Background
Autism spectrum disorder (ASD) is a neurodevelopmental disorder with a complex and poorly understood etiology [
1‐
3]. Behavioral and imaging studies have been valuable for defining deficits in affected individuals and characterizing alterations at the level of the brain. However, we are extremely limited in our ability to acquire or experimentally manipulate human brain tissue from living patients or post-mortem brain slices. This has hampered efforts to study cellular and molecular abnormalities that accompany ASD, both during and after fetal and post-natal development. Notably, both the relative integrity of brain structures in affected individuals and the diversity of ASD genetics suggest that convergent mechanisms that contribute to affectation in ASD may operate at the level of the cell [
3,
4]. These may be identifiable in experimental models derived from affected individuals. In particular, ASD appears to frequently involve abnormal development and/or function of two major classes of neurons in the cerebral cortex, glutamatergic excitatory projection neurons (cExNs) and GABAergic inhibitory interneurons (cINs) [
3,
5,
6]. In vitro differentiation-based models of these neuronal cell types can identify cellular and molecular deficits associated with ASD and provide a tractable platform to screen for pharmacologic agents that can rescue these deficits.
In recent years, such cellular models of ASD have been generated, either by deriving induced pluripotent stem cell lines (iPSCs) from affected individuals or by using CRISPR/Cas9-based gene editing to engineer ASD-associated mutations into wild-type PSCs [
7‐
19]. Most of these studies have focused on syndromic forms of ASD, or on monogenic, de novo cases, where causality is attributed to the mutation of a single ASD-linked gene [
10,
11,
13‐
16]. These forms of ASD are attractive for cellular modeling, as they streamline study design and reduce many potential confounding variables. Other studies have included individuals with an unknown genetic cause of ASD, but with subject selection based upon a shared phenotypic characteristic, such as macrocephaly [
18‐
20]. Together, these models have been informative, revealing both cellular and molecular alterations associated with affectation. These include shared and model-specific disruptions of gene expression in ASD-derived neurons, frequently involving altered expression of genes in key developmental signaling pathways, and genes that control cellular proliferation and growth [
7,
8,
13‐
20]. In addition, differences were observed in neural precursor cell (NPC) proliferation and differentiation [
9,
19], neurogenesis, [
9,
11,
18,
20], synaptogenesis [
8,
10,
17,
18], or functional neuronal activity [
7‐
9,
11,
16,
17]. Altered expression of ASD genes, in which a mutation is linked to ASD causation or risk, is also frequently observed [
7,
8,
13‐
16,
18‐
20].
These cellular modeling studies have revealed potential contributors to affectation and, in some cases, have identified targets amenable to pharmacological rescue in vitro [
10,
18]. However, they do not encompass the range of contributors to ASD burden in the general population: no single gene mutation accounts for more than 1% of overall ASD cases with predicted monogenic causality [
21], while the majority of genetic risk appears to be polygenic or idiopathic [
2,
22‐
28]. Polygenic ASD risk can involve both common and rare variation in protein-coding and non-coding regions of the genome, which may act in a combinatorial manner [
2,
27,
29,
30]. Furthermore, ASD exhibits pronounced heritability in families (estimated at 50–90% [
31]), none of which can be accounted for by de novo (germline) events. Even within each multiplex ASD family, there is often a considerable range in the extent of affectation among individuals, and in most multiplex autism families, a single causative gene mutation cannot be identified [
31].
While ASD burden in the general population predominantly involves polygenic or idiopathic risk, heritability, and variable affectation [
2,
21,
27,
29‐
31], these genetically complex forms of the disorder have been largely neglected in cellular modeling studies. Therefore, we deemed it important to determine whether cellular modeling of these complex but prevalent forms of ASD could also reveal affectation-related deficits. To test this, we focused on a multiplex ASD family with variable affectation among family members. We generated iPSC lines from three first degree relatives (a male and a female with differing degrees of affectation and their unaffected mother), as well as an unrelated unaffected female. We used these lines to perform differentiation into both cExN and cIN neural progenitors and/or neurons. Models from the affected individuals exhibited compromised cellular responses to differentiation cues and had disrupted gene expression profiles. This included altered expression of many ASD-associated genes, genes with roles related to behavior, cognition, and learning, and genes involved in nervous system development and function, including cell adhesion molecules and ion channels.
This is, to our knowledge, the first cellular modeling study of multiplex ASD, including graded affectation among family members. This work demonstrates that even genetically complex forms of ASD have discernable cellular and molecular abnormalities that track with affectation, some of which overlap those identified in prior modeling of syndromic, monogenic, and de novo forms of ASD. Therefore, this novel study design highlights the potential for cellular modeling to identify convergent hallmarks across the broad diversity and genetic complexity of pathways to affectation.
Methods
Phenotyping of the multiplex family
The nuclear family consisted of working professional non-consanguineous parents, whose firstborn child was a daughter with DSM-5 autism spectrum disorder (ASD), Level I (requiring support, meeting DSM-5 criteria for Asperger syndrome) who was very high functioning and ultimately attended college, followed by a pair of monozygotic twin boys with ASD, Level III—one more fluently verbal than the other but both severely impaired and requiring very substantial support (see below)—followed by a third son with very subtle autistic traits and predominantly affected by attention deficit hyperactivity disorder, which improved substantially with stimulant medication treatment. Trio exome sequencing (ES) of one of the twins and his parents revealed a variant of uncertain significance (VUS) in GPD2, which was inherited by all of the children from the mother, who is of above-average intelligence with no dysmorphism and no history of developmental problems. All pregnancies were uncomplicated, except for the post-natal hospital course of the twins.
The daughter was born at term with no complications or dysmorphia. Her language and motor development were typical and she was able to read at an early age. By age five she was reading at a fifth-grade level. She has been described as talented in writing and drawing. According to her parents, she exhibited social oddities from an early age, mainly in communication, and has somewhat intense/restricted interests in fantasy games. She has strong language abilities, and currently attends a 4-year college, but at times uses odd phrases and the rhythm of her speech includes irregular pauses. She has described feeling alienated and “different”, and was the victim of bullying in middle school, with few close friends. In late adolescence, she developed major depressive disorder with moderate severity, which brought her to first psychiatric contact. She is cognizant of some degree of social awkwardness, which leads to feelings of anxiety and self-consciousness. The social anxiety inhibits her from activities such as eating in the cafeteria and pursuing job opportunities for which she is otherwise well-qualified. She has a history of becoming emotionally dysregulated and overwhelmed in times of stress, which has led to self-injurious behaviors. She has had ongoing struggles with depressive decompensation and suicidal ideation. She has above-average intelligence but has struggled academically in college due to depression and anxiety. She is medically healthy with the exception of supraventricular tachycardia secondary to atrioventricular node reentry, which was treated with ablation and resulted in subsequent normalization of her electrocardiogram.
The twin boys were born at 35 weeks, had breathing problems at birth, and spent 10 days in the newborn intensive care unit. Neither child has any dysmorphic features nor congenital medical abnormalities, and brain imaging studies were negative. Likewise, neither child has a history of confirmed seizures; however, there are concerns for possible absence epilepsy. There is no history of abnormal neurological examination or macro-/microcephaly. Development of both siblings was delayed, but neither had appreciable regression. The more severely affected twin (designated as the affected proband, AP, and from whom the induced pluripotent stem cell (iPSC) model of severe ASD affectation was acquired) began to exhibit delays in development by 9 months of age. He was speaking single words at 14 months and was ultimately diagnosed with autism at 3.5 years old. Research confirmation of the diagnosis was obtained using the Autism Diagnostic Observation Schedule. Compared to his twin brother, he has had more perseverative interests on odd objects. Psychological testing at the age of nine revealed an IQ of 65 using the Leiter International Performance Scale. Now in late adolescence, he has the ability to engage in reciprocal and meaningful verbal exchanges, although his language is often echolalic and repetitive. He is socially motivated and develops superficial friendships with peers. Functionally, he is able to complete most self-care, dress himself, prepare food and feed himself, and count money. He participates in a vocational program at school and is able to complete rudimentary tasks assigned to him. His monozygotic twin was also diagnosed with ASD at 3.5 years old and is less severely affected, but still requires significant support. Selected clinical characteristics of the family members studied are summarized in Table
1.
Table 1
Selected clinical characteristics and mutational status of several individuals in the multiplex ASD pedigree
Age of ASD diagnostic confirmation | 3.5 years | 18 years | N/A |
Social responsiveness scale-2 | 83 T | 72 T | 56 T (spouse-report) |
Depression and anxiety | Yes | Yes | No |
Seizure history | No | No | No |
Developmental delay | Yes | No | No |
Eye contact | Poor | Fair | Good |
Repetitive behavior | Yes | No | No |
Abnormal sensory sensitivities | Yes | No | No |
IQ | 65 | 102 (Raven) | 108 (Raven) |
Speech delay | Yes | No | No |
ASD | Severe | Moderate | No |
Intellectual disability | Yes | No | No |
Mutation location | GPD2 | GPD2 | GPD2 |
chr2:157352686 (hg19) G>A | chr2:157352686 (hg19) G>A | chr2:157352686 (hg19) G>A |
p.G78E, c.233G>A | p.G78E, c.233G > A | p.G78E, c.233G>A |
Genotyping of the multiplex family
Standard trio ES was performed by the clinical diagnostic laboratory GeneDx for the unaffected mother (UM), the AP, and the unaffected father. As described by GeneDx, exons were captured and sequenced on an Illumina platform with at least 100 base pair read length, followed by alignment to human genome build GRCh37/hg19 and subsequent variant identification. GeneDx’s custom-developed tool, Xome Analyzer, was used for variant analysis, as described [
32]. This process involves comparing the sequences of each individual to a number of resources, such as published reference sequences, other family members, and control individuals, including the 1000 Genomes database, NHLBI Exome Sequencing Project, ExAC, gnomAD, OMIM, PubMed, and Clinvar. Variant annotations include evolutionary conservation scores, results of in silico prediction tools, and references from the published literature. A phenotype-based approach is also used to generate candidate gene lists. This information was interpreted by GeneDx experts according to the American College of Medical Genetics and Genomics Guidelines. ES was performed by The Genome Technology Access Center (GTAC) at Washington University on the intermediate phenotype sister (IS), and variant-specific testing was performed by GeneDx on the trait-affected brother (TB) to confirm the presence of the identified
GPD2 variant.
iPSC generation
iPSC lines were generated by the Genome Engineering and iPSC Center (GEiC) at Washington University. Biomaterials for reprogramming were only available from the UM, IS, and AP. Briefly, renal epithelial cells were isolated and cultured from fresh urine samples and were reprogrammed using a CytoTune-iPS 2.0 Sendai Reprogramming kit (Thermo Fisher Scientific), following the manufacturer’s instructions. At least three clonal iPSC lines were derived for each subject, and one or two of these clonal lines (clones 1 and 2) were used for all experimentation involving the UM, IS, and AP. The UC line was previously derived by the GEiC, and one clonal line was available for use in all experiments involving this cell line. All clones (clones 1 and 2) used in experiments were assessed for karyotypic abnormalities by the Washington University School of Medicine Cytogenetics and Molecular Pathology Laboratory, and were also characterized for pluripotency by immunocytochemistry (ICC) and RT-qPCR. Each statistically significant experimental finding reported here was made in experiments that used two different clonal lines per individual (except for the UC, where only one clonal line was available), with at least three independent biological replicate experiments performed per clonal line. Statistical comparisons were made by one-way ANOVA or unpaired
t test. Documentation of the clone used for each replicate experiment, the replicates performed, and the statistics for each finding is detailed in Additional file
1: Table S1.
iPSC maintenance and differentiation
iPSC lines were grown under feeder-free conditions on Matrigel (Corning) in mTeSR1 (STEMCELL Technologies). cExN and cIN differentiation of iPSCs was performed using previously described protocols [
33]. Briefly, for cExN differentiation, iPSCs were dissociated to single cells with Accutase (Life Technologies) and 40,000 cells were seeded in V-bottom 96-well non-adherent plates (Corning). Plates were spun at 200×
g for 5 min to generate embryoid bodies (EBs) and were incubated in 5% CO
2 at 37 °C in cExN differentiation medium with 10 μM Y-27632 (Tocris Biosciences). cExN differentiation medium components include Neurobasal-A (Life Technologies), 1X B-27 supplement (without Vitamin A) (Life Technologies), 10 μM SB-431542 (Tocris Biosciences), 100 nM LDN-193189 (Tocris Biosciences). On day four of differentiation, EBs were transferred from V-bottom plates to Poly-
l-Ornithine- (20 μg/ml) and laminin- (10 μg/ml) coated plates. Media (without Y-27632) was replenished every other day, and on day 12, Neural Rosette Selection reagent (STEMCELL Technologies) was used to select neural progenitor cells (NPCs) from within neural rosettes, per the manufacturer’s instructions. cExN NPCs were grown as a monolayer using cExN differentiation media for up to 15 passages. cIN differentiation media contained the same components as cExN differentiation media, while also including 1 μM Purmorphamine (Calbiochem) and 2 μM XAV-939 (Tocris Biosciences). EBs were generated as described for cExNs. At day four of differentiation, the EBs were transferred to non-adherent plates and were placed on an orbital shaker (80 rpm) in an incubator with 5% CO
2 at 37 °C. The media was replenished every other day and, at day ten, EBs were transferred to Matrigel- and laminin- (5 μg/ml) coated plates. Y-27632 was included in media until day eight of differentiation. On day 12 of differentiation, NPCs were dissociated with Accutase and maintained as a monolayer for up to 15 passages. For both cIN and cExN NPC growth analysis, an equal number of cells were seeded on Matrigel- and laminin- (5 μg/ml) coated plates and total cells were counted 4 days later when the cells reached 70–80% confluence.
For differentiation of cExN NPCs into neurons for maturation, 40,000 cells per well were seeded in V-bottom 96-well non-adherent plates. Plates were spun at 200×g for 5 min and incubated in 5% CO2 at 37 °C in maturation medium with Y-27632. Maturation medium components include Neurobasal-A, 1X B-27 supplement (without Vitamin A), 200 μM cAMP (Sigma), 200 μM Ascorbic acid (Sigma), and 20 ng BDNF (PeproTech). After 2 days, EBs were transferred to Matrigel- and laminin- (5 μg/ml) coated plates and media was replenished every other day (without Y-27632). On day 12 of neuronal differentiation and maturation, cells were dissociated with Accutase and seeded in an eight-well chamber for ICC.
For neurosphere size measurement analysis, P values *P < 0.05, **P < 0.01, ***P < 0.001 were determined by one-way ANOVA.
Sanger sequencing
DNA was isolated from cell lines using the PureLink Genomic DNA Kit (Invitrogen). Primers were designed to amplify a 248 base pair region of GPD2 flanking the identified point mutation (forward primer: AAGCAGCAGACTGCATTTCA, reverse primer: CACCATGGCACACACTTACC). Sanger sequencing was performed on this PCR amplified fragment using either the forward or reverse primer. CodonCode Aligner software was used to analyze sequencing results.
Immunocytochemistry (ICC) and immunoblotting
For ICC, cells were plated on eight-well chamber slides coated with Matrigel and laminin (5 μg/mL). After 1 day, cells were washed once with PBS without calcium and magnesium (PBS - Ca
2+/Mg
2+) and fixed in 4% paraformaldehyde for 20 min, followed by washing with PBS + Ca
2+/Mg
2+. Cells were blocked with blocking buffer (10% donkey serum, 1% BSA, and 0.1% TritonX-100 in PBS + Ca
2+/Mg
2+) for at least 1 h and incubated with primary antibodies overnight (Additional file
1: Table S1) in antibody dilution buffer (1% donkey serum, 1% BSA, and 0.1% TritonX-100 in PBS + Ca
2+/Mg
2+). After overnight incubation, cells were washed three times with wash buffer (0.1% Triton X-100 in PBS + Ca
2+/Mg
2+). Cells were incubated with corresponding secondary antibodies (Additional file
1: Table S1), along with DAPI (1 mg/mL; ThermoFisher Scientific), diluted in antibody dilution buffer for 1 h. Following secondary antibody incubation, cells were washed twice with wash buffer and once with PBS + Ca
2+/Mg
2+. Slides were mounted with Prolong Gold anti-fade agent (Life Technologies). Images were obtained using a spinning-disk confocal microscope (Quorum) with MetaMorph software and were processed using ImageJ. For immunoblotting, cell lysate was extracted and 30 μg of protein was used per lane. Antibodies used are listed in Additional file
1: Table S1.
FACS analysis
For each experiment, approximately one million cells were pelleted, washed with PBS – Ca2+/Mg2+, resuspended in PBS – Ca2+/Mg2+ and fixed by adding 70% ice-cold ethanol dropwise while vortexing. Cells were stained with 10 μg/mL propidium iodide (Sigma) and 200 μg/mL RNase A (Fisher Scientific) in FACS buffer (PBS – Ca2+/Mg2+, 0.2% BSA, 1 mM EDTA). FACS was performed on single-cell suspensions and the cell cycle analysis function of FlowJo was used to analyze cell cycle composition for each sample, based on propidium iodide staining to detect DNA content in each cell. P values *P < 0.05, **P < 0.01, ***P < 0.001 were determined by one-way ANOVA.
RNA-seq and RT-qPCR
Total RNA was collected from iPSC-derived day 12 cExN and cIN NPCs using the NucleoSpin RNA II kit (Takara) per the manufacturer’s instructions. RNA was quantified using a NanoDrop ND-1000 spectrophotometer (Thermo Scientific), and the integrity of RNA was confirmed with an Agilent Bioanalyzer 2100 to ensure a RIN value above eight. RNA-sequencing (RNA-seq) library preparation and Illumina sequencing were performed by the GTAC at Washington University. Single-end 50 base pair reads were obtained using an Illumina HiSeq 3000 sequencer, obtaining an average of ~30 million uniquely aligned reads per sample. For RT-qPCR, 1 μg total RNA was reverse transcribed using iScript Reverse Transcription Supermix (Bio-Rad). Equal quantities of cDNA were used as a template for RT-qPCR, using the Applied Biosystems Fast Real-Time quantitative PCR system. RPL30 mRNA levels were used as endogenous controls for normalization. P values *P < 0.05, **P < 0.01, ***P < 0.001 were determined by an unpaired t test.
RNA-seq data analysis was performed as described in [
33] to curate differentially expressed gene (DEG) lists. In summary, RNA-seq reads were aligned to the human genome (assembly hg38) with STAR version 2.5.4b [
34]. Gene counts were derived from the number of uniquely aligned unambiguous reads by Subread:featureCount, version 1.6.3, with GENCODE gene annotation (V27) [
35,
36]. All gene-level transcript counts were then imported into the R/Bioconductor package DESeq2 [
37]. Genes expressed below a CPM of 1.0 in more than half the samples were excluded from further analysis. DEG cutoffs were set at a log2-fold change of > 1.0 and a Benjamini and Hochberg FDR of < 0.05.
To uncover the biological significance of DEGs, network analysis was performed with the data interpretation tool Ingenuity Pathway Analysis (IPA) (Qiagen). IPA’s Ingenuity Knowledge Base uses network-eligible DEGs to generate networks and to define connections between one and more networks. Based on the number of eligible DEGs, IPA defines network scores as inversely proportional to the probability of finding the network and defines significant networks (P ≤ 0.001). Within each network, red symbols indicate upregulated genes and green symbols indicate downregulated genes, where the color intensity represents the relative degree of differential expression.
Co-expression and variance analysis
For this correlation analysis, the read counts matrix for DEGs in either the cIN or cExN samples was log2-CPM transformed. A similarity matrix for these genes was created by calculating the Euclidian distance among the genes from the log2-CPM matrix in R. The WGCNA R package “adjacency.fromSimilarity” function with arguments power=12, type=’signed’ was used to create an adjacency matrix from the similarity matrix [
38]. Hierarchical clustering was performed using the hclust function in R on the distance matrix transformation of the adjacency matrix using the ward.D2 method. The dendrogram of this gene tree was split into
k = 3 clusters using the cutree function in the stats package of R. The read counts matrix for all genes in either the cIN or cExN samples was CPM transformed. Genes expressed at < 1 CPM in over half of the samples were excluded. Average CPM among the four sample types was calculated for each gene, and genes with expression ratios of < 1.25 between the highest and lowest expressed sample type were excluded from further analysis. The filtered gene CPM matrix was log2 transformed, and the Pearson correlation matrix for the remaining genes was calculated using the cor function from the stats package in R. A gene was considered to be correlated to a DEG cluster if it had a Pearson correlation coefficient > 0.7 for over half the genes in the DEG cluster. Genes with expression correlated to a cluster of genes from the DEG clustering were merged with those genes and fed into the ToppFun GO analysis of ToppGene [
39]. For each correlated cluster, up to five terms with the most significant Benjamini and Hochberg FDR values were retained for each biological process, cellular component, and molecular function GO terms. The top three pathway and disease terms were also retained for each cluster. To assess the impact of different covariates on expression, the R/Bioconductor package variancePartition was utilized [
40]. The parameters assessed were cell type (cExN vs. cIN), subject (UC, UM, IS, AP), age (young vs. old), or sex (male vs. female).
Discussion
In recent years, the genetic structure of autism spectrum disorder (ASD) risk in the general population has been clarified. This work has confirmed that while, in some cases, deleterious, single gene variants are significant contributors to ASD, the vast proportion of population attributable risk is polygenic [
2,
51]. Furthermore, this risk is highly heritable, and individuals within a multiplex family typically exhibit variable degrees of affectation [
31]. Here, we modeled cellular and molecular correlates of ASD within one such multiplex family, performing cortical neural differentiation of iPSCs derived from several family members with differential affectation. In this family, both polygenic liability and a shared variant of uncertain significance (VUS) may contribute to risk. In cells derived from the affected individuals, we identified compromised responses to differentiation cues and altered gene expression profiles during iPSC differentiation into cortical excitatory (cExN) and inhibitory (cINs) neurons, compared to related and unrelated unaffected controls. This work demonstrates that iPSC-based modeling can be used to characterize these more genetically complex but prevalent forms of ASD, in addition to modeling simplex and monogenic forms, which have been the focus of most studies to date. Moreover, these data provide information on physiologic and transcriptomic signatures of multiplex autism, with which cellular models derived from other families and other combinations of inherited susceptibility factors can be compared in future work.
Our phenotypic analysis of these four iPSC-based models of cortical neural development included assays conducted in the stem cells, during neural specification, in the proliferating NPCs, and during neuronal differentiation. During cExN NPC specification and during cIN NPC propagation, models from both affected individuals exhibited elevated fractions of cells with sub-G1 DNA content, relative to control-derived models. These data suggest that models derived from the affected individuals are less resistant to stressors, such as induction of differentiation, with these stressors increasing the propensity for cells to undergo apoptosis. While the molecular trigger for the induction of apoptosis here is unclear, expression of stress and apoptosis-related genes, such as
CHCHD2, ANXA1, and
SPATA18 are dysregulated in these models [
52‐
56]. These findings are reminiscent of some observations made in prior work, in which schizophrenia subject-derived iPSCs exhibited reduced neurosphere size [
57] and increased apoptosis was observed in Williams-Syndrome iPSC-derived NPCs [
58]. Interestingly, few studies report cellular alterations observed prior to the NPC stage, often focusing predominantly on phenotypes seen in NPCs and mature neurons [
7‐
19]. While this may reflect a lack of earlier phenotypic changes in some models, our findings highlight the importance of tracking neurodevelopmental alterations from their earliest onset. A recent report underscores the value of using cellular modeling approaches that aim to recapitulate some aspects of in vivo neurodevelopment [
20]. This study found that direct conversion of iPSCs into neurons masked ASD-associated cellular phenotypes, which were observable during the directed differentiation of iPSCs [
20].
In our study, transcriptomic analysis of neural progenitor cells revealed dysregulated expression in affected individuals compared to controls of gene networks related to behavior, psychological disorders, and neuronal development and disease. Genes encoding transcription factors were among the neurodevelopment-related genes with reduced expression in both affected individuals. For example, ARX is required for normal telencephalic development and is associated with syndromic autism and other neurodevelopmental disorders [
59], while EMX1 and FOXB1 also play important roles in neural development [
60‐
62]. Behavioral misregulation is a key trait of ASD, and gene networks related to the GO term ‘behavior’ exhibited dysregulated expression in both affected individuals. Genes in these networks include
COMT, ADCYAP1, CNR1, HTR2C, GRIK2, and
RGS4, all of which are implicated in behavior-related phenotypes in humans and/or mice [
63‐
71]. ASD genes were also dysregulated in these affected individuals, relative to controls. Mutation of these genes in other individuals is implicated in autism risk or causation. These include adhesion-related genes (
PCDHA1,
PCHDHA6,
PCDHGA11, PCDH8, PCDH9, PCDH10 [
72‐
76]
, KIRREL3 [
77],
CNTN3,
CNTN4 [
78],
CNTNAP4 [
79], and
THBS1 [
80]), receptor and channel genes (
CACNA2D3 and
SCN9A [
81]
, GRIK2 and
GRIK3 [
82‐
85],
KCNJ2 [
86,
87]
, and
GRIA1 [
88,
89]), and genes associated with central nervous system development and axon guidance (
ERBB4 [
90,
91],
NTNG1 [
92],
TSHZ3 [
93],
EBF3 [
94]
, MYT1L [
95,
96], and
ANXA1 [
54‐
56])
. Altered expression of ASD-associated genes has also been observed in cellular models derived from affected individuals in other studies [
8,
13‐
16,
19,
97]. Therefore, these findings suggest that misregulated expression of suites of ASD-associated genes may contribute to risk or affectation, and may do so by altering neurodevelopment and/or neuronal function in these affected individuals.
A unique aspect of this study is the use of iPSC-based directed differentiation into both cExNs and cINs, enabling us to identify neural cell-type-specific alterations associated with affectation. Although DEGs identified in affected individuals in both neural cell types were associated with many similar functions and diseases (e.g., behavior), the specific DEGs obtained often varied by cell type. For example, cIN DEGs included many more ASD-associated genes and protocadherin genes, the latter of which control neuronal migration, axonal growth, and synapse formation [
74,
75]. Human post-mortem cortical tissue from individuals with ASD has been shown to exhibit disrupted expression of cIN-associated genes, evidence that this cell type may commonly be disrupted in affected individuals in vivo [
98]. These findings suggest that extending cellular modeling studies to multiple disease-relevant neuronal cell types, including cINs, may reveal additional neurodevelopmental disruptions related to affectation.
To define cellular and molecular perturbations commonly related to affectation, we compared our findings to other studies that modeled ASD by directed differentiation of iPSCs into cExNs. We identified subsets of overlapping DEGs in comparisons with studies involving idiopathic autism cases vs. controls (26 shared DEGs [
7]), syndromic ASD involving macrocephaly (31 shared DEGs [
19]), and modeling of mutation of the syndromic ASD gene
CHD8 (32 shared DEGs [
13]) (Additional file
7: Table S6). Data for such comparisons is limited at present because iPSC-based models have been generated for a relatively small number of individuals and mutations, and these almost exclusively characterize cExNs or cerebral organoids [
7,
13,
18,
19].
The multiplex pedigree studied here was subjected to clinical exome sequencing, as it was hypothesized that a single, shared, genetic contributor might mediate autism risk and differential phenotypic expression in this family. In this sequencing analysis, a thread of shared genetic liability among all children was a VUS in the ASD- and ID-associated gene,
GPD2 [
99‐
101]
, which was inherited from their mother. However, there is variable ASD expressivity among these individuals, ranging from absent, to intermediate, to severe. In addition, both males and females in the pedigree are variably affected, indicating the presence of other significant contributors to variation in severity of affectation within this family. This observation is consistent with recent evidence that genetic liability for ASD is prevalently polygenic, and that, even in multiplex pedigrees where a significant monogenic contributor has been identified, additional polygenic risk can contribute to affectation [
51]. Moreover, this multiplex family was prototypic in reflecting the most severe form of affectation occurring in a male.
We hypothesized that it might be possible to identify graded cellular phenotypes that correlated with the level of severity of phenotypic expression. In general, we instead observed many cellular and molecular alterations that were shared by the cellular models derived from the affected individuals, while not being observed in those derived from the unaffected individuals. However, we did define some proband-specific DEGs, not present in the less severely affected sister, many of which relate to behavior and nervous system development. A subset of these DEGs had graded expression, exhibiting intermediate expression levels in the intermediate phenotype sister, between her unaffected mother and her severely affected brother. These findings suggest that both the degree of dysregulation of expression and the number and identity of DEGs within these networks may contribute to the level of affectation. While further experimentation might reveal additional graded phenotypes, particularly in mature neurons, ex vivo cellular modeling cannot recapitulate many aspects of fetal and post-natal neurodevelopment that may have been perturbed to contribute to the graded affectation observed in these individuals.
Limitations
This work highlights several considerations for ongoing scientific efforts to model this complex but prevalent form of ASD in future studies. First, since the unique characteristics of any multiplex ASD pedigree present challenges for cellular modeling, it is important to control for sex and variation in affectation in subject and family selection, study design, and analysis. Related to this point is the importance of modeling affected females in such studies. Most ASD cellular modeling to date has been restricted to affected males [
7‐
10,
13,
14,
16,
18,
19], given the increased prevalence of ASD among males, and the fact that constraint to a single sex simplifies some modeling considerations. In particular, sex chromosome dosage effects do not need to be accounted for in male cells, while female-derived iPSC models cannot currently recapitulate the process of random X-chromosome inactivation that occurs in developing somatic tissues, including the brain [
102,
103]. However, the transcriptomic differences that we observed here were not driven by sex chromosome-linked gene expression: very few DEGs in any potential pairwise sample comparison (whether between same or opposite sex models) were sex chromosome-linked and/or potential contributors to sex-biased gene expression in the human brain [
104,
105]. Therefore, this work supports the feasibility of identifying DEGs associated with affectation by cellular model cross-comparisons, even when these models are derived from both female and male subjects.
Furthermore, studying both unaffected and affected male and female individuals within a multiplex family, as in this pedigree, necessitates consideration of how the so-called female protective effect may contribute to affectation [
106‐
108]. Multiplex ASD families often exhibit differences in phenotypic expression of additive genetic liability, some of which appear to be related to sex. While the most affected individual in this family is male, we cannot fully differentiate the contributors to variable phenotypic expression of ASD in this pedigree, which also includes an unaffected male and affected and unaffected females. However, this experimental system could identify affectation-linked phenotypes that may have involved a sex-based contributor, since both the cellular and behavioral phenotypes of the females studied here differed from that of the affected male. Future studies in strategically-selected samples such as these provide an opportunity to assess how sex influences phenotypic expression of ASD genetic liability; if such mechanisms exist at the level of the cell, this approach may yield insights into sex-specific interventions.
Another consideration for iPSC-based modeling of ASD is genetic background, which can be a confounding variable for cross-comparisons [
42]. In this pedigree, ASD risk was polygenic, such that it was not possible to engineer a correction of a single genome variant to create pairs of isogenic mutant versus wild-type iPSC lines with an identical genetic background for study. In such cases, modeling of first degree relatives may serve as the best control, and modeling of multiple related individuals with varying affectation provides additional opportunities for identifying potential contributors to these differences in affectation. Including unrelated controls and performing comparisons with other studies can further highlight which phenotypic and transcriptomic alterations track with affectation, even by comparison with models derived from individuals with an unrelated genetic background.
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