Abstract

Pathologically elevated serum levels of fibroblast growth factor-23 (FGF23), a bone-derived hormone that regulates phosphorus homeostasis, result in renal phosphate wasting and lead to rickets or osteomalacia. Rarely, elevated serum FGF23 levels are found in association with mosaic cutaneous disorders that affect large proportions of the skin and appear in patterns corresponding to the migration of ectodermal progenitors. The cause and source of elevated serum FGF23 is unknown. In those conditions, such as epidermal and large congenital melanocytic nevi, skin lesions are variably associated with other abnormalities in the eye, brain and vasculature. The wide distribution of involved tissues and the appearance of multiple segmental skin and bone lesions suggest that these conditions result from early embryonic somatic mutations. We report five such cases with elevated serum FGF23 and bone lesions, four with large epidermal nevi and one with a giant congenital melanocytic nevus. Exome sequencing of blood and affected skin tissue identified somatic activating mutations of HRAS or NRAS in each case without recurrent secondary mutation, and we further found that the same mutation is present in dysplastic bone. Our finding of somatic activating RAS mutation in bone, the endogenous source of FGF23, provides the first evidence that elevated serum FGF23 levels, hypophosphatemia and osteomalacia are associated with pathologic Ras activation and may provide insight in the heretofore limited understanding of the regulation of FGF23.

INTRODUCTION

Skin and skeletal lesions appearing in localized patterns are generally mosaic disorders that result from embryonic somatic mutations (1–3). The distribution and pattern of lesions are determined by the type of progenitor cell affected and the timing of mutation during embryogenesis (1,2). Epidermal nevi and congenital melanocytic nevi (EN and CMN) appear in a wide range of sizes from small subcentimeter lesions to extensive segmental lesions affecting a large fraction of the skin surface area (4,5). While both are typically limited to the skin, in rare syndromic forms including epidermal nevus syndrome (ENS), they can be accompanied by abnormalities in other organ systems, including the eyes, brain and bone (5–8). The additional systemic abnormalities almost exclusively occur in the setting of extensive skin surface area involvement, consistent with early embryonic somatic mutation in a multipotent progenitor (6–8). Since somatic mutations can occur at any point postfertilization, timing determines the relative potential of the affected cell (9,10). Mutations prior to germ layer specification, for example, could result in progeny capable of ectodermal, mesodermal or endodermal fates as is postulated to occur in ENS (11). Later embryonic mutations affect cells with more limited potential, and thus tend to be smaller and involve a single germ layer as in the skin limited form of EN and NS (11,12).

EN, nevus comedonicus and giant CMN have been found in association with precocious puberty, cognitive dysfunction, arterial stenosis and skeletal malformations (7,13–15). Depending on the combination of cutaneous lesions and associated abnormalities, various nomenclatures such as Schimmelpenning–Feuerstein–Mims syndrome (16,17), Becker's nevus syndrome (18) and ENS have been applied, but these disorders have significant clinical overlap, suggesting a common pathogenesis.

Hypophosphatemic rickets rarely occurs in EN (27 reported cases) and CMN (1 case) (14,19–39). All reported affected individuals had skin lesions over 10–60% of the body surface area with involvement of other organ systems variably reported. In addition, each exhibited skeletal dysplasia, and in all cases in which it was assessed, low levels of blood phosphate, phosphaturia and elevated blood fibroblast growth factor-23 (FGF23) (26,32,35). FGF23 is a bone-derived hormone that regulates phosphorus homeostasis and vitamin D metabolism at the level of the renal proximal tubule cell by reducing expression of sodium-phosphate cotransporters types IIa and IIc (NPT2a, NPT2c), inhibiting 1-alpha hydroxylase activity, and enhancing 24-hydroxylase activity (40,41). Thus both renal phosphate reabsorption and synthesis of 1,25 (OH)2 vitamin D3, which mediates the gastrointestinal absorption of phosphate, are impaired by elevated FGF23, collectively contributing to the development of rickets and osteomalacia (40,42). In the reported cases of hypophosphatemic rickets with skin lesions, skin involvement was widespread and abnormalities in other organ systems were frequently identified (14,27,28,30,33,36). Given the broad spectrum of cutaneous and systemic findings, we have designated this disorder cutaneous-skeletal hypophosphatemia syndrome (CSHS). The generally large size of these skin lesions and presence of systemic findings are consistent with a shared origin from an early somatic mutation.

Isolated and syndromic keratinocytic and sebaceous nevi, as well as melanocytic nevi have been shown to result from somatic activating mutations in codons 12, 13 and 61 of RAS gene family members, which cause cell proliferation via the MAP kinase pathway (43–46). Skin lesions from one case of Schimmelpenning syndrome with hypophosphatemic rickets were also found to have somatic activating mutations in HRAS (47). The Mendelian forms of hypophosphatemic rickets typically present in childhood and can be transmitted in an autosomal dominant, recessive or X-linked fashion (48–51). Almost all feature either elevated FGF23 or inappropriately normal FGF23 levels in the setting of hypophosphatemia. Autosomal dominant hypophosphatemic rickets results from missense mutations in FGF23 that affect the C-terminal RTHR cleavage site, blocking degradation and leading to increased circulating FGF23 levels (51). X-linked hypophosphatemia is caused by inactivating mutations in the endopeptidase-encoding PHEX gene (49), while autosomal recessive hypophosphatemic rickets 1 and 2 (ARHR1, ARHR2) result from mutations in DMP1 and ENPP1, respectively (48,50). Reports of activating mutations in FGF receptor 1 (FGFR1), as well as a single de novo translocation causing increased circulating levels of cleaved α-Klotho, a cofactor for FGF23 binding to FGFR1 in the renal tubules, have also been found to increase levels of FGF23 resulting in hypophosphatemic rickets (52,53).

The etiology of hypophosphatemia in the setting of skin lesions has remained unknown. Some reports have described improvement of phosphate homeostasis upon destruction of skin lesions via excision or ablative techniques, suggesting the skin to be a source of a phosphatonin (19,28,54). However, qRT–PCR using RNA extracted from affected skin did not demonstrate FGF23 expression in the skin, suggesting that the skin-derived phosphatonin may be an unknown substance or that the excess FGF23 is produced elsewhere (32). Others have postulated that the focal bone lesions may be the source of FGF23 in ENS (25). We have employed exome and Sanger sequencing of blood and tissue samples from a cohort of five patients, to determine the genetic mechanism of the pathophysiology of CSHS.

RESULTS

To identify the genetic basis of hypophosphatemia, elevated FGF23 levels, osteomalacia and skin lesions in CSHS, we investigated five such cases, four with EN affecting multiple developmental segments of the skin and a single case with a giant CMN (Table 1, Fig. 1). All cases exhibited skeletal findings that included foci of dysplastic bone and fractures; most had rickets, and available lesional bone tissue demonstrated osteomalacia (Fig. 2). Histologic examination of dysplastic bone revealed fibroblast-like spindle-shaped cells surrounded by a relatively dense collagen matrix, an appearance consistent with cells in the osteogenic lineage (Fig. 3). In addition, pathologic lesions were observed in other tissues (Supplementary Material, Fig. S1). Affected individual CSHS101 had a brainstem lipoma, a thyroid nodule and splenic hemangiomas; CSHS102 manifested subaortic valve stenosis; CSHS103 had an eccrine poroma; CSHS104 displayed a left intraventricular choroidal mass and a non-pigmented mass in the medial canthus of the right eye; CSHS105 had brain abnormalities including colpocephaly and periventricular white matter paucity. These findings were consistent with somatic mosaicism for mutations contributing to multisystem disease.

Table 1.

Elevated FGF23 and hypophosphatemia in the CSHS patient cohort

Patient Cutaneous findings Age Sex FGF23 (RU/ml) Phosphate (mg/dl) Calcium (mg/dl) ALKP (U/l) PTH (pg/ml) 
CSHS101 EN 276 2.0 9.1 007000 45.5 
CSHS102 EN 12 279 2.3 9.7 003660 47.4 
CSHS103 EN 15 527 1.5 9.3 006510 90.0b 
CSHS104 Giant CMN 795 1.5 8.5 01081 88.5b 
CSHS105 EN 16 104.5a 2.2 8.7 003460 47.9 
Patient Cutaneous findings Age Sex FGF23 (RU/ml) Phosphate (mg/dl) Calcium (mg/dl) ALKP (U/l) PTH (pg/ml) 
CSHS101 EN 276 2.0 9.1 007000 45.5 
CSHS102 EN 12 279 2.3 9.7 003660 47.4 
CSHS103 EN 15 527 1.5 9.3 006510 90.0b 
CSHS104 Giant CMN 795 1.5 8.5 01081 88.5b 
CSHS105 EN 16 104.5a 2.2 8.7 003460 47.9 

Cutaneous findings, age at time of reported laboratory values, sex and values confirming the diagnosis of CSHS are presented. All patients show elevated FGF23, hypophosphatemia, normocalcemia, elevated alkaline phosphatase (ALKP) and normal 25(OH)D (calcidiol). 1,25(OH)2D (calcitriol) levels are not shown since they were confounded by the concomitant treatment with oral calcitriol. Tubular reabsorption of phosphate (TRP%) values ranged from 73 to 85.7%, consistent with renal phosphate wasting.

Reference normal values: FGF23, average for <18 years: 25–140 RU/ml; phosphate: 3.0–4.5 mg/dl; calcium: 9.0–10.5 mg/dl; alkaline phosphatase (ALKP): 30–120 U/l; parathyroid hormone (PTH), 10–60 pg/ml (115–117).

aCSHS105 had the intact FGF-23 measured, which is recorded in pg/ml, rather than the intact + C-terminal FGF-23, measured in RU/ml. The normal range of intact FGF-23 is 8–54 pg/ml.

bCSHS103 and 104 presented with low vitamin D when values were measured. Given their normal calcium levels, the elevated PTH represents secondary hyperparathyroidism, the normal physiological response to low vitamin D.

Figure 1.

Clinical and histological features of CSHS. All photomicrographs are at ×10 magnification. (A and B) Affected individual CSHS101 is a 7-year-old Caucasian female who presented at birth with linear epidermal nevi restricted to the left side of her body distributed from the neck to the calf. Histopathology shows thickening of the epidermis (acanthosis) and papillomatosis. (C and D) CSHS102 is a 12-year-old Caucasian female with nevus sebaceus on the left side of her head and neck. Histology of lesional skin from the cheek shows sebaceous hyperplasia, thickening of the stratum corneum (hyperkeratosis) and papillomatosis. (E and F) CSHS103 is a 15-year-old black female with widespread keratinocytic epidermal nevi on the torso and sebaceous nevi on the scalp and cheek, with brown verrucous papules and plaques covering the scalp, face, torso and extremities, as well as linear white plaques on the scalp and torso. Histological examination shows marked sebaceous hyperplasia, hyperkeratosis and papillomatosis. (G and H) CSHS104 is a black female who presented with a giant melanocytic nevus covering the entire posterior torso at birth, extending across the flanks to the anterior chest. Round, raised, hairy and pigmented plaques determined to be satellitosis of the congenital nevi were located on her extremities. There was a 0.5 × 12 cm linear tan epidermal nevus on the left forearm (not shown). Histology of tissue from her back tissue shows melanocytes infiltrating the full thickness of the dermis, melanin deposition and hyperkeratosis. She died at 4 years of age from a large pericardial effusion that occurred during sedation for an MRI. (I and J) CSHS105 is a 16-year-old Caucasian male born with whorls of raised pink to tan plaques across the central right chest and nipple, as well as lesions extending around the flank towards the right back, all consistent with keratinocytic epidermal nevi. Histology of lesions shows hyperkeratosis, acanthosis and papillomatosis.

Figure 1.

Clinical and histological features of CSHS. All photomicrographs are at ×10 magnification. (A and B) Affected individual CSHS101 is a 7-year-old Caucasian female who presented at birth with linear epidermal nevi restricted to the left side of her body distributed from the neck to the calf. Histopathology shows thickening of the epidermis (acanthosis) and papillomatosis. (C and D) CSHS102 is a 12-year-old Caucasian female with nevus sebaceus on the left side of her head and neck. Histology of lesional skin from the cheek shows sebaceous hyperplasia, thickening of the stratum corneum (hyperkeratosis) and papillomatosis. (E and F) CSHS103 is a 15-year-old black female with widespread keratinocytic epidermal nevi on the torso and sebaceous nevi on the scalp and cheek, with brown verrucous papules and plaques covering the scalp, face, torso and extremities, as well as linear white plaques on the scalp and torso. Histological examination shows marked sebaceous hyperplasia, hyperkeratosis and papillomatosis. (G and H) CSHS104 is a black female who presented with a giant melanocytic nevus covering the entire posterior torso at birth, extending across the flanks to the anterior chest. Round, raised, hairy and pigmented plaques determined to be satellitosis of the congenital nevi were located on her extremities. There was a 0.5 × 12 cm linear tan epidermal nevus on the left forearm (not shown). Histology of tissue from her back tissue shows melanocytes infiltrating the full thickness of the dermis, melanin deposition and hyperkeratosis. She died at 4 years of age from a large pericardial effusion that occurred during sedation for an MRI. (I and J) CSHS105 is a 16-year-old Caucasian male born with whorls of raised pink to tan plaques across the central right chest and nipple, as well as lesions extending around the flank towards the right back, all consistent with keratinocytic epidermal nevi. Histology of lesions shows hyperkeratosis, acanthosis and papillomatosis.

Figure 2.

Radiographic features of CSHS. (A) A CT scan of the jaw of patient CSHS103 shows large areas of dysplastic bone (asterisks) that appear as lytic lesions. The skeletal disease in the mandible displays an aggressive expansion that displaces and resorbs the tooth roots. (B) A radiograph of the femurs of patient CSHS104 demonstrates healing fractures due to osteomalacic bone (solid arrows), evidence of rickets (ill-defined, frayed and widened growth plates, dotted arrows), and dysplastic bone with mixed lytic and sclerotic changes (asterisk). (C) A radiograph of the femur of patient CSHS101 at age 7 reveals a stretch of dysplastic bone with a primarily sclerotic appearance at this age (solid arrows). At a younger age, the same lesion was more lytic in nature (not shown). As a result of medical therapy, the growth plate is normal, with no evidence of rickets (dotted arrows). (D) A radiograph of the arm of patient CSHS102 shows multiple unhealed fractures (arrows). Dysplastic lesions with lytic changes are seen in some of the phalanges (asterisks).

Figure 2.

Radiographic features of CSHS. (A) A CT scan of the jaw of patient CSHS103 shows large areas of dysplastic bone (asterisks) that appear as lytic lesions. The skeletal disease in the mandible displays an aggressive expansion that displaces and resorbs the tooth roots. (B) A radiograph of the femurs of patient CSHS104 demonstrates healing fractures due to osteomalacic bone (solid arrows), evidence of rickets (ill-defined, frayed and widened growth plates, dotted arrows), and dysplastic bone with mixed lytic and sclerotic changes (asterisk). (C) A radiograph of the femur of patient CSHS101 at age 7 reveals a stretch of dysplastic bone with a primarily sclerotic appearance at this age (solid arrows). At a younger age, the same lesion was more lytic in nature (not shown). As a result of medical therapy, the growth plate is normal, with no evidence of rickets (dotted arrows). (D) A radiograph of the arm of patient CSHS102 shows multiple unhealed fractures (arrows). Dysplastic lesions with lytic changes are seen in some of the phalanges (asterisks).

Figure 3.

Dysplastic skeletal lesions from the rib of CSHS104. (A) Low power (×2) view from the pleural of the side of the rib shows a thin layer of cells that underlie the periostium along the length of the section of rib depicted (arrows). Asterisk denotes focus of dysplastic cells. (B) Higher power (×4) H&E of the area of dysplastic tissue marked by the asterisk in section (A) is shown. The lesion consists of a collection of fibroblast-like spindle-shaped cells in a relatively dense collagen matrix that appears to arise from the layer of dysplastic cells demonstrated in (A). (C) A Masson's trichrome stain of the same region seen in (B) displays both blue-staining collagen in the matrix surrounding the dysplastic fibroblast-like cells and an extensive area of osteoid surrounding the collection of dysplastic cells (asterisk). (D) A higher power (×10) view of the smaller area dysplastic cells seen in (B and C) show a collection of cells similarly arising from the layer of dysplastic cells seen in (A) as well as active osteoclast resorption of adjacent lamellar bone (arrows), suggesting that the dysplastic cells of CSHS can induce osteoclastogenesis. (E) A high power (×40) view of the region seen in (C) demonstrates collagen bundles connecting the osteoid with the pericellular matrix. The collagen bundles are parallel to the mineralizing surface (Sharpey's fibers) and interdigitate between the atypical but osteoblast-like cells that appear to have produced the adjacent osteoid.

Figure 3.

Dysplastic skeletal lesions from the rib of CSHS104. (A) Low power (×2) view from the pleural of the side of the rib shows a thin layer of cells that underlie the periostium along the length of the section of rib depicted (arrows). Asterisk denotes focus of dysplastic cells. (B) Higher power (×4) H&E of the area of dysplastic tissue marked by the asterisk in section (A) is shown. The lesion consists of a collection of fibroblast-like spindle-shaped cells in a relatively dense collagen matrix that appears to arise from the layer of dysplastic cells demonstrated in (A). (C) A Masson's trichrome stain of the same region seen in (B) displays both blue-staining collagen in the matrix surrounding the dysplastic fibroblast-like cells and an extensive area of osteoid surrounding the collection of dysplastic cells (asterisk). (D) A higher power (×10) view of the smaller area dysplastic cells seen in (B and C) show a collection of cells similarly arising from the layer of dysplastic cells seen in (A) as well as active osteoclast resorption of adjacent lamellar bone (arrows), suggesting that the dysplastic cells of CSHS can induce osteoclastogenesis. (E) A high power (×40) view of the region seen in (C) demonstrates collagen bundles connecting the osteoid with the pericellular matrix. The collagen bundles are parallel to the mineralizing surface (Sharpey's fibers) and interdigitate between the atypical but osteoblast-like cells that appear to have produced the adjacent osteoid.

Exome sequencing was performed on DNA samples from affected skin and peripheral blood leukocytes of individuals CSHS101, 102, 103 and 105, and on DNA extracted from archival affected skin from CSHS104 (Supplementary Material, Table S1). Genetic variants were annotated and compared with identify somatic mutations present solely within affected tissue and not in germline (blood) DNA. In the four samples with matched blood and skin lesion DNA, we found a single heterozygous somatic mutation, in each case a known activating mutation in either HRAS or NRAS (55–63). In addition, we found a known activating RAS mutation in the lesion that did not have matched blood DNA; in this case, the mutation was absent in DNA prepared from normal bone marrow, demonstrating its somatic origin. The mutations included c.182A>G, p.Q61R in NRAS; c.182A>G, p.Q61R in HRAS; c.37G>C, p.G13R in HRAS (Table 2). All these mutations were confirmed by direct Sanger sequencing (Supplementary Material, Fig. S2). No germline mutations were found in other genes implicated in congenital disorders of phosphate metabolism (FGF23, FGFR1, FGF2, FGF7, DMP1, DMP4, ENPP1, KL, MEPE, PHEX, SLC34A1, SLC34A3, CLCN5, VDR, OPN, SFRP4, GALNT3 and GALNT8 genes) (64–98). Furthermore, germline samples were searched for genes harboring novel protein-altering variants in more than one subject, and none were found (see Materials and Methods). Finally, plots of differences in B-allele frequency between tumor and blood did not deviate from 0.5 across the genome, excluding large segments with loss of heterozygosity in tissue (Supplementary Material, Fig. S3). Collectively, these findings suggest that somatic RAS mutation alone is sufficient to cause CSHS.

Table 2.

Somatic mutations in HRAS and NRAS identified by exome sequencing

Sample Gene Base change Protein change No. of reads in tissue
 
No. of reads in blood
 
P-value
 
  Ref. Non-ref. Ref. Non-ref. 
CSHS101 NRAS A>G Q61R 097 033 129 1.3 × 10−11 
CSHS102 HRAS G>C G13R 188 093 159 9.0 × 10−22 
CSHS103 HRAS A>G Q61R 119 054 034 1.2 × 10−5 
CSHS104 NRAS A>G Q61R 235 152 NA NA NA 
CSHS105 HRAS G>C G13R 029 015 069 1.3 × 10−6 
Sample Gene Base change Protein change No. of reads in tissue
 
No. of reads in blood
 
P-value
 
  Ref. Non-ref. Ref. Non-ref. 
CSHS101 NRAS A>G Q61R 097 033 129 1.3 × 10−11 
CSHS102 HRAS G>C G13R 188 093 159 9.0 × 10−22 
CSHS103 HRAS A>G Q61R 119 054 034 1.2 × 10−5 
CSHS104 NRAS A>G Q61R 235 152 NA NA NA 
CSHS105 HRAS G>C G13R 029 015 069 1.3 × 10−6 

The P-values denoting the significance of the differences in reference (ref.) and non-references (non-ref.) reads in tissue versus blood were calculated using a one-tailed Fisher's exact test. In all cases, the respective RAS mutation carried the lowest P-value among all SNVs, and was the only variant with a quality score of ≥50 and a P-value of <1.0 × 10−3. After correction for multiple testing, a value of 1.7 × 10−6 is considered the threshold for genome-wide significance. Given the biopsy technique, which samples both normal and affected tissue, admixture accounts for reduced mutant allele fraction in some samples.

To further examine the etiology of elevated FGF23 and hypophosphatemia in CSHS patients, we examined dysplastic bone from cases CSHS102 and CSHS104 using Sanger sequencing, and found the same HRAS G13R and NRAS Q61R mutations present in the patients' EN and CMN, respectively. The mutations were absent in normal bone (Fig. 4). While nevus tissue of individual CSHS 104 was strongly positive via immunohistochemical staining for MelanA, a transmembrane protein expressed by melanocytes, the dysplastic bone tissue was MelanA negative, confirming that it arose independently (Fig. 5).

Figure 4.

NRAS Q61R is present in dysplastic, but not normal, bone in CSHS102 and 104. (A) The region within the dotted line on a ×2 view of an affected rib from patient CSHS104 shows a representative region of dysplasia. DNA was extracted from cores of tissue excised from this region. (B) A ×20 view of the same region shown in (A) demonstrates an area of fibrous tissue composed of spindle-shaped cells set in a dense collagen matrix and in intimate association with the adjacent lamellar bone. (C) Sanger sequencing of DNA from affected bone tissue from patient CSHS104 shows the same heterozygous NRAS Q61R mutation that was found in the patient's cutaneous lesion (left panel). Sequence of DNA extracted from the patient CSHS104′s unaffected rib is wild type (right panel). (D) Sanger sequencing of DNA from a bone sample appearing grossly sclerotic from patient CSHS102 shows the heterozygous HRAS G13R mutation that was found in the patient's nevus (left panel). DNA from a region near the cortical bone is wild type (right panel).

Figure 4.

NRAS Q61R is present in dysplastic, but not normal, bone in CSHS102 and 104. (A) The region within the dotted line on a ×2 view of an affected rib from patient CSHS104 shows a representative region of dysplasia. DNA was extracted from cores of tissue excised from this region. (B) A ×20 view of the same region shown in (A) demonstrates an area of fibrous tissue composed of spindle-shaped cells set in a dense collagen matrix and in intimate association with the adjacent lamellar bone. (C) Sanger sequencing of DNA from affected bone tissue from patient CSHS104 shows the same heterozygous NRAS Q61R mutation that was found in the patient's cutaneous lesion (left panel). Sequence of DNA extracted from the patient CSHS104′s unaffected rib is wild type (right panel). (D) Sanger sequencing of DNA from a bone sample appearing grossly sclerotic from patient CSHS102 shows the heterozygous HRAS G13R mutation that was found in the patient's nevus (left panel). DNA from a region near the cortical bone is wild type (right panel).

Figure 5.

Affected skin, but not affected bone, shows staining for MelanA. Skin (A) and bone (B) specimens from patient CSHS104 confirmed to carry the NRAS Q61R mutation were sectioned and immunostained with melanocyte marker MelanA, to confirm the affected bone to be osteogenic and independent. Nuclei of abundant dermal melanocytes within a congenital nevus stain strongly brown for MelanA (A: ×10, ×20 insert), while a cellular region of dysplastic bone shows no staining (B: ×10, ×20 insert). A hematoxylin counterstain highlights nuclei in both panels.

Figure 5.

Affected skin, but not affected bone, shows staining for MelanA. Skin (A) and bone (B) specimens from patient CSHS104 confirmed to carry the NRAS Q61R mutation were sectioned and immunostained with melanocyte marker MelanA, to confirm the affected bone to be osteogenic and independent. Nuclei of abundant dermal melanocytes within a congenital nevus stain strongly brown for MelanA (A: ×10, ×20 insert), while a cellular region of dysplastic bone shows no staining (B: ×10, ×20 insert). A hematoxylin counterstain highlights nuclei in both panels.

Given prior reports suggesting the skin lesions could be a source of elevated serum FGF23 in CSHS, we next examined affected skin tissue from four cases in our cohort (101,102,103,104) for FGF23 expression using immunohistochemical staining. Immunostaining was performed using normal skin as a negative control and a tumor-induced osteomalacia (TIO) sample as a positive control. In addition, a sporadic nevus sebaceus sample arising from a somatic HRAS G13R mutation was examined (45). None of the skin samples demonstrated expression of FGF23, while the TIO specimen demonstrated strongly positive staining (Supplementary Material, Fig. S4).

DISCUSSION

We have described a cohort of five cases of hypophosphatemia, skeletal dysplasia with osteomalacia and cutaneous lesions in developmental mosaic patterns, accompanied by various associated clinical findings. We define CSHS as a multisystem skin-bone disorder in which all patients have elevated circulating levels of FGF23. Further, we present the first exome sequencing analysis of CSHS in which we identified somatic activating RAS mutations in skin lesions of all five cases and in affected dysplastic bone of two subjects.

A similar pattern of mosaicism is found in McCune–Albright syndrome (MAS), a condition in which affected individuals demonstrate elevated FGF23, focal dysplasia of the bone, and cutaneous café-au-lait lesions (99,100). Somatic activating mutations in GNAS, which encodes the signaling protein Gsα, cause MAS and have been found in the fibrous dysplasia lesions, affected skin and other affected tissues. In MAS, in vitro expression assays of mutant Gsα in osteogenic cells suggest a role for Gsα activation in FGF23 production and/or processing (101–105). Furthermore, just as the degree of clinical findings in MAS are dependent on the developmental timing and location of the postzygotic GNAS mutation during embryogenesis, the timing and location of the postzygotic RAS mutation in CSHS may determine the type, extent and distribution of mutated progeny cells in the body, and thus the spectrum of clinical phenotypes. Our data suggest that other rare clinical findings in patients with congenital nevi may also be attributable to somatic RAS mutations.

Our discovery of recurrent activating RAS mutations in tissues derived from different germ layers provides evidence that embryonic mutations in a common progenitor of skin and bone lead to CSHS, and suggests that the extracutaneous findings in CSHS and other ENS s result from the pleiotropic effects of RAS mutations acting in different tissues. Similar to postzygotic RAS mutations in a multipotent progenitor giving rise to both sebaceous and melanocytic nevi in phacomatosis pigmentokeratotica, RAS mutations found in CSHS are likely acquired during early embryogenesis in a cell destined to differentiate into both the skeletal and cutaneous systems (106). The absence of additional intersecting mutations in the exome data from our index cases and the presence of an activating RAS mutation in bone suggest that an activating RAS mutation is sufficient to drive hypophosphatemic rickets via increased production of FGF23. Support for the role of Ras in FGF23 regulation includes recent findings in a pre-osteoblast cell line and primary cultures of bone marrow stromal cells, which show that Ras is activated in response to hyperphosphatemia, an established stimulus for FGF23 secretion in vivo (107–109). Further support for the role of RAS mutations in FGF23 regulation is the recent finding of an activating somatic KRAS mutation (G12V) in a hepatic metastasis of an adenocarcinoma that was accompanied by elevated FGF23 levels and showed FGF23 expression on immunohistochemistry (110). This suggests similarity between CSHS and oncogenic osteomalacia (a.k.a.TIO), a disorder in which primarily mesenchymal tumors secrete sufficient amounts of FGF23 to cause hypophosphatemia (41). These reports implicate Ras proteins in the regulation of FGF23, and our findings in CSHS provide the first evidence, suggesting that activating RAS mutations in bone cause elevated FGF23, hypophosphatemia, focal skeletal dysplasia and osteomalacia.

MATERIALS AND METHODS

Human subjects

All subjects provided written consent to the study protocol, which was approved by the Yale Human Investigation Committee. Each subject provided punch biopsies of affected skin as well as a blood sample. For one deceased case, archival tissue was obtained. One biopsy was used for direct DNA preparation in parallel with blood, and the other embedded for subsequent frozen sectioning.

DNA isolation

Genomic DNA from EN was obtained from fresh skin biopsies or 1 mm cores of formalin-fixed paraffin-embedded (FFPE) specimens, using DNeasy Micro (Qiagen, Valencia, CA, USA) with added deparaffinization performed for FFPE tissue. Fresh bone samples were powderized in liquid nitrogen and digest overnight in 10% (w/v) extraction buffer (EDTA 0.5 m, 1% n-lauroyl sarcosinate) and 10% (w/v) proteinase K (Qiagen, Valencia, CA, USA), and DNA was extracted using a standard phenol–chloroform protocols. FFPE samples were deparaffinized before proceeding as done for fresh bone. Blood DNA was also isolated via a standard phenol–chloroform protocol.

Whole-exome sequencing

Bar-coded libraries of sheared DNA isolated from tissue and blood were prepared, and whole-exome capture was performed (EZ Exome 2.0, Roche, Nutley, NJ, USA) by the Yale Center for Genome Analysis. Illumina HiSeq 2000 and 2500 instruments were used for sequencing blood samples pooled 6 per lane and tissue samples pooled 4 per lane with 75 bp paired end reads. Resulting reads were aligned to hg19 human reference using Efficient Large-scale Alignment of Nucleotide Databases software (ELAND, Illumina). Aligned reads were subsequently processed via a Perl script, which trimmed sequence to the targeted intervals and removed PCR duplicates. Single nucleotide variants (SNVs), deletions and insertions were identified using SAMtools software (111), and all variants were annotated for functional impact using a Perl script (112). We filtered our data to examine tissue variants with ≥2 non-reference reads in tissue and ≤4 non-reference reads in blood. The resulting data was filtered in Excel to exclude variants in 1000 Genomes (release 05/2011) and the National Heart, Lung and Blood Institute exome database (release ESP6500) from further analysis. Tissue variants in each sample were then filtered to examine coding mutations (missense, nonsense, splice site SNVs and insertions and deletions) with SAMtools quality scores ≥50 and coverage ≥8 and which were not present in 2577 control exomes. We then employed a one-tailed Fisher's exact test to compare mutant and wild-type read numbers in tissue and blood (genome-wide threshold for significance ∼1.7 × 10−6, after Bonferroni correction for multiple testing of ∼30 000 genes). Aligned reads were examined with the Broad Institute Integrative Genomics Viewer (IGV) (113) to exclude variants resulting from alignment error.

Analysis of exome data for LOH

Exome CNV was employed to identify LOH segments of ≥1 Mbp with a P-value of ≤0.015 (114). LOH data were plotted by dividing the number of B-allele (non-reference) reads by the total number of reads independently for both tissue and blood at each SNV position. The difference in B-allele frequency values between tissue and blood was plotted against the genomic location.

Sanger sequencing

Verification of RAS mutations identified by exome sequencing was performed via standard PCR using Kapa 2G Fast polymerase (Kapa Biosystems, Woburn, MA, USA) and Sanger sequencing. The primers used are as follows:

  • HRAS Exon 2F: AGGTGGGGCAGGAGACCCTGTAG

  • HRAS Exon 2R: AGCCCTATCCTGGCTGTGTCCTG

  • HRAS Exon 3F: AGAGGCTGGCTGTGTGAACTCCC

  • HRAS Exon 3R: ACATGCGCAGAGAGGACAGGAGG

  • NRAS Exon 3F: GGGACAAACCAGATAGGCAGAAATGG

  • NRAS Exon 3R: TGGTAACCTCATTTCCCCATAAAGATTC

Immunohistochemistry

In order to exclude that the focus of dysplastic bone was the result of melanocytic metastasis and to confirm its osteogenic origin, the bone lesion was stained for melanocytic marker MelanA. Five-micrometer sections from FFPE tissue were deparaffinized using a xylene–ethanol gradient, rehydrated and rinsed in phosphate-buffered saline (PBS). Heat-induced antigen retrieval was performed by immersion in a modified pH 6.0 citrate buffer at 95°C for 30 min (Dako, Carpenteria, CA, USA). Slides were cooled, rinsed in PBS and endogenous peroxidase activity blocked by incubation in 3% H2O2 for 10 min, followed by washing, blocking in 10% normal goat serum and incubation with a 1:50 dilution of anti-MelanA primary antibody (IR633, Dako, Carpenteria, CA, USA). Slides were washed in PBS and incubated with an anti-mouse HRP secondary antibody for 1 h prior to substrate detection with 3,3′-diaminobenzidine tetrahydrochloride employed as a chromagen (LSAB+, Dako, Carpenteria, CA, USA). Hematoxylin with 1% ammonium hydroxide posttreatment was employed as a counterstain.

Investigation of the skin as potential source of FGF23 was done via staining affected skin tissue with FGF23 antibody (Immunotopics, San Clemente, CA, USA). Six-micrometer sections of frozen or FFPE lesional skin tissue were either fixed using a 1:1 mixture of acetone and methanol and rinsed in PBS or deparaffinized using a xylene-ethanol gradient, rehydrated and rinsed in PBS, respectively. For FFPE sections, heat-induced antigen retrieval was performed as previously described. Slides were cooled, rinsed and blocked in 10% normal donkey serum and 1% bovine serum albumin, prior to incubation for 2 h at room temperature with a 1:100 dilution of anti-FGF23 primary antibody (IR633, Dako, Carpenteria, CA, USA). Slides were washed in PBS and incubated with an anti-goat Cy3 fluorescent secondary antibody for 1 h at room temperature, and washed again. Slides were incubated with 1× DAPI diluted in PBS for 3 min and mounted using Mowiol.

SUPPLEMENTARY MATERIAL

Supplementary Material is available at HMG online.

FUNDING

This work was supported by a Doris Duke Charitable Foundation Clinical Scientist Development Award to K.A.C. and by the Yale Center for Mendelian Genomics (NIH U54 HG006504). Y.H.L. was supported by the Paul H. Lavietes, M.D. Student Research Fellowship from the Yale University School of Medicine. D.O. was supported by the “la Caixa” Fellowship Program. M.T.C. and R.I.G. were supported by the Division of Intramural Research, National Institutes of Dental and Craniofacial Research, NIH.

ACKNOWLEDGEMENTS

We thank Lynn Boyden for critical review of the manuscript, and Jing Zhou, Li Tian, Britt Craiglow, Carol Nelson-Williams, Gerald Goh, Vincent Klump and Brian Zhao for technical assistance. We also thank Thomas Carpenter, Veraragavan Eswarakumar and Karl Insogna for insightful correspondence on endocrinology and bone biology.

Conflict of Interest statement. None declared.

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Supplementary data