Mandibulofacial dysostosis with alopecia results from ETAR gain
Christopher T. Gordon Yasunobu Uchijima Ryo Sugiyama Taro Kitazawa Akiyasu Iwase Risa Kotani Rieko Asai Véronique Pingault Mitsunori Ikeguchi Jeanne Amiel
/assets/jci-favicon-378ea4dd43f03bc78136c6a261d9b28f6811fe59d12db426eae78cf0691b0008.ico
MFDA in a patient with the EDNRA mutation p.E303K. We investigated a girl presenting with symmetrical dysplastic ears (Figure 1, A and B); asymmetric lower eyelid coloboma that required surgery on the right side (Figure 1C); a squared nasal tip (Figure 1D); widely spaced, thin, horizontal eyebrows (Figure 1E); and sparse scalp hair with persistent frontal balding (Figure 1F). She has hearing aids for conductive hearing loss and generalized hypopigmentation compared with first degree relatives. Craniofacial 3D CT scans showed hypertrophic, dysplastic, asymmetric malar bones, abnormal orientation of the orbital floor, and an abnormal temporomandibular articulation on the left side (Figure 1, G–J). CT scans also showed abnormalities of the middle ear, including hypoplasia of the long process of the incus (Figure 1, K and L). The abnormal bones are all cranial NCC derivatives. Further detailed information of this patient is described in Supplemental Table 1; supplemental material available online with this article; https://doi.org/10.1172/JCI151536DS1 Next-generation sequencing revealed that the peripheral blood DNA of the patient was 8%–11% mosaic for the mutation c.907G>A; p.E303K in EDNRA (NM_001957.3) (Figure 1M). Low-level mosaicism of the mutation was confirmed by Sanger sequencing of a PCR product amplified from blood DNA (Figure 1N) and from PCR products amplified from bacterial colonies transformed with the cloned PCR product from the blood (Figure 1O). The variant was not detected in the mother or father (Figure 1N), indicating a postzygotic mutation. These results establish that p.E303K is a second-recurrent mutation causing MFDA, with structural changes of the jaw similar to those caused by p.Y129F.

Figure 1

A patient with MFDA harboring the EDNRA mutation p.E303K, that occurred postzygotically. (A–F) Dysmorphic facial features and alopecia of the patient (see text for details). (G–J) High resolution craniofacial 3D CT-scans of the patient with MFDA (G and I) and an age-matched control (H and J). (G and H) Frontal views show hypertrophic, dysplastic, and asymmetric malar bones and an abnormal orbital floor in the patient (G) compared with the control (H). (I and J) Volume-rendering sagittal reconstructions of the temporomandibular articulations show a novel, tripartite temporo-mandibulo-malar articulation on the left side (red arrow) with increased malar bone volume (blue star) in the patient (I) compared with the control (J). (K and L) Oblique axial reconstructions of CT scans of the incus and malleus showing hypoplasia of the long process of the incus in the patient (K) compared with a control (L). (M) Next-generation sequencing reads showing mosaicism for the EDNRA variant c.907G>A (8%–11% of reads, in green) in the patient’s peripheral blood DNA. (N) Sanger sequencing of EDNRA amplified by PCR from leukocyte DNA of the patient and her parents, demonstrating low -evel mosaicism for the variant c.907G>A (green arrow) in the patient. (O) Sequencing of PCR products amplified from individual bacterial colonies, following transformation with cloned EDNRA PCR fragments from the patient, indicating 3/32 colonies (9 %) harboring the variant (green arrow) while 29 colonies harbored the WT allele (black arrow). Abbreviations: orb, orbital floor; mal, malar bone; cond, condyle of the mandible; ZPT, zygomatic process of the temporal bone; mall, malleus; incus lp, long process of the incus. An asterisk indicates structures that are dysmorphic compared with normal.

ETAR-Y129F and E303K mutations in mice recapitulate craniofacial manifestations of MFDA in humans. The residues mutated in MFDA and their flanking sequences are highly conserved between humans and mice. Therefore, in order to investigate the developmental mechanism, we generated mouse models of MFDA by substituting Y129 or E303 with phenylalanine or lysine, respectively, using the CRISPR/Cas9 system. Mice carrying either heterozygous mutant allele (EdnraY129F/+ or EdnraE303K/+) showed over-folded pinnae, which was more severe for EdnraE303K/+ than EdnraY129F/+, and some mutant mice developed skin erosion around the eyelids, with sparse and coarse coat worsening with age (Figure 2A and Supplemental Figure 1A). Some EdnraY129F/+ and almost all EdnraE303K/+ mice had open eyelids at birth (Table 1 and Supplemental Figure 1A). Skeletal analysis revealed similar abnormalities in NCC-derived maxillary components in both mutants. The zygomatic arch, composed of the jugal and zygomatic processes of the maxilla and squamosal, was deformed into a rod-shaped bone with a cartilaginous proximal end (Figure 2, B and C), on the top of which a vestigial jugal-like bone was also observed (Figure 2C blue asterisk). This rod-shaped bone appears similar to the dentary and condylar process, suggesting a partial transformation of the upper jaw toward mandibular identity. The incus was also deformed in the mutants with an appearance similar to the malleus (Figure 2D). The skeletal components affected in the mutants are illustrated in Figure 2J. No major abnormalities of the mandibular components were observed except for a slightly reduced angular process in EdnraE303K/+ mice (Figure 2B arrowhead).

Figure 2

Craniofacial phenotypes of Ednra mutant mice are rescued by Edn3 deletion. (A–D) Facial appearance and skeletal morphology of WT and heterozygous mutant (EdnraY129F/+and EdnraE303K/+) mice. Mutant adult mice show over-folded pinnae and skin erosion around the eyelid (A). E18.5 bone and cartilage stains show transformation of the upper jaw components into a rod-shaped bone in the mutants (arrows in B and C) and deformity of the incus (arrowheads in D). The mandible is almost normal, except the angular process, which is slightly reduced (arrowhead in B). (E) Facial abnormalities in the EdnraY129F/+ and EdnraE303K/+ mutants are variably rescued by heterozygous deletion of the Edn3 gene. (F and G) Transformation of the upper jaw components in the mutants is rescued by heterozygous Edn3 deletion (arrows). (H) deformity of the incus (arrowheads) is not rescued by heterozygous Edn3 deletion. (I) The incus of the mutants is normalized by homozygous loss of Edn3 (arrowheads). (J) Schematic diagrams indicating the skeletal components affected in the Ednra mutants (green). (K) Comparison of Edn1 and Edn3 mRNA levels at E9.5. 4 pairs of RNA samples were extracted from pools of mandibular or maxillary arches, in each of which 7 to 14 WT littermate samples were combined and were subjected to q-RT-PCR. *P < 0.05 by Mann-Whitney U test (n = 4). (L) Schematic diagrams summarizing the effect of ET signals in normal and pathogenic states. Red lines indicate ET3-dependent activation of mutant receptors. (M) Dlx5 and Dlx6 expression at E9.5 is restricted to the mandibular region of PA1 in WT embryos, whereas the expression of both genes extends into the maxillary region of PA1 in EdnraY129F/+and EdnraE303K/+ embryos (arrowheads). n = 3–6. agp, angular process; dnt, dentary; gn, gonial; in, incus; jg, jugal; ma, malleus; Mc, Meckel’s cartilage; mx, maxilla; sq, squamosal. Scale bars: 1 mm (B, C), 500 µm (D), and 100 µm (M)

Table 1

Summary of mutant phenotypes

As previously described (24), ectopic activation of ETAR by forced ET1 expression resulted in the replacement of the ala temporalis with a Meckel’s-like cartilage, indicating a dorsal-to-ventral transformation of the first pharyngeal arch (PA1). EdnraY129F/+ and EdnraE303K/+ embryos demonstrated intact formation of the ala temporalis (Supplemental Figure 2); however, the expression of Dlx5 and Dlx6, homeobox transcription factors that act downstream of ET1-ETAR signaling to determine ventral identity in PA1, extended dorsally in mutant embryos (Figure 2M). These results suggest that the Ednra mutations had an ectopic gain-of-function effect in the dorsal PA1, although weaker than that observed upon forced expression of ET1.

The phenotype associated with either homozygous Ednra mutation was similar to that of the heterozygous mutants, except that the condylar process was reduced in EdnraE303K/E303K mice (Supplemental Figure 3C red arrow). Importantly, the homozygous mutations did not lead to a mandibular-to-maxillary transformation, which is a hallmark of inactivation of ET1-ETAR signaling in mice and of ACS in humans (Supplemental Figure 3, C and D, and Table 1).

Deletion of Edn3 rescues the phenotype associated with EdnraE303K and EdnraY129F. A previous report suggested that the Y129F mutation of EDNRA can significantly increase affinity for ET3 (32, 34), which is expressed in the maxillary and mandibular processes of E10–12 mouse embryos (32, 35). To confirm both ET1 and ET3 expression in the developing face at a stage critical for the determination of maxillomandibular identity, we performed quantitative reverse–transcription PCR (q-RT-PCR) on embryos at E9.5. In contrast to the preferential expression of ET1 in the mandibular process, ET3 expression was detected at similar levels in both maxillary and mandibular processes (Figure 2K).

The above results suggest that the MFDA phenotype is due, at least for EdnraY129F, to ectopic activation of mutant ETAR by ET3 in the maxillary region. To test this possibility, we introduced an Edn3-null allele into both mutant mice using the CRISPR/Cas9 system. Remarkably, the heterozygous Edn3-null allele mostly rescued both mutant phenotypes, except for the deformity of the incus and the eyelid dysplasia (Figure 2, E–H, Supplemental Figure 3, A and B, and Table 1). When both Edn3 alleles were deleted, the morphology of the incus was completely normalized in each heterozygous Ednra mutant (Figure 2I and Table 1), while open eyelids at birth was incompletely rescued by Edn3–/– in the ETAR-E303K mutants (Table 1). Edn3–/– also rescued most phenotypes associated with each homozygous Ednra mutant (Supplemental Figure 3, C and D, and Table 1). Our results indicate that the gain-of-function phenotypes associated with the ETAR mutants are almost entirely dependent on ET3 (Figure 2L), but that the E303K mutation may increase basal or ET3-independent ETAR activity, particularly in the context of eyelid development (Supplemental Figure 1B and Table 1).

ETAR-Y129F and -E303K enhance ET3-induced ERK phosphorylation and intracellular Ca2+ mobilization in HeLa cells. We showed previously that Gq/11 is responsible for the specification of mandibular identity by ET1-ETAR signaling (36). Using HeLa cells overexpressing WT or mutant ETARs, we next examined the effects of ET ligands on phosphorylation of ERK, which is a key effector for signaling downstream of G q/11. Western blotting revealed that both ETAR-Y129F and E303K increased the sensitivity to ERK phosphorylation to ET3 by more than 2 orders of magnitude (Figure 3A left). Remarkably, the E303K mutation also enhanced ET1-induced ERK phosphorylation by approximately 3-fold, whereas the ETAR-Y129F did not affect the sensitivity to ET1 (Figure 3A, right). ERK phosphorylation can also be attributed to activation of other G proteins and recruitment of arrestin during GPCR signaling (37–39). We therefore evaluated Ca2+ mobilization and phospholipase C-mediated hydrolysis of inositol phospholipids, which represent more specific downstream readouts of Gq/11 signaling. To estimate inositol hydrolysis, we measured concentrations of inositol 1 phosphate (IP1), which is a stable metabolite of inositol 1, 4, 5-triphosphate (IP3). ETAR-Y129F and E303K showed enhanced intracellular Ca2+ and IP1 concentrations in response to ET3, as estimated by the EC50 (Figure 3B left and Supplemental Figure 4A), whereas no difference was found in the dose response to ET1 (Figure 3B right and Supplemental Figure 4B). The response of ETAR -E303K to ET1 enhancement in the phosphorylated ERK assay compared with ETAR-WT and ETAR-Y129F, and no difference in response to the observation of ET1 in the Ca and IP1 assays, may suggest enhanced GPCR kinase (GRK) phosphorylation and arrestin recruitment for ETAR-E303K. Regarding the effects of other substitutions at E303, arginine (R) enhanced the response to ET3 as much as K, whereas aspartic acid (D) did not (Supplemental Figure 5, A and B). Replacement of E303 by neutral amino acids alanine (A) or glutamine (Q) resulted in partial enhancement (Supplemental Figure 5, A and B). These results indicate that mutations at Y129 and E303 can enhance Gq/11-mediated signaling in response to ET3, and that the charge of the residue at position 6.32 may be critical.

Figure 3

Gain-of-function properties of the ETAR mutants are caused by their ligand-binding affinity and Gq activation. (A) ERK phosphorylation assay. Representative Western blots (upper) and dose-response profiles (lower) of phosphorylated ERK (p-ERK) relative to total ERK, stimulated with ET3 (left) or ET1 (right). n = 3 (B) Intracellular calcium mobilization assay. Dose-dependent elevation of intracellular calcium concentrations stimulated by ET3 (left) or ET1 (right). n = 4. RFU; relative fluorescent unit. (C) Competitive ligand–binding assay. Competition-equilibrium binding was performed on crude membrane extracts using [125I] ET1 and the cold competitor agonist ET3 (left) or ET1 (right). Each value represents the mean ± SEM (also see Supplemental Table 4) from 2–3 independent experiments performed in duplicate. (D) Gq activation assay. Gq protein and Flag-tagged ETAR were reconstituted in rHDL particles and [35S] GTPγS binding was measured in the presence of ET3 (left) or ET1(right). Each value represents the mean ± SEM from 2–3 independent experiments performed in duplicate. (E) Measurement of basal activity. Intracellular calcium concentrations (left, n = 11) and IP1 concentrations (right, n = 6) in WT or mutant ETAR–expressing HeLa cells. Steel-Dwass’s multiple comparison test, *P < 0.05.

ETAR-Y129F and E303K confer high affinity ET3 binding on ETAR. The above findings in mouse models and in vitro experiments indicate that, in addition to Y129, which is near the ligand-binding pocket, E303 may also serve as a determinant of ligand binding affinity even though it is located on the intracellular side of TM6. To explore this possibility, we performed ligand binding assays using crude membrane extracts from HEK293 cells with WT or mutant expression vectors. Competitive binding assays showed similar IC50 values for ET1 binding in WT and mutant receptors (Figure 3C). By contrast, the binding affinity to ET3 was increased in ETAR-Y129F and ETAR-E303K by approximately 27 and 10 fold, respectively (Figure 3C). To minimize the effect of associated proteins such as trimeric G proteins, we performed competitive ligand binding assays using WT and mutant receptor preparations incorporated into a reconstituted high-density lipoprotein (rHDL) phospholipid bilayer particle (Nanodisc). The ETAR-Y129F and ETAR-E303K mutants increased ET3 binding affinity by approximately 27 and 5 fold, respectively, in qualitative accordance with experiments using crude membrane extracts (Supplemental Figure 6).

Differential effects of ETAR-Y129F and E303K on G protein activation and basal activity. That the above findings showed similar activation of downstream signaling pathways in both mutants, although ETAR-E303K increased ET3 binding affinity less effectively than ETAR-Y129F, led us to speculate that ETAR-E303K might potentiate G protein activation upon ligand binding. To test this possibility, we incorporated WT and mutant receptor proteins into rHDL particles together with Gαq and βγ subunits. After normalization of rHDL-ETAR quantities by measuring binding to equivalent amounts of radioactively labeled ET1, we evaluated G protein activation by [35S]GTPγS binding. ETAR-E303K largely increased Gαq activation efficacy upon stimulation by ET-1 as well as ET-3, whereas, to a lesser extent, ETAR-Y129F increased only ET3-induced Gαq activation (Figure 3D). Thus, the increased ET3-dependent Gαq activation of ETAR-Y129F mainly reflects enhanced ET3 binding affinity, whereas that of ETAR -E303K reflects both enhanced ET3 affinity and a high intrinsic tendency toward activation.

To further test the gain-of-function properties of ETAR-Y129F and ETAR-E303K, we evaluated basal activity by measuring intracellular Ca2+ and IP1 levels without ligand, as an index of Gq activity. ETAR-E303K and, to a lesser extent, ETAR-Y129F, exhibited an increase in intracellular IP1 and/or Ca2+ concentrations compared with ETAR -WT without any difference in maximum response (Figure 3E and Supplemental Figure 7), indicating increased basal Gq activity. Taken together, these results suggest that both mutants have gain-of-function properties with different dependence on ligand-binding affinity.

MD simulations reveal dynamic conformational changes in ETAR-E303K. Structural analysis could shed light on the mechanisms underlying the gain-of-function properties of mutant ETAR, but no experimental structures are currently available for the WT receptor. As the conformation of membrane proteins fluctuates constantly regardless of ligand binding, we established a mouse ETAR homology model under membrane–water layer conditions, using the crystallographic structure of human ETBR (PDB 5GLI) as a template, in order to perform MD simulations (Supplemental Methods). The amino acid identity between mouse ETAR and human ETBR is 62 % as a whole and 71.5% from helices 1–8, which is considered sufficient for prediction of target structures, given that reliable homology modeling requires sequence identity higher than 35%–40% between target and template (40, 41).

Analysis of apo ETAR-WT (i.e., ETAR-WT without ligand) MD simulation revealed that most of the hydrogen bonds (HBs) between the intracellular half of TM6 and other TMs were formed through E3036.32. In particular, it forms HBs with S3738.47 and/or K3748.48 located at the junction of TM7 and helix 8 (H8) with high probability (Figure 4, A and B, and Supplemental Figure 8A). On the other hand, the positively charged mutant K3036.32 had little probability of forming HBs with other TMs. When the TM6–TM7 distance on the intracellular side was evaluated as the distance between T3076.36Cα atom and V3727.56Cα atom (hereafter referred to as TMin), it showed a unimodal distribution (5–7 Å) for ETAR-WT, and a multimodal distribution (5–7 Å and > 7 Å) for each mutant (Figure 4, A and C). When HBs were formed between E303 and other TMs, they were restricted within the short TMin peak (5–7 Å) for ETAR-WT and ETAR-Y129F (Figure 4C pink). Thus, the intracellular end of TM6 harboring E303K becomes more flexible because of the reduction of HBs due to the substitution of lysine by glutamic acid. Accordingly, upon deletion of the HB between E303 and S373 by replacing the serine of the latter with glutamic acid, the helix also became more flexible, with a bimodal distribution of TMin as seen for E303K (Supplemental Figure 8, B and C), indicating the importance of E303-associated HBs for regulating the flexibility of TM6.

Figure 4

Loss of hydrogen bonds increases flexibility of TM6/7 in ETAR-E303K. (A) Homology modeling of the ligand-free inactive form of mouse ETAR (apo ETAR). 3D visualizations were generated using PyMOL. The cytoplasmic TM6/7 and H8 regions of ETAR-WT and ETAR-E303K are magnified in boxes below. The cytoplasmic half of TM6 (E2966.25 to C3186.47) is shown in pink. TM2 is denoted in lime. TMin, TMpkt and TMex represent distances between T3076.36Cα and V3727.56Cα, between D1332.57Cα and W3196.48 Cα, and between L1412.65Cα and K3296.58 Cα, respectively. Broken pink lines indicate HBs between the side chain of the 303 residue (E or K) and other TM residues, with width reflecting each HB probability. (B) Hydrogen bonding between cytoplasmic half of TM6 or E303 and other regions. Probability of HB formation between the cytoplasmic half of TM6 (from E296 to C318) and other TMs (left columns, grey), or between the 303 residue (E or K) and other TMs (right columns, colored differently according to HB partners), for ETAR-WT, -Y129F and -E303K. The probability is defined as the number of snapshots having HBs in a total of 3,000 shots (%). (C) Probability distribution of TMin. HBs formed between the cytoplasmic half of TM6 (from E296 to C318) and other TMs (light gray) and between the 303 residue and other TM residues (pink) are superimposed on total TMin (dark gray). (D) Probability distribution of TMex. Total TMex (light gray), and TMin > 7 Å (yellow) are superimposed. (E) Probability distribution of TMex < 22 Å (lime) superimposed on total TMin distribution (light gray).

The distance between TM2 and TM6 on the extracellular side, represented by the distance from L1412.65Cα to K3296.58Cα (hereafter referred to as TMex), showed a unimodal distribution for ETAR-WT and ETAR-Y129F, but was bimodal for ETAR-E303K (Figure 4, A and D). When TMin was greater than 7 Å for ETAR-E303K, it was distributed mainly in the shorter peak of TMex (Figure 4D yellow), while TMex less than 22 Å was distributed mainly in the larger peak of TMin (Figure 4E lime), indicating that the conformation of the intracellular side influenced that of the extracellular side in ETAR-E303K. Importantly, the wide TMin state is predicted to be capable of coupling to G protein without ligand binding during the dynamic and flexible motion of mutant ETAR, resulting in increased basal activity (3) (Figure 5). To assess the broader relevance of this, we estimated the distance corresponding to TMin (amino acid positions 6.36–7.56) in several class A GPCR crystal structures from the Protein Data Bank and observed that the active and inactive states of GPCRs appear to correspond to long (7.7–11.5 Å) and short (5.5–6.7 Å) distances, respectively (Supplemental Table 2). These data suggest that ETAR-E303K preferentially adopts the active state regardless of ligand availability, allowing G protein binding and increased basal activity, which itself allosterically affects the conformation of the ligand binding domain (Supplemental Figure 9).

Figure 5

Presumed mechanism for enhanced affinity of ETAR-E303K to ET3. (i) Increased flexibility of TM6 due to E-to-K substitution at residue 303. (ii) Increased basal activity. (iii) G protein–mediated enhancement of agonist affinity; upon entering the ligand binding pocket, ET3 is prevented from egressing due to the narrowing of the ligand binding pocket, leading to increased ET3 affinity and sustained Gq activation.

Reduced water network contributes to the acquired ET3 affinity in ETAR-Y129F. Next, we examined how ETAR-Y129F increases ligand binding affinity. Y1292.53 in ETAR is located in the putative Na+/water pocket under the orthosteric ligand binding pocket. As Y1292.53 is expected to form HBs with water molecules through its hydroxyl group (Supplemental Figure 10A), substitution with uncharged phenylalanine may disrupt the HB network in the Na+/water pocket and the conformation of adjacent helices. To evaluate the state of water molecules in the interhelical space under the ligand binding site, it was divided into 3 sections of 8 Å each; the G protein binding area (area 1), the D1262.50-area (area 2), and the Y1292.53-area (area 3) (Figure 6A). MD simulation showed a decreased number of water molecules in area 3 of ETAR-Y129F compared with that of ETAR-WT and -E303K (Figure 6B). Y1292.53 forms a HB with D1332.57 1 turn above Y129 at about 50% frequency in the simulation thereby restricting the movement of the tyrosine side chain (Figure 6C). However, the phenylalanine side chain does not form HB and is more flexible because it moves widely (Supplemental Figure 11). It may compete with water molecules in the Na-water pocket lumen. The narrowed distance between Y/F1292.53 and W3196.48 in ETAR-Y129F indicates that the pocket tends to shrink (Figure 6C). These results suggest that the bulky hydrophobic side chain of F129 also reduces the number of water molecules.

Figure 6

Apo ETAR-Y129F takes on a form intermediate between inactive and active states due to a reduced water network. (A) Division of the interhelical cavity beneath the ligand binding site into 3 regions of 8 Å each. Area 1 is the G protein binding area; area 2 contains D1262.50; and area 3 contains Y1292.53. (B) Probability distribution of the numbers of water molecules in area 3. The dashed vertical line indicates the point at which the water number is 7. (C) Distance between apo Y/F1292.53CZ atom and W3196.48 CH2 atom. The representative side chains are superimposed (right). (D) Probability distribution of TMpkt for apo ETAR-WT, -Y129F and -E303K. The distributions when area 3 contains ≤ 7 water molecules are overlaid (sky blue). The lowest panel is a stack of TMpkt distributions for ETAR-WT (black), -Y129F (blue), and -E303K (red). (E) TMpkt level structure. A representative apo ETAR-Y129F with area 3 = 7 water molecules (sky blue) is superimposed on Na-bound ETAR-Y129F (inactive state; grey). (F) PCA for the conformation of the extracellular half of TMs 2, 6, and 7 of ETAR-Y129F in different states. A snapshot of the 3D representation of the PC2-PC3-PC5 axis. This is available in a 3D viewer file at https://kurihara-utokyo.github.io/yk2021-et/ (G) Probability distribution of TMpkt for ETAR-Y129F (upper) and ETAR- WT (lower), in the apo forms (dark grey) or bound to ET3 (pink or light grey). (H) Probability distribution of TMex for ETAR-Y129F (upper) and ETAR- WT (lower), in the apo forms (dark grey) or bound to ET3 (pink or light grey).

When the distance of the main chain between D1332.57Cα and W3196.48Cα (hereafter referred to as TM pocket; TMpkt, Figure 4A) was regarded as an index of the conformation of the Na+/water pocket, it showed unimodal distribution for ETAR-WT or ETAR-E303K, and a bimodal distribution for ETAR-Y129F (Figure 6D). The narrow TMpkt peak for ETAR-Y129F largely corresponded to a small number of water molecules (≤ 7) in area 3 (Figure 6D), demonstrating a shrinkage of the TMpkt compared with the Na+-bound inactive state (Figure 6E). By contrast, TMin or TMex of apo ETAR-Y129F showed no apparent correlation with water molecule numbers (Supplemental Figure 10B).

We then performed further MD simulations with Na+-bound ETAR-Y129F (an inactive state) and ET3-bound ETAR-Y129F (an active state), compared with Na+-free apo ETAR-Y129F, and analyzed the movement of the extracellular portions of TM2, TM6, and TM7 which are reported to move inward upon ligand binding in ETBR (42–44), by principal component analysis (PCA) (Figure 6F). The validity of this PCA was confirmed by examining apo WT and mutant apo ETARs in a Na+-free state (Supplemental Figure 12, A–C). Apo ETAR-Y129F conformations with decreased number of water molecules (≤7) in area 3 (Figure 6F) are considered to be an intermediate state because they are located separately from the active or inactive forms. The ET3-bound forms of ETAR-Y129F and ETAR-WT have a narrower TMpkt (Figure 6G), which is similar to that of apo ETAR-Y129F with decreased water molecules (≤7) in area 3 (Figure 6D). The TMex is also narrower in ET3/ETAR-Y129F (Figure 6H) and in ET3/ ETAR-WT (Figure 6H), indicating that a narrow TMex is a feature of the active state, in addition to a narrow TMpkt.

In order to further examine the effect of a narrow TMex, we measured the distance between ET3 and ETAR, represented by the distance from D18 Cα of ET3 to R326 Cα of ETAR (Figure 7A). It was shorter in ETAR-Y129F than in ETAR-WT (Figure 7B), and the probability of HB formation between ET3 and ETAR-Y129F was high, in particular for the D18-R326 and W21-R326 interactions (Figure 7C and Supplemental Figure 13, A and B). This would have a significant effect on ET3 binding affinity, in other words, substituting tyrosine with phenylalanine narrows the TMpkt due to the decreased number of water molecules in area 3, which leads to narrowing of TMex and formation of HBs. This could prevent the dissociation of ET3 from ETAR, thereby increasing ET3 affinity (Figure 7F). To verify the importance of HBs between R3266.55 in ETAR and residues in ET3 for increased ET3 affinity in ETAR mutants, we substituted R326 with glutamine in ETARs in order to eliminate those HBs. As expected, R326Q substitution in ETAR -Y129F and ETAR-E303K led to a loss of HBs between Q326 and either D18 or W21 of ET3 (Supplemental Figure 14A), resulting in ET3 affinity that was decreased to the level observed in ETAR-WT. Moreover, R326Q substitution in ETAR-WT almost abrogated responsiveness to ET3 (Figure 7D, upper). On the other hand, R326Q substitution did not affect intracellular Ca2+ mobilization in response to ET1 in ETAR -Y129F and ETAR -E303K, despite the loss of R326 HBs (Figure 7D, lower and Supplemental Figure 14B). These results indicate that HBs involving R326 are less critical in binding to ET1 than to ET3. The narrow peak of the R326Cα-D18Cα distance corresponds to ETAR activation by ET3 (Figure 7E, upper). However, in the case of ET1, this distance does not correlate with the activity; that is, HBs between R326 and ET1 are not essential for activation by ET1 (Figure 7E, lower).

Figure 7

ETAR-Y129F prevents dissociation of ET3 by increasing probability of reduced distance between helices. (A) Comparison of representative structures of ET3 bound to ETAR-Y129F (active, pink) and Na-bound ETAR-Y129F (inactive, grey). Dashed line connects D18Cα of ET3 and R326Cα of ETAR-Y129F. (B) Probability distribution of distances between D18 of ET3 and R326 of ETAR-WT or -Y129F (grey and light pink, respectively), overlaid with the distribution of probabilities of hydrogen bonding between these residues in each case (dark pink for ETAR-Y129F and black for ETAR-WT). (C) Schematic representation of HBs between ET3 residues (orange ovals) and TM residues of ETAR (rectangles). The solid lines between pairs of residues indicate the presence of HBs, and the thickness of the line represents the probability of the HB. HB probabilities of less than 5.00% are omitted, including 4.00% for R326-W21 and 2.27% for R326-D18 for ETAR-WT. Yellow dashed line shows the main chain of the ET3 peptide. (D) Intracellular calcium mobilization assay in HeLa cells. R326Q mutants are shown in bold lines. Changes in calcium concentrations in response to each ET3 (upper) or ET1 (lower) concentration are plotted as mean ± SEM (also see Supplemental Table 4) from more than 4 independent experiments. RFU, relative fluorescent unit. (E) The distance between D18 Cα of ET3 (upper) or ET1 (lower) and R/Q326 Cα of ETARs. R326Q mutants are shown in bold lines. (F) Presumed mechanism for the enhanced affinity of ETAR-Y129F for ET3. The number of water molecules within the water network around Y129 decreases upon Y-to-F substitution, thereby increasing the probability of a state with a narrowed ligand pocket, leading to increased hydrogen bonding with ET3.

MFDA is due to ETAR gain-of-function mutations. Following the identification of 2 different ETAR missense variants, Y129F and E303K, in MFDA, we report here another patient with MFDA harboring the E303K mutation in a mosaic state and mouse models carrying the ETAR-Y129F or -E303K mutations, which mostly recapitulate morphological manifestations of MFDA. Strikingly, the majority of features observed in the mutant mice were rescued by heterozygous loss of Edn3, while the incus required homozygosity for the Edn3-null allele to recover its normal morphology. The variable gene dosage effect may reflect a regional difference in the intensity of ET3 expression or sensitivity to activation for normal development. This result, taken together with our previous finding on G protein selectivity (36), suggests that the ETAR-Y129F and -E303K phenotypes are caused by ET3-dependent gain-of-function mechanisms through the activation of Gαq/11. Indeed, both mutations enhanced signaling pathways downstream of Gαq in response to ET3, as assayed by intracellular calcium concentration, to a similar extent. By contrast, ligand binding assays and G protein activation assays indicated different mechanisms underlying the enhanced signaling for each mutation.

Some differences between patients and mouse models are noted. The most significant difference is alopecia in patients with MFDA but not in mouse models. Also, the mandible (especially the proximal region) is more dysplastic in patients with MFDA than the mutant mice (32). These differences may indicate that the extent and/or effect of ET3-induced ETAR hyperactivation is different between species. In addition, it is surprising that some manifestations of the patient harboring a p.E303K mutation in the present report are no less severe than the mouse phenotype in spite of low-level (8–11%) mosaicism, as estimated from peripheral blood DNA. Similarly, no clear differences in phenotypic severity were noted between patients with MFDA harboring a mosaic versus heterozygous p.Tyr129Phe mutation (32). Although tissue mosaicism rates can be variable in different tissues (45), it suggests that ETAR activation in only a small number of cells may be sufficient for the manifestation of gain-of-function phenotypes.

Mechanism of gain-of-function of the ETAR-E303K mutation. Our MD analysis revealed different mechanisms underlying gain-of-function in the E303K and Y129F ETAR mutants. ETAR-E303K is characterized by the combination of increased ET3 binding affinity and facilitated Gαq activity, to which changes in the intrahelical HBs are likely to contribute. In some class A GPCRs, an acidic residue at position 6.30 interacts with R3.50 within the conserved aspartate-arginine-tyrosine (DRY) motif, creating a HB-mediated ionic lock to stabilize the inactive state, and this ionic lock is broken during receptor activation (6, 7, 46–48). ETAR cannot make this TM3-TM6 ionic lock because the residue at position 6.30 is basic arginine (R301) (Supplemental Table 3), so R1833.50 forms a HB primarily with D1823.49(39%; data not shown). Instead, ETAR-WT forms the nonclassical ionic lock between TM6 and the TM7-H8 linker (S3738.57 or K3748.58, Figure 4, A and B), and the disruption of this ionic lock in ETAR-E303K produces a bimodal distribution of TMin (Figure 4C) which most likely reflects transitions between populations of inactive (intact ionic lock) and active states (broken ionic lock) of the receptor. This propensity to reside in the active state would explain the enhanced sensitivity in all the signaling assays (Figure 5).

In addition, lysine substitution at position 6.32 increases the positive charge of the Gα-binding pocket (Supplemental Figure 15A). Basic residues at this position are involved in the interaction with Gα through HBs, for example, bonds between R6.32 of the μ-opioid receptor and H5-L25 of the α5-helix in Gαi, and between K6.32 of β2AR and H5-E24 of Gαs (4, 50, 51). Considering the structural similarity of the H5 domain between Gαq and other Gα proteins (Supplemental Figure 15B), K3036.32 may strengthen the interaction with Gαq, leading to increased basal activity. Regarding the increased affinity to ET3, our MD simulation provides insights by showing a reciprocal linkage between the intracellular and extracellular sides of TM6 in ETAR-E303K, which is considered an allosteric effect on ligand binding. In the ternary-complex model of GPCR dynamics (Supplemental Figure 9), GPCRs adopt closed-active, high-affinity structures that hinder dissociation of the bound ligand in the presence of an agonist. G protein or active nanobody coupling to GPCRs also influences ligand binding affinity by inducing allosteric conformational changes of the orthosteric ligand binding site, leading to a closed-active conformation with narrowing of the extracellular side of TMs, even without ligands (3, 6, 7, 51). The E3036.32K substitution may increase the affinity of the orthosteric ligand binding site through a similar allosteric effect, which is potentiated by the increased basal G protein binding and activation. Thus, ET3/ETAR-E303K signaling may involve the mechanism of G protein–mediated enhancement of agonist affinity.

Mechanism of gain-of-function of the ETAR-Y129F mutation. Several class A GPCRs have a Na+/water pocket composed of ionic binding between D2.50 and a Na+ and surrounding water molecules (8, 9, 11). Y1292.53 is located 1 turn above D1262.50 and participates in the Na+/water network that maintains an inactive state. In our MD simulation with Na+, the sodium ion is retained by ionic bonds with D126, S362, and/or T172 (Supplemental Figure 16). The triple ionic bonds are decreased in ETAR-Y129F, but at least 1 or 2 ionic bonds are almost always present in this mutant, so phenylalanine substitution may not affect sodium retention. On the other hand, removing the hydroxy group through phenylalanine substitution decreases the number of water molecules constituting the Na+/water pocket, which increases the propensity to collapse the Na+/water pocket. This water decrease is accompanied by a lateral-inward shift of the extracellular portions of the TMs, increasing ET3 binding affinity by inhibiting dissociation from the binding pocket (Figure 7F). However, we could not rule out the possibility of the outward shift of the intracellular half of the TMs because we used the intermediate rather than the active state of ET3-bound ETBR (PDB:6IGK) as an available ligand-bound template. This template is restricted in the movement of the intracellular side of the TMs because of the large T4-lysozyme protein within intracellular loop 3 and several mutations for thermal stabilization. Although MD simulations of ETARs were performed excluding the T4-lysozyme protein, it remains possible that the movement of the intracellular side of active ETAR bound to ET3 was not adequately simulated.

Another caveat for the inference that a HB between Y1292.53 and D1332.57 could contribute to the water retention by removing bulky side chain from the water pocket space is the use of the ETBR template in which D1542.57 (corresponding to D1332.57 of ETAR) is substituted by alanine, which may be a limitation when the side chain of D1332.57 is simulated. Despite these limitations, MD simulations and experimental verification support the conclusion that the inward shifts of the extracellular portion of the TMs were considered to favor ET3 binding through interacting forces such as van der Waals forces and Coulomb’s force, given the relatively short distance between D18 of ET3 and R3266.55 of ETAR-Y129F.

ETAR-Y129F and ETAR-E303K showed increased affinity and responsiveness to ET3 due to allosteric influences from outside the ligand pocket, whereas ET1 showed no further pharmacological affinity enhancement. ET1 binding is possibly affected by the lid consisting of the N-terminal and extracellular loop as shown in ETBR(42–44), since the affinity for ET1 was well maintained upon the loss of HBs with R326, which, in contrast, is critical for ET3-ETAR binding. Thus, ET1 and ET3 clearly differ in the mechanism of affinity for the same receptor.

Our findings provide insight into how allosteric communication within GPCRs can affect ligand binding affinity, and how alteration of normal GPCR function can cause human diseases. Furthermore, the molecular details of ETAR structure and function uncovered here provide a basis for rational drug design aimed at manipulating endothelin receptor activity relevant to more frequent diseases, such as pulmonary hypertension.