| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
ARTICLES |
Department of Oral Biology (M.A., J.O.), and Department of Periodontics (J.H., W.X., J.B.), Indiana University School of Dentistry; Department of Medicine (H.L.), and Department of Anatomy (J.B.), Indiana University School of Medicine, Indianapolis, Indiana 46202; and Endocrinology (J.O., J.H.), Lilly Research Laboratories, Indianapolis, Indiana 46285
Address all correspondence and requests for reprints to: Joseph P. Bidwell, Indiana University School of Dentistry, DS235, 1121 West Michigan Street, Indianapolis, Indiana 46202. E-mail: jbidwell{at}iusd.iupui.edu
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
1(I) and one 2(I)
polypeptide chains encoded by two separate genes, COL1A1 and COL1A2
(1). The molecular mechanisms that underlie the regulation of COL1A1
expression in the osteoblast are not fully understood. Recent research
has focused on the identification of cis-regulatory elements of the
COL1A1 promoter responsible for mediating tissue-specific and
hormone-responsive expression (3, 4, 5). Cis-regulatory elements of the rat (COL1A1) promoter required for high levels of expression in bone, tendon, and skin of transgenic mice, as determined by chloramphenicol acetyltransferase (CAT)-expression vectors, are between -2.3 and -1.67 kilobase pairs (kb) but lie within -3.5 and -2.3 kb in cultured bone cells (3, 4). It has been suggested that this change in promoter regulatory site could be a consequence of differences in microarchitecture of cells in vivo and in vitro (3). This hypothesis, linking the regulatory promoter sites to cell structure, presents a unique opportunity to investigate the role of the nuclear matrix, or nucleoskeleton, in osteoblast gene expression.
The nuclear matrix is operationally defined as the proteinaceous
substructure that resists both nuclease digestion and high salt
extraction (6). This unique biochemical fraction comprises numerous
proteins that are both cell- and phenotype-specific (7, 8). Although
most nuclear matrix proteins (NMPs) remain to be identified, recent
data suggest roles in DNA replication (9), RNA processing (10), and
transcription (11). Topoisomerase II- and -ß are NMPs that mediate
changes in DNA topology and are involved in organizing the genes into
loop domains specific to cell and phenotype (12). NuMA is a NMP
required for cell division that organizes the mitotic spindle apparatus
during mitogenesis; however, its role during interphase, when it
exhibits a punctate nuclear distribution, is not known (13)
Recent evidence indicates that NMPs play a role in osteoblast growth, development, and phenotypic expression (14, 15, 16, 17). The NMP composition of osteoblasts derived from fetal rat calvariae was specific to successive stages of differentiation in vitro (14), and specific differences in NMP composition distinguished normal osteoblasts from osteosarcoma cells (15). The nuclear matrix, DNA-binding proteins NMP1 (YY1) and NMP2 (a variant AML/PEBP2/runt domain) associated with sequence specificity to the osteocalcin promoter in proximity to a vitamin D-responsive element (VDRE) and an AP-1 site (16, 17). Finally, PTH-responsive NMPs have been identified in the rat osteosarcoma cell ROS 17/2.8 (18).
The nuclear matrix is typically represented as a nuclear scaffold upon which the DNA is organized (7, 8). This nucleoskeleton is physically linked to the cytoskeleton and the cell periphery through the tissue matrix, a network of interconnecting structural proteins within and between the extracellular and intracellular domains (7, 8). This network includes the extracellular matrix, cell-cell and cell-substrate receptors on the cell surface, microfilaments, microtubules, and intermediate filaments within the cytoplasm and lamins, and the nuclear matrix within the nucleus (7, 8). Therefore, a change in microarchitecture induced by culturing cells on plastic for several passages could result in alterations of the interactions between the nuclear matrix and the promoter of an active gene.
In this study, we identified osteoblast NMPs that associate with sequence specificity to the rat COL1A1 promoter. Gel mobility shift analysis, using end-labeled fragments of the promoter, was used to identify DNA-binding proteins in the nuclear matrix and in the nonmatrix subfractions of rat osteoblasts from the primary spongiosa of the distal femur and from the rat osteosarcoma cells ROS 17/2.8 and UMR 106-POL. A NMP-DNA interaction (NMP3), common to both the primary osteoblasts and osteosarcoma cells, was observed within the region of the promoter that controls in vivo expression. Additionally, a NMP-DNA interaction, (NMP4), was observed in the promoter region that regulates in vitro expression of COL1A1. These data suggest that the link between osteoblast structure and the transcriptional regulation of this extracellular matrix protein may be specific NMPs.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
and ß (a generous gift from J. Holden,
University of Utah, UT). The peroxidase conjugates to antimouse IgG and
antirabbit IgG (Amersham Life Science, Amersham, Buckinghamshire, UK)
were used as the secondary antibodies for chemoluminescent detection
(ECL, Amersham, Buckinghamshire, UK). Reagents for the extraction
buffers were obtained from Fisher Chemical Co. (Pittsburgh, PA) and
Sigma. For the preparation of whole cell lysates, we used the protease
inhibitors aprotinin, leupeptin, pepstatin, phenylmethylsulfonyl
fluoride (Boehringer Mannheim, Indianapolis, IN), and
benzamidine · HCl (Aldrich Chemical Co., Milwaukee, WI).
Cell culture
Primary osteoblasts were derived from the trabecular spongiosa
of the distal femur metaphysis of young, male rats (Charles River
Laboratories, Boston, MA). Muscle and connective tissue were cut away
from femurs of Sprague-Dawley rats (approximately 70–90 g); the
epiphyseal cap was removed to obtain the subjacent 3-mm section of
metaphyseal bone. This section of bone was minced and digested with
trypsin for 1 h at 37 C. The released cells were pelleted and
resuspended into -MEM supplemented with 100 IU/ml penicillin, 100
µg/ml streptomycin, 25 µg/ml amphotericin, 2 mM
L-glutamine (GIBCO-BRL, Grand Island, NY), and 20% FBS
(Sigma). The cells derived from four femurs were seeded into a single
T-150 flask (Corning, Corning, NY). Cells were harvested for protein
analysis 7–8 days post seeding.
The rat osteosarcoma cells, ROS 17/2.8, a generous gift from Drs. Gideon and Sevgi Rodan (Merck Research Laboratories, West Point, PA) and the OK opossum kidney cells, a gift from Dr. James McAteer (Department of Anatomy, Indiana University, Indianapolis, IN) were grown in Ham’s F12 (GIBCO-BRL) supplemented with 2.36 g/liter NaHCO3, 0.118 g/liter CaCl2•2H2O, 6.106 g/liter HEPES, 100 IU/ml penicillin, 100 µg/ml streptomycin, 25 µg/ml amphotericin, 2 mM L-glutamine, and 10% FBS. The rat osteosarcoma, UMR-106 POL (St. Louis, MO), and the NIH3T3 mouse fibroblasts (Eli Lilly Co, Indianapolis, IN) were grown in MEM medium (GIBCO-BRL) supplemented with penicillin, streptomycin, amphotericin, glutamine, and FBS as described above. All cells were maintained in humidified 95% air/5% CO2 at 37 C.
RT-PCR
Primary spongiosa cells were assayed for gene expression at 0,
4, 8, and 24 h and 2, 4, 6, and 8 days post seeding. ROS 17/2.8
cells were assayed at confluence (day 8, post seeding). Primary
spongiosa cells collected from 0–24 h were predominantly nonadherent
but had attached to the culture flask surface by day 2. Our protocol
for RT-PCR analysis of osseous tissue has been described previously
(19) with modifications noted here. Total RNA was extracted from cells
at each time point using RNAzol B (CINNA/BIOTEX Laboratories, Inc.,
Houston, TX) and reverse transcribed using Super Script
Preamplification System for First Strand cDNA Synthesis (GIBCO-BRL).
Aliquots (5 µl) of this cDNA mixture were probed with specific
primers by PCR (Gene Amp PCR System 9600, Perkin Elmer, Norwalk, CT).
Samples were amplified for 30 cycles of 30 sec at 95 C, 30 sec at 55 C,
and 1 min at 72 C with a final 10 min extension at 72 C. To correct for
possible contamination, a sample consisting of PCR reaction cocktail
and water, in place of the RNA samples, was included in each
amplification (labeled as Negative control)(Fig. 2B
). PCR primer sets
were derived from previously published sequences (Table 1). Expected sizes of PCR transcripts were confirmed by
gel electrophoresis and the use of DNA fragment markers (19).
|
|
Northern analysis
Total cellular RNA from experimental cell preparations was
isolated by lysing the cells with RNAzol B and extraction of the lysate
with chloroform. Aliquots of equal amounts of RNA (20 µg) were
denatured and electrophoresed on a 1% agarose/6.6% formaldehyde gel
and transferred onto a nylon membrane (Hybond-N). The RNA was
cross-linked to the blot with UV light and the membrane prehybridized
in Rapid-hyb Buffer (Amersham) for 2 h at 65 C. Hybridizations
were performed using randomly primed
[-32P]deoxy-CTP-labeled cDNA probes (SA > 1
x 109 dpm/µg) of COL1A1. Human GAPDH was used to
normalize the signals from these probes. The membranes were hybridized
with the probes for 2 h at 70 C in the Rapid-hyb Buffer.
Subsequent to hybridization, the blots were washed with 2x SSC/0.1%
SDS at room temperature for 1h, followed by two 15 min washes with
0.2x SSC/0.1% SDS at 65 C with a final wash with 2x SSC at room
temperature for 5 min. Blots were autoradiographed overnight at -80 C
using Kodak (Eastman Kodak, Rochester, NY) X-AR film with intensifying
screens.
Alkaline phosphatase activity and in vitro mineralization
Alkaline phosphatase activity was demonstrated histochemically
in our primary spongiosa cultures using Sigma Diagnostics Alkaline
Phosphatase Kit (Sigma). Mineralized matrix was identified
morphologically as a nodule overlying a cell cluster and confirmed by
staining nodules with Von Kossa stain (27).
Protein extraction
Nuclear, nonmatrix proteins were isolated from the primary
osteoblasts and the ROS 17/2.8 cells using the protocol originally
described by Dignam et al. (28).
NMPs were obtained from a standard sequential extraction protocol (29). Briefly, adherent cells were washed with ice-cold PBS. Soluble and membrane proteins were extracted by adding CSK Buffer (100 mM NaCl, 300 mM sucrose, 10 mM PIPES [pH 6.8], 3 mM MgCl2, 1 mM EGTA, 1.2 mM phenylmethylsulfonyl fluoride (PMSF), 0.5% Triton X-100). This treatment was followed by extraction of the cytoskeletal proteins using RSB-Majik Buffer (10 mM NaCl, 3 mM MgCl2, 10 mM Tris [pH 7.4], 1.2 mM PMSF, 1% Tween-40, 0.5% sodium deoxycholate). Cells were scraped and incubated in Digestion Buffer (50 mM NaCl, 300 mM sucrose, 10 mM PIPES [pH 6.8], 3 mM MgCl2, 1 mM EGTA, 0.5% Triton X-100, 1.2 mM PMSF, 100 µg/ml DNaseI, 50 µg/ml RNAse A) at room temperature for 20 min. Chromatin proteins were extracted by the addition of ammonium sulfate to a concentration of 0.25 M. After centrifugation and removal of the supernatant (chromatin), the nuclear matrix-intermediate filament complex was further fractionated by disassembly (8 M urea, 20 mM MES [pH 6.6], 1 mM EGTA, 0.1 mM MgCl2, 1.2 mM PMSF, and 1% 2-mercaptoethanol) and dialysis overnight at room temperature against Assembly Buffer (150 mM KCl, 25 mM imidiazole-HCl [pH 7.1], 5 mM MgCl2, 0.125 mM EGTA, 2 mM dithiothreitol (DTT), 0.2 mM PMSF). These conditions promote reassembly of the intermediate filaments that were separated from the solubilized NMPs by ultracentrifugation. The NMPs were snap frozen and stored at -80 C. NMPs were obtained from rat kidney and liver using the same sequential extraction protocol with the following modifications. The organs were minced into approximately 2-mm pieces with a clean scapel, rinsed with ice cold PBS, and dounce homogenized in the CSK buffer. The slurry was poured through cheesecloth and processed as described above.
Whole-cell lysates were obtained by washing the adherent cells with PBS followed by scraping with extraction buffer (8 M urea, 2% NP-40, 2% ß-mercaptoethanol, 0.15 µM aprotinin, 1 mM benzamidine·HCl, 5.26 µM leupeptin, 1.52 µM pepstatin, and 1.2 mM PMSF). The lysate was clarified by brief centrifugation, snap frozen in liquid nitrogen, and stored at -80 C.
Protein analysis
Protein extracts were analyzed by gel mobility shift or by
Western blotting (30). Measurements of total protein were made using
the Coomassie Plus Protein Assay Reagent (Pierce, Rockford, Ill). For
immunoblotting, proteins were transferred onto a PVDF membrane
(Millipore Corp., Bedford, MA) by the semidry method (30). For NuMA and
ß-actin, the blot was blocked (5% milk in TBS-T, overnight at room
temperature), incubated with antibody (1:1000 NuMA; 1:100,000
ß-actin, in TBS-T, 1h, at room temperature), followed by incubation
with secondary antibody (1:1000). For topoisomerase II- and ß, the
blots were blocked (5% milk/3% BSA in TBS-T, for 1 h, at 37 C),
incubated with antibody (1:500–1000, in TBS-T/1%BSA, overnight, at 4
C). Immunoblots were developed using chemoluminescence (ECL).
We used fragments of the rat 1(I) collagen promoter (a generous gift
from A. Lichtler, B. Kream, and D. Rowe, The University of Connecticut
Health Center, Framington, CT) as probes in gel mobility shift assays.
These fragments were labeled with [-32P]deoxy-NTP. The
20-µl binding reactions (75 mM KCl, 10% glycerol, 0.15
mM EDTA, 500-2500 ng poly (dI) · poly(dC), 0.1
mM DTT, 17 mM HEPES, 2 µg nuclear protein,
and 0.5 nM of labeled promoter fragment) were incubated for
30 min at room temperature before electrophoretic fractionation of the
mixtures at 4 C in a 5% polyacrylamide gel (80:1 acrylamide:N,
N'methylbisacrylamide) in TGE buffer (30, 31).
For the bidirectional deletion analyses (32), the COL1A1 promoter fragment spanning [-2214 to -1995 nucleotide (nt)] was labeled on either the 3' or 5' end followed by digestion with restriction enzymes to successively shorten the probes from the unlabeled end. These labeled probes then were used in a series of gel shift assays to further localize the NMP-binding site.
Densitometry analysis
Protein-DNA bands detected by 32P-autoradiography,
or protein bands detected by antibody/chemoluminescence, were analyzed
for intensity, area, and molecular weight using the gel documentation
system Foto/analyst II and the software Collage (Fotodyne Inc.,
Hartland, WI). Images of protein and protein-DNA signals were obtained
with a CCD camera and stored as a TIFF file on an Apple MacIntosh
Centris 650. The reported molecular weights of our antigenic signals
are approximations based on migration against calibration standards and
fitting the data to various regression models (linear, quadratic,
cubic, logistic, and cubic spline).
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
A NMP-DNA interaction, NMP3, was detected within the in vivo
regulatory region (-2214 to -1995 nt) in both the ROS 17/2.8
osteosarcoma cells and osteoblasts (Fig. 1). A second
NMP-DNA-binding activity, NMP4, was observed in the in vitro
regulatory region (-3518 to -3406 nt) in the ROS 17/2.8 cells and was
detected inconsistently in the osteoblasts (Fig. 1). NMP4 was comprised
of two prominent doublet protein-DNA bands.
|
). Similarly, RT-PCR analysis indicated that the
osteoblasts and osteosarcoma preparations exhibited comparable levels
of RNA transcripts for alkaline phosphatase, osteopontin, osteocalcin,
histone H4, c-fos, and the PTH receptor at the time of
harvest for NMP3 and NMP4 (Fig. 2B).
The trabecular spongiosa cells rapidly and spontaneously developed the
mature osteoblast phenotype in vitro (Fig. 2B). RNA
transcripts were detected for histone H4, c-fos, COL1A1,
osteopontin, and osteocalcin at all time points from h 0 to day 8 post
seeding (Fig. 2B). At day 4 post seeding, RNA transcripts for PTH
receptor and alkaline phosphatase were detected in the osteoblast
cultures and observed to increase by approximately 60% and 70%,
respectively, by day 8 (Figs. 2B). Consistent with the RT-PCR data,
osteoblast cultures exhibited heavy staining for alkaline phosphatase
protein at day 8 post seeding (Fig. 2C). Mineralized matrix, identified
morphologically as a nodule overlying a cell cluster and positive
staining with Von Kossa, was observed in the osteoblast cultures
between days 6–8 post seeding (Fig. 2D).
Western analyses indicated that the expression of the known NMPs, NuMA,
topoisomerase II-, and topoisomerase II-ß were upregulated
severalfold in the osteosarcoma cells as compared with the osteoblasts
(Fig. 3). However, the expression of ß-actin was
similar in both cell preparations (Fig. 3).
|
and 5). NMP3 binding to the
219-bp-labeled promoter fragment (-2214 to -1995 nt) was attenuated
with the addition of the same unlabeled fragment to the binding
reaction (Fig. 4A). Binding was not decreased by the addition of the
unlabeled fragment containing the binding sequence for NMP4 (-3518 to
-3406 nt) (Fig. 4B). An unlabeled oligonucleotide containing the
consensus sequence for NMP2, a variant AML/PEBP2/runt domain
NMP that binds to the osteocalcin promoter (16), did not compete for
NMP3 binding (Fig. 4C). Similarly, an unlabeled oligonucleotide
containing the consensus sequence for NMP1 (YY1) (17), a
multifunctional transcription factor that may mediate nuclear
matrix-gene interactions, did not attenuate NMP3 binding to the
219-bp-labeled promoter fragment (data not shown).
|
|
). The intensity of the lower
prominent doublet, henceforth designated as NMP4, was decreased by the
unlabeled fragment. The addition of the unlabeled promoter fragment
containing the NMP3-binding site (-2214 to -1995 nt) did not modulate
the intensity of NMP4 activity (Fig. 5B) nor did the addition of the
unlabeled oligonucleotide containing the NMP2 site (Fig. 5C).
Competition with the unlabeled oligonucleotide containing the NMP1
(YY1) site did not attenuate NMP4 binding to the labeled promoter
fragment (-3518 to -3406 nt) (data not shown).
Bidirectional deletion analysis
We performed a bidirectional deletion (stairway) assay to further
localize the NMP3-binding site within the 219-bp (-2214 to -1995 nt)
promoter fragment (Fig. 6A). Using the sequentially
shortened probes in a series of gel shift assays, we determined that
the NMP3-binding activity lay mostly within the 43-bp fragment between
-2149 and -2106 nt (Fig. 6A). Using a labeled oligonucleotide
containing the sequence between -2149 to -2106 nt, we recovered the
NMP3 activity in the ROS 17/2.8 cells (Fig. 6B).
|
). We detected relatively high NMP3- and NMP4-binding
activities in nuclear matrix extracts of ROS 17/2.8 and UMR-106 POL rat
osteosarcoma cells. No NMP3 and NMP4-binding activity was observed in
the nuclear matrix fractions of OK opossum kidney cells or rat kidney.
A weak signal was detected inconsistently in the NMP fractions of the
transformed mouse fibroblasts, NIH3T3, and the nuclear matrix fraction
obtained from rat liver (Fig. 7).
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
The interactions among NMP3, NMP4, and the COL1A1 promoter may optimize the 3-dimensional conformation of the gene for transcriptional regulation. The DNA loop containing the gene is anchored to the nuclear matrix through matrix attachment regions (MARs), which are usually between 200–800 bp in length and AT-rich (35, 36). MARs often act as chromatin boundary elements (36) and typically have a high affinity for topoisomerases (see Ref.38), providing a mechanism for altering DNA loop organization according to cell or phenotype (8, 37). The DNA loop containing the active gene may extend along the nuclear matrix scaffold by multiple cooperative interactions between short stretches of DNA and sequence-specific, DNA-binding NMPs (38). NMP1, NMP2 (16), NMP3, and NMP4 may fulfill this role and serve to modulate local nuclear structure, thereby optimizing the 3-dimensional environment for gene regulation, consistent with their proximity to regulatory elements (39).
The organization of the DNA loop conformation mediated by MARs and
NMPs, described in this work and other studies (16, 31, 40), may
contribute to tissue-specific nuclear architecture. The protein NMP2
was determined to be osteoblast-specific (16), and our own data
indicate that NMP3 and NMP4 are expressed selectively in osteoblast- or
osteoblast-like cells. The absence of these proteins in the kidney,
another major target of calciotropic hormones, is particularly
intriguing. All of these proteins were found to be comparatively
overexpressed in osteosarcoma cells and may indicate a general
upregulation of NMP expression in tumors because the ROS 17/2.8 cells
expressed higher levels of the NMPs NuMA, topoisomerase II-, and
ß. Nevertheless, in situ analysis of osseous and
nonosteoid tissue for NMP3 and NMP4 activity is necessary for a
definitive evaluation on the relative expression of these proteins in
diploid osteoblasts.
Our hypothesis that NMPs optimize local nuclear architecture for transcriptional regulation is consistent with recent data demonstrating the influence of cell structure on gene expression (7, 8, 40). The regulation of tissue-specific gene expression in situ requires the cooperative interactions between soluble factors, e.g. growth factors, hormones, and transcription factors, and alterations in cell structure, often induced by the extracellular matrix (7, 8, 40). Several studies have demonstrated that the extracellular matrix can alter cell morphology and function and that cells cultured on plastic often fail to acquire the 3-dimensional structure required for full phenotypic expression (41, 42, 43, 44).
Because the tissue matrix physically links the genes with the cell periphery, involving both nuclear matrix and cytoskeletal proteins (7, 8, 40) alterations in microarchitecture that affect gene expression, may involve changes in both transcriptional and posttranscriptional domains (45, 46). In a recent study, the control of integrin expression by the extracellular matrix was demonstrated to be exerted at both transcriptional and posttranscriptional levels (46). In K1735-M2 melanoma cells, transcription of the ß-1 integrin gene was influenced by the substratum, although the levels of integrin protein remained similar (46). In mammary epithelial cells, the rates of ß-1 integrin gene transcription were similar, but mRNA and protein levels were higher in cells cultured on plastic than those on basement membrane (46).
Both the in vivo and in vitro domains of the
COL1A1 gene, as defined by CAT-expression vectors, may contribute to
the transcriptional regulation of COL1A1 in situ,
i.e. in the context of the 3-dimensional conformation of the
native gene. Although the use of CAT-expression vectors is appropriate
for localizing promoter regulatory elements, this technique may
neutralize the contribution of local nuclear structure to gene
expression. The dichotomy in the regulation of COL1A1-CAT expression
between cells in vivo and in vitro (3) may have
resulted from the response of an artificial gene construct to changes
in the organization of the tissue matrix. Recently, the use of a
2954-bp MAR from the chicken lysozyme locus was used in a plasmid
construct of the mouse pro-1(I) collagen promoter to increase the
ratio of transgenic mice expressing the reporter gene (47). This
suggests that structural contributions to gene expression become
evident only in constructs that closely approximate the native
conformation of the gene. Similarly, the potential importance of NMP3
and NMP4 may be lost in the restricted context of the CAT-expression
vector.
Nuclear matrix DNA-binding proteins may link cell structure and gene expression (48). The study of the specific function of these proteins will require unique experimental approaches. The use of substrates other than plastic for in vitro cell culture has revealed the importance of the extracellular matrix in gene expression. Similarly, the use of more sophisticated gene constructs and antisense RNA techniques will be required to elucidate the role of nuclear structure in transcriptional regulation.
|
Footnotes |
|---|
Received May 23, 1996.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
1(I)
collagen promoter in osseous and fibroblastic cells. J Cell Biol 116:227–2361(I)
collagen promoter activity in cultured calvariae from neonatal
transgenic mice. Mol Endocrinol 7:399–408[Abstract]
1(I) collagen
promoter. J Biol Chem 264:3072–30771(I) collagen promoter via a transforming growth
factor-ß activation element. J Biol Chem 268:13625–136311(I) collagen mRNA stability and
transcription in mouse fibroblasts. J Biol Chem 266:8470–84751(I) collagen promoter that confers high-level osteoblast
expression in transgenic mice and that binds a protein selectively
present in osteoblasts. Proc Natl Acad Sci USA 93:1027–1031This article has been cited by other articles:
 |
|
 |
|
  K. Cartharius, K. Frech, K. Grote, B. Klocke, M. Haltmeier, A. Klingenhoff, M. Frisch, M. Bayerlein, and T. Werner MatInspector and beyond: promoter analysis based on transcription factor binding sites Bioinformatics, July 1, 2005; 21(13): 2933 - 2942. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Shah, M. Alvarez, D. R. Jones, K. Torrungruang, A. J. Watt, N. Selvamurugan, N. C. Partridge, C. O. Quinn, F. M. Pavalko, S. J. Rhodes, et al. Nmp4/CIZ regulation of matrix metalloproteinase 13 (MMP-13) response to parathyroid hormone in osteoblasts Am J Physiol Endocrinol Metab, August 1, 2004; 287(2): E289 - E296. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. Barseguian, B. Lutterbach, S. W. Hiebert, J. Nickerson, J. B. Lian, J. L. Stein, A. J. van Wijnen, and G. S. Stein Multiple subnuclear targeting signals of the leukemia-related AML1/ETO and ETO repressor proteins PNAS, November 26, 2002; 99(24): 15434 - 15439. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. Torrungruang, M. Alvarez, R. Shah, J. E. Onyia, S. J. Rhodes, and J. P. Bidwell DNA Binding and Gene Activation Properties of the Nmp4 Nuclear Matrix Transcription Factors J. Biol. Chem., May 3, 2002; 277(18): 16153 - 16159. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y. L. Ma, R. L. Cain, D. L. Halladay, X. Yang, Q. Zeng, R. R. Miles, S. Chandrasekhar, T. J. Martin, and J. E. Onyia Catabolic Effects of Continuous Human PTH (1-38) in Vivo Is Associated with Sustained Stimulation of RANKL and Inhibition of Osteoprotegerin and Gene-Associated Bone Formation Endocrinology, September 1, 2001; 142(9): 4047 - 4054. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. R. Miles, J. P. Sluka, D. L. Halladay, R. F. Santerre, L. V. Hale, L. Bloem, K. Thirunavukkarasu, R. J. S. Galvin, J. M. Hock, and J. E. Onyia ADAMTS-1: A Cellular Disintegrin and Metalloprotease with Thrombospondin Motifs Is a Target for Parathyroid Hormone in Bone Endocrinology, December 1, 2000; 141(12): 4533 - 4542. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. R. Miles, J. P. Sluka, R. F. Santerre, L. V. Hale, L. Bloem, G. Boguslawski, K. Thirunavukkarasu, J. M. Hock, and J. E. Onyia Dynamic Regulation of RGS2 in Bone: Potential New Insights into Parathyroid Hormone Signaling Mechanisms Endocrinology, January 1, 2000; 141(1): 28 - 36. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 G. S. STEIN, A. J. VAN WIJNEN, J. L. STEIN, J. B. LIAN, S. H. POCKWINSE, and S. MCNEIL Implications for interrelationships between nuclear architecture and control of gene expression under microgravity conditions FASEB J, May 1, 1999; 13(9001): 157 - 166. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. Zeng, S. McNeil, S. Pockwinse, J. Nickerson, L. Shopland, J. B. Lawrence, S. Penman, S. Hiebert, J. B. Lian, A. J. van Wijnen, et al. Intranuclear targeting of AML/CBFalpha regulatory factors to nuclear matrix-associated transcriptional domains PNAS, February 17, 1998; 95(4): 1585 - 1589. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. H. Thomas, J. H. Collier, C. S. Sfeir, and K. E. Healy Engineering gene expression and protein synthesis by modulation of nuclear shape PNAS, February 19, 2002; 99(4): 1972 - 1977. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||
