Arteriosclerosis and emphysema develop in individuals with Schimke immuno-osseous dysplasia (SIOD), a multisystem disorder caused by biallelic mutations in SMARCAL1 (SWI/SNF-related, matrix-associated, actin-dependent regulator of chromatin, subfamily a-like 1). However, the mechanism by which the vascular and pulmonary disease arises in SIOD remains unknown.
We reviewed the records of 65 patients with SMARCAL1 mutations. Molecular and immunohistochemical analyses were conducted on autopsy tissue from 4 SIOD patients.
Thirty-two of 63 patients had signs of arteriosclerosis and 3 of 51 had signs of emphysema. The arteriosclerosis was characterized by intimal and medial hyperplasia, smooth muscle cell hyperplasia and fragmented and disorganized elastin fibers, and the pulmonary disease was characterized by panlobular enlargement of air spaces. Consistent with a cell autonomous disorder, SMARCAL1 was expressed in arterial and lung tissue, and both the aorta and lung of SIOD patients had reduced expression of elastin and alterations in the expression of regulators of elastin gene expression.
This first comprehensive study of the vascular and pulmonary complications of SIOD shows that these commonly cause morbidity and mortality and might arise from impaired elastogenesis. Additionally, the effect of SMARCAL1 deficiency on elastin expression provides a model for understanding other features of SIOD.
Schimke immuno-osseous dysplasia (SIOD, OMIM 242900) is an autosomal recessive disorder associated with arteriosclerosis
[1, 2]. It is characterized by prominent skeletal dysplasia, renal failure, T-cell immunodeficiency, facial dysmorphism, and hyperpigmented macules
[3–7]. Other features include osteoporosis, joint degeneration, hypothyroidism, abnormal dentition, bone marrow failure, thin hair, corneal opacities, atherosclerosis, cerebrovascular events (CVEs), and migraine-like headaches
[2, 6–10]. Severely affected patients usually die before 15 years of age from renal failure, infection, bone marrow failure, lung disease, or CVEs
SIOD is caused by loss of function mutations in the gene encoding for the chromatin remodeling enzyme SWI/SNF-related, matrix-associated, actin-dependent regulator of chromatin, subfamily a-like 1 (SMARCAL1)
. SMARCAL1 functions as an annealing DNA helicase at single to double strand transitions in DNA
 and as a DNA stress response protein
[13, 14]. It also interacts with replication protein A, participates in the resolution of stalled DNA replication forks, and modulates transcription
[14–18]. Despite advances in our understanding of the SMARCAL1 enzyme, the mechanism by which SMARCAL1 deficiency leads to SIOD remains undefined.
As renal transplantation and dialysis have prolonged the longevity of SIOD patients, cerebral ischemia from arteriosclerosis has increasingly contributed to morbidity and mortality
[7, 19]. Although treatment with anticoagulant or hemorheological medications can transiently decrease the frequency and severity of CVEs and transient ischemic attacks (TIAs), the vascular disease ultimately progresses
 and is not associated with detectable alterations in nitric oxide production or mitochondrial dysfunction
[20, 21]. Focal atherosclerotic plaques, generalized hyperplasia of the tunica media, and splitting and fraying of the internal elastic layer characterize the arterial pathology
Potential contributors to the arteriosclerosis include hypertension, hyperlipidemia, renal disease, and immune dysfunction
[3, 7, 22, 23]. However, the arterial pathology observed by Clewing et al. is most similar to that reported for osteopontin deficiency or for impaired elastogenesis
[1, 24–26]. Osteopontin is a cytokine that is induced by the WNT signaling cascade
, a cellular pathway that participates in the regulation of vascular smooth muscle cell proliferation
. Impaired elastogenesis arises either from mutations of ELN or from impaired function or expression of enzymes that process or bind elastin
. Mice heterozygous for Eln gene deletions show many features in common with SIOD patients, including systemic hypertension, pulmonary hypertension, aortic valve disease and frequent inguinal hernias
[7, 30, 31]. Further highlighting the possibility of impaired elastogenesis, the postmortem lungs of two SIOD patients showed enlarged air spaces or emphysematous changes, a common feature in disorders of elastogenesis
[7, 30, 31].
Given these observations, we hypothesized that osteopontin deficiency and/or impaired elastogenesis were the primary causes of the vascular and pulmonary disease associated with SIOD. To test these hypotheses and to determine the prevalence of vascular and pulmonary disease among SIOD patients, we reviewed the records of SIOD patients with identified SMARCAL1 mutations, delineated the arterial and pulmonary pathology and profiled gene expression in postmortem artery and lung. We identify reduced elastin expression and synthesis as a possible basis of the arteriosclerosis and pulmonary emphysema of SIOD patients.
Patients referred to this study signed informed consent documents approved by the Institutional Review Board of Baylor College of Medicine (Houston, TX, USA) or the University of British Columbia (Vancouver, BC, Canada). Clinical data for 65 SIOD patients were obtained from questionnaires completed by the attending physician and from medical records and summaries provided by that physician. Autopsy tissues were obtained according to the protocol approved by the University of British Columbia. The SMARCAL1 mutations of SIOD patients are listed in Table
Summary of pulmonary and vascular findings in SIOD patients withSMARCAL1mutations
Age at death
Cause of death
Age at onset
Lung dysfunction or pathology
Age at onset
Mild panlobular emphysema with dyspnea, pulmonary hypertension, restrictive lung disease
Abbreviation: +, feature present; -, feature not present; BMT, bone marrow transplant; CVE, cerebrovascular event; CMV, cytomegalovirus; EBV, Epstein-Barr virus; F, female; HSV, herpes simplex virus; M, male; NA, not applicable; NR, not reported; TIA, transient ischemic attack.
A[?] represents alleles with noncoding SMARCAL1 mutations as described by Clewing et al.
BNo bacteriologic or viral proof.
CInfection of peritoneal dialysis fluid.
The propositus was a 5.4-year old boy with Schimke immuno-osseous dysplasia (Additional file
1). He was born at 35 weeks gestation to healthy non-consanguineous parents by Cesarean section for intrauterine growth retardation and oligohydramnios. At 1 year of age, he underwent surgery to repair an inguinal hernia. At 3.2 years of age, he had surgery for bilateral hip dysplasia. Beginning in his third year, he developed recurrent migraine-like headaches with aura, vomiting and hemiplegia. By 4 years of age, he manifested recurrent TIAs, although he had normal brain magnetic resonance imaging and angiography as well as normal electroencephalogram studies. At 4.5 years of age, he developed nephrotic syndrome that progressed to end-stage renal disease requiring dialysis. At 5.4 years, he was admitted to hospital for fever, anemia, hypoxia, and respiratory distress. His respiratory status rapidly worsened despite antibiotics and mechanical ventilation. He died from respiratory failure without identification of its cause. Postmortem studies showed alveolar damage, alveolitis, bronchitis, and dilated air spaces as well as arteriosclerosis, atherosclerosis, and cardiac left ventricular hypertrophy.
Patients SD60 and SD84
SD60 was a 13.7-year old boy and SD84 was a 23-year old man. Both have been described previously
The propositus is a 36-year old man with mild SIOD (Additional file
1); he was described by Gilchrist et al. at 16 years of age
. Since then, he has had bilateral hip and aortic valve replacement and respiratory insufficiency requiring oxygen supplementation. He has no history of smoking or exposure to cigarette smoke. His spirometry and diffusion studies show signs of both restrictive and obstructive pulmonary disease (Additional file
2). The former is consistent with his skeletal dysplasia and the latter is explained by the mild panlobular emphysema identified by computed tomography (Figure
1A, B). The severity of his respiratory distress has been disproportionate to his lung pathology, and is explained by Type I pulmonary arterial hypertension detected on right heart catheterization.
RNA isolation and reverse transcription
For cultured cells, RNA was extracted from 1 X 107 cells using the RNeasy Mini Kit (Qiagen, Mississauga, ON, Canada). For tissues, RNA was extracted from flash frozen tissue pulverized with a Bessman tissue pulverizer and lysed with TRIzol reagent (Invitrogen, Burlington, ON, Canada) according to the manufacturer’s specifications. Subsequently, the RNeasy Mini Kit (Qiagen, Mississauga, ON, Canada) was used to purify the RNA. Residual genomic DNA was removed by DNase I digestion.
Control aorta RNA pooled from 4 unaffected individuals ranging in age from 27–45 years was purchased from Clontech (636546, Lot no. 9052725A, Mountain View, CA, USA). Control lung RNA pooled from 3 unaffected individuals ranging in age from 32–61 years was purchased from Clontech (636643, Lot no. 8101369A, Mountain View, CA, USA).
RNA from formalin-fixed paraffin-embedded umbilical cord was isolated using the Ambion RecoverAll Total Nucleic Acid Isolation Kit (AM1975, Life Technologies, Burlington, ON, Canada) according to the manufacturer’s specifications.
Reverse transcription was performed with the qScript™ cDNA Synthesis Kit (Quanta Biosciences, Gaithersburg, MD, USA) or the RT2 First Strand Kit (SABiosciences, Mississauga, ON, Canada) using 500 ng of RNA per reaction according to the manufacturer’s specifications.
Following reverse transcription, 1.5 μl of cDNA served as template for each reaction and was amplified with the HotStarTaq Master Mix Kit (Qiagen, Toronto, ON, Canada). The following conditions were used for amplification: 1 cycle of 95°C for 15 min, followed by 30 cycles of 94°C for 30 s, 55°C for 30 s, 72°C for 1 min, and a final extension at 72°C for 10 min. PCR was performed using the primers listed in Additional file
Gene expression array
The Atherosclerosis (PAHS-038) RT2 Profiler™ PCR Array from SABiosciences (Mississauga, ON, Canada) was used to assess differences in gene expression between control and SIOD aortic mRNA according to the manufacturer’s instructions.
SsoFast EvaGreen Supermix (Bio-rad Laboratories, Mississauga, ON, Canada) or RT2 Real-Time™ SYBR Green/Rox PCR master mix (SABiosciences, Mississauga, ON, Canada) was used with the ABI 7500 Fast Real-Time PCR System for quantitative PCR. The primer sequences are listed in Additional file
ELN mutation analysis
Genomic DNA was extracted from the aorta of SD120 using the DNeasy Tissue Kit (Qiagen, Toronto, ON, Canada) according to the manufacturer’s specifications. The 34 exons of ELN were amplified with the HotStarTaq Plus Master Mix Kit (Qiagen, Toronto, ON, Canada). The following conditions were used for amplification: 1 cycle of 95°C for 5 min, followed by 35 cycles of 94°C for 30 s, 55°C or 60°C for 30 s, 72°C for 45 s, and a final extension at 72°C for 10 min. PCR was performed using the primers listed in Additional file
3. Unincorporated primers and nucleotides were removed using ExoSAP-IT reagent (USB, Cleveland, OH, USA).
Sanger capillary sequencing was used to sequence the PCR products (Macrogen, Seoul, Korea), and the sequences were aligned and analyzed using Sequencher v.4.10.1 (Gene Codes, Ann Arbor, MI, USA). Mutation interpretation analysis was conducted using Alamut 2.0 (Interactive Biosoftware, San Diego, CA, USA).
Aortic smooth muscle cells (AoSMCs, CC-2571, Lonza, Walkersville, MD, USA) were grown in smooth muscle basal medium (SmBM) supplemented with 5% fetal bovine serum (FBS), epidermal growth factor (EGF), basic fibroblast growth factor (FGF-B), insulin, gentamicin, and amphotericin B (SmGM-2 BulletKit, CC-3182, Lonza, Walkersville, MD, USA).
Human iliac artery endothelial cells (HIAECs, CC-2545, Lonza, Walkersville, MD, USA) were grown in endothelial basal medium (EBM-2) supplemented with 5% FBS, EGF, FGF-B, vascular endothelial growth factor (VEGF), R3 insulin-like growth factor 1 (R3-IGF-1), hydrocortisone, ascorbic acid, gentamicin, and amphotericin B (EGM-2-MV BulletKit, CC-3202, Lonza, Walkersville, MD, USA).
Aortic adventitial fibroblasts (AoAFs, CC-7014, Lonza, Walkersville, MD, USA) were grown in stromal cell basal medium (SCBM) supplemented with 5% FBS, FGF-B, insulin, gentamicin, and amphotericin B (SCGM BulletKit, CC-3205, Lonza, Walkersville, MD, USA).
Normal human lung fibroblasts (NHLFs, CC-2512, Lonza, Walkersville, MD, USA) were grown in fibroblast basal medium (FBM) supplemented with 2% FBS, FGF-B, insulin, gentamicin, and amphotericin B (FGM-2 BulletKit, CC-3132, Lonza, Walkersville, MD, USA).
Immunostaining of cultured cells was performed as previously described
. 5 x 105 cells were grown overnight on a coverslip in a 6-well plate. With the exception of the aortic smooth muscle cells (AoSMCs), all cells were fixed with 4% paraformaldehyde (PFA) for 15 min at room temperature and permeabilized with 0.5% Triton X-100 for 15 min at room temperature. AoSMCs were fixed with 4% PFA and 0.15% picric acid for 20 min at room temperature, and permeabilized with 0.1% Triton X-100, 1% bovine serum albumin (BSA), and 10% normal horse serum in 1X phosphate buffered saline (PBS). All cells were blocked overnight with Blocker Casein in PBS (Pierce, Rockford, IL, USA) containing 10% normal horse serum at 4°C. The cells were then incubated with anti-SMARCAL1 (1:200)
, anti-α-smooth muscle actin (1:20, 1A4, Dako, Mississauga, ON, Canada), anti-VE-cadherin (1:100, 33E1, Leica, Richmond Hill, ON, Canada), anti-prolyl 4-hydroxylase (1:50, 5B5, Abcam, Cambridge, MA, USA), or anti-α-tubulin (1:400, DM 1A, Sigma-Aldrich, Oakville, ON, Canada) diluted in blocking buffer at 4°C for 24 h. Cells then were gently washed 4 times with PBS and incubated with Alexa Fluor-conjugated secondary antibodies Alexa 488 and Alexa 555 (1:1000, Molecular Probes, Burlington, ON, Canada) for 1 h at room temperature. Cells next were washed 4 times with PBS and mounted in Vectashield containing 4’,6-diamidino-2-phenylindole (DAPI, Vector Laboratories, Burlington, ON, Canada). Images were acquired using a 100×/1.30 oil Plan-NEOFLUAR objective lens, a Zeiss Axiovert 200 inverted microscope, a Zeiss AxiocamMR camera, and the Zeiss Axiovision imaging system.
Immunoblot analysis on cell lysates was performed as previously described
. Cell lysates were fractionated by 12% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to a polyvinylidene fluoride (PVDF) membrane. The membrane was blocked overnight at 4°C, using gentle agitation, in PBS containing 0.2% I-Block (Applied Biosystems, Foster City, CA, USA) and 0.1% Tween 20 overnight. Anti-SMARCAL1 (1:2000)
 and anti-glyceraldehyde 3-phosphate dehydrogenase (GAPDH, 1:2000, 6C5, Advanced ImmunoChemical Inc., Long Beach, CA, USA) were used as primary antibodies. Alkaline phosphatase-conjugated secondary antibodies (1:10000, Bio-rad Laboratories, Mississauga, ON, Canada) were used to detect the primary antibodies. The bound antibody was detected by chemiluminescence using CDP-Star (Applied Biosystems, Streetsville, ON, Canada) according to the manufacturer’s specifications. GAPDH was detected as a loading control.
Immunoblot analysis on human tissue was performed as previously described
. Anti-elastin binding protein (EBP, 1:200, a kind gift from Dr. Amelia Morrone, University of Florence, Florence, Italy)
[34, 35] and anti-GAPDH were used as primary antibodies. EBP expression in the aortas of two SIOD patients was compared to that of a control aorta protein medley pooled from 49 unaffected individuals ranging in age from 15–65 years and purchased from Clontech (635310, Lot no. 5110079, Mountain View, CA, USA). EBP expression was normalized to expression of GAPDH for each sample. Densitometry of three independent replicates was conducted using the Kodak 1D Image Analysis Software version 3.6.
Tissue immunohistochemistry and staining
Formalin-fixed, paraffin-embedded sections were cut at 5 microns. Following deparaffinization and rehydration, heat induced epitope retrieval was conducted with sodium citrate buffer (10 mM sodium citrate, 0.05% Tween 20, pH 6) or tris-ethylene diamine tetraacetic acid (EDTA) buffer (10 mM Tris base, 1 mM EDTA, 0.05% Tween 20, pH 9). For immunohistochemical detection of elastin, proteolytic induced epitope retrieval was conducted with 0.4% pepsin at 37°C for 15 min. Endogenous peroxidases were inactivated by preincubating the sections with 0.3% H2O2 in methanol. Non-specific protein binding was blocked by preincubation at 4°C with blocking buffer (10% horse serum in TBS-T (10 mM Tris–HCl, 150 mM NaCl, 0.01% Tween 20, pH 7.4)) for 24 h. Sections were then incubated with anti-SMARCAL1 (1:200)
, anti-CD3 (1:50, MRQ-39, Cell Marque, Rocklin, CA, USA), anti-CD20 (1:50, L26, Cell Marque, Rocklin, CA, USA), anti-CD68 (1:250, KP1, Dako, Mississauga, ON, Canada), anti-α-smooth muscle actin (1:500, 1A4, Dako, Mississauga, ON, Canada), or anti-elastin (1:50, BA-4, Abcam, Cambridge, MA, USA) diluted in blocking buffer at 4°C for 24 h. Sections were then washed 5 times with TBS-T and incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies (EnVision+ System, Dako, Mississauga, ON, Canada) for 30 min at room temperature. Sections were then washed 3 times with TBS-T and 3,3’-diaminobenzidine (DAB, EnVision+ System, Dako, Mississauga, ON, Canada) was subsequently used as an HRP substrate. Sections were counterstained in Mayer’s Hematoxylin (Sigma, Oakville, ON, Canada).
Histochemical stains on tissue sections included a modified Verhoeff van Geison elastic stain (HT25A, Sigma, Oakville, ON, Canada) for elastic fibers and a periodic acid-Schiff stain (395B, Sigma, Oakville, ON, Canada) for neutral glycosaminoglycans. Images were acquired using a 5x/0.15 Plan-NEOFLUAR, 10x/0.45 Plan-APOCHROMAT, 20x/0.75 Plan-APOCHROMAT, 63x/1.4 oil Plan-APOCHROMAT, or 100x/1.30 oil Plan-NEOFLUAR objective lens on a Zeiss Axiovert 200 inverted microscope, a Zeiss AxiocamHR camera, and the Zeiss Axiovision imaging system.
Fastin elastin assay
The elastin content of arterial tissue was quantified using the Fastin Elastin Assay Kit (F2000, Bicolor Life Science Assays, United Kingdom). Tissue samples were flash frozen and pulverized with a Bessman tissue pulverizer, weighed and digested with 0.25 M oxalic acid at 95°C for six 1 h time periods. Elastin concentration in pooled supernatants was calculated from the elastin standard curve and the total elastin per wet weight of each sample was determined according to the manufacturer’s specifications.
Arterial thickness analysis
Analysis of the aortic intimal and medial thickness was carried out using the Zeiss Axiovision imaging system and software to measure the width of the tunica intima and the tunica media. Four random images of each sample were taken and 5 random measures were taken for both the tunica intima and tunica media for each image. Measures of the tunica intima were taken from the luminal edge of the endothelium to the internal elastic lamina perpendicular to the internal elastic lamina; measures of the tunica media were taken from the internal elastic lamina to the boundary between the tunica media and the tunica adventitia perpendicular to the internal elastic lamina. Ratios were calculated comparing the widths of the tunica intima and tunica media of SIOD patient aortas to that of age-matched control aortas.
A transthoracic echocardiogram was obtained by a Philips iE33 echocardiography machine using an S8 phased array ultrasound transducer probe with the patient in the supine and left lateral position. M-mode, 2D, color Doppler, pulse wave Doppler and continuous wave Doppler were obtained. Standard views including long axis view, short axis view, four chambers, subcostal and suprasternal notch views were obtained.
The following measurements of the aorta in systole were obtained in accordance to the American Society of Echocardiogram guidelines in real time and confirmed postmortem
: the aortic valve, the aortic root at the sinus of Valsalva, the sinotubular junction and the ascending aorta. Body surface area and Z scores were calculated offline using the Haycock and Halifax formula, respectively
Pulmonary function testing
Patient SD16 performed complete pulmonary function tests that met the American Thoracic Society (ATS) criteria for acceptability
. This included spirometery, lung volumes measured via plethsysmography, and diffusion capacity measured via nitrogen washout.
Quantitative data are presented as the mean ± 1 standard deviation calculated from a minimum of 3 independent replicates. Data were analyzed by the paired 2-tailed Student’s t-test or the one-way analysis of variance (ANOVA) followed by the Tukey post hoc test where appropriate. A p-value of less than 0.05 was considered statistically significant.
Pulmonary and vascular disease is common in SIOD
Among SIOD patients with SMARCAL1 mutations, 22 of 51 (43.1%) patients had lung disease (Table
1). Obstructive lung disease was present in 7 (13.7%) patients, including 3 (5.9%) with asthma or reactive airway disease, 1 (2.0%) with bronchiectasis, and 3 (5.9%) with emphysematous changes (Table
Regarding the vascular disease, 32 of 63 (50.8%) patients had clinical symptoms of cerebral ischemia and 6 of 51 (11.8%) patients had documented pulmonary hypertension. Twenty-five of 58 (43.1%) patients had CVEs and 27 of 59 (45.8%) patients had TIAs (Table
1). For 7 patients, the onset of cerebral ischemia preceded the development of renal disease or hypertension, and among patients who received a renal transplant, cerebral ischemia worsened despite renal transplantation (data not shown).
Of the 65 patients with SMARCAL1 mutations, 47 have died. Six (12.8%) died from pulmonary complications, and 7 (14.9%) died from vascular disease (Table
Histopathology of the SIOD aorta shows fragmented elastin fibers and hyperplasia of the tunica intima and media
To define better the histopathology of SIOD blood vessels, we analyzed postmortem arterial tissue from three individuals with SIOD (SD60, SD84 and SD120). Verhoeff van Gieson staining showed fragmented elastin fibers compared to age-matched controls (Figure
2 and Additional file
4). The aorta, common iliac, and pulmonary arteries of SD60, SD84 and SD120 had marked intimal and medial hyperplasia accompanied by an increased number of elastic lamellae compared to age-matched controls (Figure
2 and Additional file
4). The aortic tunica intima of SD60 and SD120 were 2.6-fold (p-value = 3.5 × 10-13) and 1.4-fold (p-value = 2.1 × 10-3) thicker than age-matched controls, respectively; the tunica media of SD60 and SD120 were 1.3-fold (p-values = 5.6 × 10-21 and 7.2 × 10-15, respectively) thicker than age-matched controls. By echocardiogram, SD120 had increased diameter of the sinotubular junction compared to normal using the Halifax formula, although measures at other levels of the aortic root were within the normal range (Additional file
5). Immunostaining for α-smooth muscle actin showed an increased number of positive cells suggesting smooth muscle cell hyperplasia in the aortic intima of SD60 and SD120 (Additional file
Inflammation is not increased in the SIOD aorta
Since SIOD patients have an immune disorder and the inflammation of atherosclerosis causes smooth muscle cell hyperplasia
[40, 41], we also looked for evidence of arterial inflammation. Using CD68 as a macrophage marker, CD3 as a T-cell marker, and CD20 as a B-cell marker, immunostaining did not detect an inflammatory infiltrate within the arterial walls (Additional file
7) except for the CD68+ macrophages within the atherosclerotic lesions of SD84 (Additional file
Histopathology of the SIOD umbilical cord shows a fragmented internal elastic lamina
As longstanding hypertension, renal failure and hyperlipidemia of individuals SD60, SD84 and SD120 could be a cause of the arterial disease and are not present in individuals with SIOD at birth, we tested this hypothesis by studying the umbilical artery of a 15-week gestation fetus (SD133b) with biallelic SMARCAL1 mutations (Table
1). Compared to age-matched controls, the fetal umbilical arteries of SD133b had interrupted circumferential expression of tropoelastin and elastin suggesting an intrinsic problem with elastogenesis (Figure
3). Moreover, analysis of ELN mRNA expression in the umbilical cord of SD133b and two age-matched controls showed that the umbilical cord of both controls had 1.75- to 2.95-fold higher ELN mRNA expression compared to that of SD133b (Additional file
SMARCAL1 is expressed in the vascular smooth muscle, endothelial, and adventitial fibroblast cells of the arterial wall
The above findings suggest a local or cell autonomous basis for the arteriosclerosis in SIOD, and consistent with this mechanism, arteriosclerosis does not recur in the transplanted kidneys of SIOD patients
[1, 2]. As a first requirement for a cell autonomous mechanism, SMARCAL1 must be expressed within the affected tissues, and indeed, SMARCAL1 was expressed in the adventitial fibroblasts, smooth muscle cells, and endothelium of the normal human aorta, common iliac and pulmonary arteries (Figure
4A-C). It was also expressed in the nuclei of cultured aortic smooth muscle cells (AoSMCs), iliac artery endothelial cells (HIAECs), and aortic adventitial fibroblasts (AoAFs) (Figure
4D-K and Additional file
9). Given these findings, we hypothesized that the cell autonomous mechanisms of osteopontin deficiency or impaired elastogenesis could give rise to arteriosclerosis.
Osteopontin expression is not decreased in the SIOD aorta
Osteopontin is a direct target of the WNT pathway and deficient WNT signaling can lead to osteopontin deficiency
. Therefore, since the histopathology of aortas from Opn−/−;Ldlr−/− mice resembles that of SIOD patients
, we measured levels of SPP1 mRNA, which encodes osteopontin. Contrary to our hypothesis, SPP1 mRNA levels were increased by 2.6-fold in the aorta of SD120 compared to controls (Figure
5A and Additional file
Elastin binding protein is not decreased in the SIOD aorta
Based on the preceding, we hypothesized that the arteriosclerosis primarily arose from a defect of elastogenesis and was accentuated by hypertension, hyperlipidemia, and renal failure. One mechanism for impaired elastin fiber assembly is a reduction in the protective chaperone elastin binding protein (EBP)
[42, 43]. Elevated levels of glycosaminoglycans, which are found in the mucopolysaccharidoses like Morquio syndrome, Costello syndrome and Hurler’s disease, induce premature shedding of EBP and lead to impaired elastin fiber assembly
[44–46]. Although later studies have not confirmed mucopolysacchariduria as a consistent feature of SIOD
, we tested EBP levels since chondroitin-6-sulphaturia was initially described as a feature of SIOD by Schimke et al.. Immunoblotting showed that EBP levels in the aortic lysate from SD60 and SD120 were comparable to those of controls (Additional file
11), and PAS staining did not show evidence of the increased deposition of neutral glycosaminoglycans that is characteristic of mucopolysaccharidoses
[46, 47] (Additional file
Elastin mRNA and protein are markedly reduced in the SIOD aorta
To test whether the elastin fiber pathology directly arises from altered expression of ELN mRNA, we profiled its expression using the Atherosclerosis RT2 ProfilerTM PCR Array. ELN mRNA levels were 121-fold reduced in the aorta of SD120 (p-value = 0.0033; Figure
5A and Additional file
10), and total elastin protein, including soluble and insoluble elastin, was reduced by 63.7% in the aortic tissue lysate of SD120 (Figure
SMARCAL1 is expressed in lung myofibroblasts and ELN expression is markedly reduced in SIOD lung
Given that impaired elastogenesis may serve as a potential primary and cell autonomous cause of the arteriosclerosis in SIOD, we hypothesized that it might also be a predisposing factor for the emphysematous changes or enlarged air spaces observed in the lungs of SIOD patients. SMARCAL1 was expressed in pneumocytes and lung myofibroblast cells and in the nuclei of cultured normal human lung myofibroblasts (NHLFs) (Figure
4L-P and Additional file
9). Consistent with observations in the aorta, ELN mRNA was decreased 156-fold in the lung of SD120 compared to that of unaffected controls (p-value = 0.0023, Figure
ELN gene mutations are not the cause of the reduced elastogenesis in SIOD
To determine whether the decreased ELN mRNA in SD120 arises from mutations in ELN, we sequenced the ELN gene in the aorta of patient SD120. Among the 34 exons of the ELN gene, none were found to have pathogenic mutations. A heterozygous non-synonymous change was found in exon 20 (c.1264G>A, p.Gly422Ser). However, the nucleotide and amino acid of interest are weakly conserved; Align-GVGD and SIFT algorithms predict this variant unlikely to be pathogenic, and this variant has been reported as a single nucleotide polymorphism (SNP, rs2071307) in dbSNP XML build 135 with an average heterozygosity of 0.41. A homozygous intronic change was found in intron 20 (c.1315+17C>T), however this change was not predicted to alter splicing and has been reported as a SNP (rs2856728) with an average heterozygosity of 0.38.
Expression of ELN transcription factors is significantly altered in SIOD aorta and lung
We hypothesized that SMARCAL1 deficiency altered the expression of ELN either by direct effects on the ELN promoter or by alteration of ELN transcription factor expression. Compared to controls, the expression of ELN repressors MYBL2, JUN, and TNF was increased 9.9-, 4.2- and 3.0-fold, respectively, in the aorta of SD120 (Figure
6A and Additional file
12). Also, in the lung, the expression of all tested ELN activators was decreased 1.3- to 5.0-fold and the expression of the negative regulator FOSL1 was increased 3.4-fold (Figure
6B and Additional file
We have shown that SIOD patients have clinical and histopathological features of impaired vascular and pulmonary elastogenesis. This pathology correlates with decreased ELN gene expression and altered expression of ELN transcriptional regulators.
Considering the biochemical role of SMARCAL1
, we hypothesized that the altered expression of ELN arises from direct and indirect effects of SMARCAL1 deficiency on the ELN gene. SMARCAL1 deficiency could directly affect ELN gene expression by altering the local DNA structure of the ELN gene. As an annealing helicase
, SMARCAL1 might maintain the local DNA structure of transcribed regions and thereby regulate gene expression by modulating transcription factor binding
. Alternatively, SMARCAL1 deficiency could indirectly alter ELN gene expression by modulating the expression of upstream transcriptional regulators of the ELN gene. Consistent with the latter, we observed altered expression of negative and/or positive regulators of ELN transcription within the aorta and lung of patient SD120.
The histopathology of the arteries from SIOD patients revealed increased elastic lamellae, increased aortic wall thickness, and fragmented elastin fibers in SIOD. Since SIOD is a multisystem disease characterized by immune deficiency, hypertension, hyperlipidemia, and renal disease
, we explored cell non-autonomous mechanisms for the basis of the vascular disease. However, as suggested by the lack of recurrence of vascular disease in transplanted kidneys
[1, 2], we observed onset of vascular ischemia prior to the onset of hypertension and renal failure in some SIOD patients, no detectable inflammatory infiltrate in the aortic wall, and altered distribution of tropoelastin and elastin in a 15-week gestation SIOD fetus. We concluded therefore that a local or cell autonomous mechanism was the most likely cause of the arteriosclerosis.
Such potential mechanisms included osteopontin deficiency, EBP deficiency and impaired elastogenesis. Contrary to the first two hypotheses, the expression of osteopontin mRNA was increased and EBP levels were unaltered in SIOD arteries. Consistent with the third hypothesis, however, we found significantly decreased elastin expression in the aortic tissue of two SIOD patients, which was not a consequence of pathogenic mutations in the ELN gene. Based on these findings, we conclude that a primary defect in elastogenesis is a parsimonious mechanism of the vascular and pulmonary disease observed in SIOD.
Elastogenesis is critical for arterial and lung development and maintenance
[49, 50]. Besides being required for the proper development of the arterial wall, elastin fibers maintain the tensile and elastic integrity of blood vessel walls and regulate the proliferation, migration, and maturation of vascular smooth muscle cells
[51, 52]. In the lung, elastin fibers are also required for proper development and for elastic recoil
[53, 54]. Haploinsufficiency for ELN causes arterial stenosis and hypertension in supravalvular aortic stenosis and Williams-Beuren syndrome (WBS) as well as mild respiratory symptoms in WBS
[55–57]. Elastin deficiency also causes vascular disease, bronchiectasis, and emphysema in cutis laxa, a more severe defect of elastogenesis
. Similarly, mice heterozygous for deletion of the Eln gene have systemic and pulmonary hypertension, aortic valve disease, and frequent inguinal hernias
[30, 31]; all of which are observed with increased frequency in SIOD patients
. The increase in the number of elastic lamellae and the increased thickness of the tunica media observed in WBS and of mice heterozygous for deletion of the Eln gene were also seen in the SIOD arteries
[52, 58]. Of note, consistent with the later onset of arterial and lung disease in SIOD, the pathology in the SIOD tissue is milder than that typically observed for WBS; nonetheless, these pathological correlations suggest that impaired elastogenesis is a mechanism warranting further investigation as the cause of the arteriosclerosis and emphysematous pulmonary changes of SIOD.
Vascular and pulmonary disease are common causes of morbidity and mortality in SIOD, and consequently, individuals with SIOD should be evaluated and monitored for the development of vascular and pulmonary disease. Regarding the molecular basis of this pathology, we find that SMARCAL1 deficiency is associated with altered expression of ELN transcriptional regulators, severely decreased expression of elastin mRNA and protein and impaired elastogenesis. These observations suggest a mechanism by which SMARCAL1 deficiency affects the pathogenesis of SIOD and await confirmation in additional patients.
We are grateful to all of our patients and family members who have contributed to this study. We thank Theresa Sturby, Barbara A. Antalffy and Pauline Grennan for preparation of tissue. This work was supported in part by grants from the March of Dimes (6-FY02-136 to CF Boerkoel), the Gillson Longenbaugh Foundation (CF Boerkoel), the Dana Foundation (CF Boerkoel) and the New Development Award, Microscopy, and Administrative Cores of the Mental Retardation and Developmental Disabilities Research Center at Baylor College of Medicine (CF Boerkoel), the Burroughs Wellcome Foundation (1003400 to CF Boerkoel), the National Institute of Diabetes, Digestive, and Kidney Diseases, National Institutes of Health (R03 DK062174 and R21DK065725 to CF Boerkoel), New Investigator Award from the SickKids Foundation – Canadian Institutes of Health Research: Institute of Human Development, Child and Youth Health (CF Boerkoel), the Michael Smith Foundation for Health Research (CI-SCH-O1899(07–1) to CF Boerkoel), the Association Autour D’Emeric et D’Anthony (CF Boerkoel) and The Little Giants Foundation (CF Boerkoel). CF Boerkoel is a scholar of the Michael Smith Foundation for Health Research and a Clinical Investigator of the Child & Family Research Institute.
Provincial Medical Genetics Program, Department of Medical Genetics, Children’s and Women’s Health Centre of BC
Rare Disease Foundation
Department of Pathology, University of Oklahoma Health Sciences Center
Department of Pathology, Oregon Health and Science University
Department of Molecular and Human Genetics, Baylor College of Medicine
Department of Pathology, University of Washington
Department of Anatomic Pathology, University of British Columbia and Children’s and Women’s Health Centre of British Columbia
Department of Medicine, University of Alberta
Department of Pediatrics, University of Oklahoma Health Sciences Center
Néphrologie Pédiatrique, Hôpital d’Enfants, Centre Hospitalier Universitaire de Nancy
Department of Endocrinology & Metabolism, Kanagawa Children’s Medical Center
Department of Pediatric Nephrology, Nephro-Urology Research Center, Shiraz University of Medical Sciences
Institute of Mother and Child Healthcare of Serbia
Department of Pediatric Nephrology, VU University Medical Center
Département de Génétique, Centre Hospitalier Universitaire d’Angers
Medizinische Hochschule Hannover, Kinderklinik
Division of Genetics, Birth Defects and Metabolism, Children’s Memorial Hospital
Centre de Référence des Maladies Rènales Rares, Hospices Civils de Lyon and Université de Lyon
Serviço de Genética, Hospital Santa Maria, Centro Hospitalar Lisoboa Norte
Département de Pédiatrie, Hôpital Robert Debré
Department of Internal Medicine, Division of Endocrinology and Metabolism, Cerrahi Hospital
Pediatric Immunology & Hematology Unit, Necker Hospital
Department of General Pediatrics, Pediatric Nephrology, University Children’s Hospital Münster
Department of Medical Genetics, “Aghia Sophia” Children’s Hospital, Athens University Medical School
Unidad de Genética Médica, Servicio de Pediatría, Hospital Universitario Virgen de la Arrixaca
Oregon Institute on Disability & Development, Child Development and Rehabilitation Center, Oregon Health & Science University
Medical Genetics, Mayo Clinic
Department of Genetics, Kaiser Permanente
Mercy Pediatrics and Adolescent Clinic
Department of Pediatric Nephrology, University Hospitals Leuven
Department of Pediatrics, Immunology Program and Institute for Immunity, Transplantation, and Infection, Stanford University
Divison of Nephrology, Bambino Gesù Children’s Hospital and Research Institute
Department of Medical Genetics, Alberta Children’s Hospital
Department of Nephrology, Birmingham Children’s Hospital
Service de Pédiatrie, Centre Hospitalier Régional Universitaire Hôpital Saint-Jacques
Consulta de Genética, Hospital Pediátrico de Coimbra
Department of Medical Genetics, Pamukkale University Hospital
Division of Nephrology, Department of Pediatrics, Kosair Children’s Hospital, School of Medicine, University of Louisville
Cape Breton Regional Hospital
Institut für Humangenetik, Martin-Luther-Universität Halle-Wittenberg
Department of Neuropediatrics, Children’s Hospital, Ruhr-University Bochum
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