Department of Medicine, Duke University Medical Center, Durham, North Carolina 27710
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ABSTRACT |
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X-linked hypophosphatemia (XLH) is caused by inactivating mutations of Phex, a phosphate-regulating endopeptidase. Further advances in our knowledge of the pathogenesis of XLH require identification of the biological function of Phex and its physiologically relevant substrates. We evaluated several potential substrates using mouse recombinant wild-type Phex proteins (rPhex-WT) and inactive mutant Phex proteins (rPhex-3'M) lacking the COOH-terminal catalytic domain as controls. By Western blot analysis, we demonstrated that Phex is a membrane-bound 100-kDa glycosylated monomer. Neither casein, a substrate for the related endopeptidase thermolysin, human stanniocalcin 1 (hSTC-1), an osteoblast-derived phosphate-regulating factor, nor FGF-23 peptide (amino acid 172-186), comprising the region mutated in autosomal dominant hypophosphatemia, was cleaved by rPhex-WT. In addition, membranes expressing rPhex-WT, rPhex-3'M, and the empty vector hydrolyzed parathyroid hormone-(1-34), indicating the lack of Phex-specific cleavage of parathyroid hormone. In contrast, rPhex-WT did display an EDTA-dependent cleavage of the neutral endopeptidase substrate [Leu]enkephalin. Further studies with wild-type and mutant rPhex proteins should permit the identification of physiologically relevant substrates involved in the pathogenesis of XLH.
X-linked hypophosphatemia; hypophosphatemia; rickets; parathyroid hormone; FGF-23; [Leu]enkephalin
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INTRODUCTION |
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THE ABBREVIATION Phex is used for the phosphate-regulating gene with homologies to endopeptidases located at the HYP loci on the X chromosome (10, 21). The Phex gene product is a type II membrane-bound zinc metalloendopeptidase that is closely related to neutral endopeptidase (NEP), endothelin-converting enzyme, and Kell (25). The clinical consequences of inactivating mutations of Phex indicate that this endopeptidase is involved in regulating phosphate and mineral homeostasis. Loss-of-function mutations of Phex cause X-linked hypophosphatemic rickets (XLH), a clinical disorder characterized by defective calcification and hypophosphatemia (10, 21). A similar phenotype occurs in the Hyp mouse homolog of XLH, where a 3' deletion of Phex removes its COOH-terminal catalytic domain (1, 9). Despite identification of the disease gene, the pathogenesis of phosphate wasting and impaired mineralization remains poorly understood, in part because the physiologically relevant Phex substrates have not been identified.
The search for candidate substrates has been guided by the hypothesis that inactivating Phex mutations leads to the accumulation of unknown phosphaturic and/or mineralization inhibitory factors (16, 19, 21) and the knowledge that related endopeptidases may have multiple substrates that are coexpressed within an organ/cell type-specific fashion (25). For example, the related endopeptidase NEP is constitutively expressed in a variety of tissues, where it metabolizes different substrates, including atrial natriuretic factor in the kidney, substance P in the lungs, and enkephalins, such as [Leu]enkephalin, in the brain (24). Thus Phex may also have multiple substrates that are expressed in the same tissues as Phex.
Several observations suggest that a physiologically relevant Phex substrate may be produced in the skeleton. First, Phex is predominantly expressed in osteoblasts in association with genes regulating extracellular matrix production and mineralization (6). Second, osteoblasts derived from Hyp mice secrete factors that inhibit mineralization and renal tubular phosphate transport (4, 20, 30). Third, bone marrow transplantation can partially rescue the phenotype of Hyp mice (18). Recently, a mammalian homolog of stanniocalcin 1 (STC-1), a calcium- and phosphate-regulating glycoprotein hormone in fish secreted by the corpuscles of Stannius, has been identified in osteoblasts (11, 26). Because of its colocalization with Phex and ability to regulate systemic phosphate homeostasis, STC-1 is a candidate substrate for Phex. There are no published studies evaluating whether STC-1 is cleaved by Phex.
Phex transcripts have been detected at several extraskeletal sites, including ovary, spleen, B cells, fetal lung, and parathyroid glands (1, 2), as well as in tumors causing oncogenic hypophosphatemic osteomalacia (OHO) (21, 22), indicating the possible presence of other substrates and physiological functions of Phex. Parathyroid hormone (PTH) has been considered a Phex substrate on the basis of Phex expression in the parathyroid glands (2) and the important role of PTH in inhibiting renal phosphate reabsorption. Indeed, recent unconfirmed studies report that recombinant Phex (rPhex) proteins generated in COS-7 cell membranes can metabolize PTH in vitro (12). The specificity of PTH cleavage by rPhex, however, has not been independently confirmed. Moreover, the physiological significance of PTH hydrolysis by Phex is not clear, since PTH is not the humoral factor responsible for phosphaturia in XLH or Hyp mice (16).
The investigation of autosomal dominant hypophosphatemia (ADH) (28) and OHO (29) confirms the existence of novel phosphaturic factors and provides additional clues to possible Phex substrates. In this regard, mutations in FGF-23, a new member of the growing family of FGF proteins (28), are responsible for ADH, which has many similarities to XLH. In addition, FGF-23 and Phex are coexpressed in tumors causing OHO that cause phosphaturia (29). These associations raise the possibility that FGF-23 may be a Phex substrate as well as a phosphaturic factor. If this is so, the pathogenesis of OHO might be explained by excessive production of FGF-23 that overwhelms the normal capacity of Phex, whereas ADH might be caused by the failure of Phex to degrade mutant FGF-23 (21). For this model to be valid, however, FGF-23 would need to be a substrate for Phex, and the R176Q and R179Q mutations of FGF-23 that cause ADH (28) should prevent cleavage of mutant FGF-23 by Phex.
As an initial step in the systematic examination of Phex structure and function, we generated recombinant wild-type (Phex-WT) and 3'-truncated Phex (Phex-3'M) proteins and assessed the ability of membranes expressing rPhex to specifically degrade a panel of possible physiological substrates, including casein, PTH-(1-34), FGF-23-(172-186), and human STC-1 (hSTC-1). In addition, we tested the ability of rPhex to cleave casein and [Leu]enkephalin, respective substrates for the related thermolysin and NEP.
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METHODS |
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Materials. Oligonucleotide primers were synthesized at the Duke University DNA Core Facility. The Bac-to-Bac baculovirus expression system was purchased from Life Technologies (Baltimore, MD). Spodoptera frugiperda (Sf9) cells were obtained from Dr. Patrick Casey (Duke University). The modular retroviral vector pLuvGM was provided by Dr. Clay Smith (Duke University) (8). We contracted with HTI Bio-Products (Ramona, CA) to raise Phex-specific rabbit antiserum against a keyhole limpet hemocyanin-conjugated synthetic peptide (H-CGGPRNSTMNRGADS-OH) corresponding to 734-745 amino acids in the COOH-terminal end of mouse Phex. Other antibodies used in these studies included a mouse anti-V5 monoclonal IgG2a antibody (Invitrogen, Carlsbad, CA), an anti-mouse IgG (whole molecule) conjugated to alkaline phosphatase (Sigma, St. Louis, MO), and anti-rabbit IgG antibodies conjugated to horseradish peroxidase (Santa Cruz Biotechnology, Santa Cruz, CA). Anti-nitro blue tetrazolium and bromochloroindolyl phosphate, used in the detection of alkaline phosphatase, were obtained from Roche (Indianapolis, IN). The enhanced chemiluminescence detection kit (NEN Life Science Products, Boston, MA) was used to detect horseradish peroxidase. Nitrocellulose membranes (0.45 µm) and other chemicals used for SDS-PAGE and Western blotting were purchased from Bio-Rad Laboratories (Hercules, CA). Thermolysin and rat synthetic PTH-(1-34) were purchased from Sigma. Human FGF-23 peptide (172-PIPRRHTRSAEDDSE-186) and the mutant peptide (172-PIPRQHTQSAEDDSE-186) that substitutes glutamine for arginine at positions 176 and 179 were synthesized by Genemed Synthesis (San Francisco, CA). [Leu]enkephalin, consisting of the sequence YGGFL, was obtained from Bachem Biosciences (King of Prussia, PA). The EnzChek protease kit was purchased from Molecular Probes (Eugene, OR). We cloned the human stanniocalcin full-length cDNA and generated recombinant hSTC-1 protein in our laboratory (see below).
Construction of epitope-tagged Phex-WT and Phex-3'M.
The Phex full-length cDNA coding sequence
(Phex-WT) was amplified from mouse osteoblast total
RNA by RT-PCR. Using oligonucleotides M 5F
(5'-TTCTGATGGAAGCAGAAACAGGGA-3'), M + 930R
(5'-GGGAATCATAGCGCTGAGTTCTGA-3'), M + 786F
(5'-TAATAGCTCTCGAGCTGAACATGA-3'), and M + 2253R
(5'-AGCTACCAGAGTCGGCAAGAATCT-3'), we amplified overlapping sequences
from
5 to 930 and from 786 to 2253, respectively, that were
ligated to generate the full-length Phex cDNA. The
Phex sequence was confirmed by direct sequencing. The
Phex cDNA was subcloned into the NotI and
ApaI sites of pMT/V5-His vector to create a cassette
containing Phex in frame with V5 and histidine epitope tags
at its COOH-terminal end (pMT-Phex-WT-V5-His). The
polyhistidine tag provides an anchor for purification on the affinity
matrix, Ni2+-nitrilotriacetate-agarose beads.
Isolation and cloning of human stanniocalcin from an osteoblast cDNA library. To clone hSTC-1, we used DNA from human bone cDNA library as the template to perform PCR with primers specific to stanniocalcin (26). The primer pairs were 5'-ATTCTTGCGGCCGCTCAGAGAATGCTCCAAAACT-3' as sense primer (enzyme NotI site was added to the end) and 5'-ATTCTTGGGCCCAGCACTCTCATGGGA-3' as antisense primer (enzyme ApaI site was added to the end). The 750-bp PCR fragment corresponding to the full length of the stanniocalcin coding region was amplified. After NotI and ApaI restriction digestion, the DNA product was subcloned into the same site of pFastBac1-V5-His vector in frame with V5-His. The DNA sequence was confirmed by sequencing.
Production of epitope-tagged rPhex proteins and hSTC-1 in
Sf9 cells.
Insect Sf9 cells were grown in suspension in Sf-900
II serum-free medium (Life Technologies) supplemented with gentamicin (25 µg/ml) at 27°C under constant stirring at 50-60 rpm.
Recombinant baculoviruses were produced with the pFastBac-1 vector and
the protocol provided in the Bac-to-Bac baculovirus expression system kit. The pFastBac-V5-His, containing no insert, and Phex-WT,
Phex-3'M, and hSTC-1 cDNAs were separately transformed into
DH10BAC Escherichia coli cells for transposition into the
bacmid. Four days later, colonies containing recombinant bacmids were
identified, and their DNA was used to transfect Sf9
insect cells. After 48 h of incubation, cells and culture
supernatants were removed and centrifuged at 1,000 rpm for 5 min. The
cell pellets from membrane-associated Phex-WT and
Phex-3'M and the supernatant from secreted hSTC-1 were
analyzed by immunoblotting analysis using anti-V5 antibody. The
supernatant was used to generate additional viral stocks by successive
amplification cycles in Sf9 cells. For production of recombinant proteins, Sf9 cells in log-phase growth were
diluted to 1 × 106 cells/ml and infected with
recombinant baculoviruses at multiplicity of infection of 5-10.
Cells producing Phex-WT or Phex-3'M were harvested 48 h after infection and resuspended in 150 mM NaCl and
20 mM Tris · HCl at pH 7.6. In all cases, cells were disrupted by sonication, nuclei and debris were removed by centrifugation at 300 g for 15 min at 4°C, and membranes were pelleted by
centrifugation at 30,000 g for 50 min. The crude membranes
were solubilized with 1%
n-dodecyl--D-maltoside in the lysis buffer.
Solubilized membranes were collected after centrifugation at 10,000 g for 15 min. The protein content of each sample was
determined by the NanoOrange protein quantitation kit (Molecular
Probes). Final protein concentrations of the membrane suspensions were
1-5 mg/ml. The suspensions were stored at 70°C in multiple
aliquots. The recombinant V5-His-tagged rPhex-WT and
rPhex-3'M were purified from solubilized membrane fractions
under native conditions by Ni2+-affinity resin
chromatography. The bounded rPhex proteins were eluted with
a high concentration of imidazole (250 mM) in the lysis buffer
(3).
Production of Phex proteins in E86 cells. The Phex retroviruses pLuvPhex-WT and pLuvPhex-3'M were created by cloning in frame the full-length, non-epitope-tagged Phex cDNA or the 3'Phex deletion mutant containing the V5-His epitope tag between the NcoI site and BamHI site of the pLuvM. E86 cells, an ecotropic packaging cell line, were transfected with the empty retrovirus vector pLuvM, pLuvPhex-WT, or pLuvPhex-3'M along with pSV2Neo using the TransFast reagent (Promega, Madison, WI). Cells were selected by incubation with 700 µg/ml G418 for 2 wk, and membranes were isolated as described below.
SDS-PAGE and Western blot analyses. Immunoblot analysis was carried out by modifications of previously described methods (17). The specified amount of membrane proteins was dissolved in SDS gel loading buffer (500 mM Tris · HCl, pH 6.8, 8.5% SDS, 27.5% sucrose, 0.03% bromphenol blue) with and without 100 mM dithiothreitol to produce reducing and nonreducing conditions, respectively. Separated proteins were transferred to a nitrocellulose membrane (0.45 µm; Bio-Rad, Chicago, IL) over a 30-min period at 2.5 mA/cm2 at room temperature using a semidry blotting system (Millipore, Chicago, IL). Immunoblotting was performed using the mouse anti-V5 monoclonal antibody diluted 1:5,000 in Tris-buffered saline-Tween 20 (TBST) and 1% bovine serum albumin (final concentration 220 ng/ml; Invitrogen, Carlsbad, CA) or affinity-purified rabbit anti-Phex polyclonal antiserum (1:2,000 dilution). For anti-V5 antibody, blots were washed with TBST for 60 min and incubated with anti-mouse IgG (whole molecule) conjugated with alkaline phosphatase (diluted to 1:5,000) (Sigma) for 60 min at room temperature. After washes with TBST, immunoreactivity was detected by colorimetric reaction using 66 µl of 50 mg/ml nitro blue tetrazolium and 33 µl of bromochloroindolyl phosphate (50 mg/ml) in alkaline phosphatase buffer (50 mM NaHCO2 and 10 mM MgCl2) as previously described (17). For studies using the Phex antiserum, the protocols were identical, except the secondary antibody was goat anti-rabbit IgG conjugated to horseradish peroxidase (diluted 1:5,000). In addition, after the blots were washed three times with TBST at room temperature for 40 min each, immunoreactivity was detected by a chemiluminescence system (NEN Life Science Products).
Deglycosylation of Phex with peptide N-glycosidase F. The membrane proteins were treated with or without peptide N-glycosidase F as described elsewhere (17). Briefly, the specified amount of protein was denatured with 1% 2-mercaptoethanol at 37°C for 15 min and then incubated with or without 2.5 U of peptide N-glycosidase F (Roche, Indianapolis, IN) with 1% Triton X-100. The digestions were carried out at 30°C overnight.
Preparation of membranes containing endogenous Phex.
Stock cultures of osteoblast cell lines, including human MG63, U2Os,
SaOs, and MC3T3-E1 osteoblasts (6), were used in these studies. To isolate membranes, the cell pellet was resuspended in lysis
buffer containing 150 mM NaCl and 20 mM Tris · HCl, pH 7.6, cells were disrupted by sonication, and nuclei and debris were removed
by centrifugation at 300 g for 15 min at 4°C. Crude membranes were then pelleted by centrifugation at 30,000 g
for 50 min and solubilized with 1%
n-dodecyl--D-maltoside in the lysis buffer.
Solubilized membranes were collected after centrifugation at 10,000 g for 15 min. The protein content of each sample was determined by the NanoOrange protein quantitation kit.
Immunoprecipitation of native and recombinant Phex. Endogenous Phex in osteoblasts or recombinant V5-His-tagged Phex from Sf9 cells was immunoprecipitated using our rabbit anti-Phex polyclonal antibody. Osteoblasts at various stages of maturation and Sf9 cells expressing rPhex were used for these studies. Briefly, the medium was removed and the cell culture plate was rinsed with PBS at room temperature. All the following steps were performed on ice. RIPA buffer (1 ml), consisting of 150 mM NaCl, 1% NP-40, 0.5% deoxycholate, 0.1% SDS, and 50 mM Tris (pH 8.0), was added to the cell culture and then removed with a cell scraper. The cell lysate was transferred to a fresh tube, passed through the 21-gauge needle several times to shear the DNA, and incubated on ice for 30 min. The insoluble material from the lysate was removed by centrifugation at 10,000 g for 10 min. To remove the nonspecific binding, the soluble lysate was precleared by addition of 20 µl of Phex prebleed serum, incubated at 4°C overnight, and then incubated with 70 µl of protein A-Sepharose for 1 h. Beads were pelleted by centrifugation at 1,000 g for 5 min at 4°C. The supernatant was transferred to a fresh tube. Affinity-purified anti-Phex antibody (10 µg) was applied to the supernatant, and it was incubated overnight at 4°C. Protein A-Sepharose (70 µl) was then added, and the supernatant was incubated for 2 h at 4°C. The immune complex was collected by centrifugation at 1,000 g for 5 min at 4°C and washed four times with RIPA buffer. The Phex-anti-Phex complex was eluted in 100 µl of RIPA buffer with SDS loading buffer and assessed by Western analysis with affinity-purified anti-Phex antibody (7).
Assessing Phex enzyme activity. Phex endopeptidase activity was assessed using rat synthetic PTH-(1-34) (Sigma-Aldrich, St. Louis, MO), hSTC-1, FGF-23 peptide, BODIPY TR-X-casein (Molecular Probes), or [Leu]enkephalin (Bachem Biosciences). To assess PTH, FGF-23, and [Leu]enkephalin peptide hydrolysis, we used modifications of previously described methods (12). PTH-(1-34) (10 µg), FGF-23 peptide (25 µg), or [Leu]enkephalin (14 µg) was incubated in a total volume of 30 µl with up to 60 µg of solubilized membrane preparations derived from mock-infected Sf9 cells, Sf9 cells infected with the baculovirus vector alone, Phex-WT, or Phex-3'M, or affinity-purified rPhex-WT. The reaction was incubated at 37°C for 30 min in a buffer containing 150 mM NaCl and 20 mM Tris · HCl (pH 7.4) and terminated by the addition of 0.1% trifluoroacetic acid. In some studies, membrane preparations were incubated in the presence of 20 mM EDTA before addition of the substrate. Degradation of the peptides was monitored by reverse-phase HPLC. All samples were dissolved in a 10 mM trifluoroacetic acid injection solvent. The samples were separated by HPLC using an HPLC system (model 840, Waters, Milford, MA) equipped with a Pecosphere HS3-C18 base-deactivated column (Perkin-Elmer Biosystems, Foster City, CA) and an ultraviolet detector (model 481, Waters) set for 214 nm. Substrate hydrolysis products were eluted with a linear gradient from 100% 10 mM trifluoroacetic acid to 60% 10 mM trifluoroacetic acid-40% acetonitrile over 60 min at 1.5 ml/min. For all experiments, standards were chromatographed immediately before the chromatography of experimental samples to verify retention times.
For analysis of hSCT-1 hydrolysis, we performed in vitro and in vivo (i.e., cell culture) cleavage reactions. For the in vitro studies, 8.5 µg of hSTC-1 collected from media of hSTC-1-producing Sf9 cells were incubated with 30 µg of solubilized Sf9 membranes expressing rPhex-WT or rPhex-3'M. The reaction was incubated at 37°C for 30 min in a buffer containing 150 mM NaCl and 20 mM Tris · HCl (pH 7.4) and terminated by the addition of SDS loading buffer. For the in vivo cleavage studies, we coinfected into Sf9 cells either recombinant baculovirus containing hSTC-1 and Phex-WT or hSTC-1 and Phex-3'M. After 48 h, we collected supernatant from the coinfected Sf9 cultures and monitored secreted hSTC-1 by Western blot analysis using anti-V5 antibody. To determine whether casein is a substrate for Phex, we used the EnzChek protease kits. Different concentrations of thermolysin or membrane protein were prepared in 100 µl of digestion buffer (10 mM Tris · HCl, pH 7.8, 0.1 mM sodium azide) and added with 100 µl of casein (10 µg/ml), which was heavily labeled with pH-insensitive red fluorescent BODIPY TR-X dye. After 1 h of incubation at room temperature and protection from light, the fluorescence was measured by microplate reader (excitation 485 nm, emission 530 nm). The related zinc metalloproteinase thermolysin was used as a positive control.PTH-stimulated cAMP production.
PTH-stimulated cAMP production was performed in HEK cells stably
transfected with the rat PTH receptor cDNA. To generate these cells, a cDNA encoding the rat PTH receptor was obtained by
RT-PCR from ROS 17/2.8 cells using the following primer pair:
CCCCGAGGGACGCGGCCCTAGGAATTCGCGATGGGGGCCGCCCGGATCGCACCC and TCCATCTGTCCAGGTACCCAGGCCAGCAGT. The PCR product was
ligated into the vector pBK-CMV(lacZ) in frame with the sequences
encoding the 12CA5 epitope (27) and stably transfected
into HEK-293 cells by the calcium phosphate method and G418 selection.
HEK cells expressing PTH receptors were plated at an initial density of 2-5 × 104 cells/ml in six-well plastic culture
dishes (9.5 cm2/well; Costar, Cambridge, MA) and, after
reaching confluence, were incubated in serum-free medium containing
[3H]adenine (2 µCi/ml; Amersham, Arlington Heights, IL)
for 3 h. Before agonist stimulation, cells were incubated for 10 min with 100 µM 3-isobutyl-1-methylxanthine in a buffer consisting of
Hanks' balanced salt solution (without Ca2+ and
Mg2+) containing 10 mM HEPES (pH 7.4). Generation of cAMP
was measured after 5 min of stimulation by modifications of the method
of Salomon et al. (23). We stimulated cAMP production by
incubation with various concentrations of hydrolyzed or nonhydrolyzed
PTH-(1-34). To ensure complete PTH hydrolysis,
sequential dilutions of PTH were incubated with membranes derived from
baculovirus Sf9 membranes expressing
rPhex-WT. PTH incubated with membranes from mock-transfected cells and PTH not incubated with membranes served as controls.
Statistical analysis. Analysis of variance was performed with the Statgraphics software package (Statistical Graphics, Princeton, NJ).
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RESULTS |
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Baculovirus-based expression of Phex-WT and
Phex-3'M in Sf9 insect cells.
A strategy was developed for heterologous expression and purification
of mouse Phex using Sf9 insect cells. The
entire coding region for Phex-WT or Phex-3'M was
integrated into an engineered baculovirus genome via a recombinant
transfer plasmid (see METHODS). For these constructs,
a V5 epitope tag was placed at the COOH terminus of Phex-WT
and Phex-3'M. A polyhistidine tag follows the V5 tag (Fig.
1A) to
provide an anchor for purification on the affinity matrix,
Ni2+-nitrilotriacetate-agarose beads (see below). In
membranes prepared from Sf9 cells expressing
rPhex-WT, we identified a predominant band of ~100 kDa
using the anti-V5 monoclonal antibody under reducing conditions (Fig.
1B). Because Phex has six N-linked glycosylation sites, we determined whether the 100-kDa band represents glycosylated Phex. To accomplish this, we treated rPhex
protein isolated from Sf9 membranes with
N-glycosidase F before SDS-PAGE electrophoresis and Western blot
analysis (Fig. 1C). Treatment of membranes with N-glycosidase shifted the apparent molecular mass of the 100-kDa protein to ~86 kDa, indicating that the rPhex protein
generated in Sf9 cells has N-glycosylation sites. The
molecular mass of 86 kDa is close to the theoretical mass deduced from
the Phex cDNA sequence. In some studies, we also observed
the confluence of several products that produced a broad band at ~200
kDa as well as a slightly smaller band of ~86 kDa (Fig.
1B). In membranes prepared from Sf9 cells
expressing rPhex-3'M, we identified a ~60-kDa band,
consistent with the truncation of the COOH-terminal region (Fig.
1B). No products were observed in mock-infected
Sf9 membranes or membranes treated with vector alone.
The epitope-tagged rPhex-WT was partially purified from
solubilized membrane fractions under native conditions by
Ni2+-affinity resin chromatography (Fig. 1D).
The expression of rPhex was enriched in membrane fractions
compared with total cell lysates, consistent with its designation as a
transmembrane endopeptidase (Fig. 1D).
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Detection of Phex protein using polyclonal anti-Phex antiserum.
We also examined the ability of our polyclonal Phex
antiserum to detect Phex overexpressed in the baculovirus
expression system in Sf9 cells and the retroviral
expression system in E86 cells as well as endogenously expressed
Phex in various cell lines and selected tissues. First, we
documented the ability of our antiserum to detect Phex by
immunoblot analysis of rPhex-WT in Sf9
cells. The Phex antiserum (Fig.
2A) detected the same product
as the anti-V5 antibody (Fig. 1B) in Sf9
cells expressing rPhex-WT but failed to detect products in
Sf9 control cells infected with vector alone.
Similarly, immunoblot analysis with polyclonal anti-Phex antiserum detected the ~100-kDa product (as well as a smaller band
likely representing a differentially glycosylated product) in E86
membranes transduced with the retroviral construct containing the
nontagged Phex-WT (Fig. 2A). Next, we used this
antiserum to confirm Phex expression in osteoblasts. We were
able to immunoprecipitate the correct-size ~100-kDa product from MG63
osteoblasts with the anti-Phex antiserum (Fig.
2B). The detection required immunoprecipitation, since we
did not see bands by Western blot analysis in any osteoblast cell line,
including MC3T3-E1, Saos, MG63, or U2Os, when we used 150 µg of
protein per lane (Fig. 2C). This likely reflects the low
abundance of native Phex in osteoblasts.
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Failure to confirm PTH as a physiological substrate
for rPhex-WT protein.
Because of recent reports suggesting that PTH is a substrate for
rPhex (12), we examined PTH hydrolysis by
uninfected Sf9 membranes (mock), membranes derived
from Sf9 cells infected with the engineered
baculovirus vector (vector alone), Sf9 membranes expressing rPhex-WT or rPhex-3'M, and
affinity-purified recombinant Phex-WT protein. Endopeptidase
activity was determined by analysis of PTH-(1-34)
cleavage by reverse-phase HPLC (Fig.
3A). Although we observed no
degradation of PTH-(1-34) in mock-infected
Sf9 membranes, as indicated by the single larger peak
at ~50 min (Fig. 3A, top), endopeptidase
activity was evident after incubation with Sf9
membranes expressing rPhex-WT (Fig. 3A,
bottom). Further analysis, however, found that membranes
from Sf9 cells infected with the engineered
baculovirus vector alone as well as the inactive rPhex-3'M
construct also hydrolyzed PTH-(1-34) (Fig.
3B). Membranes treated with vector alone and
rPhex-3'M membranes resulted in disappearance of the
PTH-(1-34) peak and its replacement by several smaller peaks identical to that observed with rPhex-WT.
Moreover, affinity purification of rPhex-WT resulted in the
loss of endopeptidase activity (Fig. 3B), rather than the
increase in activity that is expected from purification of
Phex (Fig. 1C). EDTA, which should inhibit the
Phex zinc metalloproteinases, also failed to inhibit PTH
cleavage by baculovirus-expressing membranes (Table
1). Rather, the endopeptidase activity of
membrane preparations from vector-alone and Phex-expressing
membranes was inhibited by a cocktail of protease inhibitors (Table 1).
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STC-1 also is not a Phex substrate.
We originally obtained the full-length human PHEX cDNA coding sequence
from screening a human bone library (6). Using the same
library, we isolated and cloned the full-length coding sequence of
hSTC-1 using a PCR method. Sequencing of our PCR product revealed 100%
identity to the published STC-1 sequence (accession no. U467768). The
full-length cDNA encoding hSTC-1 was successfully cloned in pFastBac-V5-His vector in frame with V5 and His tag at its COOH terminus, and recombinant protein was secreted into the media of
infected Sf9 cells. By Western blot analysis using
anti-V5 antibody (Fig. 4), we identified
a broad band at 30 kDa, which comprises three distinct bands derived
from differential glycosylation of hSTC-1, as previously reported
(32). We performed in vitro (Fig. 4A) and in
vivo (Fig. 4B) analysis of rPhex-WT cleavage of
hSTC-1. Incubation of isolated rhSTC-1 with
rPhex-WT-containing membrane preparations did not result in
hydrolysis of hSTC-1 (Fig. 4A). Similarly, coinfection of
rPhex-WT with rhSTC-1 in Sf9 cells did not
result in the appearance of degradation products in the supernatant of
cultured, coinfected Sf9 cells (Fig. 4B).
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Casein and FGF-23 peptide also are not Phex
substrates.
We tested the ability of mock-infected Sf9 membranes,
membranes treated with vector alone, and Sf9
membranes expressing rPhex-WT or rPhex-3'M, as
well as affinity-purified recombinant Phex-WT protein to
hydrolyze a fluorescent quenched casein (Table
2). We chose to evaluate casein, since it
is a substrate for several zinc metalloproteinases (13).
For these studies, we also evaluated the effects of thermolysin, a
related zinc metalloprotease, as a positive control. Whereas
thermolysin resulted in a dose-dependent proteolysis of BODIPY
TR-X-conjugated casein, we observed no hydrolysis of this substrate by
any of our baculovirus-infected Sf9 membranes or
rPhex protein.
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Cleavage of [Leu]enkephalin by rPhex.
Because the above-mentioned studies did not establish the activity of
our membrane-associated Phex endopeptidase, we tested whether rPhex-WT degrades [Leu]enkephalin
(Tyr-Gly-Gly-Phe-Leu), a substrate for the related NEP and thermolysin.
NEP and thermolysin cleave [Leu]enkephalin at the
Gly3-Phe4 bond, resulting in a faster-migrating
Tyr-Gly-Gly peptide by HPLC analysis (31).
[Leu]enkephalin alone migrated as a single peak (Fig.
6A). As expected, we observed
a faster-migrating peak, likely representing the Tyr-Gly-Gly peptide,
after incubating [Leu]enkephalin with thermolysin (Fig.
6B). A similar product was observed after treatment of
[Leu]enkephalin with rPhex-WT (Fig. 6D) but not
after incubation with membrane preparations transfected with vector
alone (Fig. 6C). In addition, the cation-chelating agent
EDTA completely inhibited the ability of rPhex-WT to
hydrolyze [Leu]enkephalin (Fig. 6D), consistent with the
fact that Phex is a zinc metalloproteinase.
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DISCUSSION |
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In this report, we evaluated experimental systems to overproduce rPhex. Our studies on heterologously expressed epitope-tagged rPhex in Sf9 insect cells by using baculovirus-based vectors (Fig. 1) and nontagged rPhex in E86 cells using a retroviral vector (Fig. 2) indicate that this endopeptidase is a ~100-kDa membrane-associated glycosylated protein. After deglycosylation, the molecular mass decreases to 86 kDa (Fig. 1C), which is the size predicted for the protein deduced from the cDNA sequence. In addition, we demonstrated that the Phex protein is expressed in MG63 osteoblast membranes by immunoprecipitation-Western blot using polyclonal antiserum against a synthetic peptide corresponding to the COOH-terminal region of Phex. The fact that immunoprecipitation was required to detect Phex in osteoblasts indicates that the endogenous endopeptidase is expressed in low abundance in cell culture models.
We used these recombinant proteins to evaluate potential substrates. We found that rPhex-WT cleaves the [Leu]enkephalin (Fig. 6). The HPLC profiles of [Leu]enkephalin degradation in the presence of rPhex-WT, the similar cleavage by thermolysin, and the sensitivity of this activity to ion chelation with EDTA strongly suggest that we have successfully overexpressed a functional rPhex protein in our Sf9 membrane preparations. To our knowledge, this is the first published report that a substrate for NEP can also be cleaved by the related Phex endopeptidase. This substrate should be useful for further studies evaluating Phex function.
We also evaluated whether PTH is a substrate for Phex. Although we demonstrated PTH cleavage, we found that PTH hydrolysis was not specific for rPhex. Rather, membranes derived from Sf9 cells infected with the baculovirus alone, rPhex-WT, and rPhex-3'M displayed similar activity (Fig. 3B). Endopeptidase activity toward PTH also was lost after affinity purification of Phex-WT and renewal of membranes (Fig. 3B). Moreover, nonspecific cleavage also was observed with membranes expressing non-epitope-tagged rPhex-WT as well as inactive rPhex-3'M. In addition, EDTA, which should inhibit the Phex zinc metalloproteinases, failed to inhibit PTH cleavage by baculovirus-expressing membranes (Table 1). Finally, the membrane-associated endopeptidase activity from membranes treated with vector alone and those expressing Phex was inhibited by a cocktail of protease inhibitors (Table 1). These findings suggest that the observed cleavage of PTH is due to nonspecific endopeptidase activity derived from membrane preparations rather than the heterologously expressed Phex protein.
Our results contrast with another report showing that membranes from COS-7 cells transiently expressing Phex degrade PTH (12). These studies, however, did not characterize the requirement for metal ions, evaluate the effects of inhibitors on endopeptidase activity, determine the cleavage site in PTH, attempt to evaluate the activity of purified Phex, or use inactive mutant rPhex as controls. There also are no follow-up studies that confirm Phex cleavage of PTH-(1-34). Nevertheless, by showing that hydrolysis of PTH can occur as a result of contaminating enzymes in membrane preparations, we raise the possibility of nonspecific cleavage. We also failed to identify any alterations of the bioactivity of PTH after hydrolysis of membrane preparations (Fig. 3C), indicating that the observed cleavage does not alter PTH function, whatever the responsible enzyme. These findings, as well as previous studies that indicate that PTH is not responsible for phosphaturia in XLH (16), indicate that PTH may not be a physiologically relevant Phex substrate.
Phex is expressed in osteoblasts, albeit in low abundance (Fig. 2B), and is associated with genes regulating extracellular matrix production and mineralization (9, 19). Therefore, Phex in bone may degrade local regulators of phosphate homeostasis and mineralization (21). On the basis of this reasoning, we evaluated whether the phosphate regulatory factor STC-1, which is expressed in bone, is a substrate for Phex. We confirmed previous reports (11) of the presence of STC-1 in bone by isolating and cloning its full coding sequence from a human osteoblast library from which we originally isolated PHEX (6). However, we were unable to demonstrate cleavage of our recombinant hSTC-1 by rPhex under the conditions studied (Fig. 4). Moreover, Phex cleavage of STC-1 would be paradoxical to our current view of the pathogenesis of XLH. Because STC-1 stimulates phosphate transport, its accumulation would cause hyperphosphatemia rather than hypophosphatemia. This, as well as our negative findings, suggests that hSTC-1 is not a physiologically relevant substrate for Phex.
The possibility that XLH and ADH may share a common pathogenesis gives rise to the notion that FGF-23 may represent a Phex substrate (21). Moreover, it is well established that other members of the related matrix metalloproteinase family modulate the activity of several growth factors, including FGF (14). The mutation of FGF-23 that causes ADH has been proposed to disrupt the cleavage site in FGF-23 for Phex (21). Although we have not evaluated the full-length FGF-23 protein, we have demonstrated that the peptide encompassing the region mutated in ADH is not a substrate for Phex (Fig. 5).
In conclusion, the investigation of the pathogenesis of XLH, ADH, and OHO indicates a role for the Phex endopeptidase and novel factors regulating phosphate homeostasis. In an effort to provide evidence for a common pathogenesis for these disorders, we have created wild-type and mutated recombinant Phex proteins and used them to evaluate various candidate substrates. We have shown that the insect cell-based baculovirus system and retroviral-mediated overexpression are well suited for expression and glycosylation of rPhex. We have demonstrated that [Leu]enkephalin is a substrate for rPhex, but we were unable to show that FGF-23, hSTC-1, PTH, and casein are substrates for our rPhex protein. Rather, we have demonstrated the potential of coisolated enzymes in membrane preparations to confound interpretations of enzyme specificity. In the larger context of understanding the pathophysiology of these hypophosphatemic disorders, however, our negative studies are important. With regard to ADH, our findings suggest alternatives to a simple enzyme-substrate hypothesis where FGF-23 is a Phex substrate. Rather, our studies suggest a more complex model whereby FGF-23 is not a PHEX substrate but causes phosphaturia indirectly by altering the production of the PHEX substrate (i.e., phosphatonin) and/or the activity of PHEX itself. Possible support for this is derived from recent studies demonstrating regulation of Phex expression in bone by insulin-like growth factor I (33). Our inability to demonstrate cleavage of the FGF-23 peptide may also mean that ADH and XLH cause renal phosphate wasting through distinct pathways. With regard to XLH, we have excluded PTH and hSTC-1 as physiological substrates for Phex. On the basis of our findings of Phex protein expression in osteoblasts, the possibility remains that Phex substrates may exist in bone. Nevertheless, our results indicate that the pathogenic mechanisms underlying ADH and XLH are more complex than originally anticipated. Further advances in our knowledge of the biological function of FGF-23 as well as identification of Phex substrates are needed to piece together the pathogenesis of these phosphate-wasting disorders.
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ACKNOWLEDGEMENTS |
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We thank Dr. Patrick J. Casey for assistance with the baculoviral expression of Phex and Patrick Flannery for technical support.
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FOOTNOTES |
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This work was supported in part by National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant AR-45955.
Address for reprint requests and other correspondence: L. D. Quarles, Box 3036, Duke University Medical Center, Durham, NC 27710 (E-mail: Quarl001{at}mc.duke.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 30 January 2001; accepted in final form 17 May 2001.
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