Genetic abnormalities in synaptic proteins are common in individuals with autism; however, our understanding of the cellular and molecular mechanisms disrupted by these abnormalities is limited. SHANK3 is a postsynaptic scaffolding protein of excitatory synapses that has been found mutated or deleted in most patients with 22q13 deletion syndrome and about 2% of individuals with idiopathic autism and intellectual disability. Here, we generated CRISPR/Cas9-engineered human pluripotent stem cells (PSCs) with complete hemizygous SHANK3 deletion (SHANK3+/–), which is the most common genetic abnormality in patients, and investigated the synaptic and morphological properties of SHANK3-deficient PSC-derived cortical neurons engrafted in the mouse prefrontal cortex. We show that human PSC-derived neurons integrate into the mouse cortex by acquiring appropriate cortical layer identities and by receiving and sending anatomical projections from/to multiple different brain regions. We also demonstrate that SHANK3-deficient human neurons have reduced AMPA-, but not NMDA- or GABA-mediated synaptic transmission and exhibit impaired dendritic arbors and spines, as compared to isogenic control neurons co-engrafted in the same brain region. Together, this study reveals specific synaptic and morphological deficits caused by SHANK3 hemizygosity in human cortical neurons at different developmental stages under physiological conditions and validates the use of co-engrafted control and mutant human neurons as a new platform for studying connectivity deficits in genetic neurodevelopmental disorders associated with autism.
Accumulating evidence suggests that synaptic deficits may be responsible for the behavioral abnormalities in autism spectrum disorders (ASD) [1, 2]. Indeed, brain imaging studies have consistently identified altered functional connectivity in the brain of individuals with ASD , and well-powered sequencing studies have found multiple genetic abnormalities in various genes encoding synaptic proteins [4, 5]. However, it remains to be determined whether and how the disruption of specific synaptic genes influences the synaptic properties of human neurons under physiological conditions to alter brain connectivity.
SHANK3 is a postsynaptic scaffolding protein of excitatory synapses [6, 7]. It has been found mutated or deleted in about 0.5% of patients with idiopathic ASD, 2% of patients with idiopathic ASD and intellectual disability (ID) , and most patients with Phelan–McDermid syndrome (PMDS), a syndromic form of ASD caused by the microdeletion of chromosome 22q13.3 and is characterized by profound ID, developmental delay, severely impaired speech, and autistic behaviors [9, 10]. The most common genetic abnormality found in PMDS patients is a complete hemizygous SHANK3 deletion . However, no overt deficits have been detected in mice with complete heterozygous Shank3 deletion, which were generated to mimic patient-related genetic abnormality . Therefore, it remains unclear whether and how SHANK3 hemizygosity alters the synaptic properties of human neurons.
Synaptic deficits induced by partial homozygous Shank3 deletions or heterozygous mutations have been extensively investigated in animal models [12,13,14,15,16,17,18,19,20,21,22,23,24,25,26]. Although all studies reported deficits, there is some inconsistency between the specific synaptic and behavioral phenotypes observed in different studies, which is likely due to the loss/ectopic gain of particular isoforms of Shank3 and/or differences in the genetic backgrounds, cell types, and developmental stages examined in those studies . As a result, it remains unclear which of the reported deficits are relevant to patients, as human and rodent neurons have very different developmental trajectories as well as genetic and epigenetic profiles .
More recent studies on SHANK3-deficient human neurons obtained from patient-induced pluripotent stem cells (iPSCs) and CRISPR/Cas9-engineered embryonic stem cells (ESCs) reported more severe deficits in excitatory synaptic transmission and new deficits in intrinsic excitability and morphological development [29,30,31,32]. However, these studies were conducted on stem cell-derived neurons obtained from PMDS patients with large deletions or rare point mutations or on engineered neurons with partial SHANK3 deletions. In addition, stem cell-derived human neurons were investigated in vitro under non-physiological conditions, which may have influenced the severity and manifestation of the deficits. Therefore, it remains unclear whether and how the synaptic deficits detected in human neurons in vitro are related to those in patients.
In the present study, we developed a novel approach to transplant human stem cell-derived cortical neurons into the mouse prefrontal cortex (PFC) and used a state-of-the-art trans-synaptic tracing technique and patch-clamp electrophysiology to investigate their synaptic and morphological properties. The results show that human neurons survive in the mouse PFC for many months post transplantation, acquire cortical laminar identities, and establish anatomical connections with mouse neurons in multiple brain regions. To characterize the synaptic deficits induced by SHANK3 hemizygosity, we generated a CRISPR/Cas9-engineered stem cell line with complete hemizygous SHANK3 deletion and co-transplanted both isogenic control (iCtrl) and SHANK3+/– human stem cell-derived cortical neurons into the mouse PFC. We found that SHANK3-deficient human neurons have impaired AMPA-, but not NMDA- or GABA-mediated synaptic transmission and deficient dendrites and dendritic spines. This study identifies specific deficits induced by SHANK3 hemizygosity in human cortical neurons and validates the use of the co-xenotransplantation model to study synaptic and connectivity deficits in neurodevelopmental disorders associated with ASD.
Materials and methods
Stem cell culture and neural differentiation
The stem cell lines used in this study (H9, EYQ2-20 [SHANK3+/–], 2242–5, and EP215 [SHANK3–/–]) were cultured in E8 medium (Life Technologies, Cat#A1517001) on 6-well plates (VWR, Cat#29442–036) coated with 1% Matrigel (Corning, Cat#354277) in humidified incubator with 37 °C/5% CO2. Stem cells were passaged when reached 70–80% confluency using Dispase (1 U/mL, STEMCELL Technologies, Cat#07923). During passaging a ROCK Inhibitor, Y-27632 (2–4 µg/mL, Hello Bio, Cat#HB2297) was added to the medium to improve cell survival.
Stem cells were differentiated using a modified version of the dual-SMAD inhibition protocol . Briefly, stem cell colonies were re-plated on Matrigel-coated 6-well plates and allowed to reach approximately 95–100% confluency in E8 medium. On day 0, approximately half of the E8 medium was removed and neuroectodermal fate was induced by a 6–10-day-long incubation in neural differentiation medium (ND) containing of 1:1 mixture of N2 medium (DMEM/F-12 [Invitrogen, Cat#11330–032], 1% N2 Supplement [Thermo Fisher Scientific, Cat#17502048], 1% MEM–NEAA [Invitrogen, Cat#11140050], 2 mg/mL heparin [STEMCELL Technologies, Cat#07980], and 1% Pen/Strep) and B-27 medium (Neurobasal-A [Invitrogen, Cat#10888022], 2% B27 Supplement with vitamin A [Thermo Fisher Scientific, Cat#17504044], 1% GlutaMAX [Invitrogen, Cat#35050061], and 1% Pen/Strep) containing inhibitors of SMAD signaling, 10 µM SB431542 (Selleck Chemicals, Cat#S1067) and 4 µM dorsomorphin (Selleck Chemicals, Cat#S7306). On days 6–10, cells were dissociated into large clumps using dispase and transferred to Matrigel-coated plates in ND medium. The day after re-plating, half the medium was replaced with ND medium containing 20 ng/mL EGF (STEMCELL Technologies, Cat#780006.1) and 20 ng/mL FGF (STEMCELL Technologies, Cat#78003). On days 5–10, cells were cultured with EGF, but without FGF, to allow the formation of neural rosettes. On day 10, clusters of rosettes were dissociated using Dispase, transferred to wells precoated with Matrigel, and cultured in ND medium with EGF. On day 15, neural rosettes were dissociated to single cells with Accutase and propagated on Matrigel coated plates in ND medium without growth factors. On days 35–40, stem cell-derived neurons were transduced with different lentiviruses.
Generation of engineered stem cell lines
For generating stem cells with hemizygous and homozygous deletions, guide RNAs were designed using an online design tool (crispr.mit.edu) to target different regions of the SHANK3 genomic locus. Each individual sgRNA was synthesized as a DNA fragment and cloned into the plasmid pSpCas9(BB)-2A-Puro (Addgene, Cat#48139) which co-expresses the gRNA with the SpCas9 protein in mammalian cells . To assess the cutting efficiency, each of the plasmids expressing the sgRNA was transfected into HEK293 cells and genomic DNA of the cells was collected at 72 h post transfection. Each sgRNA-targeting region was amplified using PCR, and the amplicons were sequenced on the MiSeq instrument (Illumina, San Diego CA) to directly determine the frequency of mutations. Cutting efficiency was calculated as the highest deletion frequency of the base pair positions within the target region. Then, HEK293 cells were transfected with different combinations of gRNAs, two at a time, to test whether they can make any deletions. PCR was used to screen deletions between the two gRNA-binding regions and to select the best pairs of gRNAs.
Hemizygous SHANK3 deletion was introduced into the H9 human ESC line (WiCell Research Institute). Two sgRNA sequences, gRNA #1 (ATACCACAAGAAGGACGTCCGGG) and gRNA #4 (TTTCTCAGGGGTCCCCGGTGGGG), designed to flank the entire SHANK3 gene for creating a 69.5-kb deletion, were introduced using the same technique previously described. A total of 64 single colonies were screened for the deletion using PCR with two pairs of forward and reverse primers. Primers 1 (CAGACACTTCACGAAAGAAGC) and 4 (GGGAGCAAAAAGGAAACCTC) were amplified if deletion was successful. Primers 2 (GGAACTCAGCCCATGCCTTC) and 3 (TGCAGACGCAGAGCCTAACA) were amplified if no deletion occurred. Out of the 64 colonies, 4 were found to have heterozygous SHANK3 deletion. We selected one of the four lines for downstream experiments. The deletion was verified by Sanger sequencing.
Homozygous SHANK3 deletions were introduced into the 2242–5 human iPSC line (obtained from Dolmetsch’s laboratory). Two sgRNA sequences, gRNA #8 (TGAGCCGCTATGACGCTTCAGGG) and gRNA #4 (TTTCTCAGGGGTCCCCGGTGGGG), designed to flank exons 21–22 and the stop codon for creating an ~14-kb deletion, were introduced using electroporation together with the Cas9 sequence under the control of the EF-lα promoter. Upon electroporation of 5 µM of gRNA-Cas9 plasmids into ~3 × 106 cells using the Amaxa Nucleofector System (program A023), iPSCs were resuspended in E8 medium with ROCK inhibitor. On the following day, the cells were exposed to 0.25 µg/mL puromycin for 48 h. Antibiotic-resistant colonies were picked and expanded on Matrigel-coated plates in E8 medium. A total of 82 single colonies were manually picked, propagated, and screened for the deletion using conventional PCR and a set of one forward and two reverse primers (Primer F [yw3]: CTCAGGGCCTGCTTGATGAC; Primer R [yw4]: GTGCGCTCCTGAAGGACAAT; and Primer R [yw7]: GGTCTTGCATCGAGGTGCTC), which recognize different regions in the proximity of exons 21–22. Out of the 82 colonies, one was identified to have homozygous SHANK3 deletion encompassing exons 21–22. The deletion was verified using PCR product purification, subcloning into a TOPO TA cloning vector (Thermo Fisher, Cat# 450030), and sequencing.
Vectors and viruses
The following lenti vectors were used in this study: pBOB-synP-HTB (Addgene, Cat#30195) ; CamKIIα:smFP-TVA-RVG (constructed in the lab by subcloning the following fragments into the pLenti_CaMKIIα_GFP vector [Addgene plasmid #96941]; smFP-Flag fragment from the pCAG_smFP-Flag_dark vector [a gift from Megan Williams] ; F2A-TVA-T2A fragment from the pBOB_synP-HTB vector (Addgene, Cat#30195); oG fragment from the pCAG_oG vector [Addgene plasmid #74288] ); CamKIIα:GFP-TVA-RGV (constructed in the lab using the CamKIIα:smFP-TVA-RGV vector and substituting the smFP-Flag fragment with smFP-Flag_bright fragment from the pCAG_smFP-Flag_membrane_bright vector [a gift from Megan Williams] ). The lentiviruses were produced and titrated as previously reported . Rabies ENVAΔG-mCherry virus (Addgene, Cat#32636) was purchased from the Salk Institute Gene Transfer, Targeting and Therapeutics core facility.
Animals, transplantation, and rabies virus injection
All work with animals was performed in accordance with guidance by the University of Utah office of Institutional Animal Care & Use Committee. Both female and male NOD scid gamma mice purchased from The Jackson Laboratory (NSG, Cat#005557) were used for experiments. Neurons were dissociated using Papain enzyme (Worthington Biochemical, Cat# LS003119) and resuspended in ND medium supplemented with DNAseI (0.05 mg/mL, Sigma-Aldrich, Cat# D4527–40KU) to prevent cell clumping. Neurons were concentrated to a final concentration of 10,000–50,000 cells/μL. Cells were kept on ice before transplantation. Human neurons were transplanted into the prelimbic cortical area (AP 3.6, ML 0.3, and DV 0.9) of mouse neonates (P0–P3) using a stereotaxic injection system (Stoelting Co.). Hypothermia was used to anesthetize the animals. For this, pups are placed in a latex glove and immersed up to the neck in crushed ice and water (2–3 °C) for about 10 min. The depth of anesthesia is monitored based on the lack of spinal reflex after toe pinch. Anesthetized pups are placed under a stereoscope. A midline incision is made to open the skin, and a glass pipet is used to penetrate the skull and deliver the human cells into the brain (0.5 μL of dissociated human iPSC-derived neurons were delivered using a thin glass needle with <100 μm tip diameter at a 0.1 μL/min flow rate driven by a micro-syringe pump controller). Upon completion, the incision is closed, and the skin is sutured from side to side with sterile nylon 5–0 sutures. The entire surgical procedure lasted no longer than 10 min. After surgery, pups were kept at room temperature for 5 min and then transferred on pre-warmed Deltaphase isothermal pads for recovery. Once fully recovered, responsive, and mobile, pups were returned to their home cage and then to the animal facility. Animal were removed from the study and euthanized using methods consistent with AVMA guidelines if we saw any signs of pain or distress or rejection of transplanted pups by the dam.
Rabies ENVAΔG-mCherry virus was injected into the mPFC of adult animals in an animal biosafety level 2 environment. Briefly, animals were placed on a stereotaxic frame and kept under anesthesia by 1–3% isoflurane inhalation. A small incision was made in the skin to access the skull. The skull above the injection site was opened with a drill. The virus was loaded in a Hamilton syringe and injected at 0.1 μL/min into the adult mPFC following the stereotactic coordinates AP 1.7, ML 0.45, and DV 1.5. The opened skin was sutured after the surgery, and buprenorphine analgesic was provided to the animal every 12 h for 3 days. Rabies virus (RV) was allowed to spread for 7 days before tissue clearing and immunohistochemical analysis.
Acute slices for electrophysiology experiments were obtained from 2- to 15-month-old transplant recipient mice. Mice were anesthetized using isoflurane and, when non-responsive to a toe pinch, transcardially perfused with ice-cold “cutting” solution (~60 mL, 1 mL/s), containing (in mM/L): 2.5 KCl, 7 MgCl2, 1.25 NaH2PO4, 105 Choline-Cl, 25 NaHCO3, 25 D-glucose, 3 Na+-pyruvate, 11 Na+-L-Ascorbate, and. 0.5 CaCl2. Afterwards, the brain was quickly dissected and placed in the ice-cold “cutting” solution. Coronal slices (300–350 µm) were prepared using a Leica VT1200 vibratome in ice-cold “cutting” solution. Slices were allowed to recover for 30 min at 37 °C in standard ACSF, containing (in mM): 124 NaCl, 2.5 KCl, 2 CaCl2, 1 MgCl2, 1.25 NaH2PO4, 25 NaHCO3, 25 d-glucose, 1 Na+-pyruvate, and 1 Na+-l-Ascorbate and then kept at room temperature for the duration of the experiment (5–7 h). Both “cutting” and ACSF solutions were oxygenated using carbogen gas (95% O2/5% CO2).
Whole-cell voltage-clamp experiments were performed using an upright Ziess Axioskop 2 microscope and Multi-Clamp 700B amplifier (Molecular Devices, Palo Alto, CA, USA) at room temperature. Briefly, cells in slices were visualized under a 40× water-immersion objective using an infrared differential interference contrast filter, a digital camera (Ximea), and a fluorescence LED lamp. Recordings were obtained at room temperature in ACSF using patch pipettes pulled from borosilicate capillary glass (BF150–86–10, Sutter Instruments) with resistances 3–5 MΩ when filled with intracellular solution, containing (in mM): 120 CsMeS, 5 CsCl, 10 HEPES, 0.5 CaCl2, 2 EGTA, 3 Na-Ascorbate, 4 MgCl2, 5 d-glucose, 10 Na2-phosphocreatine, 0.4 Na2GTP, 4 Na2ATP, and 0.3% Biocytin (Osmolarity: 306 mOsm). Signals were filtered using 2 kHz Bessel filter and digitized at 50 kHz using Axon DigiData 1500 A (Molecular Devices). The Axon pClamp 10 (Molecular Devices) software was used to control the experiment. Spontaneous excitatory postsynaptic currents (EPSCs) and miniature excitatory and inhibitory postsynaptic currents (mEPSCs and mIPSCs, respectively) were recorded from fluorescent neurons positioned in the deep cortical layers. mEPSCs and mIPSCs were recorded using bath solution containing with 1 µM tetrodotoxin at −60 mV (the reversal potential for mIPSCs) and 0 mV (the reversal potential for mEPSCs), respectively. Events were collected from 3 min of recordings using a template-based approach in Clampfit 10. Templates were generated from control traces with clearly visible postsynaptic events. Thirty randomly selected events from individual recordings were used for statistical analysis in EXCEL (Microsoft) and Prizm (GraphPad). Evoked α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptors (AMPAR)- and N-methyl-D-aspartic acid receptors (NMDAR)-EPSCs were obtained from fluorescent neurons positioned in the deep cortical layers using a bipolar stimulating electrode placed in the layers II/III (at least 200 μm from the recorded cells) in response to 1 ms long current pulses of different intensity (0–0.5 mA) delivered every 5 s using an external stimulator (A365, WPI). In these experiments, the intracellular solution contained 5 mM QX-314 to suppress sodium channels and action potentials in the recorded neuron, and the bath solution contained 100 µM picrotoxin to inhibit IPSCs. In some experiments, 2 µM 2-Chloroadenosine was added to the bath solution to suppress seizure-like activity and population spikes. The amplitudes of the smallest responses (average of two traces), measured at the peak using holding potential −70 mV for AMPA-EPSCs and 100 ms post stimulation using holding potential +40 mV for NMDA–EPSCs, were used for analysis. The analysis was performed using the Clampfit 10 software and custom Python scripts.
Western blot analysis
Neurons were harvested and lysed with Laemmli Buffer (Bio-Rad, 1610737). Samples were loaded to a sodium dodecyl sulfate–polyacrylamide gel electrophoresis chamber, transferred to a polyvinylidene difluoride membrane overnight at 4 °C and blocked with 5% milk in phosphate-buffered saline (PBS) with 0.1% Tween 20 (PBST) for 1–2 h with rocking at room temperature prior to primary antibody incubation. Membranes were then incubated with anti-SHANK3 (1:350, Santa Cruz, [sc-301903]) or anti-TUJ1 (1:38,000, Covance [MMS-435P]) antibody in 5% milk in PBS containing Tween 20 (PBST) and slowly rocked at 4 °C overnight. Membranes were washed three times for 10 min each time with PBST and then incubated with horseradish peroxidase (HRP)-conjugated secondary antibody anti-rabbit or anti-mouse at 1:15,000–1:25,000 in 5% milk in PBST for 1–2 h at room temperature. Chemiluminescent HRP substrate (EMD Millipore, Cat#WBKLS0100) was used for visualizing proteins. Membranes were imaged with the ChemiDoc XRS+ System with Image Lab Software (Bio-Rad, 1708265) and analyzed using ImageJ (NIH).
Tissue clearing, immunohistochemistry, and imaging
The mice were anesthetized by intraperitoneal injection of 10 mg/kg xylazine and 100 mg/kg ketamine (Sigma-Aldrich, Cat#K113–10ML) and perfused with cold 16% paraformaldehyde (PFA, Electron Microscope Science, Cat# 15710) and 4% GA (glutaraldehyde, Electron Microscope Science, Cat# 16220) in 1X PBS, at a steady state rate of 100 mmHg2. After perfusion, the brain was dissected and kept in fixative solution for 24 h at 4 °C. To facilitate image analysis, the mouse brain was sectioned in 2.5–3-mm coronal slices for tissue clearing processing (as described in the SWITCH clearing technique by Murray et al. 2015 ).
Immediately after electrophysiology experiments, tissue sections (300 μm thick) were transferred to a 24-well plate containing 4% PFA for fixation. After 12 h of fixation, slices were transferred to 1X PBS for storage prior to free-floating immunostaining. Sections were washed three times for 10 min each in 1X PBS, blocked with 10% blocking solution (0.3% Triton X-100, 3% fetal bovine serum [FBS] in 1X PBS) for 6 h at room temperature. Brain sections were incubated with different combinations of primary antibodies (anti-eGFP, anti-CTIP2, anti-SATB2, anti-mKate, and anti-GAD67) in blocking solution for 24 h at 4 °C. Sections were then incubated with secondary antibodies (Alexa fluor 488-mouse, Alexa fluor 647-rat, Alexa fluor 405-mouse, Alexa fluor 568-rabbit, Alexa fluor 647-mouse, and streptavidin Alexa fluor 647conjugate) in blocking solution for 24 h at 4 °C in the dark. Sections were mounted in an aqueous-based mounting medium (Polysciences, Cat#18606–20) with a glass slide and coverslip. Confocal imaging (ZEISS LSM 880 with Airyscan) was performed with excitations at 405, 488, 594, and 647 nm using 5X, 10X, 20X, and 63X oil objectives.
For both the characterization of engrafted GFP-positive neurons and axon tracing, mice were perfused, and fixed brains were cryosectioned at 50-μm thickness. Brain slices were stained with anti-GFP antibody, and confocal imaging (ZEISS LSM 880 with Airyscan) was performed with an excitation wavelength of 488 nm using 5X, 10X, 20X, and 63X oil objectives.
Cells were fixed in 4% PFA (15 min at room temperature), washed (3×) with PBS 1X, permeabilized, and blocked in blocking solution (10% FBS, 0.1% Triton X-100, and 1% bovine serum albumin) for 1 h at room temperature. Primary antibodies were diluted 1:1000 in blocking solution and applied overnight at 4 °C. The following primary antibodies were used: chicken anti-eGFP (1:1000, Novus Biologicals [NB100–1614]), rabbit anti-mKate (1:1000, Axxora [AB234]), mouse anti-FLAG-m2 (1:500, Millipore Sigma [F1804]), rat anti-HA (1:500, Roche ), rabbit anti-TBR1 (1:200, Abcam [ab31940]), rat anti-CTIP2 (1:500, Abcam [ab18465]), mouse anti-SATB2 (1:250, Abcam [ab51502]), mouse anti-GAD67 (1:500, EMD Millipore [MAB5406]), mouse anti-TUJ1 (1:1000, Covance [MMS-435P]), and chicken anti-MAP2 (1:1000, Novus Biologicals [NB300–213]). Secondary antibodies and Hoechst 33342 were diluted 1:1000 in blocking solution and applied for 2 h at room temperature. The following secondary antibodies were used: Alexa-405, -488, -594, and -647. Neurons were imaged with a 20X objective on an Airyscan confocal microscope (Zeiss 880).
Rabies virus tracing in vitro
iCtrl and SHANK3+/– neurons transduced with CamKIIα:GFP-TVA-RGV lentivirus were co-plated with non-transduced human neurons of the same genotype from the same round of differentiation at 1:100 ratio (total 1 × 105 cells/well in 24-well plates) on the bed of rat astrocytes grown on poly-d-lysine precoated 15 mm glass cover slips in ND medium as previously described . One-week post plating, neurons were transduced with ENVAΔG-mCherry RV and fixed 48 h post transduction for immunostaining with anti-GFP and DAPI antibodies. Endogenous mCherry fluorescence was used for quantification. In the experiments with human neurons co-cultured with primary mouse cortical neurons, iCtrl or SHANK3+/– neurons transduced with pBOB-synP-HTB or CamKIIα:smFP-TVA-RGV lentiviruses were plated onto the bed of mouse cortical neurons at 1:100 ratio. One-week post plating, neurons were transduced with ENVAΔG-mCherry RV and fixed 48 h post transduction for immunostaining with anti-GFP, anti-FLAG, and Hoechst 33342. Endogenous mCherry fluorescence was used for quantification. The cultures of primary mouse cortical neurons were prepared from E18.5 mouse as previously described in . Briefly, mouse cortices were dissected, gently triturated, dissociated, and plated at 1 × 105 cells/well on poly-l-lysine (Sigma-Aldrich, cat#P9155) precoated 15 mm glass cover slips in Neurobasal medium (Neurobasal-A [Invitrogen, Cat#10888022], 2% B27 Supplement with vitamin A [Thermo Fisher Scientific, Cat#17504044], 1% GlutaMAX [Invitrogen, Cat#35050061], 25 μM 2-Mercaptoethanol [Gibco, cat#21985–023], and 1% Pen/Strep). Two days later, human neurons were added to the rodent cultures. Neurons were visualized using a Nikon Widefield microscope equipped with 405/488/543 lasers using Nikon NIS-Elements multi-platform acquisition software. For each coverslip, a wide-field-of-view image was acquired using a 20X objective and Perfect Focus System.
Neurite tracing and spine analysis
For analysis of neurite morphology, the 300-μm thick coronal sections previously used in the electrophysiology experiment and filled with biocytin were imaged with a 20X objective on an Airyscan confocal microscope (Zeiss 880). Basilar dendritic arbors of selected neurons were traced using Neurolucida (MBF Bioscience, Williston, VT, USA), and arbor complexity was quantified by Sholl analysis using Neurolucida Explorer software. The outcome measures were total dendritic length, number of primary dendrites, maximum number of dendritic intersections within Sholl radii, and branching points (calculated as maximum number of dendritic intersections/number of primary dendrites). Apical dendrites (100 μm in length, located ~10 μm away from the cell soma) were imaged using a 60X objective and 5X electrical zoom on Airyscan confocal microscope (Zeiss 880). For dendritic spine quantification, confocal microscopy images were analyzed using the function Autospine in Neurolucida software.
Data acquisition and statistical analysis
All experiments were performed using human stem cell-derived neurons generated in at least three independent differentiations. Pairs of isogenic lines were simultaneously differentiated and used for the experiments. Both male and female mice were used for transplantations and pooled together for analysis. Animals and cells were randomly selected for the experiments. Investigators were blinded for the genotypes of transplanted cells in animals that were used for the experiments and during analysis. The sample sizes were estimated using 3 G*Power software and the results obtained in our previous study and preliminary experiments. The sample size and statistical tests used for comparisons are reported in the figure legends and tables. Samples were excluded from analysis only if they were clear outliers (identified visually and confirmed using the ROUT method [Q = 2%]). The exclusion criteria were pre-established. We used a bootstrap test of the difference between the means to confirm that the experimentally determined differences between control and SHANK3-deficient cells were not stochastic due to an increased experimental variability.
Connectivity deficits in human stem cell-derived cortical neurons with complete hemizygous SHANK3 deletion in vitro
To investigate synaptic deficits elicited by SHANK3 hemizygosity in human neurons, we generated engineered human PSCs with complete hemizygous SHANK3 deletion (SHANK3+/–) using CRISPR/Cas9 technology and widely used H9 human ESCs (Fig. 1A–C). We employed two single guide RNAs (sgRNAs), one targeting the first exon and the second one directed against the stop codon, to achieve a complete deletion of the genomic region spanning the entire SHANK3 coding sequence (Fig. 1A). Out of 64 selected colonies, a complete hemizygous SHANK3 deletion (69.5 kB) was confirmed in one line using conventional gel electrophoresis and Sanger sequencing (Fig. 1B, C).
We next differentiated iCtrl and SHANK3+/– stem cells into neurons using a modified dual-SMAD inhibition protocol (Supplementary Fig. 1A), which allows the generation of human neurons with telencephalic identities (Supplementary Fig. 1B, C). We confirmed that at 6 weeks post induction, MAP2-expressing cells expressed typical deep-layer cortical markers TBR1 (~40%) and CTIP2 (~30%) and the superficial layer marker SATB2 (~15%) (Fig. 1D, E). We observed very similar percentages of these cells among both iCtrl and SHANK3+/– neurons (Fig. 1E), suggesting that SHANK3 hemizygosity produces no major effects on the specification of cortical excitatory neurons in vitro.
We performed western blot analysis to characterize the expression levels of SHANK3 proteins in iCtrl and SHANK3+/– cultures (Fig. 1F, G). Interestingly, we found that complete hemizygous SHANK3 deletion caused an approximately 50% reduction in the expression levels of the longest (~250 kDa) and shortest (~160 KDa) isoforms of SHANK3 proteins, but not in the expression levels of two intermediate isoforms (~200 and 180 kDa) (Fig. 1G). This result suggests that SHANK3 hemizygosity in human neurons triggers an ectopic SHANK3 expression from alternative internal promoters. This is consistent with the results of previous studies on Shank3-deficient mice and monkeys [19, 23, 40], suggesting the presence of conservative compensatory feedback mechanisms for regulating the level of SHANK3 expression [41, 42].
We next investigated whether SHANK3 hemizygosity in human neurons affects their synaptic connectivity, as was previously reported for SHANK3-deficient neurons from PMDS patients with large 22q13 microdeletions and engineered ESCs with partial SHANK3 deletion [29, 31]. To assess the levels of synaptic connectivity in the cultures of iCtrl and SHANK3+/– neurons, we used trans-synaptic RV tracing  (Fig. 1H). This approach has been widely used for mapping synaptic connectivity in the mouse brain  and in human stem cell-derived neurons [44, 45]. In contrast to conventional synaptic electrophysiology or immunostaining, this approach allows to assess the level of synaptic connectivity based on the proportions of starter cells (transduced to express the avian tumor virus receptor A (TVA), which is required for RV entry into the cells) and presynaptically connected traced cells (acquired RV through synaptic connections with the starter cells) in a robust and high-throughput fashion. Using RV tracing, we found that SHANK3+/– neurons establish significantly fewer connections than iCtrl neurons in vitro (Fig. 1I, J).
To determine whether connectivity deficits in SHANK3+/– cultures arise cell-autonomously as a result of deficits in postsynaptic neurons as we observed in our previous study on PMDS neurons , we co-cultured iCtrl and SHANK3+/– neurons with dissociated mouse cortical neurons to provide the same presynaptic inputs to both iCtrl and SHANK3+/– neuron (Supplementary Fig. 2A). We observed that SHANK3+/– neurons established significantly fewer connections with co-cultured mouse cortical neurons than did iCtrl neurons (Supplementary Fig. 2B, C). This result indicates that hemizygous deletion of SHANK3 in the postsynaptic neurons impairs synaptic connectivity.
To further validate RV tracing for studying synaptic connectivity in vitro and connectivity deficits detected in the cultures with SHANK3+/– neurons, we differentiated human neurons from another pair of iCtrl and SHANK3-deficient stem cell lines with homozygous deletions of the last two exons and the stop codon of SHANK3 (Supplementary Fig. 3A, B). We observed no SHANK3 expression in neurons with homozygous SHANK3 deletion (Wang et al., in review). We co-cultured iCtrl and homozygous SHANK3-deficient neurons with dissociated mouse cortical neurons and assessed synaptic connectivity using RV tracing (Supplementary Fig. 3C). Consistent with the results of our experiments with hemizygous SHANK3-deficient neurons, we found significantly fewer connections to homozygous SHANK3-deficient neurons, as compared to iCtrl neurons (Supplementary Fig. 3E). As expected, the connectivity deficits in homozygous SHANK3-deficient neurons were more pronounced than those in hemizygous SHANK3-deficient neurons. Finally, the transduction of mouse cortical neurons with EnvA-pseudotyped RV alone resulted in no mCherry-expressing cells (negative control, Supplementary Fig. 3F), confirming that RV transduction relies exclusively on the presence of starter cells expressing TVA receptors . Together, these results indicate that SHANK3 hemizygosity in human neurons leads to impaired excitatory synaptic connectivity, likely due to cell-autonomous deficits in the postsynaptic neurons induced by the loss of the longest and/or shortest isoforms of SHANK3.
Engraftment of human stem cell-derived neurons into the mouse PFC for studying connectivity
The main caveat of connectivity studies on human iPSC-derived neurons in vitro is the lack of a physiologically relevant brain environment. Engrafted human stem cell-derived neurons have been used for studying the maturation of human neurons and cellular deficits in human neurological and neurodegenerative disorders [47,48,49,50,51,52]. We investigated whether human stem cell-derived cortical neurons can be transplanted into the mouse PFC for studying synaptic deficits caused by SHANK3 hemizygosity. We chose the PFC region for transplantation because the expression pattern of high-confidence ASD-associated genes converges on the PFC [53, 54], and because altered functional connectivity in this region has frequently been identified in individuals with ASD [55,56,57].
We transduced control human stem cell-derived neurons with lentiviruses carrying GFP under the control of the CamKIIα promoter, which primarily labels functionally mature cortical excitatory neurons , and transplanted them into the PFC of neonatal immunocompromised mice (P0–P5) to promote their survival and integration into developing mouse neural circuits (Supplementary Fig. 1). We observed that the majority of transplanted animals (65%) survived and possessed GFP-expressing cells in the PFC (Fig. 2A, B and Supplementary Table 1). Two weeks post transplantation, all GFP-expressing cells were found localized within a 300-μm range surrounding the center of the graft (Fig. 2C), and exhibited relatively immature neuronal morphology with processes primarily oriented towards the midline (Fig. 2B), which is similar to the orientation of mouse pyramidal cortical neurons in the corresponding brain region. However, we also noticed that the numbers and distributions of GFP-expressing cells varied from animal to animal (Fig. 2D).
We next investigated how transplanted human neurons become anatomically integrated in the mouse brain. To map neuronal inputs onto the transplanted human neurons, we used RV-mediated retrograde trans-synaptic tracing [43, 58] and 3-mm thick cleared brain slices using the SWITCH approach . For this, hESC-derived neurons expressing TVA receptor and rabies virus glycoprotein were transplanted into the PFC of immunocompromised neonatal mice and then transduced with RV for connectivity tracing 4–8 months post transplantation (Fig. 2E). Interestingly, we found that RV-transduced neurons expressing mCherry were distributed to multiple brain regions proximal and distal to the transplantation site, including the secondary motor area (MOs: ~56% of all mCherry-expressing cells, Fig. 2Fa, H), medial PFC (mPFC: ~10%, Fig. 2Fb, H), anterior cingulate cortex (ACC: ~10%, Fig. 2H), primary somatosensory cortex (SSp: ~4.5%, Fig. 2H), thalamus (TH: ~3.5%, Fig. 2Fd, H), contralateral primary somatosensory cortex (cSSp: ~3%, Fig. 2H), agranular insula (AI: ~2%, Fig. 2Fc, H), contralateral agranular insula (cAI: ~2%, Fig. 2Fe, H), basolateral amygdala (BLA: ~1.7%, Fig. 2Fd, H), hippocampus (HI: ~1.7%, Fig. 2H), pallidum (PAL: ~1.4%, Fig. 2Ff, H), primary motor area (MOp: ~1.2%, Fig. 2Fa, H), contralateral anterior cingulate cortex (cACC: ~1.2%, Fig. 2H), midbrain brain (MB: ~0.5%, Fig. 2H), orbital cortex (ORB: ~0.3%, Fig. 2H), contralateral mPFC (cmPFC: ~0.1%, Fig. 2Fe, H), and taenia tecta of the olfactory cortex (TT: ~0.1%, Fig. 2H). These results suggest that human neurons transplanted into the mouse neonatal cortex are capable of receiving anatomical inputs from multiple regions in the mouse brain, which is consistent with the connectivity pattern observed for excitatory neurons in the mouse PFC [43, 59]. A caveat of this experiment, however, is that we were unable to use immunostaining with the cleared mouse brain tissue to map the starter cells and to identify the subtypes of traced presynaptic neurons in different brain regions.
To map outputs from the transplanted human neurons expressing GFP under the control of the CamKIIα promoter (CamKIIα:GFP), we counted the numbers of fluorescent processes in different mouse brain regions that are known to receive anatomical projection from the PFC . We found GFP-expressing processes in the thalamus (TH: ~38%, Fig. 2Gb, I), primary and secondary motor area (MOs: ~32%, Fig. 2Ga, I), striatum (STR: ~20%, Fig. 2Ga, I), primary somatosensory cortex (SSp: ~5%, Fig. 2Ga, I), and corpus callosum (CC: ~5%, Fig. 2Gc, I). This distribution pattern of GFP-expressing processes is similar to that observed for mouse PFC neurons .
Together, these results indicate that transplanted human neurons become anatomically integrated in the mouse PFC and to a large extent mimic the connectivity pattern of the host mouse neurons in the corresponding brain region. However, the transplantation experiments also revealed that the connectivity pattern and distribution of transplanted human neurons vary substantially from animal to animal (coefficient of variability: 57–283%, Supplementary Table 2), which may prohibit the use of this approach for identifying specific anatomical connections disrupted in ASD. To overcome this limitation, we adapted the dual-color labeling scheme  and co-transplantation to deliver human iCtrl and SHANK3+/– neurons into the same brain region for co-development under the same experimental conditions (Fig. 3).
Impaired AMPAR-mediated excitatory synaptic transmission in developing SHANK3 +/− human neurons
To determine whether hemizygous SHANK3 human neurons develop excitatory synaptic deficits in vivo, we transduced iCtrl and SHANK3+/– neurons with lentiviruses carrying RFP or GFP, respectively (or vice versa), under the control of the CamKIIα promoter and co-transplanted them together into the PFC of neonatal immunocompromised mice (Fig. 3A). Consistent with the results of our experiments in vitro, we observed that iCtrl and SHANK3+/– neurons acquired similar cortical laminar identities in the mouse brain (Fig. 3B–E). Specifically, we found that about 55% of engrafted human neurons expressed the deep-layer cortical marker CTIP2 (Fig. 3B, D) and approximately 30% expressed the superficial layer cortical marker SATB2 (Fig. 3B, E). These results suggest that SHANK3 hemizygosity in human neurons produces no overt effects on the specification of deep and superficial cortical neuronal identities in vivo.
We next investigated the functional synaptic properties of co-transplanted iCtrl and SHANK3+/– neurons in the PFC slices using patch-clamp electrophysiology (Fig. 3F–M). To determine whether basal excitatory synaptic transmissions was altered in SHANK3+/– neurons, we measured the amplitudes and frequencies of spontaneous EPSC at −70 mV in the presence of picrotoxin, an antagonist of GABAA receptors 3–8 months post transplantation (Fig. 3G and Supplementary Fig. 4). We found that amplitude but not frequency of spontaneous EPSC was significantly reduced SHANK3+/– neurons, as compared to co-transplanted iCtrl neurons (Fig. 3H, I). The reduced EPSC amplitude in SHANK3+/– neurons was observed at both 3 and 8 months post transplantation (Supplementary Fig. 4). This result is consistent with the result of a previous study on induced stem cell-derived human neurons with heterozygous SHANK3 deletion in vitro . It suggests that SHANK3 deficiency in developing human neurons affects the strength of excitatory synapses.
Excitatory synapses in the mammalian brain express two major subtypes of glutamatergic receptors: AMPARs and NMDARs . We investigated whether the impaired excitatory synaptic transmission detected in SHANK3+/– neurons is associated with reductions in both AMPAR- and NMDAR-mediated excitatory postsynaptic currents (AMPAR-EPSCs and NMDAR-EPSCs, respectively) (Fig. 3J–M). To assess AMPAR- and NMDAR-EPSCs in transplanted human neurons, we recorded evoked postsynaptic currents from co-transplanted iCtrl and SHANK3+/– human neurons in layer 5 induced by electrical stimulations with an electrode positioned in layers 2/3 to activate both local and distal inputs (Fig. 3J). Interestingly, we detected a significantly reduced amplitude of AMPAR-EPSCs, but not NMDAR-EPSCs (Fig. 3K, L), and, as a result, a significantly reduced AMPA to NMDA ratio in SHANK3+/– human neurons as compared to co-transplanted iCtrl neurons (Fig. 3M). This result suggests that excitatory synaptic deficits in SHANK3-deficient human neurons are associated with reduced expression of functional AMPARs, but not NMDARs, at excitatory synapses. This is consistent with the results of our previous study , in which we observed that only the AMPAR-EPSCs were completely rescued by exogenous SHANK3 in iPSC-derived neurons from PMDS patients.
Reduced excitatory synapses, spines, and dendrites in mature SHANK3 +/− human neurons
An advantage of the xenotransplantation model is that it supports prolonged development of xenografted human neurons to allow their functional and morphological maturation [51, 52]. It has been shown that the properties of xenografted stem cell-derived human cortical neurons resemble those of mature human cortical pyramidal neurons only after 10–11 months post transplantation . Therefore, we investigated the synaptic and morphological properties of co-transplanted iCtrl and SHANK3+/– neurons at 13–17 months post transplantation (Fig. 4). To assess the basal excitatory and inhibitory synaptic transmissions, we measured the amplitudes and frequencies of miniature excitatory and inhibitory postsynaptic currents (mEPSC and mIPSC, respectively) in the presence of TTX at different holding membrane potentials in co-transplanted iCtrl and SHANK3+/– neurons (Fig. 4A–F). Surprisingly, we found a similar mEPSC amplitude but significantly reduced mEPSC frequency in SHANK3+/– neurons, as compared to iCtrl neurons (Fig. 4A–C). Assuming that co-transplanted iCtrl and SHANK3+/– neurons interact with the same presynaptic inputs in the brain, this result suggests that SHANK3+/– neurons have reduced numbers of excitatory synapses, as compared to co-transplanted iCtrl.
We observed no differences in either the amplitude or frequency of mIPSCs (Fig. 4D–F), suggesting that co-transplanted iCtrl and SHANK3+/– neurons receive similar inhibitory synaptic inputs. This result is consistent with the results of our previous in vitro study on patient-derived SHANK3+/– neurons . It indicates that a balance of excitatory and inhibitory synaptic inputs on the individual human neurons may be disrupted as a result of SHANK3 deficiency.
Mature AMPAR-containing excitatory synapses predominantly reside on dendritic spines in cortical excitatory neurons [62,63,64,65]. We investigated whether reduced mEPSC frequency detected in SHANK3+/– neurons is associated with defects in dendritic spines. We reconstructed the dendritic morphologies of most recorded neurons filled with biocytin (Fig. 4G) and analyzed the density and morphology of spines on apical dendrites (Fig. 4H, I). We found that the density of dendritic spines on iCtrl neurons was similar to that detected in another study on transplanted stem cell-derived human neurons  and comparable to that observed in neonatal human brain tissue [66,67,68,69,70]. Interestingly, SHANK3+/– neurons exhibited significantly reduced spine density as compared to iCtrl neurons (Fig. 4I), particularly those with mushroom and thin morphologies that have been associated with more mature synaptic contacts [58, 59]. This result is consistent with the idea that SHANK3 is an important regulator of spinogenesis in neurons  and that reduced number of dendritic spines could be responsible for excitatory synaptic deficits in more mature SHANK3+/– human neurons.
In addition, we found that SHANK3+/– neurons had dramatically underdeveloped dendritic arbors, including significantly reduced dendritic length and number of branches, as compared to iCtrl neurons (Fig. 4B, C). This result is consistent with the results of the previous studies on SHANK3-deficient human neurons that detected morphological deficits in vitro . It suggests that SHANK3 is an important regulator of the overall morphological development and maturation in human neurons.
In this study, we established a co-xenotransplantation model for studying synaptic deficits in genetically engineered human neurons. We demonstrated that a complete hemizygous SHANK3 deletion, which is the most common genetic abnormality identified in PMDS patients, causes impaired AMPAR-mediated excitatory synaptic transmission, reduced dendritic spine density, and underdeveloped dendritic arbors in human cortical neurons engrafted in the mouse PFC. Interestingly, we also found that SHANK3 deficiency predominately affects the strength of excitatory synapses at early developmental time points and leads to the losses of excitatory synapses and spines later in development. These results are consistent with the idea that SHANK3 hemizygosity causes AMPAR-mediated excitatory synaptic deficits and morphological abnormalities in human neurons.
The xenotransplantation approach has been previously used to investigate the maturation of human neurons under physiological conditions [52, 72,73,74] and to develop cell-based therapies for patient with neurological and neurodegenerative disorders [47,48,49,50,51, 58, 75]. Our study presents a new direction for using this approach to study synaptic connectivity deficits in neurodevelopmental disorders associated with ASD. To ensure that the phenotypes observed in human neurons are physiologically relevant and caused by the genetic abnormality and not other factors that are difficult to control in transplantation studies, including different host brain environments and shifts in transplantation coordinates, we developed a co-xenotransplantation model. In this model, both wild-type and mutant human neurons reside in the same brain environment throughout development and interact with the same presynaptic inputs from both human and rodent neurons. Therefore, the synaptic and morphological deficits detected in this study are likely to be postsynaptic cell-autonomous deficits caused by SHANK3 hemizygosity.
Multiple synaptic deficits have been detected in rodent neurons in association with Shank3 deficiency, including an impaired amplitude and/or frequency of AMPAR-mediated excitatory synaptic transmission, impaired NMDAR-mediated synaptic responses, impaired long-term potentiation (LTP) and/or long-term depression (LTD), impaired mGluR signaling and mGluR-dependent LTD, and reduced dendritic spine density. This has led to the idea that synaptic function may be disrupted in association with SHANK3 hemizygosity in humans. However, the specific deficits have been incongruent across studies. For example, five studies found a reduced frequency of EPSCs in Shank3-deficient neurons [12, 13, 19, 20, 22], whereas two studies observed no such reduction [16, 24] and two other studies found an increased frequency of EPSCs [18, 26]. Similarly, four studies reported impaired LTP in CA1 pyramidal neurons of Shank3-deficient mice [12, 13, 24, 26], but three studies found no LTP deficits [16, 19, 20]. Furthermore, two studies reported reduced mGluR5 expression [15, 76], while three studies reported either no reduction or increased mGluR5 expression [12, 13, 22]. Moreover, no overt synaptic deficits have been detected in Shank3-deficient mice with a complete heterozygous Shank3 deletion . These phenotypic inconsistencies are likely due to differences in the deleted isoforms of Shank3, the genetic backgrounds, and the developmental stages and types of investigated neurons. As a result, it remains unclear which of the detected synaptic deficits can be targeted for therapy development in the preclinical studies.
Studies on human stem cell-derived neurons obtained from patients and CRISPR/Cas9-engineered stem cell lines have provided further insights into early patient- and mutation-specific deficits induced by SHANK3 deficiency. Interestingly, it has been demonstrated that even heterozygous SHANK3 deletions in human neurons lead to profound excitatory synaptic deficits and morphological abnormalities in vitro . These results suggest either that SHANK3 is critically involved in the development of excitatory synapses in human neurons so that the loss of even a single copy of SHANK3 leads to excitatory synaptic deficits, or that synaptic deficits detected in vitro are compensated in the brain environment. Our results on co-transplanted control and SHANK3-deficient human neurons grown in the mouse PFC support the notion that SHANK3 is essential for excitatory synaptic development in human neurons.
Interestingly, we found that human neurons with hemizygous SHANK3 deletion exhibit different excitatory synaptic deficits at different developmental stages. At 3–8 months post transplantation, SHANK3 deficiency is associated with reduced amplitude but not frequency of spontaneous EPSCs and impaired amplitude of evoked AMPAR-mediated synaptic responses; while at 13–17 months post transplantation, it is associated with reduced mEPSC frequency, but not amplitude, and decreased number of dendritic spines. Reduced amplitude of spontaneous EPSCs and evoked AMPAR-mediated postsynaptic current can be attributable to reduced expression of functional AMPAR in the postsynaptic density. Reduced mEPSC frequency can be caused by presynaptic deficits and/or decreased number of excitatory synapses. In the co-xenotransplantation system, both iCtrl and SHANK3+/– neurons interact with the same pool of presynaptic inputs coming from both mouse and human neurons, suggesting that reduced mEPSC frequency is likely associated with decreased number of excitatory synapses on SHANK3+/– neurons as compared to iCtrl neurons. AMPAR-containing excitatory synapses are predominantly localized on spines . Consistent with this idea, we observed that mature SHANK3+/– neurons have fewer dendritic spines as compared to iCtrl neurons. Together, these results suggest that SHANK3 hemizygosity in human neurons grown under physiological conditions leads to defective expression of functional postsynaptic AMPA receptors early in development and decreased number of excitatory synapses and spines at later developmental stages.
It remains to be determined whether the deficits detected during different developmental stages are interrelated or part of distinct mechanisms of synapse development and/or function mediated by SHANK3. In addition, in future studies, it is important to investigate the time course for the development of functional and morphological deficits in SHANK3+/– human neurons and determine the relationship between defective synaptic connectivity and the intrinsic excitability deficits observed in vitro . Finally, it remains unclear how reduced SHANK3 expression causes these deficits in human neurons and whether the same deficits could be detected in patient-derived neurons with different genetic abnormalities or different lines of isogenic iPSC-derived neurons with hemizygous SHANK3 deletion.
In summary, this study identifies specific postsynaptic deficits induced by SHANK3 hemizygosity in human stem cell-derived cortical neurons transplanted in the mouse PFC and validates the use of the co-xenotransplantation model for detecting synaptic and morphological deficits in genetic neurodevelopmental disorders associated with ASD.
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We thank Ricardo Dolmetsch and Theo Palmer for providing iPSC lines; Yuanyuan Wu for helping with generating homozygous CRISPR/Cas9-engineered stem cells; Travis Philyaw for helping with mapping deletions; Brittney Nhem and Nico Edgar for helping with image analysis; Megan Williams, Edward Callaway, Liqun Luo, and John Naughton for plasmids; Chris Rodesch and the University of Utah Cell Imaging Core for assisting with microscopy, Jay Spampanato and Peter West for advising on intracranial injections and slice physiology; Monica Vetter, Richard Dorsky, and Sungjin Park for commenting on the manuscript. This work was supported by grants from the NIMH, NINDS, Brain Research Foundation, Brain and Behavior Research Foundation, Whitehall Foundation, Alan B. Slifka Foundation, and Utah Neuroscience and Genome Project Initiatives.
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Chiola, S., Napan, K.L., Wang, Y. et al. Defective AMPA-mediated synaptic transmission and morphology in human neurons with hemizygous SHANK3 deletion engrafted in mouse prefrontal cortex. Mol Psychiatry 26, 4670–4686 (2021). https://doi.org/10.1038/s41380-021-01023-2
Molecular Psychiatry (2021)