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Background The genetic cause of cerebral autosomal recessive arteriopathy with subcortical infarcts and leukoencephalopathy (CARASIL), which is characterized by ischemic, nonhypertensive, cerebral small-vessel disease with associated alopecia and spondylosis, is unclear.
Methods In five families with CARASIL, we carried out linkage analysis, fine mapping of the region implicated in the disease, and sequence analysis of a candidate gene. We also conducted functional analysis of wild-type and mutant gene products and measured the signaling by members of the transforming growth factor β (TGF-β) family and gene and protein expression in the small arteries in the cerebrum of two patients with CARASIL.
Results We found linkage of the disease to the 2.4-Mb region on chromosome 10q, which contains the HtrA serine protease 1 (HTRA1) gene. HTRA1 is a serine protease that represses signaling by TGF-β family members. Sequence analysis revealed two nonsense mutations and two missense mutations in HTRA1. The missense mutations and one of the nonsense mutations resulted in protein products that had comparatively low levels of protease activity and did not repress signaling by the TGF-β family. The other nonsense mutation resulted in the loss of HTRA1 protein by nonsense-mediated decay of messenger RNA. Immunohistochemical analysis of the cerebral small arteries in affected persons showed increased expression of the extra domain–A region of fibronectin and versican in the thickened tunica intima and of TGF-β1 in the tunica media.
Conclusions CARASIL is associated with mutations in the HTRA1 gene. Our findings indicate a link between repressed inhibition of signaling by the TGF-β family and ischemic cerebral small-vessel disease, alopecia, and spondylosis.
Cerebral autosomal recessive arteriopathy with subcortical infarcts and leukoencephalopathy (CARASIL) is characterized by nonhypertensive cerebral small-vessel arteriopathy with subcortical infarcts, alopecia, and spondylosis, with an onset in early adulthood.6,7,8 On neuropathological examination, arteriosclerosis associated with intimal thickening and dense collagen fibers, loss of vascular smooth-muscle cells, and hyaline degeneration of the tunica media has been observed in cerebral small arteries.7,8,9 These pathological findings resemble those seen in patients with nonhereditary ischemic cerebral small-vessel disease.7,8,9,10,11 We conducted a study to determine whether mutations in HTRA1, a gene encoding HtrA serine protease 1, cause CARASIL.
Methods
Subjects and Genetic Analysis
We enrolled a total of six probands from six consanguineous families of Japanese ancestral origin and some of their family members. The first five families were included in a linkage analysis, and one member of the fifth family and one of the sixth family underwent neuropathological examination. Ancestry was reported by the participant or family members.
We isolated genomic DNA from 11 subjects from five of the families with CARASIL: 5 probands, 3 unaffected siblings, and 3 parents. We performed a genomewide linkage analysis using 763 microsatellite markers (Applied Biosystems). Pairwise lod scores were calculated with the MLINK program of the LINKAGE software package (version 5.2) and the FASTLINK package (version 4.1).12,13 We established five new microsatellite markers — M1236, M1238, M1241, M1260, and M1264 — on the basis of simple-repeat information from the University of California, Santa Cruz, Human Genome Browser. Primer sequences of these markers are summarized in the Supplementary Appendix (available with the full text of this article at NEJM.org). We designed primer pairs for amplification of the nine coding exons of HTRA1.
We isolated genomic DNA from all participants, including healthy persons of Japanese ancestral origin, as determined by means of self-report. These control subjects were between 74 and 90 years of age and had no signs of dementia, as defined by the Mini–Mental State Examination. We obtained fibroblast specimens from four controls and from Subject II-2 in Family 1.
We obtained written informed consent from the affected persons and their family members and written informed consent from the controls. The institutional review board of Niigata University approved this study.
Assay of HTRA1 Protease Activity
To express HTRA1 in Escherichia coli as fusions with glutathione S-transferase, we subcloned wild-type or mutant HTRA1 complementary DNA (cDNA), lacking codons 1 through 140, into the vector pGEX 6P-3 (GE Healthcare). The N-terminus of HTRA1 is toxic to E. coli. Amino acid substitution of the serine protease motif S328A, which abolishes the protease activity in HTRA1, was used as a negative control.14 Glutathione S-transferase fusion proteins were overexpressed and purified. Protease activity, measured as fluorescein isothiocyanate–labeled substrate β-casein, was evaluated with the use of a QuantiCleave Fluorescent Protease Assay Kit (Pierce) and recombinant glutathione S-transferase–HTRA1. To eliminate the possibility that a deletion of the N-terminus in HTRA1 affects protease activity, we also performed the identical protease assay using the serum-free medium containing cells stably expressing full-length wild-type or mutant HTRA1 tagged with a green fluorescent protein at the C-terminus. Green fluorescent protein–tagged HTRA1 proteins were detected by means of anti–green fluorescent protein antibody (Medical and Biological Laboratories, Nagoya, Japan).
To assay the formation of a stable complex with 
1-antitrypsin, we transiently expressed 1-antitrypsin and either wild-type or mutated HTRA1 cDNA with a simian virus 5 peptide (V5) tag at the C-terminus in the human embryonic kidney cell line HEK293. These cells were grown in serum-free medium, and samples of the resultant conditioned medium were then immunoblotted with anti–V5 antibody.14
Expression of HTRA1 and NOG
Total RNA was isolated from specimens of whole blood or cultured skin fibroblasts, and cDNA was synthesized with the use of the High-Capacity cDNA Reverse Transcription kit (Applied Biosystems). We assayed the expression of HTRA1 messenger RNA (mRNA) in whole-blood samples by using gene-specific primers for HTRA1. To assay HTRA1 mRNA levels in cultured skin fibroblasts in relation to the expression of glyceraldehyde-3-phosphate dehydrogenase, we performed a real-time quantitative reverse-transcriptase polymerase-chain-reaction (RT-PCR) assay by using specific TaqMan probes and primer sets (Applied Biosystems). We assayed mRNA levels of the noggin gene (NOG) in cultured skin fibroblasts in relation to the levels of β-actin by using real-time quantitative RT-PCR and SYBR Green (Applied Biosystems) (for details, see the Methods section of the Supplementary Appendix).
Assay of Signaling by TGF-β Family Proteins
We used a site-directed mutagenesis system (GeneTailor, Invitrogen) to synthesize cDNA encoding mutant HTRA1 and cDNA encoding a constitutively active TGF-β1 proprotein (pro–TGF-β1 containing the activating amino acid mutations C223S and C225S).15 We then individually subcloned this cDNA into the vector pcDNA DEST-40 (Invitrogen). Constitutively active TGF-β1 was synthesized from pro–TGF-β1 containing the activating mutations C223S and C225S. We isolated cDNA from the SMAD family member 2 gene (SMAD2), obtained from a library of human whole-brain cDNA (Clontech), and subcloned it into the pcDNA DEST-40 vector. Luciferase assays were performed as previously described.16,17 Mouse C2C12 myoblasts (mesenchymal precursor cells, obtained from the American Type Culture Collection) were cotransfected with HTRA1-expression vectors, the pRL-TK renilla luciferase expression plasmid, and the following constructs: the (Smad binding element)4–firefly luciferase vector (TGF-β–responsive reporter vector) and vectors containing SMAD2, the SMAD family member 4 gene (SMAD4), and TGFB1 (encoding pro–TGF-β1 with the two point mutations C223S and C225S)15; the pGL3-Id985WT–firefly luciferase vector (bone morphogenetic protein [BMP]–responsive reporter vector)17 and vectors containing the SMAD family member 1 gene (SMAD1), SMAD4, and BMP-4 (encoding pro–BMP-4); and the pGL3-Id985WT–firefly luciferase vector17 and vectors containing SMAD1, SMAD4, and BMP-2 (encoding pro–BMP-2).18 Cell extracts were assayed for luciferase activity with the use of the Dual-Luciferase Reporter Assay System (Promega). The luciferase activity was corrected for transfection efficiency by dividing it by the pRL-TK renilla luciferase activity. Every sample was transfected in triplicate, and every experiment was repeated three times.
Phosphorylation of SMAD Proteins
HEK293 cells were cotransfected with vectors containing HTRA1 and the following constructs: vectors containing SMAD2, SMAD4, and TGFB1 (encoding pro–TGF-β1 with two point mutations [C223S and C225S]); vectors containing SMAD1, SMAD4, and BMP-4; and vectors containing SMAD1, SMAD4, and BMP-2.15,18 The cells were lysed in radioimmunoprecipitation assay buffer containing phosphatase inhibitor. We performed Western blotting to detect SMAD1, phosphorylated SMAD1, SMAD2, and phosphorylated SMAD2, using the corresponding anti-SMAD antibodies (Cell Signaling): anti-SMAD1, anti–phospho-SMAD1/5/8, anti-SMAD 2/3, and anti–phospho-SMAD2 antibodies.
Immunohistochemical Studies and In Situ Hybridization
We carried out immunoperoxidase staining on formalin-fixed, paraffin-embedded brain specimens obtained from two patients with CARASIL and from four controls (a 40-year-old woman with amyotrophic lateral sclerosis, an 84-year-old woman, a 62-year-old man with a stroke, and a 36-year-old woman with schizophrenia).8,9 The primary antibodies used were those against TGF-β1 (1:50, Santa Cruz), against versican (1:100, Seikagaku), and against the extra domain–A region of fibronectin (1:100, Abcam). The negative control was prepared with the use of nonimmune IgG as the primary antibody. We used cDNA encoding the extra domain–A region of fibronectin (spanning nucleotides 5404 through 5704 of fibronectin isoform 1 [region NM_212482.1 [GenBank] ]) as a template for digoxigenin-labeled antisense and sense-complementary RNA probes. The sense probe was used as a negative control. We carried out in situ hybridization on the paraffin-embedded sections by using the probes. After the sections had been washed and blocked, they were incubated with alkaline phosphatase–conjugated anti-digoxigenin antibodies, stained with 4-nitroblue tetrazolium chloride–5-bromo-4-chloro-3-indolyl phosphate solution (Roche), and counterstained with fast red.
Results
Subjects
Clinical characteristics of the six probands with CARASIL are listed in Table 1, and the pedigrees are shown in Figure 1A. The patients in Families 1 through 5 were enrolled initially, and the causative gene for CARASIL was identified. We then enrolled an additional subject with pathologically confirmed CARASIL, in a sixth family, to perform immunohistochemical analysis. On neuropathological examination of this additional subject, arteriosclerosis associated with intimal thickening and dense collagen fibers were observed in cerebral small arteries (Figure 1I).
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Genetic Analysis
From genomewide linkage analysis of the five families enrolled, we obtained maximal cumulative pairwise lod scores of 3.97 and 3.59 at D10S587 and D10S1656 (
=0.0); all patients were homozygous for these loci (Figure 1A). We then performed fine mapping of the region between D10S597 and D10S575 using D10S1483 and five established polymorphic microsatellite markers (M1236, M1238, M1241, M1260, and M1264; see the Supplementary Appendix) (Figure 1A and 1B). All probands were homozygous for the loci between M1238 and D10S1656, suggesting that the causative gene was located within this 2.4-Mb region.
We first selected HTRA1 as a candidate gene (Figure 1B) because it is expressed in the blood vessels, skin, and bone.19 We identified two homozygous nonsense nucleotide mutations: 1108C
T (resulting in a stop codon at position 370; amino acid mutation R370X) in Family 1 and 904CT (resulting in a stop codon at 302; R302X) in Family 2 (Figure 1C). We also identified two homozygous missense mutations: 889GA (predicted to result in the amino acid substitution V297M) in Families 3 and 4 and 754GA (predicted to result in the amino acid substitution A252T) in Family 5. In addition, we observed the homozygous nonsense mutation 904CT again in the proband of Family 6. The missense mutations V297M and A252T were located in the genic region encoding the serine protease domain (Figure 1C), and the amino acids predicted to be affected were either completely or largely conserved among the HTRA homologues HTRA1 through HTRA4 and among HTRA1 orthologues (Figure 1D). We did not find evidence of such mutations in the 125 controls.
Protease Activity of Mutant HTRA1
The level of protease activity in the mutant HTRA1 encoded by cDNA containing V297M, A252T, or R302X was 21% to 50% of the activity level in wild-type HTRA1. In contrast, HTRA1 encoded by a construct containing the R370X mutation had a protease activity level similar to that in wild-type HTRA1 (Figure 2A and 2B). HTRA1 attacks the reactive center loop of 1-antitrypsin, instigating the serine protease activity of 1-antitrypsin, which thereby mediates the formation of a covalent complex between the two molecules.14 We did not observe the formation of a stable complex between 1-antitrypsin and mutant HTRA1 encoded by cDNA containing V297M, A252T, or R302X. In contrast, wild-type HTRA1 and HTRA1 encoded by cDNA containing R370X did form stable complexes with 1-antitrypsin (Figure 2C).
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If a premature stop codon is located at least 50 to 55 nucleotides upstream of the exon–exon junction close to the 3' end, mRNA may become degraded through nonsense-mediated decay.20 Because the location of R370X fulfills this criterion for decay (Figure 1C), we determined whether R370X-containing HTRA1 mRNA is degraded by means of nonsense-mediated decay. The level of HTRA1 mRNA expression in fibroblasts from the patient with the R370X mutation was 6.0% of that in fibroblasts from control subjects, and treatment with cycloheximide, an inhibitor of nonsense-mediated decay, increased this level by a factor of four (Figure 3A). We did not detect HTRA1 protein in the culture medium of fibroblasts from the patient carrying the R370X mutation (Subject II-2, Family 1) (Figure 3B). Furthermore, analysis of HTRA1 in leukocytes from a heterozygous carrier of this mutation (Subject II-1, Family 1) showed the presence of wild-type HTRA1 mRNA only (Figure 3C).
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The serine protease activity of HTRA1 is necessary for inhibition of TGF-β family signaling.16 We therefore tested the ability of mutant variants of HTRA1 with missense amino acids (i.e., A252T and V297M) to repress signaling by the TGF-β family members BMP-4 and BMP-2 (Figure 4A, and Fig. 1 in the Supplementary Appendix). As expected, neither of the missense-mutated HTRA1 proteins repressed signaling by these molecules. To further investigate the ability of the mutant variants of HTRA1 to repress signaling by the TGF-β family members, we assayed phosphorylation of SMAD in these assays (phosphorylated SMAD is a downstream effector of the TGF-β–family signaling pathway), and observed that none of the mutant HTRA1 proteins repressed the subsequent phosphorylation of SMAD proteins (Figure 4B, and Fig. 2 in the Supplementary Appendix).
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Increased TGF-β signaling results in vascular fibrosis, with synthesis of extracellular matrix proteins, including the extra domain–A region of fibronectin and versican.22,23,24 In the patients with CARASIL who carried the R302X or A252T mutation, the tunica intima showed increased expression of the extra domain–A region of fibronectin (Figure 4E through 4H, and Fig. 4A in the Supplementary Appendix) and versican (Figure 4K, and Fig. 4B in the Supplementary Appendix), as compared with that in control subjects (Figure 4M, 4N, and 4O). The result was confirmed by an in situ hybridization assay that used a probe for the extra domain–A region of fibronectin (Figure 4I and 4J, and Fig. 5 in the Supplementary Appendix). Moreover, in the patients with CARASIL, the tunica media exhibited elevated expression of TGF-β1 (Figure 4L and 4P, and Fig. 4C in the Supplementary Appendix). These results indicate increased TGF-β signaling in the cerebral small arteries in patients with CARASIL.
Discussion
Signaling by members of the TGF-β family is closely associated with vascular angiogenesis and remodeling and has multifaceted roles in vascular endothelial and smooth-muscle cells, depending on the type of cell and extracellular matrix.25,26 Moreover, dysregulation of TGF-β–family signaling results in hereditary vascular disorders.26 Defective TGF-β signaling due to mutations in the TGF-β receptors leads to hereditary hemorrhagic telangiectasia, whereas activation of TGF-β signaling contributes to Marfan's syndrome and associated disorders.26 Our findings extend the spectrum of diseases associated with the dysregulation of TGF-β signaling to include hereditary ischemic cerebral small-vessel disease. In addition, the pathological findings in patients with CARASIL resemble those observed in patients with nonhereditary ischemic cerebral small-vessel disease with hypertension, suggesting that hypertension may increase TGF-β signaling.7,8,9,10,11,27 Thus, TGF-β signaling might underlie the molecular basis of nonhereditary ischemic cerebral small-vessel disease with hypertension.
Dysregulation of the inhibition of signaling by members of the TGF-β family also has been linked to alopecia and spondylosis, the other cardinal clinical features of CARASIL. Transgenic mice overexpressing BMP-4, BMP-2, and TGF-β exhibit hair loss or retardation of the development of hair follicles.28,29 Members of the BMP family are well-known regulators of bone formation, repair, and regeneration.30 Furthermore, overexpression of HTRA1 decreases BMP-2–induced mineralization, whereas reduced expression of HTRA1 accelerates mineralization.31 Although the loss of protease activity by HTRA1 on other substrates may be associated with the pathogenesis of CARASIL, our findings strengthen the hypothesis that increased signaling by the TGF-β family contributes to the pathogenesis of CARASIL.14,31,32,33 It remains unclear why disinhibition of signaling by TGF-β family members caused by mutant HTRA1 results in narrowly restricted clinical phenotypes. Possible explanations are tissue-specific regulation of signaling by the TGF-β family and tissue-specific expression of HTRA1.14,19,33,34
The molecular basis for regulation of TGF-β1 signaling by HTRA1 remains to be elucidated.16,35,36 TGF-β1 is synthesized as a proprotein (pro–TGF-β1) and is subsequently cleaved into latency-associated protein and mature TGF-β1 by proprotein convertase.26 The mature TGF-β1 is noncovalently bound to the latency-associated protein and is sequestered as a latency-associated protein–TGF-β1 complex in an extracellular matrix.26 The mature TGF-β1 is released from the latency-associated protein–TGF-β1 complex. Therefore, the TGF-β1 signaling is regulated by a balance among maturation, sequestration, and presentation. The elastin microfibril interfacer 1 protein (EMILIN1) inhibits TGF-β1 signaling by preventing the processing of pro–TGF-β1 into mature TGF-β1.37 In our study, the patients with CARASIL had increased expression of mature TGF-β1, suggesting that the HTRA1 may also prevent the processing of pro–TGF-β1 into mature TGF-β1, depending on its protease activity.
A single-nucleotide polymorphism in the promoter region of HTRA1, which is associated with elevated levels of HTRA1 expression, is a genetic risk factor for the neovascular form of age-related macular degeneration.38,39 We did not find macular degeneration in the persons with CARASIL.6,7,8 Although all our patients were younger than the typical age at the onset of the neovascular form of age-related macular degeneration, the absence of macular degeneration in the patients is consistent with the hypothesis that increased expression of HTRA1 contributes to age-related macular degeneration.6,7,8,38
Our results indicate that disinhibition of TGF-β–family signaling underlies the molecular basis of CARASIL. They also provide a basis for further investigation of therapeutic strategies for ischemic cerebral small-vessel disease, alopecia, and spondylosis.
Supported in part by grants from the Advanced Brain Science Project and Applied Genomics, the Center for Integrated Brain Medical Science, the Ministry of Education, Culture, Sports, Science, and Technology of Japan, the Japan Society for the Promotion of Science, and Niigata University.
Dr. Fukutake reports receiving lecture fees from Mitsubishi Pharma, Kissei, Sanofi, and Novartis. No other potential conflict of interest relevant to this article was reported.
We thank the patients and family members for their participation; M. Tsuchiya, Y. Hazama, Y. Koike, R. Izumita, and M. Nihonmatsu for technical assistance; and Drs. B. Vogelstein (Howard Hughes Medical Institute and Sidney Kimmel Comprehensive Cancer Center), T. Katagiri (Saitama Medical School Research Center for Genomic Medicine), and E. Schwartz (Institut für Biotechnologie, Martin Luther Universität Halle–Wittenberg) for their generous gifts of plasmids.
Source Information
From Niigata University, Niigata (K.H., A.S., H.N., A.M., A.Y., A.K., T.T., M.I., Y.Y., T.Y., T.I., R.K., M.N., O.O.); Kameda Medical Center, Kamogawa City (T.F.); Jichi Medical University, Tochigi (H.K., A.T., I.N.); National Center of Neurology and Psychiatry, Tokyo (K.A.); Nihon University School of Medicine, Tokyo (H.S.); Nagaoka-Nishi Hospital, Nagaoka (M.T.); Kashima Rosai Hospital, Kashima (Y.S.); Kasugai Municipal Hospital, Kasugai (M.H.); Minamikyushu National Hospital, Kagoshima (T.A.); Iida Municipal Hospital, Iida (S.Y.); Shinshu University School of Medicine, Matsumoto (S.I.); and University of Tokyo, Tokyo (S.T.) — all in Japan.
Dr. Hara and Mr. Shiga contributed equally to this article.
Address reprint requests to Dr. Onodera at the Brain Research Institute, Niigata 951-8585, Japan, or at onodera{at}bri.niigata-u.ac.jp.
References
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