8 mice; P > 0.05; charge transfer: WT: 19.53 ± 0.81 pC, 24 neurons from 8 mice; Mut: 19.41 ± 1.28 pC, 24 neurons from 8 mice; P > 0.05; Fig. 6, B–D). However, NR2B currents, calculated by both peak current and charge transfer, were significantly reduced in mutant CA1 neurons (peak current: WT: 197.44 ± 14.88 pA, 24 neurons from 8 mice; Mut: 145.55 ± 10.03 pA, 24 neurons from 8 mice; **P < 0.01; charge transfer: WT: 14.55 ± 1.21 pC, 24 neurons from 8 mice; Mut: 9.89 ± 0.92 pC, 24 neurons from 8 mice; **P < 0.01; Fig. 6, B–D).

Fig. 6.

Fig. 6.

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Altered NR2A and NR2B current in mutant CA1 neurons. A: experimental design to directly measure NMDAR- and NR2A-mediated current (INMDA and INR2A). B: representative traces of INMDA, INR2A, and INR2B in wild-type and mutant CA1 neurons. Note: the dotted INR2B traces were derived from subtraction of INR2A from INMDA. C: peak amplitude of INR2B (2B) was decreased, whereas peak currents of INMDA (N) and INR2A (2A) were not altered, in mutant CA1 neurons (n = WT: 24 neurons from 8 mice; Mut: 24 neurons from 8 mice). D: charge transfer of INR2B (2B) was decreased, whereas charge transfers of INMDA (N) and INR2A (2A) were not altered, in mutant CA1 neurons (n = WT: 24 neurons from 8 mice; Mut: 24 neurons from 8 mice). E: the ratios of INR2A/INMDA (2A/NMDA) and INR2B/INMDA (2B/NMDA), measured by peak current, were significantly altered in mutant CA1 neurons (n = WT: 24 neurons from 8 mice; Mut: 24 neurons from 8 mice). F: the ratios of INR2A/INMDA (2A/NMDA) and INR2B/INMDA (2B/NMDA), measured by charge transfer, were significantly altered in mutant CA1 neurons (n = WT: 24 neurons from 8 mice; Mut: 24 neurons from 8 mice). G: the ratios of INR2A/INR2B (2A/2B) measured by both peak current and charge transfer, were increased in mutant CA1 neurons (n = WT: 24 neurons from 8 mice; Mut: 24 neurons from 8 mice). **P < 0.01, ***P < 0.001.

Furthermore, the percentages of NR2A and NR2B currents, measured by both peak current and charge transfer, were significantly changed in mutant CA1 neurons (NR2A peak current: WT: 53.91 ± 1.32%, 24 neurons from 8 mice; Mut: 61.73 ± 1.42%, 24 neurons from 8 mice; ***P < 0.001; NR2A charge transfer: WT: 58.26 ± 1.95%, 24 neurons from 8 mice; Mut: 66.67 ± 1.76%, 24 neurons from 8 mice; **P < 0.01; NR2B peak current: WT: 46.09 ± 1.32%, 24 neurons from 8 mice; Mut: 38.27 ± 1.42%, 24 neurons from 8 mice; ***P < 0.001; NR2B charge transfer: WT: 41.74 ± 1.95%, 24 neurons from 8 mice; Mut: 33.33 ± 1.76%, 24 neurons from 8 mice; **P < 0.01; Fig. 6, E and F). In addition, NR2A to NR2B ratios were increased in mutant CA1 neurons (peak current: WT: 121.85 ± 7.54%, 24 neurons from 8 mice; Mut: 170.40 ± 11.04%, 24 neurons from 8 mice; ***P < 0.001; charge transfer: WT: 156.92 ± 17.04%, 24 neurons from 8 mice; Mut: 216.25 ± 13.98%, 24 neurons from 8 mice; **P < 0.01; Fig. 6G). These results indicated that although the total NMDAR-mediated current was unchanged, the contribution by NR2A increased and by NR2B decreased in mutant CA1 neurons. These findings were consistent with the idea that NR2B to NR2A switch occurred earlier in the mutant SC-CA1 synapses.

AMPA-silent synapses are characterized by the presence of NMDAR and absence of surface AMPAR (Clement et al. 2012; Durand et al. 1996; Hanse et al. 2013; Isaac et al. 1997; Kerchner and Nicoll 2008; Liao et al. 1995; Tsien 2000). Under patch-clamp conditions, these synapses respond to stimulations at a holding potential of +40 mV but not at −70 mV. Using a modified stimulation protocol as described previously (Bateup et al. 2013; Qiu et al. 2014), we determined the percentages of silent synapses in both wild-type and mutant CA1 neurons (Fig. 7A) at two developmental time points, P13–P15 and P20–P22. Wild-type and mutant CA1 neurons from P13–P15 mice showed no differences in failure rates at −70 mV (WT: 52.33 ± 1.37%, 15 neurons from 5 mice; Mut: 49.17 ± 1.16%, 18 neurons from 6 mice; P > 0.05) or +40 mV (WT: 37.33 ± 1.45%, 15 neurons from 5 mice; Mut: 38.06 ± 0.92%, 18 neurons from 6 mice; P > 0.05; Fig. 7, B–D). However, mutant CA1 neurons demonstrated a significantly smaller change in failure rates than the wild-type neurons between these two holding potentials (Fig. 7H; WT: 15.00 ± 1.09%, 15 neurons from 5 mice; Mut: 11.11 ± 0.76%, 18 neurons from 6 mice; **P < 0.01), suggesting that the percentage of AMPA-silent synapses was lower in mutant CA1 neurons than in the wild-type neurons.

Fig. 7.

Fig. 7.

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Altered profiles of AMPA-silent synapses in postnatal days 13–15 (P13–P15) mutant CA1 neurons. A: experimental design to measure AMPA-silent synapses. B and C: representative traces and scattered plots of AMPAR-mediated (−70 mV) and compound (AMPAR- plus NMDA-mediated) current in wild-type (B) and mutant (C) CA1 neurons at P13–P15. D and E: representative traces and scattered plots of AMPAR-mediated (−70 mV) and compound (AMPAR- plus NMDA-mediated) current in wild-type (E) and mutant (D) CA1 neurons at P22-P22. F and G: failure rates of wild-type and mutant CA1 neurons at potentials of −70 and +40 mV at P13–P15 (F; n = WT: 15 neurons from 5 mice; Mut: 18 neurons from 6 mice) and P20–P22 (G; n = WT: 15 neurons from 5 mice; Mut: 12 neurons from 4 mice). (The same neurons were connected by dotted lines.). H: changes in failure rates in wild-type and mutant CA1 neurons from P13-15 and P20-22 mice. there were significant differences in changes of failure rate between wild-type and mutant CA1 neurons at P13–P15 but not P20–P22. Wild-type, not mutant, CA1 neurons showed reduction of changes of failure rate between P13–P15 (n = WT: 15 neurons from 5 mice; Mut: 18 neurons from 6 mice) and P20–P22 (n = WT: 15 neurons from 5 mice; Mut: 12 neurons from 4 mice). PTX, picrotoxin. **P < 0.01, ***P < 0.001.

Similarly, wild-type and mutant CA1 neurons from P20–P22 mice showed comparable failure rates at −70 mV (WT: 51.33 ± 1.03%, 15 neurons from 5 mice; Mut: 48.75 ± 1.09%, 12 neurons from 4 mice; P > 0.05) and +40 mV (WT: 41.00 ± 0.87%, 15 neurons from 5 mice; Mut: 39.58 ± 0.96%, 12 neurons from 4 mice; P > 0.05; Fig. 7, E–G). In contrast to P13–P15, the differences in failure rates were statistically indistinguishable between wild-type and mutant neurons (WT: 10.33 ± 0.91%, 15 neurons from 5 mice; Mut: 9.17 ± 1.04%, 12 neurons from 4 mice P > 0.05; Fig. 7H), suggesting that the percentage of AMPA-silent synapses was similar in wild-type and mutant CA1 neurons at P20–P22. It is worth mentioning that the stimulation intensities used to achieve 40–60% successful responses were significantly smaller for mutant CA1 neurons than for the wild-type (Table 1; WT: 10.21 ± 0.27 µA; Mut: 9.02 ± 0.33 uA, **P < 0.01). This observation was consistent with the aforementioned results that demonstrated increased presynaptic neurotransmitter release and elevated excitability of SC-CA1 synapses (Fig. 2, 3, and 4).

Interestingly, wild-type, but not mutant, CA1 neurons demonstrated a reduction in failure rate between P13–P15 and P20–P22 (WT, P13–P15: 15.00 ± 1.09%, 15 neurons from 5 mice; WT, P20–P22: 10.33 ± 0.91%, 15 neurons from 5 mice; **P < 0.01; Mut, P13–P15: 11.11 ± 0.76%, 18 neurons from 6 mice; Mut, P20–P22: 9.17 ± 1.04%, 12 neurons from 4 mice; P > 0.05). This result suggested that while synapses onto wild-type CA1 neurons underwent maturation between P13–P15 and P20–P22 with reduction of AMPA-silent synapses, those onto mutant CA1 neurons had already reached the developmental stage of P20–P22 at the age of P13–P15 with regard to the status of AMPA-silent synapses.

Taken together, the increased A/N ratio, relatively higher percentage of NR2A subunit current, and reduced percentage of AMPA-silent synapses at the mutant SC-CA1 synapses strongly suggested that the developmental trajectory of these synapses was altered in mutant mice.

DISCUSSION

Chromosome copy number variations are frequently associated with neurodevelopmental disorders (Girirajan et al. 2013; Levy et al. 2011; Malhotra and Sebat 2012; Pinto et al. 2014). The involved chromosomal regions usually contain multiple genes with diverse functions. Characterization of mouse models carrying syntenic deletions is a crucial step toward understanding the disease mechanism.

In this study, we investigated the synaptic physiology of developing hippocampal neurons in a mouse model of human chr16p11.2 microdeletion (Horev et al. 2011). Using acute slice preparations, we determined intrinsic membrane properties, basal synaptic transmission, and functions of two predominant types of glutamatergic receptors, AMPAR and NMDAR. The results revealed three major abnormalities in CA1 neurons from chr16p11.2df/+ mice.

First, mutant CA1 neurons show increased excitatory activities. Several observations from our study support this conclusion: 1) the frequencies of both sEPSCs and mEPSCs are elevated compared with wild-type neurons; 2) the action potential frequencies, determined by the f-I curve, are increased at all injected current steps; and 3) AMPAR-mediated current is augmented under stimulated condition. These changes in excitability most likely result from both presynaptic and postsynaptic mechanisms. While increased mEPSC frequency and reduced PPRs in mutant CA1 neurons support a presynaptic contribution, increased AMPAR-mediated current is consistent with a postsynaptic mechanism. Because CA3 is one of the major afferents onto CA1, it is plausible to speculate that increased excitatory activity may also be present in CA3 neurons, thus contributing to the observed abnormalities in sEPSCs, mEPSCs, and PPR in mutant CA1 neurons. Increased EPSCs and E/I ratio (see below) has been described in the striatum of an independent mouse model of human 16p11.2 microdeletion by Portmann et al. (2014). Similarly, our unpublished study on the principal neurons of the basolateral amygdala nucleus also revealed increased excitatory tone (unpublished observations). Together, these electrophysiological studies strongly suggest that chr16p11.2 microdeletion results in increased excitation in several brain regions critical for learning, memory, and social behavior.

Second, mutant CA1 neurons exhibit perturbed E/I balance. The supporting evidence is 1) under the basal condition, sEPSC and mEPSC frequencies in mutant CA1 neurons are significantly increased whereas sIPSC and mIPSC frequencies are unaffected; and 2) under a stimulation condition, the evoked EPSC amplitude is increased, whereas the evoked IPSC amplitude is unaltered. Perturbation of E/I balance is hypothesized as a synaptic mechanism for autism, and has been supported by multiple studies (Bateup et al. 2013; Dani et al. 2005; Rothwell et al. 2014; Rubenstein and Merzenich 2003). Recently, Mullins et al. (2016) expanded this hypothesis and proposed that dysfunctional autoregulation is a general mechanism of ASD. The authors conceptualized autoregulation at various levels, including biochemical, structural, synaptic, and transcriptional. Importantly, they redefined the concept of E/I balance as E/I coordination to reflect unknown circuitry that contributes to the temporal complexity of E/I. Although the mechanism has yet to be determined, the perturbed E/I relationship seen in the hippocampal CA1 neurons from chr16p11.2df/+ mice support the hypotheses by Rubenstein and Merzenich (2003) and Mullins et al. (Mullins et al. (2016).

Third, mutant mice show accelerated maturation at the Schaffer collateral (SC)-CA1 synapses. Specifically, mutant CA1 neurons demonstrate 1) increased A/N current ratio; 2) higher NR2A/NR2B subunit current ratio; and 3) a reduced percentage of AMPA-silent synapses. These results indicate that genes within the deleted region play critical roles in the maturation of glutamatergic synapses in the hippocampus. It has been shown that a gradual increase in the A/N current ratio, NR2B to NR2A switching, and reduction of AMPA-silent synapses are the molecular events that are critical milestones for developing hippocampal glutamatergic synapses (Clement et al. 2012; 2013; Qiu et al. 2014). Disruption in the timing of these key processes, as exemplified by SynGap1 and Met mutant mice, can have profound long-term consequences on cognition and behavior (Clement et al. 2012, 2013; Muhia et al. 2012; Qiu et al. 2014; Thompson and Levitt 2015). Indeed, adult chr16p11.2df/+ mice exhibit a deficit in hippocampus-associated learning in a contextual fear-conditioning test (Tian et al. 2015), further supporting the hypothesis that alteration in the kinetics of hippocampal synaptic development is a shared pathogenic mechanism among a subset of genetic abnormalities associated with neurodevelopment and neuropsychiatric disorders.

Interestingly, a previous study using the same mouse model demonstrated normal presynaptic neurotransmitter release and postsynaptic glutamatergic response at the SC-CA1 synapses from P30–P35 mice (Tian et al. 2015). In contrast, the current study investigated synaptic functions and revealed both presynaptic and postsynaptic changes in younger mice (P20–P22). It has been reported that genetic defects can impact synaptic functions within certain developmental windows, particularly during the early stage of synapse formation and establishment (Bozdagi et al. 2012; Clement et al. 2012; Hsiao et al. 2016). The findings from our studies of chr16p11.2 microdeletion at different ages in mice are likewise in keeping with these examples, and highlight the importance of studying and understanding the effect of mutations in a developmental context.

A limitation of the current study is that we have only investigated the synaptic functions in a particular neuronal population. Whether similar abnormalities are present in other synapses in different brain regions needs further investigation. Another limitation is that this study does not aim to distinguish the primary effect of the microdeletion on the synaptic function from the abnormal compensatory responses to the initial alterations. Investigation of mice carrying a temporally and spatially modifiable microdeletion will help to differentiate these two scenarios, thus deepening our understanding of the temporal regulation of developmental processes and neural circuit origins of synaptic abnormalities associated with chr16p11.2 microdeletion. In addition, the result in mice would have to be validated in humans to establish the contributions of the microdeletion to specific pathophysiology.

In summary, this study has uncovered multiple synaptic abnormalities affecting basal neurotransmission, local network activity, and synaptic development in a mouse model of the human chr16p11.2 microdeletion. The results indicate that genes within chr16p11.2 play important roles in regulating synaptic development and function in the hippocampus. In addition, the abnormalities seen in chr16p11.2 microdeletion mice are in keeping with those found in several mouse models of syndromic and nonsyndromic forms of ASD and ID (Bhakar et al. 2012; Ebrahimi-Fakhari and Sahin 2015; Santoro et al. 2012; Südhof 2008). In this context, our studies suggest that synaptic dysfunction is potentially one of the shared mechanisms between ASD and ID associated with chromosome copy number variation and single gene mutations.

GRANTS

This study was supported by a Research Career Development Award from The Saban Research Institute of Children’s Hospital Los Angeles (to D. Tian), and awards from the Simons Foundation Research Initiative (to A. A. Mills).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

D.T. conceived the research; HC.L. and D.T. designed experiments; HC.L. performed the experiments; HC.L. analyzed data; HC.L. and D.T. interpreted results of experiments; HC.L. and D.T. prepared figures; HC.L. and D.T. drafted manuscript; HC.L., A.A.M., and D.T. edited and revised manuscript; HC.L., A.A.M., and D.T. approved final version of manuscript.

ACKNOWLEDGMENTS

We thank Drs. Pat Levitt, Allison Knoll, Arnold Heynen, and Harvey Pollack for critically reading the manuscript.

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