Introduction

Obesity is an increasing global concern: 20% of the world’s adult population is expected to become obese by 2025 if current trends persist (Collaboration, 2017). However, the complex environmental, behavioural, and genetic contributions to obesity make this a difficult condition to model and address. Monogenic obesity disorders provide a consistent system where the pathophysiology, genetic predisposition, and neurobiology underlying phenotypes can potentially provide insights that translate to polygenic obesity (Loos & Yeo, 2022).

One such monogenic disorder is Smith-Magenis syndrome (SMS; OMIM #182290) (Smith et al, 1986), a neurogenetic condition frequently associated with obesity. 80% of SMS cases are caused by interstitial deletions of the 17p11.2 chromosomal region containing a dosage-sensitive gene, retinoic acid-induced 1 (RAI1) (Slager et al, 2003). More than 90% of SMS patients are overweight (40% > 95^th percentile weight, 40% > 85^th percentile weight) and exhibit over-eating tendencies by adolescence (Alaimo et al, 2015; Burns et al, 2010; Gandhi et al, 2022). 20% of SMS patients do not have 17p11.2 deletions but instead carry RAI1 point mutations that are still frequently associated with obesity (Boot et al, 2021; Edelman et al, 2007). These clinical data indicate that RAI is critical for energy homeostasis underlying body weight regulation.

The human RAI1 protein shares >80% sequence identity with the mouse Rai1 protein, allowing us to study Rai1 function in vivo (Huang et al, 2016; Javed et al, 2020). We previously found that Rai1 encodes a transcriptional regulator that binds to the promoter or enhancer regions of its target genes, most of which are involved in neurodevelopment and synaptic transmission (Huang et al., 2016). SMS (Rai1^+/-) mice exhibit obesity associated with hyperphagia and increased adiposity (Burns et al., 2010; Huang et al., 2016). Interestingly, Rai1 may have a sexually dimorphic function, as female SMS mice exhibit more pronounced obesity than males (Burns et al., 2010; Huang et al., 2016).

Using a homozygous Rai1 conditional knockout (Rai1-cKO) mice, we found that Rai1 deletion in either subcortical excitatory neurons (vGlut2⁺) or single-minded homolog 1 (Sim1⁺) neurons induced obesity in mice (Huang et al., 2016). Sim1⁺ neurons reside in several brain regions including the nucleus of the lateral olfactory tract, the supraoptic nucleus, the medial amygdala, and the paraventricular nucleus of the hypothalamus (PVH), a region that is critical for body weight regulation (Balthasar et al, 2005). Sim1⁺ neurons in the PVH are composed of several molecularly defined cell types including those that express oxytocin, vasopressin, corticotropin-releasing hormone, somatostatin, melanocortin-4-receptor (MC4R), growth hormone-releasing hormone (GHRH), or brain-derived neurotrophic factor (Bdnf) (An et al, 2015). Deleting Bdnf from the PVH results in hyperphagic obesity and reduced locomotion (An et al., 2015). While PVH neuronal activity is reduced during feeding (Xu et al, 2019), activation of PVH^Bdnf neurons results in weight loss (Wu & Xu, 2022). We recently found that Rai1 promotes Bdnf expression in hypothalamus throughout life (Javed et al, 2021). We hypothesized that Rai1 may be connected to Bdnf-producing neurons in PVH, explaining how a neural transcription factor regulates body weight homeostasis.

Bdnf regulates energy metabolism by binding to its cognate receptor tropomyosin receptor kinase B (TrkB) (An et al., 2015) and may thus provide a potential therapeutic target for SMS. However, the link between Rai1, Bdnf, and SMS symptoms is currently unclear. Patients carrying monogenic mutations in either BDNF (Gray et al, 2006) or NTRK2 (encoding TrkB) (Yeo et al, 2004) experience severe hyperphagic obesity. Moreover, obesity and associated metabolic dysfunction such as impaired glucose and insulin sensitivity are correlated with impaired PI3K-Akt pathway, a TrkB downstream target (Su et al, 2021). Given that (1) Bdnf overexpression during early embryonic development or (2) targeting Bdnf overexpression to the PVH of adolescent SMS (Rai1^+/-) mice protected them from weight gain and metabolic defects (Javed et al., 2021), and that (3) a point mutation in RAI1 disrupts its ability to transcriptionally enhance Bdnf expression in vitro (Abad et al, 2018), pharmacological targeting of Bdnf signaling is an attractive and yet unexplored therapeutic avenue for SMS.

Building on our previous discoveries, here we examine the neurobiological functions of Rai1 in Bdnf-producing neurons, specifically those that reside in the PVH. Importantly, we evaluated the involvement of the Bdnf-TrkB pathways in SMS pathogenesis using a small molecule drug (LM22A-4) to enhance TrkB signalling. Our work indicates that PVH^Bdnf neurons depend on Rai1 to maintain normal neuronal activity and regulate body weight homeostasis, and TrkB signalling dysfunction contributes to obesity associated with SMS mice.

Results

Rai1 deficiency dysregulates specific downstream targets of neurotrophin in the mouse hypothalamus

Multiple lines of evidence indicated that SMS mice show decreased Bdnf mRNA expression in the hypothalamus (Burns et al., 2010; Huang et al., 2016; Javed et al., 2021). To determine if Bdnf protein levels were consequently altered in the hypothalamus, we performed enzyme-linked immunosorbent assay (ELISA) and found that Bdnf protein levels were significantly reduced in 7-week-old pre-symptomatic SMS mice relative to controls (Fig 1A). To quantify if molecular pathways associated with Bdnf-TrkB signalling were altered, we subjected hypothalamic tissue lysates from 7-week-old pre-obese SMS and control (Ctrl) mice to reverse phase protein array (RPPA) (Fig 1B). We found that control and SMS samples clustered separately in principle component analysis (Fig 1C).

Figure 1:

Hierarchical clustering analysis indicated that the majority of differentially expressed phosphorylated proteins (Fig 1D) and total proteins (Figure 1—figure supplement 1) were downregulated in SMS mice. We mapped all proteins, as well as phospho-proteins, that were differentially expressed in Ctrl and SMS samples onto the interconnected biological network and identified a subset of connected nodes downregulated in SMS (FDR < 10e-5) belong to the neurotrophin, PI3K-Akt, and mTOR signalling pathways (Fig 1E). These pathways are known to play a critical role in regulating body weight homeostasis and can be activated by Bdnf either directly (neurotrophin pathway) or indirectly (PI3K-Akt and mTOR pathways) (Cota et al, 2006; Schultze et al, 2012). The expression levels of PIK3CA, a common node in these pathways, were significantly reduced in the SMS mice compared to controls (Fig 1F). Moreover, the expression levels of phosphorylated NF-κB (nuclear factor-kappa B), a downstream target of the neurotrophin pathway, were significantly reduced in the SMS group (Fig 1G). Together, these results suggest that Rai1 loss impairs specific pathways, including neurotrophin signalling, and indicate that dysfunction of Bdnf-producing cells could contribute to SMS pathogenesis.

Bdnf-producing cells depend on Rai1 to regulate body weight homeostasis

To target Bdnf-producing cells, we used a Bdnf^Cre/+mouse line that carries a viral 2A oligopeptide and Cre recombinase immediately upstream of Bdnf’s endogenous STOP codon (Tan et al, 2016). We crossed the Bdnf^Cre/+ mice with tdTomato-Ai9 reporter mice and found that Ai9 signals faithfully reflected endogenous Bdnf protein expression in the PVH (Figure 2— figure supplement 1A-E). More than 50% of Bdnf-producing cells within the PVH were positive for phosphorylated TrkB (p-TrkB) (Figure 2—figure supplement 1F). Moreover, 20% of p-TrkB⁺ PVH cells express Bdnf, suggesting that Bdnf could have an autocrine mode of action in this cell type (Figure 2—figure supplement 1F-J). We also found that Rai1 is expressed in PVH^Bdnf cells that are distinct from larger magnocellular cells such as the PVH oxytocin neurons (Figure 2—figure supplement 2A-I). Specifically, 55% of PVH^Bdnf neurons express Rai1, whereas only 2% of PVH^Oxytocin neurons express Rai1 (Figure 2—figure supplement 2I). These data suggest that Rai1 is enriched in Bdnf-producing neurons in the mouse PVH.

To determine if Bdnf-producing cells depend on Rai1 to regulate body weight, we deleted one or both copies of Rai1 using the Bdnf^Cre/+ and the conditional Rai1^fl/fl mice (Huang et al., 2016) (Fig 2A, Figure 2—figure supplement 3A). Conditional knockout mice (cKO: Bdnf^Cre/+; Rai1^fl/fl) showed a significant reduction in Rai1 expression in PVH^Bdnf neurons compared to controls (Fig 2B-C, Figure 2—figure supplement 3B). The total number of Bdnf-producing cells in the PVH remained similar in Ctrl and cKO mice (Figure 2—figure supplement 3C), indicating that Rai1 loss did not affect PVH neurogenesis or survival.

Figure 2:

We monitored body weight weekly and found that male and female cKO mice showed a significant increase in body weight beginning at 15 weeks of age when compared to controls (Ctrl: Bdnf^Cre/+) (Fig 2D, Figure 2—figure supplement 4A). However, conditional Rai1 heterozygous mice (Bdnf^Cre/+; Rai1^fl/+) did not show a pathological increase in body weight (Figure 2—figure supplement 3D, Figure 2—figure supplement 4B), suggesting that non-Bdnf-producing cells also contribute to SMS pathogenesis. EcoMRI found that 26-week-old cKO male and female mice showed an increased proportion of fat mass but not lean mass (Fig 2E, Figure 2—figure supplement 4C). Further dissection of fat tissues from different organs showed an increased mass of subcutaneous inguinal (S.C. Ing.) fat in cKO mice of both sexes and increased mass of epididymal adipose tissues (eWAT) in female cKOs (Fig 2F, Figure 2— figure supplement 4D).

In contrast, the total mass of brown adipose tissue (BAT) remained unchanged between groups (Fig 2F, Figure 2—figure supplement 4D). In both male and female cKO mice, we found hypertrophic eWAT adipocytes (Fig 2G, Figure 2—figure supplement 4E) and elevated blood leptin levels (Fig 2H, Figure 2—figure supplement 4F). However, there was no significant alteration in blood lipids such as triglycerides (TG), high-density lipoprotein (HDL), low-density lipoproteins (LDL), and very low-density lipoprotein (VLDL) in either male or female cKO mice (Figure 2—figure supplement 3E-G, Figure 2—figure supplement 4G-I). Altogether, these data suggest that obesity caused by ablating Rai1 from Bdnf-producing cells is associated with increased fat mass, fat deposition, and leptin levels.

Obesity could result from increased food intake, reduced energy expenditure, or both. Similar to SMS mice, male and female cKO mice did not have defects in respiratory exchange rates when measured by indirect calorimetry (Figure 2—figure supplement 3H, Figure 2—figure supplement 4J). Interestingly, cKO mice showed sex-specific differences in energy expenditure. During the dark phase, female cKO mice showed a reduction in energy expenditure (Fig 2I) and decrease in total locomotor activity (Fig 2J). In contrast, total food intake remained unchanged (Fig 2K). Male cKO mice became hyperphagic (Figure 2—figure supplement 4K), showed increased energy expenditure during the dark phase (Figure 2—figure supplement 4L), and exhibited normal locomotor activity when compared to controls (Figure 2—figure supplement 4M). These results suggest that obesity in male cKO mice is attributable to overeating despite an increase in energy expenditure.

To determine if Rai1 loss from Bdnf-producing cells impacts metabolic function, a glucose tolerance test (GTT) was performed. Interestingly, cKO mice of both sexes showed normal blood glucose levels right before the test but became hyperglycemic 20 minutes (female cKO) and 45 minutes (male cKO) after glucose administration (Fig 2L, Figure 2—figure supplement 4N). This change was accompanied by a significantly higher area under the curve (AUC) for both groups (Figure 2—figure supplement 3I, Figure 2—figure supplement 4O). While the male cKO mice remained hyperglycemic throughout the GTT (Figure 2—figure supplement 4P), the glucose levels in female cKO mice returned to normal towards the end (Fig 2L). We also found increased blood insulin levels during the GTT in both sexes (Figure 2—figure supplement 3J, Figure 2—figure supplement 4P). In the insulin tolerance test (ITT), female cKO mice showed moderately elevated glucose levels 20 minutes into the test (Figure 2—figure supplement 3K). By contrast, male cKO mice showed considerably higher glucose levels than controls in the ITT (∼6.5 ng/ml for cKO and ∼2.0ng/ml for Ctrl) during the 60- to 120-minute intervals (Figure 2— figure supplement 4Q). Altogether, these experiments demonstrate that Bdnf-producing cells depend on Rai1 to regulate body weight, adiposity, glucose and insulin tolerance in mice.

Rai1 deletion reduces PVH^Bdnf neuronal excitability

Bdnf-producing neurons are widely distributed in the hypothalamus, being found in the dorsomedial hypothalamus (Unger et al, 2007), ventromedial hypothalamus (Xu et al, 2003), and PVH (An et al., 2015). To understand how Rai1 loss affects intrinsic neuronal excitability and synaptic transmission of PVH^Bdnf neurons that underlie energy homeostasis (An et al., 2015), we fluorescently labelled them with Cre-dependent tdTomato (Fig. 3A). We found that a hyperpolarized step followed by increasing the holding voltage at resting membrane potentials (Rm) was able to initiate rebound firing or rebound repetitive firing in most PVH^Bdnf neurons (Figure 3—figure supplement 1A). Hyperpolarization-induced I_H currents and low depolarization events were observed in some PVH^Bdnf neurons (Figure 3—figure supplement 1A). cKO PVH^Bdnf neurons showed reduced action potential (AP) firing frequency at a holding voltage of -60mV compared to Ctrl PVH^Bdnf neurons (Fig 3B-C). The resting membrane potential (Rm) of cKO (-52mV) neurons showed a 10mV positive shift compared to Ctrl neurons (-62mV) (Fig 3D). We also found that fewer cKO PVH^Bdnf neurons showed spontaneous firing at Rm (Fig 3E, Ctrl:10/13; cKO:5/12) and at holding voltage of -70mV (Fig 3F, Ctrl:6/12; cKO:1/12) compared to Ctrl PVH^Bdnf neurons. The threshold of the first AP initiated by a ramp current injection (1 nA/sec) from Rm was significantly increased in cKO neurons (Fig 3G-H). Because the higher the holding voltage is, the narrower the shoulder and the lower the amplitude of APs are (Figure 3—figure supplement 1B), we compared the AP waveforms at a holding voltage of -60mV, close to the Rm of both genotypes. The AP of the cKO group exhibited a lower peak amplitude (measuring from the threshold to peak) (Fig 3I), a normal threshold (Fig 3J), and half-width (Figure 3—figure supplement 1C). We found that both passive and AP properties of cKO PVH^Bdnf neurons were altered. The Rm of cKO neurons shifted to a more positive voltage than the gating threshold of voltage-gated sodium channels (6/12 were more positive than -55mV), which was unfavourable for their transition from the inactivation state to the closed state. Combined with the higher AP threshold of cKO neurons, these changes contribute to a decreased excitability in cKO PVH^Bdnf neurons.

Figure 3:

PVN neurons receive glutamatergic and GABAergic inputs and are tonically inhibited by GABAergic neurons (Colmers & Bains, 2018; Gao et al, 2017). The rightward shift of the probability of miniature inhibitory postsynaptic current (mIPSC) frequency indicated that cKO PVH^Bdnf neurons received a higher frequency of inhibitory inputs, although the average frequency (Fig 3K) was not significantly different between groups. The probability of mIPSC amplitude also showed a right shift, suggesting an increased mIPSC postsynaptic response (Fig 3L, Figure 3—figure supplement 1D). Furthermore, we observe an increased area under the curve (Fig 3M), indicating that Rai1 loss induced a larger total outward current upon the inhibitory inputs. Interestingly, we found that cKO PVH^Bdnf neurons had a smaller cellular diameter (Figure 3—figure supplement 1E), indicating that the increased inhibitory currents were unlikely due to changes in passive membrane properties associated with increased cell size. Overall, our data argue that Rai1 deletion decreases the excitability of PVH^Bdnf neurons and likely increases pre-synaptic inhibitory transmission and post-synaptic inhibitory responses, resulting in decreased AP firing.

Rai1 expression is necessary for PVH^Bdnf neurons to regulate energy homeostasis

Reduced PVH^Bdnf neuronal activity in cKO mice suggested that Rai1 loss in Bdnf-producing PVH neurons might be sufficient to induce defective energy homeostasis in vivo. To test this hypothesis by deleting Rai1 only in PVH^Bdnf neurons but not other Bdnf-producing cells, we performed PVH^Bdnf-specific Rai1 deletion using a CRISPR/Cas9 system packaged into AAV9. Two viruses were delivered; one encoding a reporter molecule (GFP) and three single guide RNAs targeting the Rai1 gene (sgRai1), and the other carrying Cre-dependent Staphylococcus aureus CRISPR-associated protein 9 (saCas9) (Fig 4A). Hereafter, our experiments focus on female SMS mice since they showed more consistent defects in fat distribution (Burns et al., 2010) and body weight regulation (Huang et al., 2016) than males. Bilateral administration of two AAVs into the PVH of female Bdnf^Cre/+mice (Exp group) resulted in a significant loss of Rai1 protein expression in PVH^Bdnf neurons when compared to Bdnf^Cre/+ mice injected with an AAV encoding the sgRai1 and GFP (Ctrl group) (Fig 4B-C, Figure 4—figure supplement 1A). Quantification of AAV-infected (GFP⁺) Bdnf neurons showed that while the total number of Bdnf⁺ (Figure 4—figure supplement 1B) and Bdnf⁺ GFP⁺ (Figure 4—figure supplement 1C) cells was not different in the two groups, the percentage of AAV-infected Bdnf⁺ cells that express Rai1 was significantly reduced in the Exp group (Fig 4C, Figure 4—figure supplement 1D-E). Deleting Rai1 from the PVH^Bdnf neurons resulted in a significant increase in body weight beginning at 17 weeks of age (Fig 4D). Similar to the cKO (Bdnf^Cre/+; Rai1^fl/fl) mice, obesity in 26-week-old Exp mice was due to increased fat mass (Fig 4E) and more specifically, increased subcutaneous inguinal (S.C. ing) and epididymal white adipose tissues (eWAT) but not BAT (Fig 4F). We also found that eWAT adipocytes became hypertrophic (Fig 4G). Similar to the cKO mice, food intake did not differ between Exp and Ctrl groups (Fig 4H), likely due to the high level of satiety signals in the brain caused by increased blood leptin levels (Fig 4I). We also found that energy expenditure (Fig 4J), respiratory exchange rate (Figure 4—figure supplement 1F), and blood lipid parameters, including TG (Figure 4—figure supplement 1G), HDL (Figure 4—figure supplement 1H), LDL and VLDL levels (Figure 4—figure supplement 1I), were similar in Exp and Ctrl groups, consistent with the cKO data (Figure 4— figure supplement 14). Interestingly, Exp mice showed significantly less locomotor activity during the dark phase than the Ctrl mice (Fig 4K). A GTT indicated that Exp mice have a hyperglycemic profile (Fig 4L), accompanied by a slightly higher area under the curve for blood glucose levels (p=0.05) (Fig 4M). Insulin levels in Exp mice were also significantly elevated 15 minutes after glucose administration (Fig 4N).

Figure 4:

While Exp mice showed a trend towards increased glucose levels in insulin tolerance tests, the change was not statistically different (Figure 4—figure supplement 1J). Altogether, these experiments indicate that conditional deletion of Rai1 from PVH^Bdnf neurons during adolescence impairs the regulation of body weight, adiposity, glucose tolerance, and locomotion. These changes recapitulate most phenotypes observed in cKO mice with Rai1 deleted in Bdnf-producing cells from early development.

Pharmacologically enhancing TrkB signalling partially alleviates obesity in SMS mice

Our findings so far indicate that multiple pathways, including neurotrophin signalling, are dysregulated in SMS mice, and that Rai1 loss from Bdnf-producing cells, including PVH^Bdnf neurons, impairs energy homeostasis in vivo. Next we sought to examine the contribution of Bdnf-TrkB dysfunction in SMS pathogenesis by pharmacologically activating downstream targets of the Bdnf-TrkB pathway.

We systematically treated mice with LM22A-4, a small molecule Bdnf loop domain mimetic (Massa et al, 2010). To measure the bioavailability of LM22A-4 in the SMS brain, we simultaneously administered LM22A-4 intranasally (5 mg/kg body weight) and intraperitoneally (50 mg/kg body weight), using the highest doses used in previous studies (Canals et al, 2004; Chang et al, 2006; Gu et al, 2022; Kron et al, 2012; Ogier et al, 2007). We then harvested forebrain and hypothalamic tissues one- or three-hours after drug administration. Spectrometric analysis found a significantly higher LM22A-4 concentration in both brain regions one-hour post-treatment, while its concentration decreased three hours after treatment (Figure 5—figure supplement 1A). Consistent with the reduced neurotrophin signalling observed in our proteomic analysis, saline-treated SMS mice showed reduced phosphorylated-Akt (p-Akt; Fig 4A, Figure 5—figure supplement 1B) levels in the hypothalamus. LM22A-4 treatment significantly increased the levels of p-Akt in the SMS mice 1-hr post-treatment (Fig 4A, Figure 5—figure supplement 1B), indicating increased signalling downstream of the Bdnf-TrkB pathway.

We next examined if chronic LM22A-4 treatment was sufficient to reverse or modify the progression of the obesity phenotype in adult SMS mice. Specifically, we treated SMS mice daily from 8 to 14 weeks of age, during which SMS mice began to gain significantly more weight. Saline-treated SMS mice (Fig 5C) were significantly heavier than saline-treated control mice from 9 weeks of age onward. Interestingly, LM22A-4-treated SMS mice showed a similar body weight to saline-treated control littermates until 13 weeks of age but became obese by 14 weeks of age (Fig 5D). Consistent with a partial rescue, LM22A-4-treated SMS mice were significantly less obese than saline-treated SMS mice throughout the treatment period (Fig 5E). Control mice treated with LM22A-4 did not show a further reduction in body weight (Figure 5—figure supplement 1C), indicating that the therapeutic effects were specific to SMS pathogenesis.

Figure 5:

We measured food intake and found that LM22A-4 did not significantly rescue increased food intake in SMS mice. (Fig 5F). SMS mice showed normal energy expenditure (Figure 5—figure supplement 1D) and respiratory exchange rate (Figure 5—figure supplement 1E), neither of which was altered by LM22A-4 treatment. Interestingly, we did observe a partial rescue of locomotor activity in LM22A-4-treated SMS mice that was not significantly different from saline-treated Ctrl or SMS mice (Fig 5G). We also found that LM22A-4 treatment partially rescued fat deposition (Fig 5H) and completely rescued HDL levels (Fig 5I) in SMS mice. During an insulin tolerance test, we found that saline-treated SMS mice showed increased baseline insulin levels (Fig 5J) and blood glucose levels during the test (Fig 5K), both of which were partially rescued by LM22A-4 treatment.

In addition to a deficit in energy homeostasis, SMS mice showed two highly reproducible behavioral deficits: increased stereotypic rearing in the open field and decreased social dominance in the tube test (Huang et al, 2018; Rao et al, 2017). Saline-treated SMS mice showed significantly more rearing behavior when compared to the saline-treated controls (Fig 5L). Interestingly, LM22A-4 treatment reduced the repetitive rearing behaviour in SMS mice (Fig 5L). LM22A-4 treatment did not change repetitive behaviors in control mice (Figure 5— figure supplement 1F), suggesting the effect is specific to Rai1 haploinsufficiency. However, we did not observe a rescue of social dominance deficits in SMS mice (Figure 5—figure supplement 2A-E). Together, these data indicate that LM22A-4 treatment delayed the onset of obesity and fully rescued repetitive behaviour in SMS mice, suggesting that Bdnf-TrkB signalling dysfunction contributes to changes in the mouse SMS brain linked to obesity and behavioral phenotypes.

Discussion

The hypothalamus, and Bdnf-TrkB signalling in this region, control body weight by regulating the intricate balance between energy intake and expenditure (Xu & Xie, 2016). The role of Bdnf-producing neurons in human obesity disorders and how the activity of these cells is regulated remained poorly studied. We found that Rai1 deficiency, linked to Smith-Magenis syndrome (SMS), regulates Bdnf expression and maintains intrinsic excitability and inhibitory synaptic transmissions in PVH^Bdnf neurons.

Consistent with this mechanism, we found that removing Rai1 from Bdnf-producing neurons or PVH^Bdnf neurons results in imbalanced energy homeostasis and results in obesity. A small molecule drug LM22A-4 led to increased Akt phosphorylation and partially rescued locomotor activity, insulin intolerance, and HDL levels, while delaying the onset of obesity in SMS mice. These findings highlight a potential role for hypothalamic Bdnf-producing neurons, specifically PVH^Bdnf neurons, in SMS pathogenesis and highlight Bdnf signalling as a potentially druggable pathway to improve energy homeostasis defects and repetitive behaviours commonly observed in SMS patients (Burns et al., 2010; Moss et al, 2009).

The involvement of different PVH subtypes in human obesity is only beginning to be elucidated. Dysfunction of oxytocin PVH neurons is linked to the Prader-Willi syndrome (Lee et al, 2012), another genetic disorder associated with obesity. Since Rai1 expression is enriched in PVH^Bdnf neurons but not PVH^Oxytocin neurons, our data suggest that distinct pathophysiological mechanisms underlie SMS and Prader-Willi syndrome.

Conditional knockout analysis allowed us to specifically map the cell-specific origin of neuropathological phenotypes, such as SMS-like symptoms in mice. Our previous work found that deleting Rai1 from cortical excitatory neurons using an Emx1^Cre allele (Gorski et al, 2002), from the GABAergic neurons using a Gad2^Cre allele (Taniguchi et al, 2011), or from astrocytes using a GFAP^Cre allele (Garcia et al, 2004) did not induce SMS-like obesity (Huang et al., 2016). In contrast, obesity could be induced by ablating Rai1 from Sim1-lineage neurons and, to a lesser extent, SF1-lineage neurons (Huang et al., 2016). This analysis suggests that Rai1 expression in PVH is more important than in ventromedial nucleus of the hypothalamus (VMH) for body weight regulation. Interestingly, embryonic deletion of Bdnf from SF1-lineage neurons including the VMH only resulted in modest weight gain (Unger et al., 2007). By contrast, deleting Bdnf from Sim1-lineage neurons including the PVH resulted in a more severe obesity phenotype (An et al., 2015). This is consistent with the idea that Bdnf expression in PVH is critical for energy homeostasis. We found that Rai1 expression in Bdnf-producing cells regulates energy homeostasis in a sexually dimorphic manner. Specifically, female cKO mice showed normal food intake, decreased energy expenditure, and reduced locomotor activity. In contrast, male cKO mice showed increased food intake with no changes in locomotor activity. Intriguingly, energy expenditure was increased in male cKO mice, likely due to the higher demand to maintain normal activity levels in obese mice. In both sexes, cKO mice showed an increased fat mass and adiposity with no change in lean mass, highlighting an important role for Rai1 in regulating fat deposition. Our data also indicate that Bdnf-producing cells are only partially responsible for obesity in SMS: male and female cKO mice become overweight by 15 weeks, in contrast to SMS mice which become obese by 9 weeks of age, suggesting that non-Bdnf neurons contribute to SMS phenotypes.

We found the metabolic phenotypes induced by deleting Rai1 from Bdnf-producing cells were specifically due to Rai1 deletion in PVH^Bdnf neurons. Therefore, our study focused on PVH^Bdnf neurons. Electrophysiological experiments showed that most wild-type PVH^Bdnf neurons display high frequency spontaneous AP firing (1-10 Hz) at the resting membrane potential. The majority of PVH^Bdnf neurons fire sharp APs held at a hyperpolarized potential and show repetitive overshoot during the current ramp test. Loss of Rai1 significantly reduced intrinsic neuronal excitability and increased inhibitory synaptic transmissions onto PVH^Bdnf neurons. Because GABAergic Rai1 loss did not induce obesity (Huang et al., 2016) and most Bdnf-producing neurons are glutamatergic (Canals et al, 2001), the increased mIPSC amplitude was likely due to postsynaptic changes in GABA receptor functions caused by Rai1 loss. The ionic changes in PVH^Bdnf neurons that precisely explain these excitability changes await future investigations. Different groups of PVH neurons show phasic bursting or slowly inactivating delayed rectifier potassium conductance, which are properties mediated by different potassium channels (Lee et al., 2012). Given that our previous RNA-seq experiment found that hypothalamic Rai1 loss induces misexpression of potassium channel subunits Kcnc2 and Kcnj11 (Huang et al., 2016), Rai1-dependent potassium channels are likely involved in altered neuronal activity. Furthermore, we found fewer low threshold depolarization events in cKO PVH^Bdnf neurons during the hyperpolarized voltage step, suggesting a potential involvement of HCN channels.

Our previous work found that Rai1 loss in the dentate gyrus resulted in neuronal hyperexcitability (Chang et al, 2022b). Here we demonstrated that Rai1 loss in the hypothalamus resulted in neuronal hypoactivity, suggesting that Rai1 regulates neuronal excitability in a cell type-specific manner. Reduced PVH^Bdnf neuronal activity upon Rai1 loss could potentially explain obesity, consistent with a previous report that found chemogenetic activation of PVH^Bdnf neurons promotes negative energy balance by decreasing feeding and increasing energy expenditure (Wu & Xu, 2022).

In future work, it would be interesting to dissect the sex-specific role of Rai1 in regulating the functional development of PVH^Bdnf neurons. In addition to a role for Rai1 in regulating neuronal activity, it is possible that Rai1 regulates other aspects of PVH^Bdnf neuronal function, including projections to other brain regions to regulate energy homeostasis. In this regard, it would be interesting to dissect how Rai1 controls circuit connectivity of PVH^Bdnf neurons and how Rai1 loss in PVH^Bdnf neurons affects nearby PVH^TrkB neurons that receive Bdnf signaling and are linked to body weight regulation (An et al, 2020).

At the molecular level, our RPPA data show that Rai1 depletion disrupted multiple intracellular signalling pathways and resulted in the hypoactivation of specific phospho-signalling proteins. Multiple early studies support the notion that Rai1 directly promotes Bdnf expression. First, hypothalamic Bdnf mRNA levels were reduced in SMS mice (Burns et al., 2010; Javed et al., 2021) or mice with Rai1 conditional deletions (Huang et al., 2016). Second, Rai1 protein directly binds to a promoter region of Bdnf (Huang et al., 2016). Finally, when Rai1 expression was induced using CRISPR activation, Bdnf mRNA levels were significantly elevated (Chang et al, 2022a).

To lend further support, here we show that SMS mice have reduced hypothalamic Bdnf protein levels, as well as reduced activation of a TrkB downstream target Akt. Reduced Akt activity is frequently associated with insulin resistance and obesity in mice (Shao et al, 2000) and children (Su et al., 2021). We also found that obesity, increased HDL levels, and insulin insensitivity in SMS mice could be partially ameliorated using a small molecule drug (LM22A-4) that promotes Akt phosphorylation. LM22A-4 penetrates the blood-brain barrier reasonably well and has been shown to ameliorate symptoms of other neurological disorders associated with Bdnf dysfunction including Rett syndrome, Huntington’s disease, Dravet syndrome, and refractory neonatal seizures (Canals et al., 2004; Chang et al., 2006; Gu et al., 2022; Kron et al., 2012; Ogier et al., 2007). The partial rescue of SMS phenotypes that we observed contrasts with our previous findings that found (1) AAV-mediated Bdnf overexpression in PVH at 3 weeks of age and (2) genetically overexpressing Bdnf from early development could fully prevent the development of obesity in SMS mice (Javed et al., 2021). It is therefore possible that further increasing TrkB signalling activation and/or earlier intervention might improve therapeutic outcomes.

Alternatively, our RPPA data showed that in addition to disruption of neurotrophin signalling, multiple pathways including mTOR were downregulated in SMS. Increasing hypothalamic mTOR activity has been shown to decrease food intake and body weight (Cota et al., 2006). Therefore, it is possible that enhancing both mTOR and TrkB signalling could achieve stronger rescue of obesity in SMS mice. Nevertheless, these data support the concept that drugging TrkB signalling may improve stereotypical repetitive behavior and partially alleviate metabolic functions in a mouse model of SMS. Delayed symptom onset could provide a window of opportunity for other interventions to further modify disease progression, an area requiring further investigation.

This study has several limitations. First, whether the PVH^Bdnf neurons depend on Rai1 to regulate adaptive thermogenesis awaits future analysis. It is known that a subset of posterior PVH^Bdnf neurons project to the spinal cord and regulate thermogenesis (An et al., 2015). However, we found the weights of brown adipocytes were similar in control and cKO mice. Moreover, SMS mice have normal respiratory exchange rate and there are currently no reports of defective adaptive thermogenesis in SMS patients. This suggests that adaptive thermogenesis is not impacted in SMS. Another limitation is that we did not test different doses, durations, and time points for LM22A-4 treatment. The precise mechanisms by which LM22A-4 improves body weight, metabolic function, and repetitive behaviours also remain unknown. Nevertheless, our work identifies Rai1 as a novel regulator of the neuronal excitability of PVH^Bdnf neurons. This regulation is critical for regulating food intake and energy expenditure. Our evidence suggests that Bdnf-producing neurons are involved in SMS pathogenesis and that enhancing Bdnf-TrkB signalling should be explored as a future therapeutic avenue for obesity and metabolic conditions associated with reduced neurotrophin signalling (Xu & Xie, 2016).

Materials and Methods

Animal Management

The animal protocol was approved by the animal care committee of the Montreal General Hospital and was performed according to the guidelines of the Canadian Council on Animal Care. Mice were group-housed in 12-hour light/12-hour dark cycles with an ad libitum supply of regular chow and water. The exception was in the fasting studies when the animals were food deprived for 5 hours and in CALMS and food intake studies when animals were single-housed in a 12-hour light/12-hour dark cycle with an ad libitum supply of regular chow and water. All mice were maintained in a C57Bl/6J background. Bdnf^Cre/Cre, Rosa-tomato^td/td(Ai9), and Actβ^Cre/Cre mice were purchased from the Jackson laboratory. The Rai1^fl/fl strain was previously generated and characterized by us (Huang et al., 2016). SMS mice (Rai1^+/-: whole-body germline Rai1 heterozygous knockout that genetically mimics SMS) were generated by crossing germline Actβ^Cre/Cre mice with Rai1^fl/+. To generate conditional knockout mice, Bdnf^Cre/+; Rai1^fl/+ mice were crossed with Rai1^fl/+ to create control and experimental groups from the same littermate.

Metabolic profiling

Metabolic profiling was conducted in three different experimental conditions: male and female conditional knockout mice, female virus-injected Bdnf^Cre/+mice, and female drug- or saline-injected SMS and control mice. Metabolic profiling included body weight and body composition analysis, food intake measurement, indirect calorimetry for locomotion activity, energy expenditure (EE), respiratory exchange rate (RER), measurement of serum lipids and leptin levels, adipose tissue histology analysis, and glucose and insulin tolerance tests. All experiments were performed as previously conducted experiments (Javed et al., 2021). A brief description is detailed below.

Body weight and body composition analysis

The body weight of animals was measured weekly from week 3 (weaning age) onward. Body composition, including total fat and lean mass, were assessed using a nuclear EchoMRI whole-body analyzer. Toward the end of experiments, animals were euthanized to collect brown adipose, visceral, subcutaneous (inguinal) and ependymal white adipose tissues and weighed using an analytical scale (Sartorius).

Food Intake Measurement

Mice were single-housed for food intake measurement. Food intake was measured over a one-week period during which food was weighed prior to placement in the cage with minimal bedding. Each morning remaining food inside the cage was weighed. The amount of total food consumed was averaged over 7 days.

Indirect Calorimetry

A Comprehensive Lab Animal Monitoring System (CLAM) (Columbus instruments) was used to measure the RER and EE (normalized over lean mass). Animals were singled-housed in the CLAMS apparatus at 21^◦C (70^◦F) in a light–dark cycle matching their housing conditions for 24 hours (acclimation), followed by 48 hours of measurement.

Serum lipid level determination

A fluorescence-based assay kit (Abcam ab65390) was used to measure serum lipid levels. The manufacturer’s instructions were followed except that we used half-area black 96-well microplates. Fluorescence measurements at 535/587 nm (Ex/Em) were recorded with the Ensight instrument from PerkinElmer. A new standard curve ranging from 0 to 10 μg/ml was produced for each microplate to accurately determine lipid levels in serum samples. Standards and samples were loaded and analyzed twice.

Histology of adipose tissue

Ependymal white adipose tissues (WAT) were excised immediately after the animals were euthanized and fixed with 10% formalin in PBS. The fixed tissues were dehydrated, embedded in paraffin, and sectioned. Two 5-micrometre thick sections were taken from each animal and stained with hematoxylin and eosin. Images of sections were then captured using an Aperio ScanScope CS slide scanner at 20X optical magnification, and the resulting images were saved in jpeg format using the ImageScope software. The images were manually cleaned to remove non-adipose tissues such as lymph nodes. An automated method using the Adiposoft plugin version 1.16 was utilized to determine adipocyte size and number from the jpeg images in Fiji version 2.1.0/1.53c. The Adiposoft settings used excluded cells on edges and set a minimum and maximum cell diameter of 20 and 300 μm, respectively. An experimental value of 2.49 μm per pixel was determined using the Aperio scale bar and used for Adiposoft. To ensure accuracy, two images from each animal were analyzed blindly, examining more than 1600 cells per mouse and more than 9100 cells per condition. Adipocyte size frequencies were calculated using the frequency function in Excel.

Glucose and insulin tolerance tests

For the glucose tolerance test, experimental mice were food deprived for 5 hours with ad libitum water supply. A bolus of glucose (1.5g/kg of lean body weight) was then administered intraperitoneally and glycemia was measured from tail vein blood samples using the Accu-chek Performa glucometer at T0 (before the injection) as well as at 15, 30, 45, 60, 90 and 120 min (after the injection). Tail vein blood samples were collected via a capillary for insulin assays at T0, 15 and 30 min. For insulin tolerance tests, experimental mice were food-deprived for 5 hours with ad libitum access to water. A bolus of insulin (1 U/kg of lean body weight) was administered via an IP injection and glycemia was measured from blood sampled at the tail vein using a Accu-chek Performa glucometer at T0 (before the injection), as well as at 15, 30, 45, 60, 90 and 120 min (after the injection). Tail vein blood samples were collected via a capillary for insulin assays at T0.

Stereotaxic Injections

Bdnf^Cre/+ mice were anesthetized using isoflurane, and the ears were barred onto a stereotaxic device (David Kopf Instruments). Adeno-associated viruses (AAVs, volume = 250 nl) were bilaterally injected into the PVH using the following coordinates relative to the Bregma: -0.6 mm anteroposterior (AP), ±0.35 mm mediolateral (ML) and -5.52 mm dorsoventral (DV). A description of the AAV constructs and titre used is detailed below:

AAV9-CMV7-DIO-saCas9 (Viral title: >1 × 10^13), Vector Biolab

AAV9-CMV7-sgRai1-GFP (Viral title: >1 ×10^9), Applied Biological Materials Inc. Three sgRNAs were pooled to target Rai1 (sgRNA1: TTCCTCGCCAGAGTAGCGCC, sgRNA2: CCCAGCCTCATGATAGGCCG, sgRNA3: CCCAGCCTCATGATAGGCCG).

Immunostaining of mouse brain tissues

Immunostaining experiments were performed as previously described (Huang et al, 2012). Briefly, mice were anesthetized and perfused with 1X PBS and 4% formaldehyde, and the brains were dissected and fixed in 4% formaldehyde overnight. Fixed brains were then transferred to 30% sucrose solution and subsequently embedded in the OCT solution. The brains were then sectioned (40µm), washed in PBS, permeabilized with 0.5% Triton X-100, and incubated with 10% normal donkey serum in PBS for 2 hours. The sections were then incubated overnight with primary antibodies (Rai1: 1:500 dilution, RRID:AB_2921229 (Huang et al., 2016); Bdnf: 1:500, RRID:AB_10862052, Abcam ab108319; p-TrkB: 1:500 dilution, RRID:AB_2721199, Millipore ABN1381; oxytocin: 1:1000 dilution, RRID:AB_11212999, Millipore MAB5296) at 4°C on a shaker. The following day, sections were washed with 1X PBS twice for 10 minutes and incubated with a secondary antibody (1:1000 dilution) for 3 hours at room temperature. Sections were washed with 1X PBS once before mounting on glass slides with the DAPI-mounting (DAPI-Fluoromount-G, SouthernBiotech) and then imaged using a confocal microscope (Olympus FV-1000 confocal laser scanning microscope) with 20x or 40x objectives (NA 0.85 and 1.3, respectively). Z-stacks were taken with a step size of 1.0 µm with a 1024 x 1024 resolution. Images were acquired as a stack with at least 13 optical sections. Image analysis was performed using the Fiji software.

Real-Time Quantitative Reverse Transcription PCR (qRT-PCR)

qRT-PCR was performed using mRNAs isolated from hypothalamic tissue samples. After isolation of total RNA, mRNAs were reverse-transcribed with the SuperScript III First-Strand Synthesis System (Thermo-Fisher). Quantitative PCR reactions were conducted using SsoFast EvaGreen Supermix on a Bio-Rad qPCR system.

Enzyme-linked immunosorbent assay (ELISA)

ELISA was performed using hypothalamic tissues from 7-week-old SMS mice to assess the mature Bdnf protein levels using a commercially available Bdnf ELISA kit (Mature BDNF Rapid ELISA Kit: Human, Mouse, Rat).

Western blotting analysis

Mice were quickly decapitated, brains were rapidly removed, and hypothalamic tissues were collected, weighed, and snap-frozen with liquid nitrogen. Hypothalamic tissues were lysed using a tissue lysis buffer containing RIPA buffer (5M NaCl, 1M PH 8.0 Tris HCl, 1% NP.40, 10% C₂₄H₃₉NaO₄, and 20% SDS) and the Halt™ Protease and Phosphatase Inhibitor Cocktail (Thermo-Fisher). Lysates were first sonicated on ice for 15 seconds followed by centrifuging at 10000 RPM at 4°C for 10 minutes. The supernatant was collected in a separate Eppendorf tube, and a BCA assay was performed to quantify protein levels. Proteins were then separated using SDS-PAGE gel and transferred onto nitrocellulose membranes. To determine the phosphorylation levels of Akt, membranes were cut and blocked in 5% milk in tris-buffered saline (TBS) for 1 hour and then incubated overnight at 4°C with anti-p-Akt (Ser473) antibody (1:1000 dilution, RRID:AB_2315049, Cell Signaling Technology 4060T) and anti-H3 antibody (internal control: 1:1000 dilution, RRID:AB_302613, Abcam ab1791). The next day, membranes were washed with TBST three times for five minutes before incubating with a HRP-conjugated secondary antibody (1:10000 dilution, Thermo Fisher 200-403-095S). Membranes were later stripped by incubating in 0.5M NaOH for 10 min and then re-probed for total Akt levels by incubating with anti-Akt (C67E7) antibody (1:1000 dilution, Cell Signaling Technology 4691T). The density of each immunoreactive phospho-protein band was expressed as a fraction of the band for the total Akt protein in the same lane.

Reverse-phase protein array (RPPA)

Reverse-phase protein array (RPPA) was used for precise high-throughput quantification of total and phospho-protein levels in different signalling pathways. Protein lysates from hypothalamic tissues were extracted and transferred to 384-well microarray plates to print individual slides. The slides were then labelled with validated antibodies that recognize total protein and specific phosphorylation sites. SYPRO Ruby staining was used to measure total protein levels. The data obtained from RPPA were normalized for individual samples. Each antibody’s raw signal intensity (SI) was determined by subtracting the background from the antibody-specific signals during image analysis. The normalized antibody SI was expressed using the formula N = A - C/T x M, where N is the normalized SI, A is the raw antibody SI, C is the negative control SI, T is the SI of total protein, and M is the median SI of total protein from a spot within the same sample group. Antibodies with SI less than 200 were filtered out, and quality control (QC) scores were established to discard poor-quality reactions. The coefficient of variation (CV) score was determined for each antibody and defined as the percentage of samples on a slide with a CV < 20%. Antibodies with a median technical triplicate CV >20% were also excluded from further analysis. T-tests were performed for each antibody to identify differentially expressed proteins in the SMS compared to control groups. PCA plots were generated for all samples, annotated with their experimental group, and hierarchical clustering was performed using a Python analysis tool. Enrichment analysis on the differentially expressed proteins was performed using an online data resource string: version 11.5.

Electrophysiology of acute brain slices

Mouse brains were removed and placed in ice-cold carbonated slicing artificial cerebrospinal fluid (aCSF), which contained (in mM) 124 NaCl, 3 KCl, 1.4 NaH₂PO₄, 26 NaHCO₃,1.3 Mg SO4, 10 D-glucose, 0.5 CaCl₂, and 3.5 MgCl₂. Coronal sections (300 µm) were cut on a vibratome. Slices were allowed to recover at 32 °C for 40 minutes and then at 30 °C for 30 minutes to 6 hours in the recording medium, carbonated aCSF (124 NaCl, 3 KCl, 1.4 NaH₂PO₄, 26 NaHCO₃,1.3 Mg SO4, 10 D-glucose, 2.4 CaCl₂, in mM, ∼300 mOsm). Signals were recorded with a 5× gain, low-pass filtered at 2 kHz, digitized at 10 kHz (Molecular Devices Multiclamp 700B) and analyzed with pClamp 11 (Molecular Devices). Whole-cell recordings were made using 3–5 MΩ pipettes filled with an internal solution that contained (in mM) 131 potassium gluconate, 8 NaCl, 20 KCl, 2 EGTA, 10 HEPES, 2 MgATP and 0.3 Na3GTP (pH 7.2, with KOH, 300–310 mOsm with sucrose). For miniature current recording, the internal solution contained (in mM) 130 gluconate acid, 8 CsCl, 1 NaCl, 2 EGTA, 0.8 CsOH, 10 HEPES, 2 MgATP and 0.3 Na3GTP (pH 7.2, with CsOH, 300–310 mOsm adjusted with sucrose). mEPSCs were recorded in the presence of 1 µM TTX at a holding potential of - 70 mV. mIPSCs were recorded at a holding potential of +10 mV. The mIPSCs were analyzed from events during a 2-minute continuous recording that contained > 15 events/neuron. Additional calculations, statistical analysis, and plotting was performed in Microsoft Excel or Prism (GraphPad).

Spectrometric analysis of LM22A-4 levels in the brain

We assessed the efficacy of LM22A-4, a small molecule pharmacological agent, in 8-week-old SMS and control mice. LM22A-4, chemical name N1, N3, N5tris (2hydroxyethyl) 1, 3, 5 benzenetricarboxamide (Molecular formula: C₁₅H₂₁N₃O₆, Formula weight: 339.3), was manufactured by Cayman Chemical with purity specification >98%. A stock solution was made by first dissolving LM22A-4 in DMSO (solubility =30mg/ml) and then diluting it with 0.9% isotonic sterilized saline solution (Sigma). To maximize brain concentrations, LM22A-4 was administered both intranasally (5mg/kg) and intraperitoneally (50mg/kg), as previously reported (Simmons et al, 2013). The bioavailability of this drug in the hypothalamus using these doses and routes of administration was assessed with the reverse phase LC triple-quadrupole tandem mass spectroscopic detection (LC-MS/MS) by Absorption Systems. Mice were euthanized, and hypothalamic tissue was collected at either 1 h or 3 h post-injection (n= 4/timepoint). Known amounts of LM22A-4 were added to the blank sample to generate a standard curve to verify test accuracy.

LM22A-4 treatment

Mice were divided into four groups: Saline-injected control, Saline-injected SMS, LM22A-4-injected SMS, and LM22A-4-injected control mice. Mice were injected daily, intra-nasally (5mg/kg) and intraperitoneally (50mg/kg), for 7 weeks. Body weight was measured between 8 and 14 weeks of age. Food intake was measured from 14 to 15 weeks of age. After 15 weeks of age, mice were euthanized to collect brown adipose, visceral, subcutaneous (inguinal) and ependymal white adipose tissues and weighed using an analytical scale (Sartorius).

Hypothalamic brain tissues were collected for western blot analysis. Metabolic profiling was performed on a separate cohort of mice injected for two weeks prior to the start of experiments as well as at specific weeks during the experiment.

Vertical rearing

To measure vertical repetitive rearing, mice were taken into the testing room and allowed to habituate for 1 h prior to testing. During the test, mice were placed in the center of a 45 cm (W) × 45 cm (D) × 40 cm (H) square arena, and their movement was recorded for 10 minutes.

Statistical analysis

Sample sizes used were based on our previously studies (Huang et al., 2016; Javed et al., 2021). The data were subjected to statistical analysis for significance by using GraphPad Prism 0.9. Significance was defined as having a p-value < 0.05. The significance levels were expressed as follows: * p< 0.05, **p<0.01, ***p<0.0001, **** p<0.0001. The statistical tests used for each analysis (e.g., Student’s t-test, Fisher’s exact test, analysis of variance [ANOVA]) are indicated in the text and legends.

Data Availability

This study includes no data deposited in external repositories.

Acknowledgements

Sehrish Javed was supported by a doctoral award from the Fonds de Recherche du Québec - Santé (FRQS). This work was supported by the Rare Diseases: Models & Mechanisms Network and Smith-Magenis syndrome Research Foundation.

Author contributions

Sehrish Javed: Conceptualization; formal analysis; investigation; data curation; visualization; writing. Ya-Ting Chang: Formal analysis; investigation; data curation; visualization. Yoobin Cho: Formal analysis; investigation. Yu-Ju Lee: Formal analysis; investigation. Hao-Cheng Chang: Formal analysis; investigation. Minza Haque: Formal analysis; investigation. Yu Cheng Lin: Formal analysis; investigation. Wei-Hsiang Huang: Conceptualization; supervision; funding acquisition; visualization; writing; project administration.

Disclosure and competing interests statement

The authors do not have any conflicts of interest to disclose relating to this work.