Background
Oxytocin (OXT) is a nonapeptide hormone primarily produced by magnocellular neurons in the paraventricular and supraoptic nuclei of the hypothalamus [
1‐
6]. Oxytocin is secreted into the systemic circulation via the hypothalamic-neurohypophyseal system and transported to exert its peripheral effects, such as uterine contractions and milk ejection [
1‐
6]. Oxytocin is released into the brain from dendrites [
7] or centrally projected axonal terminals [
8], and it acts on various areas of the central nervous system, modulating affiliative and social behaviors, emotional states, and cognitive functions in many animal species [
1‐
6].
The actions of OXT are mediated by the OXT receptor (OXTR), which belongs to the GTP-binding protein-coupled receptor family [
9]. It is well established that the OXTR is coupled to heterotrimeric G
q/11 protein and that it stimulates phospholipase C
β (PLC
β), leading to the production of inositol 1,4,5-trisphosphate (IP
3) and 1,2-diacylglycerol (DAG) [
9]. Inositol 1,4,5-trisphosphate induces an increase in intracellular calcium concentration ([Ca
2+]
i) by triggering IP
3 receptor-mediated Ca
2+ release from intracellular stores, whereas DAG activates protein kinase C (PKC), which phosphorylates downstream signaling molecules [
9]. Upon prolonged agonist stimulation, OXTR undergoes desensitization and internalization through phosphorylation by GPCR kinase 2 (GRK2) and subsequent interaction with β-arrestin 2 [
9‐
13]. The internalized OXTRs are then recycled to the cell surface via vesicles containing small GTPases, Rab4 and Rab5 [
13].
Previous studies have shown that impaired OXTR-mediated signaling causes behavioral abnormalities [
2,
6,
14]. Mice deficient in the
OXTR gene show diminished social discrimination, increased aggressive behavior, reduced cognitive flexibility, and increased susceptibility to seizures [
15,
16]. Using homozygous mutant mice lacking CD38, a transmembrane protein with ADP-ribosyl cyclase activity, we have shown that a decrease in the formation of cyclic ADP-ribose (cADPR) results in dysfunction of Ca
2+-induced Ca
2+ release for OXT secretion in hypothalamic neurons. The reduction in cADPR levels also causes marked defects in maternal nurturing and social behaviors similar to those observed in
OXT- and
OXTR-knockout mice [
17,
18]. We also demonstrated that ADP-ribosyl cyclase activity increases after OXTR stimulation, regulating OXT release in the hypothalamus and pituitary in adult male mice [
19].
In humans, OXTR has been implicated in the pathogenesis of neuropsychiatric disorders and considered a potential target for therapeutic intervention [
20]. Gregory
et al. [
21] have reported a heterozygous deletion of the
OXTR gene, located on chromosome 3p25, in a patient with autism whose mother had a putative obsessive-compulsive disorder. In addition, many studies have demonstrated that single-nucleotide polymorphisms (SNPs) of the
OXTR gene are associated with autism spectrum disorders (ASDs), social anxiety disorders, and schizophrenia [
22‐
29]. However, all SNPs reported so far reside outside the protein-coding region, and their functional importance is unclear. If allelic variations leading to amino acid substitutions in the
OXTR gene were associated with these neuropsychiatric disorders, then such variations would help to predict the etiological relevance based on protein structure-function relationship and to introduce individualized OXT-based therapy.
Therefore, we analyzed nucleotide variations in the protein-coding regions of the human
OXTR gene in ASD patients and unrelated healthy controls. Furthermore, we focused on a triallelic variation (rs35062132; c.1126C> G/T; the nomenclature of variation recommended by the Human Genome Variation Society [
30]) and investigated whether the amino acid substitution R376G/C causes any changes in OXTR-mediated cellular responses.
Methods
Participants
We recruited 132 ASD subjects (102 males, 30 females; 15.9 ± 0.7 years) from the outpatient psychiatry department of the Kanazawa University Hospital. All subjects fulfilled the DSM-IV criteria for pervasive developmental disorder. The diagnoses were made by two experienced child psychiatrists through interviews and clinical record reviews as described previously [
31], and the subjects had no apparent physical anomalies. The two experienced child psychiatrists independently confirmed the diagnosis of ASD for all patients by semi-structured behavior observation and interviews with the subjects and their parents. During each of these interviews with parents, which were helpful in the evaluation of autism-specific behaviors and symptoms, the examiners used one of the following methods: the Asperger Syndrome Diagnostic Interview [
32], the Autism Diagnostic Interview-Revised (ADI-R)[
33], the Pervasive Developmental Disorders Autism Society Japan Rating Scale [
34], the Diagnostic Interview for Social and Communication Disorders [
35], or the Tokyo Autistic Behavior Scale [
36]. Based on these evaluation methods, 97 patients were classified as autistic disorder (autism), 10 as Asperger’s disorder, and 25 as pervasive developmental disorder not otherwise specified (PDD-NOS). The 248 controls (143 males, 105 females; 31.3 ± 0.6 years) were unrelated healthy Japanese volunteers. All patients and controls were Japanese with no non-Japanese parents or grandparents. This study was approved by the ethics committees of Kanazawa University School of Medicine. All examinations were performed after informed consent according to the Declaration of Helsinki.
Re-sequencing
Genomic DNA was extracted from venous blood samples using the Wizard Genomic DNA Purification kit (Promega, Madison, WI, USA), or from nails using the ISOHAIR DNA extraction kit (Nippon Gene, Tokyo, Japan). DNA fragments containing exons 3 and 4 of the hOXTR gene were amplified by PCR in a 20-μl reaction mixture containing 2.5 ng genomic DNA, 200 μM dNTPs, 200 nM of each primer, 4 μl Ampdirect G/C (Shimadzu, Kyoto, Japan), 4 μl Ampdirect-4 (Shimadzu), and 1 unit
Ex Taq DNA polymerase (Takara Shuzo, Otsu, Japan). The amplification procedure consisted of denaturation at 96°C for 1 min, followed by 35 cycles of 96°C for 30 s, 65°C for 1 min, and 72°C for 1 min. The primers used were FWD1 or FWD4 and REV1 for exon 3, and FWD5 and REV3 for exon 4 (sequences are given in Table
1). After amplification, PCR products were purified with the High Pure PCR Cleanup Micro kit (Roche, Mannheim, Germany), subjected to cycle sequencing reaction (BigDye version 1.1; Applied Biosystems, Foster City, CA, USA) according to the manufacturer’s protocol, and analyzed on an ABI PRISM 310 Genetic Analyzer (Applied Biosystems). The primers used were FWD1–FWD5 and REV1–REV6 (Table
1).
Table 1
Oligonucleotides used in this study
FWD1: TGGACTCAGCAGATCCGTCCG | FWD8: GCTCCGCCAGCTACCTGAAG | 376G-FAM: TGAGCCATGGCAGCTCC | R376G-FWD: CCTTTGTCCTGAGCCATG GCAGCTCCAGCCAGAGG | EGFP-FWD: CCGCAGGTGCACATCTTCTC |
FWD2: CTAAGCATCGCCGACCTGGT | REV7: TGGTGGGTCACGCCGTGGAT | 376R-VIC: TGAGCCATCGCAGCTCC | R376G-REV: CCTCTGGCTGGAGCTGCC ATGGCTCAGGACAAAGG | EGFP-REV: GTGGATCCCGCCGTGGATGG |
FWD3: CCGCAGGTGCACATCTTCTC | | 376C-FAM: TGAGCCATTGCAGCTCC | R376C-FWD: CCTTTGTCCTGAGCCATT GCAGCTCCAGCCAGAGG | |
FWD4: TGGAGTCTCCAGGAGTGGA | | | R376C-REV: CCTCTGGCTGGAGCTGCA ATGGCTCAGGACAAAGG | |
FWD5: GTCTGGAAGTGGCTCCAGTG | | | | |
REV1: CCTGGACATTCTGAGGCAGC | | | | |
REV2: GATGAGCTTGACGCTGCTGAC | | | | |
REV3: GTCAGCAGCGTCAAGCTCATC | | | | |
REV4: CAGTCGAAGACGCCGTC | | | | |
REV5: ACATGAGCAGCAGCAGG | | | | |
REV6: CAGGAGCAGGATGAGAC | | | | |
Real-time PCR
Two primers, FWD8 and REV7 (Table
1), were designed to amplify a 138-bp product encompassing rs35602132 at the mRNA position 1748 (g.8734707; c.1126). Three custom-designed TaqMan minor groove binder (MGB) probes were obtained from Applied Biosystems: 376G-FAM and 376C-FAM targeted to the variant alleles, and 376R-VIC to the common allele (Table
1). PCR was performed in 20-μl mixtures containing 10 μl TaqMan universal master mix (Applied Biosystems), 200 nM of each primer, 100 nM 376R-FAM, 100 nM 376G-VIC or 376C-VIC, and 10 ng of sample DNA. Thermocycling was performed on the ViiA 7 Real-Time PCR System (Applied Biosystems). The amplification was conducted at 60°C for 30 s, 95°C for 5 min, and 40 cycles of 95°C for 15 s and 60°C for 1 min. Data were analyzed with SDS2.3 software (Applied Biosystems).
Site-directed mutagenesis
cDNAs encoding hOXTR variants at amino acid residue 376 were generated by site-directed mutagenesis as described previously [
37]. pCHOXTR [
38] harboring hOXTR-376R, a common-type receptor, was used as a template for PCR; the primers used were R376G-FWD and R376G-REV (Table
1) for the R376G substitution, and R376C-FWD and R376C-REV (Table
1) for R376C. The amplified products were treated with
EX Taq DNA polymerase (Takara Shuzo) at 72°C for 10 min and cloned into a pCR2.1-TOPO vector (Invitrogen, Carlsbad, CA, USA) to yield pCHOXTR-376G and pCHOXTR-376C. The cDNA inserts were verified by nucleotide sequencing.
Construction of expression plasmids
The 1.4-kb
Eco RI fragments from pCHOXTR [
38], pCHOXTR-376G, and pCHOXTR-376C were cloned into the
Eco RI site of the mammalian expression plasmid pcDNA3.1 (+) (Invitrogen) to yield pcDNAHOXTR-376R, pcDNAHOXTR-376G, and pcDNAHOXTR-376C, respectively.
In some experiments, hOXTRs of the common type and variants were fused to the enhanced green fluorescent protein (EGFP). Expression plasmids for these fusion proteins were constructed essentially as described previously [
38]. In brief, the 1.4-kb
Eco RI fragments obtained from pCHOXTR [
38], pcDNAHOXTR-376G, and pcDNAHOXTR-376C were subjected to PCR using primers EGFP-FWD and EGFP-REV (Table
1). The amplified fragments were cleaved with
Pst I and
Bam HI. The resulting 0.46-kb
Pst I (1334)/
Bam HI (primer) fragments and the 0.78-kb
Bam HI (556)/
Pst I (1334) fragment from pCHOXTR were ligated to
Bam HI/
Bgl II-cleaved pEGFP-N3 (Clontech) to yield pHOXTR-376R-EGFP, pHOXTR-376G-EGFP, and pHOXT-R376C-EGFP. Restriction endonucleasesites are identified by numbers (in parentheses; in accordance with the data deposited in GenBank under accession number NM_000916) indicating the 5'-terminal nucleotide generated by cleavage.
Cell culture and transfection
Human embryonic kidney HEK-293 cells and COS-7 cells were maintained in DMEM supplemented with 10% FBS at 37°C in a humidified atmosphere of 95% air and 5% CO
2. NG108-15 neuroblastoma × glioma hybrid cells were cultured as described previously [
39]. Cells were grown in culture dishes to 80 to 90% confluence and transfected with expression plasmids using FuGENE HD Transfection Reagent (Roche) following the manufacturer’s instruction.
Radioligand binding assay
COS-7 cells transfected with pcDNAHOXTR-376R, pcDNAHOXTR-376G, pcDNAHOXTR-376C, or pcDNA3.1 (+) were collected 48 h after transfection, and resuspended in homogenization buffer (25 mM Tris, pH 7.4, 1 mM EDTA, 250 mM sucrose). The cells were homogenized and centrifuged at 1,500 g at 4°C for 10 min, to pellet nuclei and intact cells. The resulting supernatants were centrifuged at 40,000 g at 4°C for 30 min, and crude membrane pellets were suspended in binding buffer containing 50 mM HEPES (pH 7.4), 5 mM MgCl2, 1 mM CaCl2, and 0.2% BSA. Protein concentration was determined by the bicinchoninic acid assay (Pierce, Rockford, IL, USA) using BSA as a standard. [Tyrosyl-3H]-oxytocin ([3H]-OXT; PerkinElmer Life Sciences, Waltham, MA, USA) was diluted in the binding buffer to concentrations of 0.05 to 4 nM. Specific binding assays were performed in 5-μg cell membrane preparations incubated at room temperature for 1 h with increasing concentrations of [3H]-OXT, in the absence or presence of 200-fold excess of unlabeled OXT. Binding reaction mixtures were filtered through a GF/C glass fiber filter (Whatman, Maidstone, Kent, UK) and washed three times with Wash Buffer (50 mM HEPES, pH 7.4, 500 mM NaCl, 0.1% BSA). Radioactivity associated with membranes retained by glass filters was quantified in a liquid scintillation counter (LSC-5100; Aloka, Tokyo, Japan). Specific binding was calculated by subtracting non-specific binding in the presence of 200-fold excess of OXT at each radioligand concentration from total binding. Affinity (Kd) and maximal binding capacity (Bmax) of saturation binding were obtained from saturation isotherm specific binding data by nonlinear regression curve analysis using the standard equation for a rectangular hyperbola fitted to one site with the GraphPad Prism 5 (GraphPad Software Inc., San Diego, CA, USA). Competition binding experiments were performed in duplicate by incubating 5 μg of cell membranes at room temperature for 1 h, in the same medium with 1 nM [3H]-OXT and increasing concentrations of unlabeled OXT (10−12 to 10−5 M). Log IC50 values were derived from nonlinear least-squares analysis using the GraphPad Prism 5 software (GraphPad Software Inc.).
Measurement of receptor recycling
HEK-293 cells were plated on poly-D-ornithine-coated glass coverslips and cultured overnight. After transfection with pcDNAHOXTR-376R, pcDNAHOXTR-376G, or pcDNAHOXTR-376C, the cells were incubated at 37°C in serum-free DMEM for different times, in the absence or presence of OXT (100 nM). Reactions were stopped by removing the medium and fixing the cells with 4% paraformaldehyde at room temperature for 10 min. The cells were then blocked with PBS containing 1% BSA and 1.5% normal horse serum for 30 min, and incubated with goat anti-OXTR antibody (N-19; 1:200; sc-8103, Santa Cruz Biotechnology, Santa Cruz, CA, USA) at 4°C overnight. After washing in PBS, the cells were incubated with donkey anti-goat IgG (H + L) antibody conjugated with Alexa Fluor 488 (1:500; Invitrogen) combined with 4',6-diamidino-2-phenylindole (DAPI; 1 μg/ml; Dojindo, Kumamoto, Japan). In some experiments, pSNAPf-ADRβ2 (New England Biolabs, Ipswich, MA, USA), an expression plasmid for β2 adrenergic receptor fused to SNAPf, was cotransfected into HEK-293 cells. The cells were labeled with SNAP-Surface Alexa Fluor 488 (2 μM; New England Biolabs), fixed with 4% paraformaldehyde, and reacted with goat anti-OXTR antibody (N-19; 1:200; Santa Cruz) at 4°C overnight, followed by incubation with donkey anti-goat IgG (H + L) antibody conjugated with Alexa Fluor 594 (1:500; Invitrogen); none of the solutions contained any cell-permeabilizing reagents. Fluorescent images were obtained using confocal laser scanning microscopes (LSM5 Pascal; Carl Zeiss, Jena, Germany; FluoView FV10i, Olympus). Data were analyzed using the FV10-ASW software (Olympus).
[Ca2+]i measurement
We measured [Ca
2+]
i using the fluorescent Ca
2+ indicator fura-2-acetoxymethyl ester (fura-2/AM). HEK-293 cells or NG108-15 cells transfected with pHOXTR-376R-EGFP, pHOXTR-376G-EGFP, or pHOXTR-376C-EGFP were loaded with fura-2/AM to a final concentration of 3 μM in complete medium and incubated at 37°C. After 30-min loading, the cells were washed three times with HEPES-buffered saline (HBS) solution (145 mM NaCl, 5 mM KCl, 1 mM MgCl
2, 20 mM HEPES-NaOH, 2 mM CaCl
2, 20 mM glucose, pH 7.4). Cells expressing EGFP-tagged OXTRs were visualized at a wavelength of 488 nm before OXT application. The fluorescence of the cells loaded with fura-2/AM was then measured at 37°C, at the determined sites, through a pinhole (10 to 20 μm in diameter). We used alternating excitation wavelengths of 340 and 380 nm in a Ca
2+ microspectrofluorometric system (OSP-3 Model; Olympus, Tokyo, Japan), as described previously [
39]. The Ca
2+ emission was detected every 10 s for 5 min after OXT application. The ratio of fluorescence at 340 nm and 380 nm (
F340/F380) was used to determine [Ca
2+]
i. All data were normalized to the baseline fluorescence (
F0) recorded 10 s before OXT addition, and given as
F/F0.
IP3 accumulation assay
HEK-293 cells were plated at a density of approximately 5 × 104 per well in a 24-well plate, and transiently transfected with pcDNAHOXTR-376R, pcDNAHOXTR-376G or pcDNAHOXTR-376C, or pcDNA3.1(+). Before treatment with OXT, the transfected cells were washed and incubated in HBS containing 2 mM CaCl2, 10 mM glucose, and 10 mM LiCl at 37°C for 10 min. After OXT treatment, the reaction was terminated at designated time points by the addition of 50 μl of 10% (w/v) perchloric acid. The acid-soluble extract was neutralized with 150 μl of 1.53 M KOH-75 nM HEPES, and perchloric acid was precipitated on ice and removed by a brief centrifugation. The IP3 aqueous extract was then examined with an Inositol-1,4,5-triphosphate [3H]-Radio Receptor Assay Kit (PerkinElmer Life Sciences).
Statistical analyses
Data obtained from genetic studies were analyzed using a contingency table and the Fisher’s exact test (GraphPad Prism 5; GraphPad Software Inc.). All data from the in vitro studies are shown as mean ± standard error of the mean. Statistical significance was determined by the Student’s t test or two-way analysis of variance (ANOVA) using an interactive fitting program (GraphPad Prism 5; GraphPad Software Inc.); P values smaller than 0.05 were considered to be statistically significant.
Discussion
The main findings of this study are as follows. (i) Triallelic non-synonymous variation (rs35062132, c.1126C>G/T; R376G/C) is observed in both ASD patients and healthy controls in a Japanese population; the frequencies of the G allele are significantly higher in the ASD patients than in healthy controls. (ii) In HEK-293 cells expressing hOXTR-376G, the agonist-induced internalization and recycling of the OXTR are faster than that in the cells expressing hOXTR-376C or hOXTR-376R. (iii) In both HEK-293 cells and NG108-15 neuronal cells, the agonist-induced increase in [Ca2+]i mediated by hOXTR-376G-EGFP or hOXTR-376C-EGFP is smaller than the increase associated with hOXTR-376R-EGFP. These results provide new insights into the genetic architecture and therapeutic aspects of ASDs.
The present study is unique in associating non-synonymous allelic variations of the
OXTR gene with ASDs. These variations at rs35062132 have not been reported in other disorders. To date, several studies have found ASD-associated SNPs in the
OXTR gene, which reside in introns (rs4564970 [
41], rs237897 [
25], rs53576 [
23,
24], rs2254298 [
23‐
25], rs2268493 [
42], rs7632287 [
42], rs11720238 [
41]), 3'-untranslated region (rs1042778) [
42], and intergenic regions (rs7632287 and rs11720238) [
41]. The functional consequences of these SNPs remain unclear. Importantly, all minor alleles of these SNPs have a frequency more than 5%, and thus can be classified as common variations. In contrast, the frequencies of the G and T alleles of rs35062132 are 0.4% and 0.6%, respectively, in controls; these minor alleles can be categorized as rare [
43‐
45]. Therefore, most previous studies favor the common disease–common variant model, in which most of the risk is caused by common genetic variations (>5% allele frequency), each allele conferring a slight risk [
43‐
47]. By contrast, our results support the common disease–rare variant model, in which the risk is mostly attributable to rare variations (<5% allele frequency), each variant conferring a moderate but readily detectable increase in the relative risk [
43‐
47]. In agreement with this model, we observed that the minor alleles clearly affected OXT-induced cellular responses. Future studies will be conducted to test whether behavioral changes caused by these variations are moderate and reasonably deleterious. Also it will be interesting to examine whether the rs35062132 variations cosegregate with the risk alleles in the non-coding region [
22‐
24,
41,
42].
The limitation of this study is that sample size is not sufficient to analyze the rare variant alleles at rs35062132. Future studies with larger sample size or family-based association testing are necessary to conclude that the G allele of the rs35062132 is a genetic risk factor for ASDs. The result reported here should be replicated in independent populations with various ethnic backgrounds.
The amino acid variation R376G/C is located in the intracellular carboxy (
C)-terminal tail of the receptor protein, which is critical for desensitization, internalization, and recycling of the OXTR, as in many other GPCRs [
9‐
13]. Arg
376 is associated with two structural components involved in these processes: (i) Arg
376 is a part of a PKC consensus motif (Ser
374-His
375-Arg
376; Figure
1A), which is thought to interact with PKC after OXT stimulation [
10]; and (ii) Arg
376 is in conjunction with one of the two Ser triplets in the
C-terminus (Figure
1A) that are primary sites of agonist-induced phosphorylation and β-arrestin-2 binding [
11]. When either one of the two Ser triplets was mutated to alanine residues, the stability of the β-arrestin-2-OXTR complex was altered, and the ability of β-arrestin-2 to internalize with the receptor was eliminated [
11]. In the vasopressin receptor V2, which is highly homologous to OXTR in structure, the equivalent Ser triplets function as a retention motif. GRK phosphorylation of these Ser residues promotes the formation of stable receptor-β-arrestin complexes, followed by cotrafficking and colocalization of the complexes to endocytotic vesicles and slow recycling to the cell surface [
48]. Therefore, it is suggested that the R376G substitution in the OXTR might suppress the phosphorylation of Ser residues in the Ser triplets or reduce the stability of β-arrestin binding to the triplets, resulting in faster receptor recycling.
We demonstrated that, both in HEK-293 cells and in NG108-15 neuronal cells, agonist-induced [Ca
2+]
i elevation mediated by variant receptors, hOXTR-376G-EGFP and hOXTR-376C-EGFP, is smaller than that by the common receptor hOXTR-376R-EGFP. Despite the marked reduction in the peak [Ca
2+]
i, the decrease in IP
3 levels was slight. Provided that Arg
376 is not located in the known site for G
q binding (Figure
1A) [
49], the G
q/11/PLC
β/IP
3-independent, unidentified signaling pathway for [Ca
2+]
i elevation might be affected by the R376G/C substitution.
Ca
2+ signaling pathways in neurons have been implicated in the pathogenesis of ASDs [
50]. However, the involvement of OXTR-mediated, PLC/IP
3-dependent Ca
2+ signaling in ASDs is still largely unknown. Recently, Ninan [
51] has demonstrated that U73122, a PLC inhibitor, reduced OXT-induced suppression of glutamatergic synaptic transmission in pyramidal neurons of the medial prefrontal cortex, possibly by inhibiting PLC-dependent increase in postsynaptic calcium. Given that this process is critical for the OXT effects on social cognition, it is conceivable that the OXTR variations may serve as a risk factor for ASDs.
We have previously demonstrated that cADPR, which may act as an intracellular second messenger downstream of the OXTR, activates Ca
2+ release from intracellular stores through the ryanodine receptor Ca
2+ release channel. cADPR also initiates Ca
2+ influx through melastatin-related transient receptor potential 2 (TRPM2) channels [
38]. Our previous SNP analysis has suggested that R140W substitution of CD38 protein, a regulator of ryanodine receptor-mediated Ca
2+-induced Ca
2+ release for OXT secretion [
17], could be a potential risk factor for a subset of Japanese ASD patients [
31]. Although this study shares a part of the ASD patients with our previous analysis [
31], the 140W allele of CD38 has not been found in the patients carrying the OXTR-376G or OXTR-376C until now (unpublished data). It is therefore conceivable that the underlying signaling process affected by the R376G/C substitution is independent of the CD38-mediated OXT release from hypothalamic neurons [
17].
Besides their relevance to the etiology of ASDs, the results presented here might contribute to the development of new pharmacological treatments. Genetic polymorphisms of many drug targets predict responsiveness to drugs [
52,
53]. It is likely that the alteration in the cellular functions mediated by the variant OXTRs could cause some individual differences in both behavioral and non-behavioral responses to OXT. Although no side effects specific to intranasal OXT administration have been reported [
54], new clinical information regarding functional OXTR variants will be helpful in the determination of individual administration protocols to maximize therapeutic benefit with least adverse effects.
Competing interests
The authors declare they have no competing interests.
Authors’ contributions
HH and SY conceived and designed the research. WJM, TM, KH, KY, MY, and SY performed the genetic analysis of the ASD patients and healthy volunteers. WJM, MH, and SY carried out the experiments on cultured cells. WJM, HH, and SY analyzed data. WJM and SY drafted the initial manuscript; and WJM, TM, HH, and SY prepared successive versions of the document. All authors read and approved the final manuscript.