Skip to main content
Device Global Header

Subscribe
or Renew

Advanced Search

Original Article

Inactivating Mutations in the 25-Hydroxyvitamin D3 1α-Hydroxylase Gene in Patients with Pseudovitamin D–Deficiency Rickets

List of authors.
  • Sachiko Kitanaka, M.D.,
  • Ken-ichi Takeyama, M.S.,
  • Akiko Murayama, M.D.,
  • Takashi Sato, M.S.,
  • Katsuzumi Okumura, Ph.D.,
  • Masahiro Nogami, M.S.,
  • Yukihiro Hasegawa, M.D.,
  • Hiroo Niimi, M.D., Ph.D.,
  • Junn Yanagisawa, Ph.D.,
  • Toshiaki Tanaka, M.D., Ph.D.,
  • and Shigeaki Kato, Ph.D.

Abstract

Background

Pseudovitamin D–deficiency rickets is characterized by the early onset of rickets with hypocalcemia and is thought to be caused by a deficit in renal 25-hydroxyvitamin D3 1α-hydroxylase, the key enzyme for the synthesis of 1α,25-dihydroxyvitamin D3.

Methods

We cloned human 25-hydroxyvitamin D3 1α-hydroxylase complementary DNA (cDNA) using a mouse 1α-hydroxylase cDNA fragment as a probe. Its genomic structure was determined, and its chromosomal location was mapped by fluorescence in situ hybridization. We then identified mutations in the 1α-hydroxylase gene in four unrelated patients with pseudovitamin D–deficiency rickets by DNA-sequence analysis. Both the normal and the mutant 1α-hydroxylase proteins were expressed in COS-1 cells and were assayed for 1α-hydroxylase activity.

Results

The gene for 25-hydroxyvitamin D3 1α-hydroxylase was mapped to chromosome 12q13.3, which had previously been reported to be the locus for pseudovitamin D–deficiency rickets by linkage analysis. Four different homozygous missense mutations were detected in this gene in the four patients with pseudovitamin D–deficiency rickets. The unaffected parents and one sibling tested were heterozygous for the mutations. Functional analysis of the mutant 1α-hydroxylase protein revealed that all four mutations abolished 1α-hydroxylase activity.

Conclusions

Inactivating mutations in the 25-hydroxyvitamin D3 1α-hydroxylase gene are a cause of pseudovitamin D–deficiency rickets.

Introduction

Hereditary pseudovitamin D–deficiency rickets, also known as vitamin D–dependent rickets type I, is characterized clinically by hypotonia, weakness, growth failure, and hypocalcemic seizures in early infancy.1 The patients also have hypocalcemia, radiologic findings typical of rickets, elevated serum parathyroid hormone concentrations, and generalized aminoaciduria.2,3 It is inherited as an autosomal recessive trait.

Two other inherited types of rickets are known,4 and the mutated genes in patients with these diseases have been identified. The diseases are hypocalcemic vitamin D–resistant rickets, also known as type II vitamin D–dependent rickets, in which the gene for the vitamin D receptor is mutated,5-7 and X-linked hypophosphatemic vitamin D–resistant rickets, in which the PEX gene (phosphate-regulating gene with homology to endopeptidases on the X chromosome) is mutated.8 In contrast, the molecular basis of pseudovitamin D–deficiency rickets has remained unclear, although the disease locus has been mapped to chromosome 12q14 by linkage analysis.9

Pseudovitamin D–deficiency rickets is distinguished from other types of hereditary rickets in that affected patients have low serum concentrations of 1α,25-dihydroxyvitamin D and normal or high concentrations of 25-hydroxyvitamin D.10,11 In these patients, physiologic doses of 1α,25-dihydroxyvitamin D3 but massive doses of vitamin D or 25-hydroxyvitamin D3 are required to cure the rickets.12 These findings suggest that the activity of renal 25-hydroxyvitamin D3 1α-hydroxylase,13,14 which converts 25-hydroxyvitamin D3 to 1α,25-dihydroxyvitamin D3, is defective in patients with pseudovitamin D–deficiency rickets. Whether this defect is due to abnormalities in the 25-hydroxyvitamin D3 1α-hydroxylase gene or to genetic abnormalities in other factors required for the full activity of this enzyme is unknown.13,15 Since direct enzymatic measurement of 1α-hydroxylase activity in these patients is difficult, the human 1α-hydroxylase gene was cloned in the present study to investigate genetic mutations in patients with pseudovitamin D–deficiency rickets.

Using a novel vitamin D receptor–mediated expression cloning method,16 we recently cloned 1α-hydroxylase complementary DNA (cDNA) from the kidneys of vitamin D receptor–deficient knockout mice in which the 1α-hydroxylase gene was overexpressed.17 In the present study, we cloned human 1α-hydroxylase cDNA, determined the structure of the gene and the chromosomal locus, and analyzed whether this gene is mutated in patients with pseudovitamin D–deficiency rickets.

Methods

Patients

Table 1. Table 1. Clinical and Biochemical Features in Four Patients with Pseudovitamin D–Deficiency Rickets.

We studied four unrelated Japanese patients with pseudovitamin D–deficiency rickets and four of their unaffected parents, one sibling, and one nephew. The diagnosis of pseudovitamin D–deficiency rickets in these patients was based on the early onset of hypocalcemia, radiologic findings characteristic of rickets, and the requirement of high doses of vitamin D and physiologic doses of 1α-hydroxyvitamin D3.2,3 The clinical data on these patients are summarized in Table 1. The diagnosis was confirmed by low serum concentrations of 1α,25-dihydroxyvitamin D determined when the patients were 10 to 20 years old, after the cessation of therapy with vitamin D2 or 1α-hydroxyvitamin D3. The parents of Patient 1 were second cousins; those of Patient 2 were first cousins; those of Patient 3 were unrelated and came from the same remote area; those of Patient 4 were unrelated and came from different areas.

The studies were approved by the appropriate institutional review committees, and all subjects gave informed consent.

Isolation of 1α-Hydroxylase cDNA and Gene

A human kidney cDNA library was prepared from poly(A)+RNA purified from normal kidney tissue.19 A total of 1×106 plaques were screened by hybridization with a mouse 1α-hydroxylase cDNA N-terminal fragment (0.7 kb).16 Two positive plaques were subcloned and sequenced automatically in a Prism 377 DNA sequencer (Applied Biosystems, Foster City, Calif.), with AmpliTaq DNA polymerase FS (Perkin–Elmer, Norwalk, Conn.) and dye terminator.

The genomic sequence of human 1α-hydroxylase was determined by sequencing the polymerase-chain-reaction (PCR) products amplified with three sets of primers specific to the exon and the untranslated region, with normal human leukocyte DNA as a template.

Fluorescence in Situ Hybridization

Three human genomic fragments covering all areas of the 1α-hydroxylase gene were used as probes. Fluorescence in situ hybridization was carried out on more than 20 chromosomes in prometaphase.20

PCR Amplification and Sequence Analysis of the 1α-Hydroxylase Gene

Genomic DNA of all patients and family members was extracted from peripheral white cells. Exons of the 1α-hydroxylase gene were amplified by PCR with specific primers derived from intronic sequences. Information about the sequence of the primers is available elsewhere (NAPS). AmpliTaq Gold (Perkin–Elmer) and its standard buffer were used in all reactions. All exons were amplified in a PCR thermocycler (Perkin–Elmer) by initial denaturation at 95°C for nine minutes, followed by 30 cycles at 95°C for one minute, 60°C for one minute, and 72°C for one minute. The corresponding PCR products were purified and sequenced directly in both directions.

Plasmid Construction

The wild-type human 1α-hydroxylase cDNA was introduced into the expression vector pcDNA3 (Invitrogen, San Diego, Calif.), and the patients' mutations were introduced with a site-directed mutagenesis kit (Quick Change, Stratagene, La Jolla, Calif.). The expression plasmid for the ligand-binding domain of the vitamin D receptor (VDR) fused to the GAL4 DNA binding domain [GAL4-VDR(DEF)] was prepared as described elsewhere.21

Assay of 1α-Hydroxylase Activity

1α-Hydroxylase activity was assayed both by vitamin D receptor–mediated transactivation assay and by high-performance liquid chromatography.16 Briefly, for the vitamin D receptor–mediated transactivation assay, COS-1 cells were transiently transfected with 0.5 μg of GAL4-VDR(DEF), 1 μg of 17M2-G-CAT reporter plasmid,22 0.2 μg each of the adrenodoxin and adrenodoxin reductase expression plasmids, 1 to 2 μg of either wild-type or mutant 1α-hydroxylase expression plasmid, and 1 μg of β-galactosidase expression plasmid pCH110 (Pharmacia Biotech, Uppsala, Sweden). Twelve hours after transfection, the ligands were added to the medium. After incubation for an additional 36 hours, cell extracts were prepared and used for chloramphenicol acetyltransferase assays.23

For analysis by high-performance liquid chromatography, COS-1 cells were transfected with 0.5 μg each of adrenodoxin and adrenodoxin reductase expression plasmids and 3 μg of either wild-type or mutant 1α-hydroxylase expression plasmid and were then incubated with tritiated 25-hydroxyvitamin D3 for six hours. The incubation medium and cells were extracted and analyzed by normal-phase and reverse-phase high-performance liquid chromatography. The eluent fractions were collected, and the radioactivity was estimated by liquid-scintillation counting.16,24,25 Authentic vitamin D derivatives were chromatographed, and the retention times were determined by ultraviolet absorption at 264 nm.

Results

Isolation of 25-Hydroxyvitamin D3 1α-Hydroxylase cDNA and Gene

A 2.4-kb cDNA clone was isolated from a normal human kidney cDNA library by using the mouse 1α-hydroxylase cDNA fragment as a probe. This cDNA contains a 1527-bp open reading frame predicted to encode a protein of 508 amino acids. The sequence of this cDNA was verified by determination of the genomic sequence.

The sequence of the coding region was 82 percent identical to that of mouse 1α-hydroxylase at both the nucleotide and the amino acid levels.16 The deduced amino acid sequence of this cDNA has substantial homology with members of the mitochondrial P450 family,26 particularly human vitamin D3 25-hydroxylase (CYP27, 40 percent),27 25-hydroxyvitamin D3 24-hydroxylase (CYP24, 32 percent),28 P450scc (CYP11A, 33 percent),29 and 11β-hydroxylase (CYP11B1, 30 percent).30

The structure of the human 1α-hydroxylase gene was determined by PCR amplification of normal human leukocyte DNA, with use of exon-specific primers designed from the cDNA sequence, on the basis of the finding that the known members of the mitochondrial P450 family usually have short introns.31 The 1α-hydroxylase gene consisted of nine exons spanning a region of approximately 4.8 kb.

Chromosomal Localization of the 1α-Hydroxylase Gene

Figure 1. Figure 1. Chromosomal Localization of the 1α-Hydroxylase Gene According to Fluorescence in Situ Hybridization.

Panel A shows fluorescence in situ hybridization with three genomic 1α-hydroxylase fragments covering the entire coding region as probes on chromosome preparations obtained from phytohemagglutinin-stimulated blood lymphocytes. The probes were labeled by nick translation with biotin-16–deoxyuridine triphosphate and detected with avidin–fluorescein (Boehringer Mannheim, Mannheim, Germany).20 Precise mapping of the gene was performed by using prophase-like chromosome spreads with higher-resolution banding. Hoechst (G-banding), propidium iodide (R-banding), and avidin–fluorescein images were merged and colored blue, red, and green, respectively. Fluorescence signals on the R-banded chromosomes are indicated by arrowheads. The inset is a larger image of chromosome 12 showing precise mapping of the gene. Panel B shows the idiogram of chromosome 12 indicating the localization of the 1α-hydroxylase gene to 12q13.3. The vitamin D–receptor gene, the gene responsible for hypocalcemic vitamin D–resistant rickets, was previously mapped to 12q13–14, as indicated in the figure.32

Fluorescence in situ hybridization revealed that the 1α-hydroxylase gene was located on chromosome band 12q13.3 (Figure 1A and Figure 1B). Since the gene responsible for pseudovitamin D–deficiency rickets has been mapped to this locus,9 this finding suggested that pseudovitamin D–deficiency rickets is caused by a defect in the structural gene encoding 1α-hydroxylase.

Tissue Expression of the 1α-Hydroxylase Gene

Figure 2. Figure 2. Northern Blot Analysis of 1α-Hydroxylase Gene Expression in Human Tissues.

The expression of the 1α-hydroxylase gene in various human tissues was analyzed by Northern blotting with 2 μg of poly(A)+RNA from each tissue (Clontech, Palo Alto, Calif.).33 The 0.7-kb cDNA fragment corresponding to the N-terminal region of human 1α-hydroxylase was used as a probe, and the levels of expression of β-actin and glutaraldehyde-3-phosphate dehydrogenase (G3PDH) served as internal controls.

Using the human 1α-hydroxylase cDNA clone as a probe, we examined its expression in various human tissues by Northern blot analysis. The 1α-hydroxylase transcript (approximately 2.4 kb) was detected only in renal tissue (Figure 2).

Homozygous Mutations in the 1α-Hydroxylase Gene in Patients with Pseudovitamin D–Deficiency Rickets

Figure 3. Figure 3. Mutation Analysis of the 1α-Hydroxylase Gene in Four Families with Pseudovitamin D–Deficiency Rickets.

Panel A shows the positions of the missense mutations detected in the four patients with pseudovitamin D–deficiency rickets. The exons are numbered, and the positions of the primers used for PCR amplification of each exon are indicated by the arrows under the diagram. Panel B shows the mutations detected in each family. The affected probands are indicated by arrows. Homozygotes are indicated by solid circles (females) or squares (males), heterozygotes by half-solid circles or squares, and the one tested family member with no mutation by an open square. Those not studied are indicated by gray symbols. Relevant portions of the sequencing chromatograms obtained from a normal subject (left) and the affected proband (right) are shown, with the position of each nucleotide mutation (arrowhead) and amino acid substitution (asterisk) indicated.

To determine whether the 1α-hydroxylase gene was mutated in patients with pseudovitamin D–deficiency rickets, we analyzed genomic DNA from the four patients. Four different missense mutations in four different positions were detected in these patients (Figure 3A and Figure 3B). Patient 1 had a mutation in exon 2, Arg107His (CGC to CAC), and Patient 2 had a mutation in exon 2, Gly125Glu (GGA to GAA). Patient 3 had a mutation in exon 6, Arg335Pro (CGG to CCG), that eliminates a restriction site for HpaII. The presence of the mutation was further confirmed by digesting the PCR product of exon 6 with the enzyme (data not shown). Patient 4 had a mutation in exon 7, Pro382Ser (CCT to TCT). All these patients were homozygous for their mutations, and the parents and one sibling who were studied were all heterozygous carriers (Figure 3B). A nephew of Patient 4 was a normal homozygote.

Functional Analysis of Wild-Type and Mutant 1α-Hydroxylase Proteins

Figure 4. Figure 4. Functional Analysis of Wild-Type and Mutant 1α-Hydroxylase Proteins.

Panel A shows the schema of the assay system for 1α-hydroxylase activity. COS-1 cells were transfected with GAL4-VDR(DEF), 17M2-G-CAT, adrenodoxin, and adrenodoxin reductase expression plasmids together with wild-type or mutant 1α-hydroxylase expression plasmid in the presence of 25-hydroxyvitamin D3 (25(OH)D3). When the expressed 1α-hydroxylase converts 25-hydroxyvitamin D3 into 1α,25-dihydroxyvitamin D3 (1α,25(OH)2D3), the activated GAL4-VDR(DEF) induces expression of chloramphenicol acetyltransferase (CAT) on the reporter gene (17M2-G-CAT) through the GAL4 upstream-activating sequence (GAL4-UAS), and 1α-hydroxylase activity can be analyzed by chloramphenicol acetyltransferase assay.

Panel B shows the 1α-hydroxylase activity of wild-type and mutant 1α-hydroxylase as determined by the assay shown in Panel A. Significant activation was not detected in the absence of 25-hydroxyvitamin D3 (lane 1) or 1α-hydroxylase expression plasmid (lane 2); however, 1α,25-dihydroxyvitamin D3 effectively induced chloramphenicol acetyltransferase activity (lanes 6 and 7). The expression of wild-type 1α-hydroxylase induced chloramphenicol acetyltransferase activity in a dose-dependent way (lanes 3 and 4), as observed in mouse 1α-hydroxylase (lane 5).16 In contrast, none of the mutants increased the activity of the reporter gene (lanes 8, 9, 10, and 11). The results of one representative chloramphenicol acetyltransferase assay and a graph corresponding to the mean (±SE) values for three independent experiments are shown.

Panel C shows the conversion of 25-hydroxyvitamin D3 to 1α,25-dihydroxyvitamin D3 by 1α-hydroxylase as determined by high-performance liquid chromatography. The results of normal-phase high-performance liquid chromatography are shown on the left and those of reverse-phase high-performance liquid chromatography on the right. The upper panels show the retention times of authentic vitamin D derivatives (1α-hydroxyvitamin D3 [1α(OH)D3], 25-hydroxyvitamin D3, 24R,25-dihydroxyvitamin D3 [24R,25(OH)2D3], 1α,25-dihydroxyvitamin D3, and 1α,24R,25-trihydroxyvitamin D3 [1α,24R,25(OH)3D3]) as determined by ultraviolet absorption at 264 nm. The lower panels show the production of 1α,25-dihydroxyvitamin D3 as determined by high-performance liquid chromatography when COS-1 cells were transfected with wild-type 1α-hydroxylase expression plasmid in the presence of tritiated 25-hydroxyvitamin D3. Note that the retention times of a converted metabolite matched those of authentic 1α,25-dihydroxyvitamin D3 in both normal and reverse phases (arrows).

Panel D shows the conversion of 25-hydroxyvitamin D3 to 1α,25-dihydroxyvitamin D3 by the wild-type (lane 2) and mutant (lanes 3, 4, 5, and 6) 1α-hydroxylase proteins. The activity of 1α-hydroxylase was determined by high-performance liquid chromatography, as in Panel C. Note that all the mutants failed to convert 25-hydroxyvitamin D3 to 1α,25-dihydroxyvitamin D3 (lanes 3, 4, 5, and 6). Values corresponding to the mean (±SE) for three independent experiments are shown.

The 1α-hydroxylase activities of the wild-type and the mutant enzymes in the four patients were assessed by examining the ligand-induced transactivation function of the vitamin D receptor. The vitamin D receptor was activated only when the 25-hydroxyvitamin D3 added to the culture medium was converted to an active ligand, possibly 1α,25-dihydroxyvitamin D3 (Figure 4A). The expression of wild-type 1α-hydroxylase activated the reporter gene (Figure 4B, lanes 3 and 4), as in mouse 1α-hydroxylase16 (lane 5). In contrast, none of the mutants (Figure 4B, lanes 8, 9, 10, and 11) increased the activity of the reporter gene. In vitro translation analysis indicated that the wild-type and the mutant plasmids were translated with similar efficiency at 55 kd in size (data not shown).

The 1α-hydroxylase activity of these mutants was assayed to determine the ability of the enzyme to catalyze the conversion of 25-hydroxyvitamin D3 to 1α,25-dihydroxyvitamin D3. When tritiated 25-hydroxyvitamin D3 was added to cells transfected with the wild-type 1α-hydroxylase expression plasmid, a metabolite was detected (Figure 4C, lower panels) at a retention time identical to that of authentic 1α,25-dihydroxyvitamin D3 (Figure 4C, upper panels). However, none of the mutants converted 25-hydroxyvitamin D3 into 1α,25-dihydroxyvitamin D3 (Figure 4D). These results indicate that the protein products of the four mutations found in the patients with pseudovitamin D–deficiency rickets had no 1α-hydroxylase activity.

Discussion

The responsible genes and their mutations have been identified for two types of hereditary rickets besides pseudovitamin D–deficiency rickets.5-8 We isolated a human cDNA encoding 25-hydroxyvitamin D3 1α-hydroxylase and identified different homozygous missense mutations in this gene in four patients with pseudovitamin D–deficiency rickets. Each of the four mutations abolished 1α-hydroxylase activity. The asymptomatic parents and sibling were heterozygous for the mutations, demonstrating the autosomal recessive inheritance of this disease. We conclude that pseudovitamin D–deficiency rickets is caused by inactivating mutations in the 1α-hydroxylase gene.

We found no correlation between phenotype and genotype; all four mutations abolished 1α-hydroxylase activity, and the clinical features in all four patients were similar. Clinical heterogeneity has been reported in pseudovitamin D–deficiency rickets,2,34 and there may be other mutations that only partially reduce enzyme activity, as is the case with other P450 enzymes.35,36 In a preliminary study of a patient who had pseudovitamin D–deficiency rickets with mild clinical features, we detected no mutations in the coding region (unpublished data), a fact that suggests that gene expression may be impaired by a mutation in the promoter region. Alternatively, there may be genetic heterogeneity in pseudovitamin D–deficiency rickets, and further analysis may reveal mutations in another gene. We found only homozygous mutations in the four patients, a result in agreement with the finding that in rare autosomal recessive diseases, homozygous mutations are frequent and compound heterozygotes are uncommon.37

The mutated residue (Pro382Ser) in one of our patients was located in a conserved region among various P450 enzymes that is assumed to be required for substrate binding,38,39 and another mutated residue (Gly125Glu) was located in the region corresponding to the proposed substrate-recognition site in the P450 family (CYP2).40 These mutations probably abolish the activity of 1α-hydroxylase by reducing its affinity for 25-hydroxyvitamin D3. We do not know how the other two mutations abolish 1α-hydroxylase activity, but because these four residues are highly conserved among several mitochondrial P450 enzymes, they are likely to be important for the enzymatic function of this protein.

With the data obtained in this study, we now know the molecular basis of all three major types of hereditary rickets. The phenotypes of pseudovitamin D–deficiency rickets and hypocalcemic vitamin D–resistant rickets are similar, except for alopecia in the latter and differences in serum concentrations of vitamin D metabolites.4 There are many endogenous vitamin D derivatives with biologic activity, 41 and therefore altered serum concentrations of vitamin D derivatives in these types of hereditary rickets may cause different changes.

Funding and Disclosures

Supported by a grant-in-aid from the Ministry of Education, Science, Sports, and Culture of Japan (to Dr. Kato).

NAPS See NAPS document no. 05444 for one page of supplementary material. To order, contact NAPS c/o Microfiche Publications, 248 Hempstead Tpk., West Hempstead, NY 11552.

We are indebted to Mr. Yoshikazu Masuhiro, Dr. Yasuo Yanagi, Dr. Seiji Fukumoto, Dr. Masato Kobori, Dr. Takehiko Yokomizo, Mr. Fumihiko Ichikawa, and Dr. Naoki Kubota for technical assistance; to Dr. Toshiyuki Yasuda and Dr. Ayako Tanae for providing data on patients; to Mr. Tatsuya Yoshizawa for helpful discussions; to Dr. Chifumi Kitanaka for valuable comments; to Dr. Yoshiyasu Yabusaki for the kind gift of adrenodoxin and adrenodoxin reductase cDNA plasmids; and to Chugai Pharmaceuticals for vitamin D–related compounds.

Author Affiliations

From the Institute of Molecular and Cellular Biosciences, University of Tokyo, Tokyo (S. Kitanaka, K.T., A.M., T.S., J.Y., S. Kato); Core Research for Evolutional Science and Technology, Japan Science and Technology Corporation, Saitama (S. Kato); the Faculty of Bioresources, Mie University, Mie (K.O., M.N.); Tokyo Metropolitan Kiyose Children's Hospital, Tokyo (Y.H.); the Department of Pediatrics, Chiba University School of Medicine, Chiba (H.N.); and the Department of Endocrinology and Metabolism, National Children's Medical Research Center, Tokyo (T.T.) — all in Japan.

Address reprint requests to Dr. Kato at the Institute of Molecular and Cellular Biosciences, University of Tokyo, Yayoi, Bunkyo-ku, Tokyo 113-0032, Japan.

References (41)

  1. 1. Prader A, Illig R, Heierli E. Eine besondere Form des primären vitamin-D-resistenten Rachitis mit Hypocalcämie und autosomal-dominantem Erbgang: die hereditäre Pseudo-Mangelrachitis. Helv Paediatr Acta 1961;16:452-468

  2. 2. Balsan S. Hereditary pseudo-deficiency rickets or vitamin D-dependency type I. In: Glorieux FH, ed. Rickets. New York: Raven Press, 1991:155-63.

  3. 3. Demay MB. Hereditary defects in vitamin D metabolism and vitamin D receptor defects. In: DeGroot LJ, Besser M, Burger HG, et al., eds. Endocrinology. 3rd ed. Vol. 2. Philadelphia: W.B. Saunders, 1995:1173-8.

  4. 4. Marx SJ. Vitamin D and other calciferols. In: Scriver CR, Beaudet AL, Sly WS, Valle D, eds. The metabolic and molecular bases of inherited disease. 7th ed. Vol. 2. New York: McGraw-Hill, 1995:3091-107.

  5. 5. Hughes MR, Malloy PJ, Kieback DG, et al. Point mutations in the human vitamin D receptor gene associated with hypocalcemic rickets. Science 1988;242:1702-1705

  6. 6. Malloy PJ, Hochberg Z, Tiosano D, Pike JW, Hughes MR, Feldman D. The molecular basis of hereditary 1,25-dihydroxyvitamin D3 resistant rickets in seven related families. J Clin Invest 1990;86:2071-2079

  7. 7. Liberman UA, Marx SJ. Vitamin D resistance. In: Weintraub BD, ed. Molecular endocrinology: basic concepts and clinical correlations. New York: Raven Press, 1995:425-44.

  8. 8. The HYP Consortium. A gene (PEX) with homologies to endopeptidases is mutated in patients with X-linked hypophosphatemic rickets. Nat Genet 1995;11:130-136

  9. 9. Labuda M, Morgan K, Glorieux FH. Mapping autosomal recessive vitamin D dependency type I to chromosome 12q14 by linkage analysis. Am J Hum Genet 1990;47:28-36

  10. 10. Scriver CR, Reade TM, DeLuca HF, Hamstra AJ. Serum 1,25-dihydroxyvitamin D levels in normal subjects and in patients with hereditary rickets or bone disease. N Engl J Med 1978;299:976-979

  11. 11. Delvin EE, Glorieux FH, Marie PJ, Pettifor JM. Vitamin D dependency: replacement therapy with calcitriol? J Pediatr 1981;99:26-34

  12. 12. Fraser D, Kooh SW, Kind HP, Holick MF, Tanaka Y, DeLuca HF. Pathogenesis of hereditary vitamin-D-dependent rickets: an inborn error of vitamin D metabolism involving defective conversion of 25-hydroxyvitamin D to 1α,25-dihydroxyvitamin D. N Engl J Med 1973;289:817-822

  13. 13. Henry HL. Vitamin D hydroxylases. J Cell Biochem 1992;49:4-9

  14. 14. Kawashima H, Torikai S, Kurokawa K. Localization of 25-hydroxyvitamin D3 1α-hydroxylase and 24-hydroxylase along the rat nephron. Proc Natl Acad Sci U S A 1981;78:1199-1203

  15. 15. Pedersen JI, Ghazarian JG, Orme-Johnson NR, DeLuca HF. Isolation of chick renal mitochondrial ferredoxin active in the 25-hydroxyvitamin D3-1α-hydroxylase system. J Biol Chem 1976;251:3933-3941

  16. 16. Takeyama K, Kitanaka S, Sato T, Kobori M, Yanagisawa J, Kato S. 25-Hydroxyvitamin D3 1α-hydroxylase and vitamin D synthesis. Science 1997;277:1827-1830

  17. 17. Yoshizawa T, Handa Y, Uematsu Y, et al. Mice lacking the vitamin D receptor exhibit impaired bone formation, uterine hypoplasia and growth retardation after weaning. Nat Genet 1997;16:391-396

  18. 18. Sasaki T, Nakajima H, Suzuki S. Studies on the pathogenesis of hypophosphatemic vitamin D refractory rickets of the simple type or phosphatdiabetes. Endocrinol Jpn 1961;8:272-278

  19. 19. Kobori M, Nojima H. A simple treatment of DNA in a ligation mixture prior to electroporation improves transformation frequency. Nucleic Acids Res 1993;21:2782-2782

  20. 20. Okumura K, Nogami M, Taguchi H, Hisamatsu H, Tanaka K. The genes for the α-type HC3 (PMSA2) and β-type HC5 (PMSB1) subunits of human proteasomes map to chromosomes 6q27 and 7p12-p13 by fluorescence in situ hybridization. Genomics 1995;27:377-379

  21. 21. Ebihara K, Masuhiro Y, Kitamoto T, et al. Intron retention generates a novel isoform of the murine vitamin D receptor that acts in a dominant negative way on the vitamin D signaling pathway. Mol Cell Biol 1996;16:3393-3400

  22. 22. Kato S, Endoh H, Masuhiro Y, et al. Activation of the estrogen receptor through phosphorylation by mitogen-activated protein kinase. Science 1995;270:1491-1494

  23. 23. Sasaki H, Harada H, Handa Y, et al. Transcriptional activity of a fluorinated vitamin D analog on VDR-RXR-mediated gene expression. Biochemistry 1995;34:370-377

  24. 24. Fujii H, Sato T, Kaneko S, et al. Metabolic inactivation of retinoic acid by a novel P450 differentially expressed in developing mouse embryos. EMBO J 1997;16:4163-4173

  25. 25. Mawer EB, Hayes ME, Heys SE, et al. Constitutive synthesis of 1,25-dihydroxyvitamin D3 by a human small cell lung cancer cell line. J Clin Endocrinol Metab 1994;79:554-560

  26. 26. Nelson DR, Kamataki T, Waxman DJ, et al. The P450 superfamily: update on new sequences, gene mapping, accession numbers, early trivial names of enzymes, and nomenclature. DNA Cell Biol 1993;12:1-51

  27. 27. Guo YD, Strugnell S, Back DW, Jones G. Transfected human liver cytochrome P-450 hydroxylates vitamin D analogs at different side-chain positions. Proc Natl Acad Sci U S A 1993;90:8668-8672

  28. 28. Chen KS, Prahl JM, DeLuca HF. Isolation and expression of human 1,25-dihydroxyvitamin D3 24-hydroxylase cDNA. Proc Natl Acad SciU S A 1993;90:4543-4547

  29. 29. Chung BC, Matteson KJ, Voutilainen R, Mohandas TK, Miller WL. Human cholesterol side-chain cleavage enzyme, P450scc: cDNA cloning, assignment of the gene to chromosome 15, and expression in the placenta. Proc Natl Acad Sci U S A 1986;83:8962-8966

  30. 30. Chua SC, Szabo P, Vitek A, Grzeschik KH, John M, White PC. Cloning of cDNA encoding steroid 11β-hydroxylase (P450c11). Proc Natl Acad Sci U S A 1987;84:7193-7197

  31. 31. Mornet E, Dupont J, Vitek A, White PC. Characterization of two genes encoding human steroid 11β-hydroxylase (P-45011β). J Biol Chem 1989;264:20961-20967

  32. 32. Labuda M, Fujiwara TM, Ross MV, et al. Two hereditary defects related to vitamin D metabolism map to the same region of human chromosome 12q13-14. J Bone Miner Res 1992;7:1447-1453

  33. 33. Takeyama K, Kojima R, Ohashi R, et al. Retinoic acid differentially up-regulates the gene expression of retinoic acid receptor α and γ isoforms in embryo and adult rats. Biochem Biophys Res Commun 1996;222:395-400

  34. 34. Cowen J, Harris F. Late presentation of vitamin D-dependent rickets. Arch Dis Child 1980;55:964-966

  35. 35. Yanase T, Simpson ER, Waterman MR. 17α-Hydroxylase/17,20-lyase deficiency: from clinical investigation to molecular definition. Endocr Rev 1991;12:91-108

  36. 36. White PC, Tusie-Luna MT, New MI, Speiser PW. Mutations in steroid 21-hydroxylase (CYP21). Hum Mutat 1994;3:373-378

  37. 37. Kitanaka S, Katsumata N, Tanae A, et al. A new compound heterozygous mutation in the 11β-hydroxysteroid dehydrogenase type 2 gene in a case of apparent mineralocorticoid excess. J Clin Endocrinol Metab 1997;82:4054-4058

  38. 38. Nonaka Y, Matsukawa N, Morohashi K, et al. Molecular cloning and sequence analysis of cDNA encoding rat adrenal cytochrome P-45011β. FEBS Lett 1989;255:21-26

  39. 39. Kawajiri K, Gotoh O, Sogawa K, Tagashira Y, Muramatsu M, Fujii-Kuriyama Y. Coding nucleotide sequence of 3-methylcholanthrene-inducible cytochrome P-450d cDNA from rat liver. Proc Natl Acad Sci U S A 1984;81:1649-1653

  40. 40. Gotoh O. Substrate recognition sites in cytochrome P450 family 2 (CYP2) proteins inferred from comparative analyses of amino acid and coding nucleotide sequences. J Biol Chem 1992;267:83-90

  41. 41. Bouillon R, Okamura WH, Norman AW. Structure-function relationships in the vitamin D endocrine system. Endocr Rev 1995;16:200-257

Citing Articles (279)

    Figures/Media

    1. Table 1. Clinical and Biochemical Features in Four Patients with Pseudovitamin D–Deficiency Rickets.
      Table 1. Clinical and Biochemical Features in Four Patients with Pseudovitamin D–Deficiency Rickets.
    2. Figure 1. Chromosomal Localization of the 1α-Hydroxylase Gene According to Fluorescence in Situ Hybridization.
      Figure 1. Chromosomal Localization of the 1α-Hydroxylase Gene According to Fluorescence in Situ Hybridization.

      Panel A shows fluorescence in situ hybridization with three genomic 1α-hydroxylase fragments covering the entire coding region as probes on chromosome preparations obtained from phytohemagglutinin-stimulated blood lymphocytes. The probes were labeled by nick translation with biotin-16–deoxyuridine triphosphate and detected with avidin–fluorescein (Boehringer Mannheim, Mannheim, Germany).20 Precise mapping of the gene was performed by using prophase-like chromosome spreads with higher-resolution banding. Hoechst (G-banding), propidium iodide (R-banding), and avidin–fluorescein images were merged and colored blue, red, and green, respectively. Fluorescence signals on the R-banded chromosomes are indicated by arrowheads. The inset is a larger image of chromosome 12 showing precise mapping of the gene. Panel B shows the idiogram of chromosome 12 indicating the localization of the 1α-hydroxylase gene to 12q13.3. The vitamin D–receptor gene, the gene responsible for hypocalcemic vitamin D–resistant rickets, was previously mapped to 12q13–14, as indicated in the figure.32

    3. Figure 2. Northern Blot Analysis of 1α-Hydroxylase Gene Expression in Human Tissues.
      Figure 2. Northern Blot Analysis of 1α-Hydroxylase Gene Expression in Human Tissues.

      The expression of the 1α-hydroxylase gene in various human tissues was analyzed by Northern blotting with 2 μg of poly(A)+RNA from each tissue (Clontech, Palo Alto, Calif.).33 The 0.7-kb cDNA fragment corresponding to the N-terminal region of human 1α-hydroxylase was used as a probe, and the levels of expression of β-actin and glutaraldehyde-3-phosphate dehydrogenase (G3PDH) served as internal controls.

    4. Figure 3. Mutation Analysis of the 1α-Hydroxylase Gene in Four Families with Pseudovitamin D–Deficiency Rickets.
      Figure 3. Mutation Analysis of the 1α-Hydroxylase Gene in Four Families with Pseudovitamin D–Deficiency Rickets.

      Panel A shows the positions of the missense mutations detected in the four patients with pseudovitamin D–deficiency rickets. The exons are numbered, and the positions of the primers used for PCR amplification of each exon are indicated by the arrows under the diagram. Panel B shows the mutations detected in each family. The affected probands are indicated by arrows. Homozygotes are indicated by solid circles (females) or squares (males), heterozygotes by half-solid circles or squares, and the one tested family member with no mutation by an open square. Those not studied are indicated by gray symbols. Relevant portions of the sequencing chromatograms obtained from a normal subject (left) and the affected proband (right) are shown, with the position of each nucleotide mutation (arrowhead) and amino acid substitution (asterisk) indicated.

    5. Figure 4. Functional Analysis of Wild-Type and Mutant 1α-Hydroxylase Proteins.
      Figure 4. Functional Analysis of Wild-Type and Mutant 1α-Hydroxylase Proteins.

      Panel A shows the schema of the assay system for 1α-hydroxylase activity. COS-1 cells were transfected with GAL4-VDR(DEF), 17M2-G-CAT, adrenodoxin, and adrenodoxin reductase expression plasmids together with wild-type or mutant 1α-hydroxylase expression plasmid in the presence of 25-hydroxyvitamin D3 (25(OH)D3). When the expressed 1α-hydroxylase converts 25-hydroxyvitamin D3 into 1α,25-dihydroxyvitamin D3 (1α,25(OH)2D3), the activated GAL4-VDR(DEF) induces expression of chloramphenicol acetyltransferase (CAT) on the reporter gene (17M2-G-CAT) through the GAL4 upstream-activating sequence (GAL4-UAS), and 1α-hydroxylase activity can be analyzed by chloramphenicol acetyltransferase assay.

      Panel B shows the 1α-hydroxylase activity of wild-type and mutant 1α-hydroxylase as determined by the assay shown in Panel A. Significant activation was not detected in the absence of 25-hydroxyvitamin D3 (lane 1) or 1α-hydroxylase expression plasmid (lane 2); however, 1α,25-dihydroxyvitamin D3 effectively induced chloramphenicol acetyltransferase activity (lanes 6 and 7). The expression of wild-type 1α-hydroxylase induced chloramphenicol acetyltransferase activity in a dose-dependent way (lanes 3 and 4), as observed in mouse 1α-hydroxylase (lane 5).16 In contrast, none of the mutants increased the activity of the reporter gene (lanes 8, 9, 10, and 11). The results of one representative chloramphenicol acetyltransferase assay and a graph corresponding to the mean (±SE) values for three independent experiments are shown.

      Panel C shows the conversion of 25-hydroxyvitamin D3 to 1α,25-dihydroxyvitamin D3 by 1α-hydroxylase as determined by high-performance liquid chromatography. The results of normal-phase high-performance liquid chromatography are shown on the left and those of reverse-phase high-performance liquid chromatography on the right. The upper panels show the retention times of authentic vitamin D derivatives (1α-hydroxyvitamin D3 [1α(OH)D3], 25-hydroxyvitamin D3, 24R,25-dihydroxyvitamin D3 [24R,25(OH)2D3], 1α,25-dihydroxyvitamin D3, and 1α,24R,25-trihydroxyvitamin D3 [1α,24R,25(OH)3D3]) as determined by ultraviolet absorption at 264 nm. The lower panels show the production of 1α,25-dihydroxyvitamin D3 as determined by high-performance liquid chromatography when COS-1 cells were transfected with wild-type 1α-hydroxylase expression plasmid in the presence of tritiated 25-hydroxyvitamin D3. Note that the retention times of a converted metabolite matched those of authentic 1α,25-dihydroxyvitamin D3 in both normal and reverse phases (arrows).

      Panel D shows the conversion of 25-hydroxyvitamin D3 to 1α,25-dihydroxyvitamin D3 by the wild-type (lane 2) and mutant (lanes 3, 4, 5, and 6) 1α-hydroxylase proteins. The activity of 1α-hydroxylase was determined by high-performance liquid chromatography, as in Panel C. Note that all the mutants failed to convert 25-hydroxyvitamin D3 to 1α,25-dihydroxyvitamin D3 (lanes 3, 4, 5, and 6). Values corresponding to the mean (±SE) for three independent experiments are shown.

      More Research