(Received for publication, February 13, 1996, and in revised form, January 13, 1997)
From the Laboratory for the Study of Skeletal Disorders and Rehabilitation, Department of Orthopaedic Surgery, Harvard Medical School, and Children's Hospital, Boston, Massachusetts 02115
Osteopontin (OPN) is one of the major secretory phosphoproteins in both calcifying and non-calcifying tissues. Evidence has accumulated for the biological importance of the phosphoproteins and, in particular, the phosphate groups in bone formation, resorption, and calcification. The precise locations of the phosphate groups in the OPN molecule were determined by metabolically labeling OPN with 32P in cultured chicken osteoblasts, followed by purification to homogeneity. N-terminal sequencing showed a single sequence of WPVSKRQHAISA, consistent with that deduced from both cDNA, and previous amino acid sequencing of the protein isolated from chicken bone. Three 32P-labeled peptides were isolated by reverse-phase high performance liquid chromatography of thrombin-digested, 32P-labeled OPN. The N-terminal sequencing of each of these thrombin fragments gave single sequences as follows: WPVSKSRQHAIS, SHHTHRYHQDHVD, and ASKLRKAARKL, with approximate molecular masses of 5, 30, and 20 kDa. These data demonstrate that 32P was incorporated throughout the N- to C-terminal sequence of the protein. Thrombin specifically cleaved chicken OPN at two sites: between Arg-22 and Ser-23, which generated the 5-kDa N-terminal end fragment, and another between Lys-138 and Ala-139, which generated the 30- and 20-kDa fragments. To further define the exact locations of the phosphorylated amino acids and the surrounding amino acid sequences, OPN was digested with trypsin, which generated seven major 32P-labeled peptides whose amino acid sequences were determined. The phosphorylated peptide regions of osteopontin were identified as amino acids 8-18 (QHAIS*AS*S*EEK), 39-54 (LASQQTHYS*S*EENAD), 150-171 (LIEDDAT*AEVGDSQLAGLWLPK), 179-191 (ELAQHQSVENDSR), 194-205 (FDS*PEVGGDSK), 214-219 (ES*LASR), and 239-248 (HSIENNEVTR). The phosphorylated amino acid sites are followed by an asterisk (*). Of the seven identified phosphorylated peptide regions, three were localized on the N-terminal end of the osteopontin molecule (with five phosphorylated serines) and contained the sequence motifs that were phosphorylated by casein kinase II type(s), whereas the remaining four peptides are concentrated toward the C-terminal half of the molecule (with five phosphorylated residues) and contained recognition motifs for other kinases as well as casein kinase II.
It has been well established that the processes of phosphorylation and dephosphorylation of proteins catalyzed by protein kinases and phosphatases, respectively, play a major role in the initiation, regulation, and termination of a wide range of intracellular biochemical processes with significant functional consequences. Such processes may also play an important role in a wide variety of intercellular mechanisms, including general cell-cell signal transduction of extracellular agonists via specific transmembrane receptors, often causing alterations of intracellular concentrations of cAMP, calcium, inositol polyphosphates, and/or diacylglycerol. These intracellular modulators in turn induce a cascade of biochemical processes by mechanisms that also involve phosphorylation of specific rate-determining enzymes or proteins by protein kinases (1-3). Phosphoproteins containing o-phosphoserine and o-phosphothreonine are not only found in the intracellular compartments of cells, but have also been identified in extracellular matrices, particularly those of normal vertebrate mineralized tissues and in experimental and human pathologically calcified tissues (4-16). There is uncertainty as to the biological functions of these proteins and the precise role of the covalently bound phosphate groups per se. The presence of these phosphorylated proteins in all of the normal and pathologically mineralized tissues of vertebrates, and their ultrastructural localization (17-23), have combined to suggest that one biological function of these phosphoproteins and specifically of the covalently bound phosphate groups is their critical role in the nucleation and growth of inorganic calcium-phosphate crystals (24-26). This view is also supported by in vitro nucleation experiments, which attempted to simulate the postulated in vivo nucleation substrate of bone tissue by cross-linking the resident phosphoproteins in situ in their native positions and comparing the efficacy (induction or lag time) of this nucleation substrate with samples containing the collagen-phosphoprotein complexes after the covalent phosphate groups were enzymatically cleaved. Not only did the decalcified bone samples containing the collagen-phosphoproteins complexes markedly decrease the nucleation induction time, but this property was lost when the phosphate groups alone were removed enzymatically, leaving the dephosphorylated collagen-phosphoprotein completely intact and, hence, pinpointing the role of the phosphate groups in this in vitro crystal nucleation event (27, 28). In addition to their postulated role in calcification, these phosphoproteins have been implicated in many other biological functions (13-16, 29).
Two major glycosylated phosphoproteins in bone and many other mineralized tissues, osteopontin (OPN)1 and bone sialoprotein (BSP), have been extensively studied in the past decade. The potential roles of these two phosphoproteins in the calcification process have not been clearly defined. Data have been reported demonstrating that these two components can induce or inhibit calcification in several different in vitro models of calcification (30-33). The results may be due to the experimental techniques and models utilized to assess crystal nucleation and the accumulation of increasing amounts of the solid crystalline phase of calcium phosphate by secondary nucleation and possibly by growth of the crystals. However, their presence in pathologically calcified soft tissue such as breast tumors, urinary stones, and atherosclerotic plagues (13-16) supports the general hypothesis that they are involved in the deposition of a solid phase of calcium phosphate in normal and pathologically calcified tissues.
OPN is one of the most characterized of the secreted phosphoproteins. It is synthesized in a highly phosphorylated form by osteoblasts (34, 35), and can be isolated from the mineralized extracellular matrix of bone tissue (12, 26). In contrast, except for milk, it is essentially unphosphorylated in non-mineralized tissues (37, 38). Clearly, such differences in the extent of phosphorylation of the same protein when expressed in widely different tissues suggests different functional properties of both the unphosphorylated and phosphorylated protein, and further suggests the presence of tissue-specific kinase(s) that modify this protein. The protein kinase activity found in the bone, which phosphorylates the endogenous proteins of bone, was first observed three decades ago (39). More recently, the presence of membrane-bound casein kinase type II activity was detected in 14-day-old embryonic chicken bone, which phosphorylated endogenous bone phosphoproteins and dephosphorylated casein (40). Further studies using cultured chicken osteoblasts as the source of protein kinase from bone cells led to the conclusion that the microsomal casein kinase II (CKII) type activity of chicken osteoblasts was the predominant enzyme that phosphorylated purified chicken OPN and recombinant mouse OPN (41, 42). In these studies assessment of this enzyme with other protein substrates, including casein and synthetic peptide substrates, indicated that, while its activity is very similar to that of casein kinase II, it also possessed unique enzymatic activities that distinguished it from that of classical casein kinase II (41, 42). This enzyme was found to be localized in the Golgi apparatus of chicken osteoblasts (43). Furthermore, in vitro studies also identified the major protein kinase responsible for the phosphorylation of both bovine OPN and BSP to be a CKII (44). Other investigators using ROS 17.28 osteosarcoma cells have also isolated a membrane-associated CKII, which phosphorylates the major dentin phosphoprotein, phosphorphoryn (45). This enzyme was also found in the endoplasmic reticulum and Golgi apparatus (46, 47). The subcellular location of the CKII to the endoplasmic reticulum and Golgi apparatus is significant mechanistically, since both OPN and BSP are secretory proteins and the enzyme participating in their post-translational modification must encounter them.
Extensive studies have been carried out in this laboratory to isolate, purify, and characterize the protein kinase(s) responsible for the phosphorylation of the secreted extracellular matrix phosphoproteins of bone. Quantitative amounts (mg) of 32P-secreted osteopontin from cultured chicken osteoblasts were isolated and purified to define both the location of the phosphorylated amino acids in the protein and the amino acid sequences adjacent to the phosphorylated residue. This paper reports on the isolation and purification of 32P-labeled peptides of OPN and the solid-phase amino acid sequencing of seven peptides, which unequivocally identify the peptide motifs and the specific sites of the phosphorylated OPN.
Materials
Sources for materials were are follows: GF/F glass filters (5.0 mm diameter) (Whatman, Ltd.); p-aminophenyltrimethoxysilane (Petrach Co.), radioactive orthophosphate
(32Pi, 3000 Ci/mmol, ICN Inc.); reverse-phase
Vydac C-4 column (15 × 0.38 cm) (Nerst Group); reverse-phase HPLC
Dynamax C-18 column (25 × 0.46 cm) (Rainin Co.); and trypsin
(tosylphenylalanyl chloromethyl ketone-treated), human thrombin (5000 units/mg), endoproteinase Asp-N, Kemptide (LRRASAV), and
cAMP-dependent kinase, water-soluble coupling agent
N-ethyl-N-(dimethylaminopropyl)carbodiimide
(EDAC) (all obtained from Sigma).
Methods
Labeling and Isolation of Secreted 32P-Labeled Osteopontin from Mineralized Cultured Chicken OsteoblastsChicken
calvarial osteoblasts were isolated by sequential collagenase/trypsin
treatment of 17-day-old embryonic calvaria. Cells were initially
maintained in minimal essential media supplemented with 10% fetal
bovine serum until confluence. They were then put into BGJb
media supplemented with 10% fetal bovine serum, 12.5 µg/ml ascorbic
acid, and 10 mM -glycerol phosphate. Cultures were grown
for 3 weeks in the latter medium to promote extracellular matrix
mineralization. To generate 32P-labeled OPN, mineralized
cultures (15 100-mm culture dishes, 15 × 106
cells/dish) were initially incubated with phosphate-free Dulbecco's modified Eagle's medium for 1 h, followed by incubation of
Dulbecco's modified Eagle's phosphate-free medium (5 ml/dish)
containing 12.5 µg/ml ascorbic acid (1 mCi of 32P/dish)
and incubated for 18 h. All protein extraction buffers and wash
buffers were ice-cooled to ~4 °C. The medium was removed, and the
mineralized cell layers were washed (three times) with 10 ml of 10 mM phosphate buffer, pH 7.2. The cells were lysed with
ice-cooled phosphate buffer (2 ml/dish), pH 7.2, containing 0.5% Tween
20, 0.05% deoxycholate, 50 mM NaCl, 1 mM
MgCl2, 50 mM NaVO3, 10 mM NaF, and 1 mM phenylmethylsulfonyl fluoride
for 10 min on ice. The lysis buffer was removed, and the mineralized cell layer was washed (three times) with 10 ml of 10 mM
phosphate buffer (pH 7.2)/dish. The mineralized cell layer was then
extracted 0.1 M HCl (adjusted to pH ~2.0 with citric
acid) 10 ml/dish over ice for 3 h. The acid extracted proteins
were lyophilized, and 32P-labeled OPN was purified by
reverse-phase HPLC on a C-4 column as described below.
The lyophilized acid extract was resuspended in ~0.5 ml of H2O, 0.1% trifluoroacetic acid (v/v), and aliquots (0.25 ml) were subjected to reverse-phase HPLC on a C-4 column (15 × 0.38 cm). After injection, the column was washed for 10 min with H2O, 0.1% trifluoroacetic acid (v/v) followed by elution using a linear gradient from H2O, 0.1% trifluoroacetic acid (v/v) to 60% CH3CN, 0.55% trifluoroacetic acid (v/v) at 60 min at a flow rate of 0.72 ml/min. Fractions of 0.72 ml were collected, and absorbance = 230 nm was recorded by a continuous on-line chart recorder/integrator. Aliquots of 0.025 ml from each fraction were counted for 32P. The radiolabeled peaks were pooled, freeze-dried, and rechromatographed under same conditions as described above. An aliquot of the purified osteopontin was then N-terminally sequenced.
Thrombin Digestion of Purified 32P-Labeled Osteopontin and Isolation of Thrombin Fragments by HPLC on a C-4 ColumnApproximately 200 µg of purified 32P-labeled OPN was incubated with bovine thrombin (1 unit/4 µg of protein) in 0.20 ml of 50 mM Tris buffer, pH 8, containing 2 mM CaCl2 for 2 h at 37 °C. The reaction products were treated with 50 µl of H2O, 0.1% trifluoroacetic acid (v/v) and HPLC-chromatographed on C-4 column (15 × 0.38 cm) as described above. The 32P-radiolabeled fragments were identified by counting for 32P, separately pooled, and freeze-dried. Aliquots of each thrombin fragment were N-terminally sequenced.
Trypsin Digestion of Purified 32P-Labeled Osteopontin and Isolation of the Labeled Peptides by Reverse-phase HPLC on a C-18 ColumnThe purified 32P-labeled OPN (~0.5 mg) from the HPLC C-4 column was digested with trypsin (2% w/w) in 0.2 ml of 50 mM NH4HCO3, pH ~8.0, for 20 h at 37 °C. The reaction products were then treated with 50 µl of H2O, 0.1% trifluoroacetic acid (v/v) and subjected to HPLC on a C-18 column (25 × 0.46 cm). After injection the column was washed for 10 min, followed by linear gradient elution from H2O, 0.1% trifluoroacetic acid (v/v) to 60% CH3CN, 0.055% trifluoroacetic acid (v/v) over 120 min, with a second gradient from 60% CH3CN to 80% CN3CN over 30 min at a flow rate of 0.5 ml/min. The absorbance at 219 nm was recorded continuously on an on-line chart recorder-integrator using a Gilson HM Holochrome detector. Fractions of 0.5 ml were collected. Aliquots of 0.025 ml from each fraction were counted for 32P radioactivity, and the radioactive fractions were then separately pooled for each peak, freeze-dried, and rechromatographed. Each purified 32P-labeled peptide was then sequenced by automated solid-phase amino acid sequencing technique described below.
N-terminal Sequencing Utilizing Biobrene-treated Glass FiltersN-terminal sequencing was carried out by Edman degradation (48) using an automated model 477A sequenator (Applied Biosystems Inc., Foster City, CA) under the same conditions as described previously (49). Glass filters coated with Biobrene were used, and the proteins or peptides were adsorbed on these filters. This approach was used for N-terminal sequence analysis in which the prime interest was to identify the N-terminal sequence, for example, of purified secreted [32P]OPN and large thrombin fragments. For identification of the 32P-labeled peptide regions and the specific sites of phosphorylation, the sequencing conditions are described below.
Automated Solid-phase N-terminal Sequencing in Identification of Sites of PhosphorylationThe sequencing method for identification of phosphorylated sites required an approach that would optimize the identification of phosphorylated residues such as Ser, Thr, or Tyr. This is because these derivatives as well as inorganic phosphate that may be released during sequencing conditions are highly hydrophilic and not extracted by the usual sequence extraction solvent (1-chlorobutane). Therefore, there is significant limitation in attaining sufficient 32P counts for identification of the site of phosphorylation. A solid-phase sequencing technique utilizing GF/F glass filters derivatized by p-aminophenyltrimethoxysilane, followed by covalent attachment of the phosphorylated peptide through its C terminus (50), was used, which allowed the use of a sufficiently hydrophilic extraction solvent (S3) without loss of the peptide, which would be the case if the "adsorptive" Biobrene method was used.
The glass GF/F Whatman filters (5.0 cm diameter) were treated with trifluoroacetic acid overnight at 37 °C. These filters were then freed of trifluoroacetic acid, air-dried in a fume hood, and treated with p-aminophenyltrimethoxysilane overnight at 37 °C. The filters were removed from the solution and extensively washed with MeOH and dried. In the present study, trimethoxysilane was used instead of "triethoxysilane" since the latter reagent was no longer commercially available. The derivatized filters were stored in dry, dark glass dishes until needed. The covalent attachment of the 32P-labeled peptides was carried out on derivatized GF/F glass filters cut in circular sizes of ~1 cm in diameter and using water-soluble carbodiimide (EDAC) (50).
The automated N-terminal sequencing of the 32P-labeled peptides were performed directly from these filters. Further modifications of the sequencing conditions included the following. The extraction solvent S3 was CH3CN instead of 1-chlorobutane and contained 0.1% thiourea, and the acid cleavage step was performed using HCl/MeOH (1 ml of acetyl chloride + 13 ml of methanol + 0.1% thiourea). Thiourea was used in the sequencing solvents instead of DTT. This is more advantageous than DTT since phosphoserines under sequencing conditions can be derivatized by nucleophilic attacks by DTT. For identification of sites of phosphorylation, one third of the ATZ products of each cleavage step were converted to PTH-derivatives and analyzed by on-line HPLC (model 120A, Applied Biosystems) and two thirds were collected as ATZ-derivatives for 32P counting. To quantify yields of sequenced peptides, initial yield (Io) and repetitive yield (R) were calculated by linear regression analysis of the observed yield (M) at each cycle (n): log10(M) = n log10(R) + log10(Io).
Evaluation of Efficiency of the Solid-phase N-terminal Sequencing of 32P-Labeled Peptides by Using Synthetic Peptide Substrate, Kemptide, 32P-Phosphorylated by cAMP-dependent KinaseKemptide (LRRASAV), 20 µg (26 nmol) was phosphorylated using cAMP-dependent kinase (1 µg) and 150 µM [32P]ATP (4 mCi/mmol) in
0.1 ml of KH2PO4/NaHPO4 buffer, pH
7.4, I (ionic strength) = 0.2 mol liter1 for
30 min at room temperature. The phosphorylated Kemptide was isolated by
HPLC on a C-18 column using a linear gradient from H2O,
0.1% trifluoroacetic acid (v/v) to 50% CH3CN, 0.055%
trifluoroacetic acid (v/v) in 40 min. An aliquot of the
32P-labeled Kemptide (~3 nmol; ~26,500 dpm) in 40 µl
of CH3CN/H2O (1:3) was applied onto a
derivatized Whatman GF/F glass filter (~1.0 cm in diameter) and
dried. 40 µl of 160 mM MES buffer, pH 4.0, containing (5 mg/ml) EDAC was added and incubated for 1 h at 37 °C. The
filter was washed with H2O and counted. 17,200 dpm remained
on the filter, which was equivalent to ~65% covalent coupling.
N-terminal sequencing was then carried out directly from the filter
with one third of the Edman degradation products converted to
PTH-derivatives and analyzed on an on-line HPLC model 120A (Applied
Biosystems), and two thirds were collected as ATZ-derivatives and
counted for 32P.
SDS-PAGE (10%) polyacrylamide
gel electrophoreses were carried out as described previously (51).
Samples of the isolated proteins from mineralized bone and
[32P]ATP-labeled proteins were prepared in 10% sample
buffer (20% sucrose, 10% SDS, 0.12 M Tris (pH 6.8),
0.25% bromphenol blue, 10% 2-mercaptoethanol, and 25% glycerol) and
heated at 100 °C for 5 min. Autoradiography was carried out on
32P-labeled proteins, which had been resolved by SDS-PAGE
after the gels had been stained and unstained. Gels were dried on a Bio-Rad dryer (model 224) for 2-3 h and exposed to X-Omat AR film (Eastman Kodak Co., Rochester, NY) at 70 °C.
Following SDS-PAGE with prestained low
Mr protein standards (Bio-Rad), the gels were
soaked in blotting buffer (48 mM Tris, 39 mM
glycine, 20% methanol, pH 9.2) with Immobilon P and 6 pieces of
Whatman filter paper for 30 min. The electrophoretic transfer of
proteins onto Immobilon was carried out at 14 V and 280 mA for 1 h
using Bio-Rad Trans-Blot SD blotter. The membranes were blocked
overnight at room temperature by 10% nonfat milk in 20 mM
phosphate buffer, pH 7.4, containing a 0.15 M NaCl and
0.1% Tween 20. The Immobilon P-bound proteins were subjected to
specific reaction with anti-chicken OPN (19) and 170 ng/ml of the same buffer without the nonfat milk antibodies for 1 h. The membranes were then washed three times with the same buffer and incubated with
horseradish peroxidase-conjugated goat anti-rabbit IgG antibodies for
1 h. After several washings, the membranes developed with 3,3,5,5
-tetramethylbenzidine substrate. Nonspecific reactions were
evaluated using primary and secondary antibodies with chicken, mouse,
and rabbit serum albumin.
Our initial attempts at large scale purification of
32P-labeled secreted OPN of mineralized cultured chicken
osteoblasts directly from the culture media were unsuccessful. However,
milligram amounts of 32P-labeled OPN were extracted from
the calcified cell layer matrix in 0.1 M HCl after 14 days
of culture and subsequently purified by two successive reverse-phase
HPLC chromatographies on a C-4 column (Fig. 1). The
purified OPN had a single N-terminal sequence (12 residues):
WPVSKRQHAISA, which was consistent with the sequence deduced from
cDNA (52) and direct amino acid sequencing of chicken OPN (18).
Most of the OPN (>90%) was recovered from the cell-calcified matrix
layer; less than 10% was found in the culture medium. Analysis of the
purified 32P-labeled OPN on SDS-PAGE gave a molecular mass
of ~58 kDa. The protein reacted with a polyclonal antibody of chicken
osteopontin (Fig. 2, panel A). OPN, isolated
and purified from mineralized chicken bone (44) and used as a substrate
for the protein kinases isolated from the osteoblast membrane fractions
(42), had the same characteristics as the secreted OPN from the
cultured osteoblasts (Fig. 2, panel B), indicating that the
cultured osteoblasts post-translationally modified the protein in a
fashion similar to that of osteoblasts in vivo.
After cleavage of the 32P-labeled OPN with thrombin, three
radiolabeled fragments were isolated by reverse-phase chromatography on
a C-4 column (Fig. 3), fractions 37, 42, and 45. N-terminal sequencing revealed a single sequence in each of the three
peptides: F37, WPVSKSRQHAIS; F42,
SHHTHRYHQDHVD; and F45, ASKLRKAARKL.
SDS-PAGE of the whole thrombin digest revealed three components with molecular masses of 5, 20, and 30 kDa (Fig. 3, inset).
The performance efficiency of the solid-phase sequencing was evaluated
using 32P-labeled synthetic peptide, Kemptide (a substrate
for cAMP-dependent protein kinase). Sequence analysis
showed that the procedures adopted and developed were effective in
identifying both the phosphorylated amino acid sequences and the site
of phosphorylation. The identification of a particular phosphorylated
residue was accomplished during sequencing by collecting two thirds of
the Edman degradation products of each cycle as ATZ-amino acid
derivative and radioactive counting for 32P radioactivity;
one third was converted to the PTH-amino acid derivative and analyzed
by on-line HPLC to identify the amino acids. 32P-Labeled
Kemptide (1.95 nmol, 17, 200 dpm), covalently attached through its C
terminus to derivatized Whatman GF/F glass filter, was sequenced as
LRRASAV, with an initial yield of I = 251 pmol and a
repetitive yield R = 82%. Counts for 32P
for each cycle of Edman degradation showed a sharp increase in
32P released at cycle 5, which corresponded to the serine
residue, indicating that the phosphate group was present in this
residue. At cycle 5 residual serine and dehydroalanine (a elimination product of phosphoserine) were identified. Thus the total
count in this cycle reflected 32Pi + 32P-Ser. Total radioactivity recovered (10,270 dpm) through
the sequencing steps was 6846 dpm from two thirds of the Edman
degradation and 3423 dpm. from the one third used for PTH-amino acids,
reflecting a recovery of ~60%. The filter retained 4390 dpm (25% of
the initial total 32P dpm) after eight cycles.
To identify the specific sequences adjacent to the phosphorylated
serine residue, 32P-labeled OPN was digested with trypsin
and the 32P-labeled peptides isolated by reverse-phase HPLC
on a C-18 column (Fig. 4). Seven major radiolabeled
peptides were isolated and N-terminally sequenced. Typically, 200-300
pmol of the 32P-labeled peptides from secreted OPN were
recovered and covalently attached to derivatized GF/F glass filters,
representing 65-85% of the total as determined from the
32P counts after washing the filters. The total
32P radioactivity on the filter ranged from 2000 dpm to
6000 dpm. 32P released during sequencing and counting of
two thirds of each cycle gave a background of 30-40 dpm, with the
cycle containing the phosphorylated residues releasing 100-300 dpm
depending on the initial 32P level in a given peptide.
Typical data are shown in Fig. 5, which demonstrates
that the repetitive yields of sequencing were approximately 85%, with
initial yields ranging between 50 and 300 pmol, depending on the amount
of a given peptide. It is worth noting that during sequencing, some of
the Ser(P) led to the formation of some dehydroalanine by
-elimination and thus the 32P counts of a specific cycle
reflected a combination of 32Pi and
32P-Ser. This method permitted simultaneous identification
of the amino acid sequence of the phosphorylated peptide and the
phosphorylated residue. While unequivocal identification of each
peptide sequence and the site of phosphorylation was possible, the size
of one peptide (Fig. 4, fraction 6) with an amino acid sequence
starting at Tyr-29 and extending to Lys-78 did not allow the
unequivocal identification of the site of phosphorylation. The
sequencing of this peptide up to 20 amino acid did not release any
32P radioactivity, indicating that the phosphate label was
not on the Thr-26, Ser-36, Ser-38, or Thr-45. The
32P-labeled peptide containing residues Tyr-29 to Lys-78
was then further digested with endopeptidase Asp-N, from which a
smaller 32P-labeled peptide (Leu-49 to Lys-78) was isolated
by reverse-phase HPLC chromotography on a C-18 column. N-terminal amino
acid sequencing revealed a peptide with an SSEE region with only 9 amino acids from its N-terminal end.
The seven phosphorylated peptide regions of OPN were identified as:
peptides 8-18 QHAIS*AS*S*EEK), 48-63 (DLASLQQTHYS*S*EENA), 149-162 (LIEDDAT*AEVGDSQLAGLWLPK), 179-191 (ELAQHQSVENDSR), 194-205 (FDS*PEVGGDSK); 214-219 (ES*LASR), and 239-248 (HSIENNEVTR),
with the specific residues phosphorylated followed by asterisks. The distribution of the phosphorylated peptide regions and the specific sites of phosphorylation relative to each other and to the complete primary amino acid sequence of chicken OPN are shown in Fig.
6. The peptides with amino acid sequences 8-18, 48-63,
and 149-162 contain sequence motifs, which are recognized only by
CKII (SEE or SXE), while the peptides 179-191, 194-205,
and 214-219 contain recognition sequences for both CKII(s) as well
as cGMP-dependent kinases (XSR(K)X).
In addition, the phosphorylated serine in peptide 194-205 had a
sequence recognition for Ca2+/Calmodulin-dependent
protein kinase (RXXS). Two peptides, 179-191 (ELAQHQSVENDSR) and 239-248 (HSIENNEVTR), were also phosphorylated. The time lapse between HPLC purification of these peptides and their
solid-phase sequence analyses resulted in a sufficient decay of
32P so that the release of 32P during
sequencing was not high enough to define which of the two serines were
phosphorylated. Based on the data obtained from the other peptides, it
appeared that the serine residues in the "SXE" sites
were the only residues phosphorylated. However, phosphorylation of the
second serines "S(T)R" in these peptides could not be excluded.
Our initial survey of the culture medium and the cell-calcified
matrix layer during the isolation and purification of quantitative (milligram) amounts of 32P-labeled secreted OPN from
cultured chicken osteoblasts indicated that 90% of the secreted OPN
was in the calcified matrix of the cell layer. Our quantitative
analysis of OPN in the medium and mineral layer during purification
indicated that 93% of the secreted 32P-labeled OPN was in
the mineral and 7% in the medium. These results were based on
calculations from the HPLC-purified material in terms of the
32P counts and protein content. Furthermore, from such data
the calculated ratio of 32P label/OPN protein for medium
and mineralized layer were comparable, suggesting that the
phosphorylated forms of OPN in these two compartments were of very
similar nature.
The unique cleavage of OPN by thrombin (19, 53-57), which cleaved chicken OPN at two specific sites (one between Arg-22, and Ser-23 and the other between Lys-138 and Ala-139) (Fig. 6), was used to establish that phosphorylated sites were distributed throughout the length of the OPN molecule, which is synthesized and secreted by cultured chicken osteoblasts. In vitro studies of the extent of phosphorylation and of the protein kinases that predominantly phosphorylate bovine bone OPN showed that bovine bone OPN was highly phosphorylated (~9 mol of phosphate/mol of OPN) by CKII, and that these phosphates were distributed on both the N-terminal and C-terminal halves of the molecule (44), consistent with the results obtained from metabolically labeled chicken OPN of cultured osteoblasts.
The present results indicate that three peptides (8-18, 48-65, and 149-162) concentrated in the N-terminal half of the molecule are phosphorylated only by a CKII, while the remaining four peptides concentrated in the C-terminal half of the molecule appear to be phosphorylated by a combination of CKII and cGMP-dependent kinase (peptide 179-191). Peptide 194-205 can be phosphorylated on Ser-196, either by an CKII or a Ca2+/calmodulin-dependent kinase, and Ser-204 of this peptide can be phosphorylated by cGMP-dependent protein kinase. The results indicate that the CKII is the predominant kinase that post-translationally modifies chicken bone OPN (6 serines and 1 threonine residue; ~70% of the total phosphorylated residues), while the remaining ~30% of the total phosphorylated residues (3 serines) are phosphorylated by other kinases. The determination of the total number of moles of phosphate/mol of OPN in the metabolically 32P-labeled OPN synthesized by cultured chicken osteoblast was not possible using 32P count and the content of the protein. This is because the specific activity of 32P during pulse-chase experiments cannot be accurately established. This is the result of the synthesis of ATP, which occurs from both unlabeled Pi and 32Pi within the cells during the pulse-chase, leading to dilution of the added 32Pi, the extent of which is unknown. However, a good estimation of the total of the content of 32P for each specific tryptic peptide and the amino acid sequences was possible from the data obtained. For example, peptides F1, F2, F3, F4, and F7 all contained a single phosphorylated residue but contained different concentrations of 32P. It can be assumed that the peptide F3, which contained the largest concentrations of 32P, represented 1 mol of phosphate/mol of peptide (i.e. 100% phosphorylation). Using this as a base line, the relative distribution of phosphates on each peptide and phosphorylated residue was calculated (Table I). It is interesting to note that metabolic phosphorylation led to a wide range of degrees of phosphorylation on different peptides (30-100%). Additionally, peptides that contained multiple juxtaposed phosphorylated serines, e.g. peptides F5 and F6, showed different extents of phosphorylation of the adjacent serine residues. In F5, the three serines were about equally phosphorylated (30% of each, i.e. a total 0.88 mol of phosphate/mol of peptide), whereas the first serine of peptide F6 was phosphorylated approximately 2-fold greater than the second serine (67% and 33%, respectively, with a combined total of 1.0 mol of phosphate/mol of peptide). The total number of moles of phosphate/mol of metabolically phosphorylated OPN calculated from the values in Table I (including an unidentified peptide between F5 and F6; see Fig. 4) was 6.1. This occurred despite the fact that there were a total of seven different phosphorylated peptides identified, containing up to 10 phosphorylated residues, as well as an eighth unidentified phosphorylated peptide. These findings indicate that not all the potential phosphorylation sites are phosphorylated on every OPN molecule in a given population. On average, most of the peptides contain approximately 0.76 mol of phosphate/mol peptide, with an average overall extent of phosphorylation per peptide per site of ~53%. The biological significance of the variable extent phosphorylation of the potential phosphorylation sites and its variation with age and rate of synthesis and the possible rate of dephosphorylation are not known at this time, but present intriguing avenues for future research.
|
In vitro phosphorylation of purified OPN obtained from
mineralized chicken bone and recombinant mouse OPN by four pure protein kinases, namely cAMP-dependent kinase, protein kinase C,
cGMP-dependent kinase, and CKII, showed that there was
essentially no phosphorylation of OPN by cAMP-dependent
kinase, minor phosphorylation by protein kinase C and
cGMP-dependent kinase (each introducing 1 mol of phosphate/mol of OPN), whereas CKII significantly phosphorylated OPN
(~9 mol of phosphate/mol of OPN) (41, 42). These data are completely
consistent with the conclusions obtained in the present study on the
extent and sites of phosphorylation in 32P-labeled OPN
synthesized by chicken osteoblasts. Similar in vitro phosphorylation data were obtained for bovine bone OPN (44). It was of
interest that in vitro, CKII phosphorylated all the potential sites (9 mol of phosphate/mol) of chicken OPN in contrast to
the phosphorylation of OPN in osteoblast cell culture, in which case
only partial phosphorylation of the potential sites for phosphorylation was observed (6.1 mol of phosphate/mol of [32P]OPN
(61%)). A possible explanation of the more complete phosphorylation that occurs in vitro is that the overall reaction conditions
are optimized in vitro. For example, the relative
concentrations of the reactants such as the enzyme, ATP, and OPN would
be very different when compared with those in the secretory pathway of
the protein in cell culture and/or the length of the reaction time.
Although there are some conflicting reports with respect to whether phosphorylation of matrix phosphoproteins occur extracellularly or intracellularly (40, 46), recent work carried out in our laboratory with OPN and BSP and a series of purified protein kinases isolated from osteoblasts, as well as the present study, strongly support the conclusions that phosphorylation of OPN and BSP occurs "intracellularly."
Recently, 27 phosphorylated serines and one phosphorylated threonine were identified in bovine milk OPN (58). The predominant sites for phosphorylation contained recognition sequences for casein kinase II. Fig. 6 compares the phosphorylated regions of chicken bone OPN obtained in the present study with those of bovine milk OPN. Although most of the phosphorylated regions in both proteins are highly conserved, bovine milk OPN was more phosphorylated than chicken bone OPN. This is probably related to the fact that chicken OPN is only ~60% homologous with the primary structure of bovine milk OPN. Of the 28 phosphorylated sites in bovine milk OPN, representing 60% of the potential sites for phosphorylation, 12 of the serines with recognition regions SXE/SXSSEE are either absent in chicken bone OPN or the serine residue is substituted by amino acids such as I, P, or A. However, the phosphorylated sites in the chicken bone OPN were similar to those found in bovine milk OPN. Therefore, it is apparent that the lower level of phosphorylation of chicken OPN is due to the absence of the specific amino acid sequence, SXE/SXSSEE, or to the substitution of serine residues by other amino acids. In view of the fact that bovine milk OPN has a very high homology with OPN from other mammalian species, it may be expected that the extent of phosphorylation in mammalian bone OPN may be similar to milk OPN. Nevertheless, although the number of potential sites for phosphorylation in mammalian OPN may be generally the same in each species (regardless of tissue origin), the extent of phosphorylation can vary due to differences in the types and numbers of protein kinases in the various tissues or organs. As a result of different levels of phosphorylation, interactions of the OPN from various tissues with other proteins and conformational changes collectively may reflect different biological functions and open new avenues of research.
As often is the case in complex biological systems, the local environment and type of matrix with which OPN is interacting may play an important role whereby a given state of phosphorylation is optimal for induction or inhibition of calcification. One may therefore intuitively postulate that OPN at a particular state of phosphorylation in different matrix environments can behave and function very differently. The variable extent of phosphorylation of OPN from different tissues or organs and the variable degree of phosphorylation in a given population of OPN molecules, all signify the possibility that an important and a wide range of biological functions of this molecule may be exerted frequently by the capacity of a given cell population to secrete OPN at a particular state of phosphorylation. Additionally, the biological functions of OPN may be further modulated in the extracellular matrix by the varying degrees of dephosphorylation that can take place by the actions of the phosphatases.
We are indebted to the Howard Hughes Medical Institute for the Applied Biosystems model 477A automated sequenator used under the direction of E. Salih.