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Dynamic Regulation of RGS2 in Bone: Potential New Insights into Parathyroid Hormone Signaling Mechanisms

Endocrine Division (R.R.M., J.P.S., R.F.S., L.V.H., G.B., K.T., J.M.H., J.E.O.) and Cardiovascular Division (L.B.), Lilly Research Labs, Indianapolis, Indiana 46285

Address all correspondence and requests for reprints to: Dr. J. E. Onyia, Bone Metabolism Research Group, 0403, Endocrine Division, Lilly Research Laboratories, Indianapolis, Indiana 46285. E-mail: JEO{at}lilly.com

	Abstract

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The initial steps involved in mediating the transduction of PTH signal via its G protein-coupled receptors are well understood and occur through the activation of cAMP and phospholipase C pathways. However, the cellular and molecular mechanisms for subsequent receptor desensitization are less well understood. Recently, a new family of GTPase activating proteins known as regulators of G protein signaling (RGS), has been implicated in desensitization of several G protein-coupled ligand-induced processes. At present, it is not known whether any of the RGS proteins play a role in PTH signaling. Using the differential display method, we screened for genes that are selectively expressed after a single sc injection of human PTH (1–38) (8 µg/100 g) in osteoblast-enriched femoral metaphyseal spongiosa of young male rats (3–4 weeks old). We found and cloned one full-length complementary DNA that encodes a 211-amino acid RGS protein and shares 97% sequence identity with mouse and human RGS2. Based on sequence similarity, we have designated this clone as rat RGS2. Northern blot analysis confirmed that the expression of RGS2 messenger RNA (mRNA) is rapidly and transiently increased by human PTH (1–38) in both metaphyseal (4-to 5-fold) and diaphyseal (2- to 3-fold) bone, as well as in cultured osteoblast cultures (2- to 37-fold). In vitro, forskolin and dibutyryl cAMP similarly elevated RGS2 mRNA. In vivo, PTH analog (1–31) [which stimulates intracellular cAMP accumulation, PTHrP (1–34), and prostaglandin E₂] induced RGS2 mRNA expression; whereas PTH analogs (3–34) and (7–34), which do not stimulate cAMP production, had no effect on expression. In tissue distribution analysis, RGS2 is widely expressed and was detected in all tissues examined (heart, spleen, liver, skeletal muscle, kidney, and testis), with significant expression in two nonclassical PTH-sensitive tissues: the brain, and the heart. After PTH injection, RGS2 mRNA expression was induced in rat bone but not in any of the other tissues examined. These findings demonstrate that RGS2 is regulated by PTH, prostaglandin E₂, and PTHrP and that regulation by PTH in bone occurs via the cAMP pathway. Additionally, these results suggest the exciting possibility that increased RGS2 expression in osteoblasts may be one of the early events influencing PTH signaling.

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
PTH CONTROLS the differentiation and function of target cells through its specific G protein-coupled receptor (PTH1R) in two principal target organs, kidney and bone (1, 2). Recently, an additional receptor (PTH2R) of unknown function has been described in the brain and other nonclassical PTH target tissues (3, 4). PTH occupation of the receptors leads to the activation of adenylate cyclase and phospholipase C pathways that results in the accumulation of multiple signal transducers including cAMP, inositol triphosphate, a transient increase in the intracellular calcium, and activation of both protein kinase A (PKA) and protein kinase C (PKC) (5, 6, 7, 8, 9, 10, 11, 12). Although these initial steps involved in mediating the transduction of PTH signal via its receptors are well understood, the cellular and molecular mechanisms that control the signal intensity, duration, and subsequent receptor desensitization are less well understood.

As has been reported for most G protein-coupled receptors, PTH binding to the receptor results in activation of G proteins by stimulating exchange of GDP for GTP on the -subunit. After the exchange, GTP-bound G dissociates from Gß, both of which function as signal transducing molecules by regulating the activities of various downstream cellular effector systems (13, 14). Termination of G protein signaling occurs by intrinsic GTPase activity of the -subunit and subsequent reassembly with ß-subunit to form the inactive -GDPß (14). Previous studies have established that, as with other G protein-coupled receptors, the intensity and duration of response is, in part, regulated at the level of the receptor by mechanisms that include agonist-dependent phosphorylation of the receptor, subsequent receptor inactivation, uncoupling from interaction with their transducing G protein, and turnover (15, 16).

Recently, another mechanism of controlling dynamic G protein signaling kinetics has been discovered, and results from the interaction of the G protein -subunit with members of a family of RGS proteins (regulators of G protein signaling) (reviewed in Refs. 17, 18, 19). RGS proteins are GTPase-activating proteins (GAPs) which function to accelerate the rate of intrinsic GTP hydrolysis by G and thereby limit the duration of G protein activation. RGS proteins were first identified as negative regulators of G protein signaling in Sacchromyces cerevisiae (Sst2p) (reviewed in Refs. 20, 21) and Caenorhabditis elegans (EGL-10) (22). To date, about 18 members of this family of proteins have been described in mammalian tissues and contain a conserved diagnostic RGS domain consisting of approximately 120 amino acids. In vitro biochemical assays indicate that most of the RGS proteins tested bind and/or stimulate the GTPase activity of G_i subfamily (17, 23, 24, 25). Some of these RGS proteins also act as GAPs toward members of the G_q subfamily (17, 24, 25, 26). GAPs for G_s and G12 subfamily members have not been detected, although binding to G_s, as well as inhibitory effects on G_s- and G12-mediated signaling pathways, has been described (27, 28, 29). Several studies have provided the much needed evidence for the regulatory effects of RGS proteins on G_i, G_q, G_s, and G12 mediated signaling in intact cells (24, 25, 27, 28, 29, 30, 31, 32). Presently, information on how RGS proteins are regulated in mammalian cells is limited. Regulation of transcription of members of the RGS family of proteins has been noted in response to p53 (33), or polyclonal activation of T and B cells (23, 34, 35, 36), forskolin (37), amphetamine response in brain (38), and neuronal activation by stimuli that evoke plasticity, such as electroconvulsive seizure and haloperidol (25). Surprisingly, there have been only two examples of altered transcription of a mammalian RGS gene in response to activation of a G protein-coupled receptor (29, 39).

In the present study, we report that a member of the RGS gene family, RGS2, is rapidly and selectively up-regulated in bone (in vivo and in vitro) in response to PTH [as well as PTH related peptide (PTHrP) and PG E₂]. RGS2 was identified as an early-response gene regulated by PTH in a differential display PCR analysis of messenger RNA involved in PTH actions in rat bone in vivo. This dynamic regulation of RGS2 suggests a potential novel mechanism in G protein signaling and response in bone.

	Materials and Methods

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Materials
PTH. Synthetic human PTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38), PTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31), and PTHrP (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34) and bovine PTH (3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34) and (7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34) (Bachem California, Inc., Torrance, CA) were prepared in a vehicle of acidified saline containing 2% heat-inactivated rat serum. Prostaglandin E₂ (PGE₂) (Sigma, St. Louis, MO) was first dissolved in 100% ethanol and further diluted in vehicle to a final ethanol concentration of 10%. Forskolin and dibutyryl cAMP were purchased from Sigma and were solubilized in dimethylsulfoxide.

Animals. Young virus-antibody-free, male Sprague Dawley rats, 60–75 g, (Harlan Laboratories, Indianapolis, IN) were housed with a 12-h light, 12-h dark cycle. Animals were fed Purina chow (calcium 1%, phosphate 0.61%; PMI Feeds, Inc., St. Louis, MO) and water ad libitum. Animal protocols were approved by the Lilly Animal Care and Use Committee.

In vivo protocols. Rats were weighed and sorted into groups of comparable mean body weight (four rats per group). The rats were injected (sc) with either the various analogs of PTH (80 µg/kg), PTHrP (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34) (80 µg/kg), or PGE₂ (6 mg/kg) and were killed using CO₂ at indicated time points. Control rats received an equal amount of the vehicle sc and were killed at the same time interval. The doses of PTH and PGE₂ were chosen from our previous work (40, 41) and reports from other laboratories (42, 43) demonstrating an effect in bone and gene expression. After death, rat femora were resected; and all connective tissue, including periosteum, was completely removed. The distal epiphysis, including the growth plate, was removed; and a subjacent 3-mm-wide band of the metaphyseal trabecular primary spongiosa, or diaphyseal middle third of the same femur, were resected and frozen in liquid nitrogen until messenger RNA (mRNA) analyses were performed (40, 41). For experiments involving PGE₂, the distal metaphysis (6 mm subjacent to the growth plate) were used for mRNA analysis.

Cell culture. Primary osteoblast cultures were derived from the rat femur metaphysis and diaphysis, as previously described (44, 45, 46). ROS 17/2.8 cells were maintained in growth medium: F-12 nutrient mixture (Life Technologies, Inc., Gaithersburg, MD) containing 10% FBS (HyClone Laboratories, Inc. Logan, UT) plus 2 mM glutamine (Life Technologies, Inc.). All cultures were maintained in a humidified 5% CO₂ atmosphere at 37 C_. For mRNA analysis, cultures (4 T150 flasks/group) of cells were grown (as described above) to 80–90% confluence and then switched into medium containing 0.1% FBS overnight. The cells were then treated with human PTH (hPTH) (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38) at a final concentration of 5 x 10^-8 M for 0, 1, 6, or 24 h. In additional experiments, ROS 17/2.8 cells were treated with indicated concentrations of either hPTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38), forskolin or dibutyryl cAMP for 1 h.

RNA isolation and complementary DNA (cDNA) synthesis. Total RNA from three independent experiments was used in the cDNA synthesis and differential display to ensure reproducibility and to reduce the false positives. For each experiment, RNA was extracted from the metaphyseal primary spongiosa of vehicle or PTH-treated rats at 1 and 24 h, as previously described (40, 41). With each experiment, samples were pooled into treated or control groups (four animals per group) for each indicated time point after treatment. Samples were removed from the animals, snap frozen, and pooled for isolation of RNA. Total RNA was extracted by homogenization in Ultraspec-II (BIOTECX, Houston, TX) using an LS 10–35 Polytron homogenizer (Brinkmann Instruments, Inc., Westbury, NY), as recommended by the manufacturer. Isolated RNA was quantitated, using spectrophotometry, by measuring the absorbance at 260 nm with the 260/280 nm ratio calculated to ensure the absence of protein contamination. To remove contaminating DNA from the RNA preparation, samples were incubated with ribonulclease-free deoxyribonuclease I (Roche Molecular Biochemicals, Indianapolis, IN) for 15 min at room temperature and then extracted with phenol/chloroform. First-strand cDNA was synthesized from 4 µg of total RNA, by oligo dT priming, using the Superscript Preamplification kit (Life Technologies), in a final volume of 40 µl.

PCR and differential display. To amplify differentially expressed bands from cDNA, arbitrary primer sets were chosen, and differential display was carried out as previously described (47, 40). The upstream (arbitrary primer) and downstream (anchored) primers that detected RGS2 were 5' TGA GCG GAC A 3' and 5' TTT TTT TTT TTT C 3'. Using cDNA diluted 1:25 or a no-cDNA template control (negative control), duplicate PCR reactions were assembled robotically (Tecan Genesis, Reading, UK) to a final concentration of 10 mM Tris-HCl (pH 8.3), 1.5 mM MgCl₂, 50 mM KCl, 2.0 mM deoxynucleotide triphosphates, 15 nM [³³P]-dATP (Amersham Pharmacia Biotech, Arlington Heights, IL), and 1 U AmpliTaq polymerase (Perkin-Elmer Corp., Foster City, CA) in a final vol of 20 µl. Reactions were then subjected to the following PCR conditions on a DNA Engine PTC-225 thermacycler (MJ Research, Inc., Watertown, MA): 1 cycle of 92 C for 2 min; 40 cycles of 92 C for 15 sec, 40 C for 2 min, 72 C for 1 min; and 1 cycle of 72 C for 5 min. Subsequently, PCR products were separated on a 6% TBE/urea sequencing gel (Sequagel, National Diagnostics, Atlanta, GA) for 3 h at 1700 V. Gels were dried and exposed to BIOMAX x-ray film (Eastman Kodak Co., Rochester, NY). The negative controls with no-cDNA template yielded no-PCR products.

Reamplification, cloning, and sequencing of cDNA. To characterize differential display products, bands of interest representing differentially expressed genes were excised from the gel, boiled for 5 min in H₂O, and purified over a Centricon 50 column (Amicon, Beverly, MA). Samples were then reamplified to confirm the size and specificity of the primer sets used in the display. Reamplified products were ligated into pCR2.1 TA cloning vector (Invitrogen, San Diego, CA) and transformed into DH10B cells (Life Technologies, Inc.). For each clone, 10 colonies were picked, amplified in LB broth, and the plasmids were isolated (Wizard Plus, Promega Corp., Madison, WI). Clones which contained inserts were submitted for automated cycle sequencing (Lilly DNA Technology Group, Indianapolis, IN). All sequences were analyzed, using BLAST and FASTA against GenBank and EMBL databases, to determine sequence identity and tissue distribution.

Generation of radiolabeled probes for Northern analysis. To generate radioactive probes for Northern analysis, the inserts containing RGS2 cDNA were released from the plasmid by restriction digest. Rat glyceraldehyde 3-phosphate dehydrogenase (GAPDH) cDNA probes were cloned using PCR with specific primer pairs as published previously (41, 44). Twenty-five nanograms of cDNA were labeled by the random primer method (Life Technologies/BRL) using -[³²P]-deoxycycidine triphosphate (Amersham Pharmacia Biotech). Free nucleotides were removed by centrifugation through a Centricon-50 column (Amicon).

Isolation of Poly A + RNA and Northern blotting. RGS2 mRNA expression was analyzed by Northern blot. Bone, various tissues, and cell culture samples were pooled into treated or control groups for each indicated time point after treatment. Total RNA was extracted from bone and various tissues by homogenization in Ultraspec-II (BIOTECX) using an LS 10–35 Polytron homogenizer (Brinkmann Instruments, Inc.) as recommended by the manufacturer. Total RNA was extracted from the osteoblast cultures by adding Ultraspec-II directly to the culture flasks. The resulting cell lysates were passed several times through a 10-mL pipette before collection. Poly A + RNA was isolated from total RNA using Oligotex (Qiagen, Santa Clarita, CA), according to the manufacturers protocol, and was quantitated by spectrophotometry. The absorbance at 260 nm was determined, and the 260/280 nm absorbance ratio was calculated to ensure the absence of protein contamination. Samples of poly A + RNA (2 µg) were denatured in 0.04 M 3-(N-morpholine) propanesulfonic acid (pH 7.0) (MOPS), 10 mM sodium acetate, 1 mM EDTA, 2.2 M formaldehyde, and 50% formamide at 60 C for 10 min and was size fractionated by electrophoresis through 1% agarose gels in 2.1 M formaldehyde and 1 x MOPS and transferred to nylon membranes (Brightstar-Plus; Ambion, Inc., Austin, TX). The membranes were air dried, and the RNA samples were cross-linked to the nylon membrane by UV irradiation in a Stratalinker (Stratagene, La Jolla, CA). Migration of 28 S and 18 S ribosomal RNA was determined by ethidium bromide staining. DNA probes were labeled by the random primer method (Life Technologies, Inc.) using -[³²P]-deoxycycidine triphosphate. Prehybridization and hybridization were carried out at 48 C in NorthernMax buffers (Ambion, Inc.). After hybridization, membranes were washed for 30 min at room temperature in buffer containing 2 x SSC and 0.1% SDS, then 30 min at 48 C in 0.2 x SSC, and exposed to Biomax MS x-ray film (Eastman Kodak Co.) at -70 C. Autoradiograms were quantitated by scanning laser densitometry (2400 Gel Scan XL, LKB, Piscataway, NJ). Labeled bands were quantitated as densitometric units and normalized to that of the GAPDH signals to correct for variations in RNA transfer and gel loading. The data were expressed as fold change vs. untreated control samples. The experiments were repeated 2–4 times for each time point to confirm findings.

Multitissue RNA analysis. To determine the distribution of the RGS2 transcript, we probed poly A + RNA from rat tissues using multiple-tissue Northern blots (CLONTECH Laboratories, Inc., Palo Alto, CA). The multiple-tissue Northern blot contained 2 µg/lane of poly A + RNA from heart, kidney, spleen, lung, liver, skeletal muscle, and testis. The specificity of PTH effect on gene expression was examined in tissue blots prepared from rats that were treated with vehicle or PTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38), as described above, for 1 h. The blots were analyzed by hybridization with radiolabeled probes, as described for Northern blot analysis.

	Results

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Identification of RGS2 as a PTH-regulated gene in rat metaphyseal bone
Using differential display PCR (DDPCR), we screened for genes that are differentially regulated by PTH in rat metaphyseal bone. cDNA derived from RNA isolated from hPTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38) treated and control femoral metaphyses, at 1 and 24 h after treatment, was differentially displayed using primer sets, as described in Materials and Methods. To ensure reproducibility and to reduce the detection of false positives, as has been reported previously (48, 49, 50, 51, 52, 53), the DDPCR was conducted on cDNA derived from 3 independent (RNA preparations) experiments. As a negative control, the DDPCR was also conducted omitting the cDNA template (no cDNA control), which, as expected, yielded no bands (Fig. 1A). Parallel display of duplicate samples from control and treated bones showed a 1.2-kb band that was rapidly up-regulated in 1 h, but returned to control levels by 24 h (Fig. 1A). This band was excised from the gel and reamplified by PCR. The PCR product was then cloned and sequenced. Sequence analysis revealed that cDNA from this band encodes a single open reading frame of 211 amino acids and shares 97% sequence identity with both mouse and human RGS2 proteins (Fig. 1B). Based on sequence similarity, we have designated this clone as the rat homolog of RGS2.

View larger version (59K): [in this window] [in a new window]   Figure 1. Identification of RGS2 as a PTH-regulated gene in rat metaphyseal bone. A, DDRT-PCR products amplified from cDNA derived from vehicle and hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 )-treated femur bones (pooled, n = 4/group) were resolved on a 6% TBE/urea sequencing gel. To eliminate false positives, cDNA derived from three independent (RNA preparations) experiments were analyzed simultaneously. A negative control omitting the cDNA template (no cDNA control) was also analyzed. Samples were run in duplicate for each time point examined. A band representing the candidate PTH-regulated gene is indicated by the arrow. This band was excised from the gel, reamplified by PCR, and cloned for sequence analysis. B, Sequence alignment of the candidate PTH-regulated gene (rat RGS2), compared with mouse and human RGS2. The enclosed table shows percent sequence identity, and mismatches are indicated by shading.

View larger version (34K): [in this window] [in a new window]   Figure 2. The effect of hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ) on RGS2 mRNA expression in rat femur metaphysis and diaphysis. A, Representative autoradiograph showing the time course of hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ) treatment on RGS2 mRNA expression in rat femur metaphysis. V represents mRNA expression in vehicle-treated control animals. B, Quantitation of RGS2 mRNA bands by densitometric scanning. The figure represents two to three independent experiments normalized to GAPDH, and the values are shown as fold induction over vehicle-treated control (depicted as 0 h). C, Comparison of basal and PTH effect on RGS2 in metaphyseal and diaphyseal bone. RNA was isolated from the femur metaphyseal and diaphyseal bone of young male rats (pooled, n = 4/group) at indicated times after a single PTH injection (80 µg/kg, sc). Two micrograms of poly A + RNA were loaded per lane and analyzed for RGS2 expression by Northern blot hybridization. GAPDH was rehybridized as a control for RNA integrity and quantification.

View larger version (54K): [in this window] [in a new window]   Figure 3. The effect of hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ), forskolin, and dibutyryl cAMP on RGS2 mRNA in osteoblast-like cells in vitro. A, Time course of hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ) on RSG2 mRNA expression in primary osteoblast cultures derived from the metaphysis (metaphyseal osteoblasts) and diaphysis (diaphyseal osteoblasts), as well as in the ROS 17/2.8 osteosarcoma cell line. Cells were treated with hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ) (5 x 10-8 M) for 0, 1, 6, or 24 h. B, Dose-dependent effect of hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ) on RGS2 mRNA was compared with the effect of forskolin and dibutyryl cAMP in ROS17/2.8 osteosarcoma cells. Cells were treated with indicated concentrations of either hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ), forskolin (FSK), or dibutyryl cAMP (db cAMP) for 1 h. RGS2 mRNA levels were determined by Northern analysis (2 µg/lane poly A + RNA). GAPDH was rehybridized as a control for RNA integrity and quantification. RGS2 mRNA values normalized to GAPDH signals are expressed as fold induction over vehicle-treated control (which is set as 1).

View larger version (44K): [in this window] [in a new window]   Figure 4. Effects of PTH analogs, PTHrP, and PGE2 on RGS2 mRNA expression in rat femur metaphysis. Poly A + RNA, isolated from the distal femur metaphysis of rats (pooled, n = 4/group) 1 h after injection (sc) with (A) PTH analogs (80 µg/kg), PTHrP (80 µg/kg), or vehicle equivalent, were analyzed for RGS2 expression by Northern blot hybridization. B, RNA isolated 1, 6, or 24 h after injection with PGE2 (6 mg/kg) or vehicle equivalent was also analyzed for RGS2 mRNA expression. Two micrograms per lane of poly A + RNA were used for Northern blot hybridization. GAPDH was rehybridized as a control for RNA integrity and quantification. RGS2 mRNA values normalized to GAPDH signals are expressed as fold induction over vehicle-treated control (which is set as 1).

Tissue expression and regulation of RGS2 mRNA
Having demonstrated RGS2 expression in unstimulated and stimulated bone, we next sought to determine whether RGS2 was detectable in unstimulated and PTH-stimulated nonosseous tissues. Tissue profiling by Northern blot analysis of poly A + RNA showed that RGS2 is widely expressed and was detected in all tissues examined (heart, spleen, liver, skeletal muscle, kidney, and testis), with significant expression in two nonclassical PTH-sensitive tissues: the brain, and heart (Fig. 5; data not shown). To efficiently examine PTH effects on RGS2 expression in nonosseous tissues, we limited our analysis to selected PTH1R-positive tissues; namely, brain, heart, kidney, liver, and spleen (61, 62). One hour after injection with vehicle or hPTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38) (80 µg/kg), poly A + RNA from these tissues, including metaphyseal and diaphyseal bone, was analyzed for PTH1R expression and RGS2 expression. As shown in Fig. 5, PTH1R was detected in bone, brain, heart, kidney, liver, and spleen. As expected, PTH treatment rapidly down-regulated PTH1R expression in the two classical target organs of PTH action, kidney and bone. However, no significant changes in RGS2 mRNA levels were observed in any of the nonosseous tissues examined.

View larger version (61K): [in this window] [in a new window]   Figure 5. Tissue expression and regulation of RGS2 mRNA. Expression of RGS2 mRNA was examined in poly A + RNA from vehicle or hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 )-treated nonosseous tissues as indicated. The effect of 1 h treatment of hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ) on RGS mRNA expression was examined only in selected PTH1R positive tissues. Vehicle treatment is indicated by the minus sign, and hPTH (1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 ) treatment is indicated by the plus sign. Expression of PTH1R was confirmed using cDNA probe for rat PTH1R. GAPDH was rehybridized as a control for RNA integrity and quantification. RGS2 and PTH1R mRNA values normalized to GAPDH signals are expressed as fold induction over vehicle-treated control (which is set as 1).

	Discussion

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Our results show that the rapid induction of RGS2 by PTH was specific to bone and not observed in other PTH receptor-positive tissues examined (brain, heart, kidney, liver, and spleen). Interestingly, in the two classical target organs of PTH action, bone and kidney, down-regulation of PTH1R mRNA, reminiscent of signal desensitization, was evident. However, in kidney, this event was not accompanied by changes in RGS2 mRNA. This selective difference in response to PTH suggests the idea that other interacting proteins may be required for different tissue responses. In all tissues examined, constitutive expression of RGS2 mRNA was detected in the uninduced or control state. This is consistent with a fundamental role of RGS2 in tightly modulating G protein signaling. The differences observed in basal RGS2 mRNA expression in the various tissues may reflect the functional output needed by the tissue to regulate the biochemical activity of G proteins. In bone, the higher level of RGS2 mRNA expression detected in the metaphysis vs. diaphysis, after PTH treatment, is consistent with the presence of more PTH-responding cells (osteoblasts) or greater responsiveness per cell in the metaphysis than in the diaphysis. This differential responsiveness in the two regions of bone is similar to results we obtained with several other PTH-responsive genes, including c-fos, c-jun, and interleukin 6 (unpublished results, J. E. Onyia). These data suggest a specific role for RGS2 protein in negatively regulating basal and PTH-regulated signaling.

Because it has been shown that PTH activates multiple signal transduction pathways that lead to generation of second messengers, including cAMP, inositol triphosphate, and intracellular calcium (5, 6, 7, 8, 9, 10, 11, 12), we evaluated which signal transduction pathways were involved in stimulation of RGS2 expression by PTH. Our studies suggest that the increase in RGS2 expression by PTH is mediated primarily by the cAMP/PKA pathway. This conclusion is based on the following in vivo and in vitro findings: 1) both forskolin, a direct activator of adenylate cyclase that enhances cAMP accumulation, and dibutyryl cAMP, a cell permeable analog of cAMP, stimulated RGS2 mRNA expression in vitro; 2) only agents that activate the cAMP/PKA pathway [PTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38), (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31), PTHrP, and PGE₂] were able to increase RGS2 expression in vivo. Unlike PTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34, 35, 36, 37, 38), which has a full spectrum of activity, PTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31) activates only cAMP/PKA, with no demonstrable effects on PKC or PLC (58, 59). This specificity of PTH (1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31) to the PKA pathway is substantiated in target cells of bone expressing endogenous PTH1R but not in transfected cells overexpressing PTH1R (58, 63). In contrast, N-terminally truncated PTH analogs (3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34) and (7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34) were ineffective in increasing RGS2 expression. These analogs activate PKC but not adenylate cyclase (55, 56, 57). However, it is important to note that although these results demonstrate that PTH responses depend on cAMP, they do not exclude a role for the PLC/PKC pathway (as suggested by the lack of response to PTH analogs 3–34 and 7–34). To fully explore the role of the PLC/PKC pathway in basal and PTH control of RGS2 expression, extensive studies employing agents that induce or block PKC activation, intracellular calcium release, and uptake of extracellular calcium will be needed.

A role for intracellular cAMP in regulating RGS2 has been previously suggested. Recently, Pepperl et al. (37) demonstrated that RGS2, but not RGS4 or 7 mRNA, was strongly induced by forskolin (adenylate cyclase activator) in pheochromocytoma (PC12) cells. Tseng et al. (29) showed that glucose-dependent insulinotropic peptide (GIP), a potent stimulator of intracellular cAMP levels, induced a small, but significant, increase in RGS2 mRNA in GIP-treated PTC3 cells at 1 h. Similarly, amphetamine induction of RGS2 within the striatum of the brain has been speculated to involve dopamine D1 receptor stimulation and to occur via elevation of cAMP levels (38). Taken together, these findings suggest that RGS2 mRNA may be induced by changes in intracellular cAMP levels, and we speculate that agonist-stimulated cAMP production induces RGS2 expression, resulting in feedback desensitization of the stimulating receptor. However, it is important to note that agents or signaling pathways other than cAMP have also been shown to regulate RGS2. Regulation of RGS2 transcript has been demonstrated in response to T cell lectin concanavalin A, cycloheximide, calcium ionophore (ionomycin), and neuronal activation by stimuli that evoke plasticity, such as electroconvulsive seizure and haloperidol (23, 25). This dynamic transcriptional control of RGS2 by multiple signals suggests an important role in modulating cellular signaling.

The role of RGS2 in mediating cellular signaling is presently unclear. Although our data suggest a primary interaction of RGS2 with the cAMP/PKA pathway, no RGS protein to date has shown detectable GAP activity toward G_s (17). The closest demonstration of an interaction between RGS2 and G_s comes from a recent study by Tseng et al. (29), showing RGS2-bound G_s protein in an in vitro system. Additionally, ectopic overexpression of RGS2 was able to inhibit, by 50%, a GIP-induced cAMP response in L293 cells engineered to overexpress GIP receptor cDNA. Clearly, these results suggest that RGS2 can attenuate Gs-adenylate cyclase signaling pathway. Although, in this study, a direct measurement of the GAP activity of RGS2 toward G_s was not examined, the possibility remains that RGS2 protein might act as an antagonist, blocking the binding of G_s to adenylate cyclase. In contrast, other studies have shown that RGS2 functions as a GAP for both G_i and G_q in reconstituted lipid vesicles (25, 26). Additionally, RGS2 can function to inhibit both G_i- and G_q-dependent responses in transfected cells (25). In these cell assays, RGS2 showed an inhibitory effect on M1 and M2 muscarinic acetylcholine receptor-dependent or interleukin-8 receptor-dependent activation of mitogen-activated protein kinase (25, 39). These findings suggest that RGS2 may play a role in cross-talk between the G_q-mediated calcium/phospholipid signaling pathway and the G_s/G_i-mediated cAMP signaling pathway. Given that PTH, PTHrP, and PGE₂ regulate these multiple signal transduction pathways in bone, we speculate that RGS2 expression and induction may modify the signaling properties of these agents by acting as a switch, to block or turn off one pathway in favor of another. Future studies are required to define which signaling pathway is most affected by RGS2 up-regulation and its consequence, if any, to bone cell function.

In summary, these results provide direct evidence that PTH, PTHrP, and PGE₂ rapidly and transiently increase the level of RGS2 mRNA in bone cells in vivo, through the activation of cAMP signal transduction. This stimulation suggests the exciting possibility that RGS2 may have important consequences in G protein signaling in bone. We conclude that the selective increase in RGS2 expression may represent one of the initiating events involved in controlling PTH actions in bone.

Received May 11, 1999.

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Top Abstract Introduction Materials and Methods Results Discussion References

 

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