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Originally published In Press as doi:10.1074/jbc.M400755200 on March 3, 2004

J. Biol. Chem., Vol. 279, Issue 19, 19764-19774, May 7, 2004
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Transcriptional Regulation of Mouse µ Opioid Receptor Gene by PU.1*

Cheol Kyu Hwang{ddagger}§, Chun Sung Kim{ddagger}, Hack Sun Choi{ddagger}, Scott R. McKercher¶, and Horace H. Loh{ddagger}

From the {ddagger}Department of Pharmacology, University of Minnesota Medical School, Minneapolis, Minnesota 55455 and The Burnham Institute, Center for Neuroscience and Aging, La Jolla, California 92037

Received for publication, January 23, 2004 , and in revised form, March 3, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
We previously reported that the 34-bp cis-acting element of the mouse µ opioid receptor (MOR) gene represses transcription of the MOR gene from the distal promoter. Using a yeast one-hybrid screen to identify potential transcription factors of the MOR promoter, we have identified PU.1 as one of the candidate genes. PU.1 is a member of the ets family of transcription factors, expressed predominantly in hematopoietic cells and microglia of brain. PU.1 plays an essential role in the development of both lymphoid and myeloid lineages. Opioids exert neuromodulatory as well as immunomodulatory effects, which are transduced by MOR. Moreover, MOR-deficient mice exhibit increased proliferation of hematopoietic cells, suggesting a possible link between the opioid system and hematopoietic development. The PU.1 protein binds to the 34-bp element of the MOR gene in a sequence-specific manner confirmed by electrophoretic mobility shift assay and supershift assays. We have also determined endogenous PU.1 interactions with the 34-bp element of MOR promoter by chromatin immunoprecipitation assays. In co-transfection studies PU.1 represses MOR promoter reporter constructs through its PU.1 binding site. When the PU.1 gene is disrupted as in PU.1 knock-out mice and using small interfering RNA-based strategy in RAW264.7 cells, the transcription of the endogenous target MOR gene is increased significantly. This increase is probably mediated through modification of the chromatin structure, as suggested by the reversal of the PU.1-mediated repression of MOR promoter activity after trichostatin A treatment in neuroblastoma NMB cells. Our results suggest that PU.1 may be an important regulator of the MOR gene, particularly in brain and immune cells.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Opioid receptors mediate the various functions of opioids and endogenous opioid peptides, including analgesia, euphoria, respiratory depression, and endocrine/immune regulation. Opioid receptors have been classified into three major types (µ, {delta}, and {kappa}) using pharmacological studies and subsequently by molecular cloning studies (1). They are mainly expressed in the central nervous system (CNS)1 (2, 3) and peripheral tissues such as immune cells (4–6). All three types of receptors belong to the superfamily of G-protein-coupled receptors (2), and the µ opioid receptor (MOR) is known to play an essential role in morphine-induced analgesia, tolerance, and dependence as indicated from pharmacological studies, i.e. MOR gene knock-out studies (7).

The MOR is expressed mainly in the CNS, where it exhibits distinct temporal and spatial patterns of distribution (1). MOR is also detected in immune cells, such as rat peritoneal macrophages (8), and a variety of human and monkey immune cells, including human CEM x174 T/B lymphocytes, human Raji B cells, human CD+ cells, human monocytes/macrophages, human neutrophils, monkey peripheral blood mononuclear cells, and monkey neutrophils (4). The existence of MOR in immune cells can explain why drug abusers (opioids and heroin) frequently become more susceptible to external pathogenic challenges, especially as observed in human immunodeficiency virus-infected opiate users (9–12). Several studies, including in situ hybridization studies and other methods (e.g. RNase protection assay and RT-PCR), indicate that expression of MOR is regulated at the transcriptional level, especially during embryonic development (13–16). Opioid receptors of human and monkey lymphocytes were up-regulated by morphine treatment at the transcriptional and translational levels (17). Interleukin-1{beta} treatment in primary astrocyte-enriched cultures was found to increase the level of MOR messenger RNA in striatal, cerebellar, and hippocampal cultures by 55–75% but not in cultures derived from the cortex or hypothalamus (18). The expression of MOR in the rat mesentery was induced by treatment with a bacterial endotoxin, lipopolysaccharide (19). This induction may be mediated through the actions of IL-1 on MOR (19). The levels of MOR transcripts in discrete brain regions are also regulated by administration of other non-opioid drugs (the dopaminergic drugs, such as cocaine and haloperidol) (20, 21), indicating that the transcriptional regulation of MOR gene is closely associated with the dopaminergic system in the brain. However, the molecular mechanisms of MOR gene regulation are still not completely understood. Studies in our laboratory show that expression of MOR is driven by two promoters, distal and proximal (22, 23). Both promoters exhibit characteristics of housekeeping genes lacking a TATA box (23, 24), and the distal promoter is known to be 20-fold less active than the proximal promoter, based on quantitative RT-PCR using adult mouse brain mRNA (23). Choe et al. (25) have found that a 34-bp cis-acting element in the 5'-distal promoter regulatory sequence possesses a strong inhibitory effect against the distal promoter transcriptional function. Our objective in this study was to identify the transcription factors that bind to the cis-acting element regulating transcription from the mouse MOR distal promoter.

A member of the ets family of transcription factors, PU.1, was isolated using the yeast one-hybrid system, which helped identify the trans-acting factors that bind to the negative regulatory cis-acting element of the MOR distal promoter. This paper describes the in vitro and in vivo data on the regulation of MOR gene expression by PU.1.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Plasmid Construction and in Vitro Translation—Constructs for the recombinant luciferase reporter gene plasmids pL1.3K/687, pL1.3K/728, Linker 3, Linker 6, and Linker 7 have been previously described (25). The mammalian expression plasmid pcFLAG-PU.1 was constructed by inserting the EcoRI-XhoI fragment of the original library clone, pACT2-PU.1, in the same site of pcDNA-5'UT-FLAG (26). For functional analysis of PU.1 protein, pJ6-PU.1 and expression vector pJ6 were gratefully obtained from Dr. M. J. Klemsz (27). The integrity of all constructs was confirmed by restriction enzyme analysis and sequencing.

In vitro translation was carried out with pcFLAG-PU.1 in a reaction mixture containing [35S]methionine (Amersham Biosciences) using a TNT quick-coupled transcription/translation system (Promega). The labeled proteins were then electrophoresed in 10% SDS-PAGE, and their sizes were compared with the predicted sizes.

Cell Culture, Transfection, and Reporter Gene Assay—Human neuroblastoma NMB cells and the murine macrophage-like RAW264.7 cell line were grown in RPMI 1640 medium supplemented with 10% heat-inactivated fetal bovine serum (Invitrogen) at 37 °C in a humidified atmosphere of 5% CO2. Rat glioma C6 cells (American Type Culture Collection) were maintained as described previously (28).

Cells were plated in 6-well dishes at a concentration of 1 x 106 cells/well and cultured overnight before transfection. Various plasmids at concentrations indicated in each figure were used with the Effectene transfection reagent (Qiagen) as described by the manufacturer. Briefly, for luciferase analysis of the distal MOR promoters, 0.5 µg of each reporter plasmid was mixed with the Effectene transfection reagent for 10 min before being added to various cells. Forty-eight hrs after transfection, cells grown to confluence were washed once with 1x phosphate-buffered saline and lysed with lysis buffer (Promega). To correct for differences in transfection efficiency, a 1:5 molar ratio of a pCH110 plasmid (Amersham Biosciences) containing {beta}-galactosidase gene under the SV40 promoter was included in each transfection for normalization. The luciferase and galactosidase activities of each lysate were determined as described by the manufacturers (Promega and Tropix, respectively). For co-transfection assays, the procedures were the same as above, except that the plasmids were a mixture of the given amount of PU.1 construct and 0.5 µg of a corresponding reporter plasmid, pL1.3K/687, pL1.3K/728, Linker 3, Linker 6, and Linker 7.

For trichostatin A (TSA, Sigma) and 5-aza-2'-deoxycytidine (5-Aza-dC, Sigma) treatments, the pL1.3K/687 plasmid was transiently co-transfected either with vector pJ6 or with PU.1 plasmid into NMB cells, and 24 h after the co-transfection the cells were treated with indicated amounts of either TSA or 5-Aza-dC. Luciferase activities were analyzed after 24 h of treatment.

Yeast One-hybrid Screening for cDNAs Encoding Negative Regulatory DNA-binding Proteins—The MATCHMAKER one-hybrid system (Clontech) was used according to the supplier's protocol. The following yeast one-hybrid screening was carried out as described in our recent paper (29) with minor modifications. Four tandem repeats of the –731 to –687-bp sequence (containing the negative regulatory 34-bp cis-acting element) of the mouse MOR distal promoter were ligated into the EcoRI-XbaI site of pHISi and EcoRI-XhoI site of pLacZi to generate pHISi-4xNM and pLacZi-4xNM, respectively (NM, negative modulator, denotes the sequence between –731 and –681). These two bait constructs were then linearized with XhoI and NcoI, respectively, and integrated into the genome of yeast strain YM4271. The resultant yeast cells with the integrated pHISi-4xNM were tested for growth on yeast growth medium lacking histidine (His–) in the presence of increasing concentrations of 3-amino-1,2,4-triazole. Background growth was inhibited in the presence of 45 mM 3-amino-1,2,4-triazole, and this concentration was then used when yeast cells were transformed with a mouse brain cDNA library for one-hybrid screening. Five positive transformants grown on His– plates were selected. To exclude false positive clones, plasmids recovered from these five clones were used to transform yeast cells harboring the pLacZi-4xNM construct. Positive transformants grown on His- and leucine-negative (Leu–) medium containing 45 mM 3-amino-1,2,4-triazole were streaked onto a nylon filter and incubated by placing the filter on the same medium at 30 °C for 2 days. The filter was then soaked in liquid nitrogen for 10 s and placed on a Whatman Whatman No. 3MM filter that had been presoaked in Z buffer (60 mM Na2HPO4, 60 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, 50 mM {beta}-mercaptoethanol, pH 7.0), containing 0.01% 5-bromo-4-chloro-3-D-galactopyranoside (X-gal) and incubated at 30 °C for 2 h. Plasmids from five positive blue clones were sequenced, and their homology was analyzed using the BLAST algorithm.

Electrophoretic Mobility Shift Assay (EMSA) and Supershift Assay— Nuclear extracts from mouse brain and RAW264.7 cells were prepared using a modification of Dignam's procedure (30, 31). The upper and lower strands of each probe were annealed, and the double-stranded oligonucleotides were then end-labeled with [{gamma}-32P]ATP. The end-labeled DNA probes were incubated with nuclear extracts or 2 µl of in vitro translated products in a final volume of 20 µl of EMSA buffer (10 mM Tris·HCl, pH 7.5, 5% glycerol, 50 mM NaCl, 1 mM dithiothreitol, 1 mM EDTA, and 0.1 mg/ml poly(dI-dC)) at room temperature for 30 min. For oligonucleotide competition analysis, a 100-fold molar excess of cold competitor oligonucleotide was added to the mixture before adding the probe. For supershift assay, samples were incubated with 1 µl of each antibody for 30 min before adding the probe. The reaction mixtures were electrophoresed in 4% polyacrylamide nondenaturing gel in 0.5x TBE buffer (45 mM Tris borate and 1 mM EDTA) at 4 °C and visualized by autoradiography. Polyclonal anti-FLAG antibody was purchased from Sigma. Anti-neuron-restrictive silencer factor, anti-PU.1 (T-21, C-terminal epitope), and anti-PU.1 (D-19, N-terminal epitope) antibodies were obtained from Santa Cruz Biotechnology. Anti-histone deacetylase 1 (HDAC1) antibody was purchased from GeneTax, Inc.

Chromatin Immunoprecipitation Assay (ChIP)—RAW264.7 and C6 cells were used for ChIP assays. ChIPs were performed by using a modified protocol from Upstate Biotechnology (Lake Placid, NY). Cells in a 10-cm dish (70% confluent) were treated for 10 min with 1% formaldehyde at room temperature. The cells were lysed in cell lysis buffer (5 mM Hepes, pH 8.1, 85 mM KCl, 0.5% Triton X-100), and the nuclei were resuspended in nuclei lysis buffer (50 mM Tris·HCl, pH 8.1, 10 mM EDTA, 1% SDS). The lysate was sonicated under conditions yielding fragments ranging from 200 to 1000 bp. One-tenth diluted lysate was used for input, and the residual lysate was subjected to the following immunoprecipitation. Samples were subsequently precleared at 4 °C with recombinant protein A-agarose beads (Upstate Biotechnology) coated with salmon sperm DNA. Precleared lysate (100 µl) diluted in immunoprecipitation buffer (0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris·HCl, pH 8.1, 167 mM NaCl) was used for overnight immunoprecipitation with 10 µg of each antibody at 4 °C. Complexes were collected for 4 h by using recombinant protein A-agarose beads coated in salmon-sperm DNA. After washing and elution, formaldehyde cross-linking was reversed with 6 h incubation at 65 °C. After the reverse cross-linking step, DNA was precipitated after phenol/chloroform extraction and dissolved in 30 µl of TE buffer (10 mM Tris·HCl and 1 mM EDTA, pH 8.0). PCR reactions contained 4 µl of immunoprecipitated chromatin sample with the following primers spanning either the rat or mouse MOR distal promoter region in 25 µl of total volume. Mouse MOR D2S and D4AS primers (produces a 245-bp PCR DNA): D2S forward, 5'-GCACATGAAACAGGCTTCTTTCAC-3', and D4AS reverse, 5'-CCCTTATACCTCAACACTCTTCCG-3'; D7S and D7AS primers (produce 142-bp PCR DNA size): D7S forward, 5'-GCTTCTTTCACAAAAATTT-3', and D7AS reverse, 5'-CCATATCTGAGGGAAAAC-3'. Rat MOR primers, same as above for D2S; for rD8AS (produces a 208-bp PCR DNA): rD8AS reverse, 5'-ATACTTAAACACTCTTCCGACTCA-3' was used. After 35 cycles of amplification, 8 µl of PCR product was analyzed on a 1.5% agarose gel.

PU.1 Knock-out Mice and RT-PCR Experiments—The generation of PU.1 null mice has been reported previously (32). The mice brain samples were obtained from the mid- and hindbrain of the animals. The brains were homogenized in 1 ml of RNA-Bee (TEL-TEST Inc.) with a Dounce homogenizer. All samples were placed immediately on dry ice and stored at –80 °C until used. Total RNA was isolated according to the supplier's protocol (TEL-TEST Inc.). The following cDNA synthesis was performed using the SuperScriptTM first-strand synthesis system (Invitrogen). The MOR primers for RT-PCR were used as follows; D2S forward (see the above primer) and P1AS reverse, 5'-CATCCCCAAAGCGCCACTCTCTGAG-3', generating a 547-bp PCR product with PCR 35 cycles of 95 °C for 45 s, 60 °C for 45 s, and 72 °C for 45 s. This PCR was performed in a PerkinElmer Life Sciences PCR machine 9600. A similar reaction was carried out using primers for {beta}-actin (33) as an internal control, except the number of cycles was reduced to 25. The resulting PCR products were analyzed on a 2% agarose gel. The DNA sequences of PCR products were confirmed by sequencing. Quantitative analyses were carried out using ImageQuant 5.2 (Amersham Biosciences) software.

Small Interfering RNA-based (siRNA) Experiments—siRNA strategy (34) was employed to silence the endogenous PU.1 in RAW264.7 cells for the MOR gene regulation. PU.1 and scrambled siRNAs were generated using Donze's procedure (34) and the T7 RiboMAX Express RNAi system (Promega). Briefly, the siRNA sequences for PU.1 and primers were chosen using the web-based tool siRNA Target Designer from Promega. The primers are: PU.1 si-SS sense (nucleotides 70–88 relative to the start codon), 5'-AAGTTGGTATAGCTCTGAATCTATAGTGAGTCGTATTAGGATCC-3', and PU.1 si-AS, antisense 5'-AAGATTCAGAGCTATACCAACTATAGTGAGTCGTATTAGGATCC-3'. The target sequences for PU.1 gene are underlined, and the remaining 3' regions of each primer are the corresponding T7 promoter sequences. T7 si primer, 5'-GGATCCTAATACGACTCACTATAG-3', was synthesized, and scrambled primers (sense and antisense) as negative siRNA control were also synthesized. The oligonucleotide-directed production of small RNA transcripts with T7 RNA polymerase, previously described (35) (34), was carried out using T7 RiboMAX Express RNAi system (Promega). Sense and antisense 21-nucleotide RNAs generated in separate reactions were annealed by mixing both transcription reactions, incubating at 70 °C for 10 min, followed by 20 min at room temperature to obtain small interfering double-stranded RNA synthesized by T7 RNA polymerase. The mixture was then purified by isopropanol precipitation followed by 70% ethanol washing, dried, and resuspended in appropriate amount of nuclease-free water.

The concentration of siRNAs was optimized to 2.5 µg in each transfection for RAW264.7 cells using RNAiFect transfection reagent (Qiagen). Forty-eight hours after the transfection, total RNA and protein samples were prepared using TRI Reagent (Molecular Research Center, Inc.). RT-PCR was performed to determine MOR expression at the mRNA level using the same PCR conditions as in "PU.1 Knock-out Mice and RT-PCR Experiments" under "Experimental Procedures." For Western blotting, equal amounts of total proteins were separated on a 10% SDS-PAGE gel and transferred to ImmobilonTM-P (polyvinylidene difluoride membrane, Millipore) membrane. The proteins were detected either with anti-PU.1 (T-21) (Santa Cruz Biotechnology.) and anti-MOR (generated from our laboratory) antibodies using ECFTM substrate (Amersham Biosciences). The membrane was scanned using a PhosphorImager (Storm 840, Molecular Dynamics).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Isolation of Transcription Factors That Interact with the Negative 34-bp Cis-acting Element Region in the MOR Distal Promoter—Studies from our laboratory have shown that expression of MOR is driven by two promoters, distal and proximal (22, 23). Choe et al. (25) have found that a 34-bp cis-acting element in the 5'-distal promoter regulatory sequence possesses a strong inhibitory effect against the distal promoter transcriptional function. To determine whether this negative cis-acting element could recruit transcriptional factors to regulate the promoter activity, EMSA experiments were carried out using nuclear extracts from mouse brain. A double-stranded oligonucleotide, SL-1 (see Fig. 2A), was prepared spanning the region from –731 to –681 of the MOR promoter, covering the region of the 34-bp-cis-acting element (–721 to –687) with the flanking sequences on both ends. The SL-1 sequence, containing the negative cis-acting element, could form sequence-specific DNA-protein complexes with the nuclear extracts of mouse brain (data not shown), suggesting the presence of endogenous transcription factors in brain.



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  		FIG. 2.
EMSA analysis of a PU.1 binding motif using mutant competitors and supershift assay using anti-PU.1 antibodies. EMSAs were performed as indicated in Figs. 1 and 2. A, double-stranded oligonucleotide sequences were used for 32P-radiolabeled probe (SL-1) and competitors. SL-mPU, the PU.1 binding site of SL-1 was mutated as underlined; PU.1, the PU.1 consensus sequence from major histocompatibility complex class II I-A{beta} promoter (39); mPU.1, the PU.1 binding site of PU.1 consensus sequence (PU.1) was mutated as underlined. B, lane 1, SL-1 probe alone; lanes 2-6, probe SL-1 and 5 µgofnuclear extracts (NE) from macrophage-like cell line RAW264.7. Lane 2 is the control reaction in absence of competitor. Lane 3 is with SL-1 self-competitor. Competitor oligonucleotides are indicated at the top of each lane and were used in 100-fold molar ratio excess (lanes 3–6). C, all lanes contained the probe SL-1 and 5 µg of nuclear extracts from RAW264.7 cells. Lane 1 is the control reaction without antibody. Lanes 2 and 5 contain PU.1 antibodies, anti-PU.1 (T-21), and anti-PU.1 (D-19), respectively. Lane 3 is preimmune serum added as a negative control. Lane 4 contains anti-neuron-restrictive silencer factor (NRSF) antibody as a nonspecific antibody control. The nuclear extracts were preincubated with PU.1 antibodies, preimmune serum, or anti-neuron-restrictive silencer factor as indicated on top of the gel. Antibodies PU.1 (T-21) and PU.1 (D-19) were generated from C-terminal and N-terminal epitopes, respectively (Santa Cruz Biotechnology).

 
To identify transcription factors that bind to the negative element of the distal promoter, we utilized the yeast one-hybrid system to screen an adult mouse brain GAL4-AD fusion cDNA library (Fig. 1, A and B). The following yeast one-hybrid procedures are very similar to what we had done in our previous paper (29), with minor modifications as described below. The SL-1 sequence (Fig. 2A) was used as bait, because it contains additional flanking sequences along with the 34-bp cis-acting element that could be important for transcription factor binding affinity. A double-stranded oligonucleotide having four tandem repeats of the SL-1 sequence was subcloned into pHISi. The resulting plasmid pHISi-4xNM was integrated into the yeast YM4271 genome. Using this yeast strain, 6 x 106 independent colonies from the library were screened. A total of five independent clones (histidine-positive clones) were selected (Fig. 1, A and B) and sequenced. The results revealed that one of the clones is identical to a PU.1 transcription factor. Some of the other positive (His+ and LacZ+) clones have been identified as COUP-TFII (36), NGFI-B (37), and TR2–11 (38). The potential regulatory role of these factors in the MOR gene transcription is currently under investigation in our laboratory.



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  		FIG. 1.
Yeast one-hybrid screening using the negative cis-acting element of the MOR distal promoter. An oligonucleotide with four tandem copies of the wild-type SL-1 sequence was used as bait in the yeast one-hybrid screening. Yeast media dishes A and B were positive for the growth of yeast colonies in the absence of histidine (His-negative yeast selection media) plus 45 mM 3-amino-1,2,4-triazole. The presence of colonies indicates that the yeast cells expressed candidate proteins (including PU.1, clone 1–7) that interact with the bait construct. The positive and negative control experiments, as indicated in dish A, has also been done as recommended by the supplier's manual (Clontech). The other positive (His+ and LacZ+) clones are: clone 1–1, COUP-TFII (36); clone 1–2, NGFI-B (37); clone 1–5, TR2–11 (38); clone 1–8, unknown gene. C, supershift assay of in vitro translated FLAG-tagged PU.1 and the SL-1 probe with anti-FLAG antibody. EMSAs were performed as indicated under "Experimental Procedures." All lanes contained 2 µl of the in vitro translated PU.1 product from 50 µl of reaction volume (Promega's in vitro translation kit). Lane 1 is the control reaction without antibody. Lane 2 shows the reaction mix with the anti-FLAG antibody. Lane 3 is preimmune serum added as a negative control. Lane 4 is self-competitor control without antibody. In vitro translated PU.1 protein was preincubated with polyclonal FLAG antibody, preimmune serum, or self-competitor as indicated.

 
To exclude the possibility that this SL-1 element may form some specific sequences with the minimal HIS3 promoter that favor binding of the PU.1 protein, we used another reporter (LacZ) yeast strain containing the pLacZi-4xNM construct. We also used a control yeast strain without repeats of the cis-acting elements but with pLacZi integrated into its genome. The cloned PU.1 plasmid was transformed into these yeast strains individually. The PU.1 clone in yeast strains containing pLacZi-4xNM was found to be a true positive after the LacZ expression test. In control yeast strains containing pLacZi transformed with PU.1 plasmid, there was no activation of the LacZ promoter (data not shown), indicating that the PU.1 protein indeed has specific strong binding activity to the four tandem repeats of the SL-1 cis-acting elements in vivo.

The ability of PU.1 cDNA to encode proteins was verified by in vitro translation, and the products were analyzed on SDS-PAGE. The in vitro translated PU.1 protein was electrophoresed and found to be the correct size as expected from the calculated molecular weights (data not shown). To confirm whether the isolated PU.1 protein can indeed bind to the negative cis-acting element, EMSAs were carried out using in vitro translated PU.1 protein product. The PU.1 protein was able to shift the target SL-1 oligonucleotide probe (Fig. 1C, lane 1). The specificity of this DNA-protein interaction was verified by competitive inhibition in the presence of cold self-competitor using the PU.1 protein (Fig. 1C, lane 4), whereas the mutated PU.1 competitor was not able to block the complex (data not shown). To further confirm the specificity of PU.1-DNA interaction, we performed supershift assay using in vitro translated FLAG-tagged PU.1 protein with anti-FLAG antibody (Fig. 1C). The PU.1-DNA complex formation was abolished with the addition of anti-FLAG antibody in lane 2 of Fig. 1C but was retained after the addition of preimmune serum, indicating the specificity of the interaction between the FLAG antibody and the complex. We conclude that the in vitro translated PU.1 protein has affinity for and binds to the PU.1-binding motif within the probe SL-1.

Nuclear Extracts from Macrophage-like RAW264.7 Cells Specifically Bind to a PU.1 Binding Motif within the Cis-acting Element—Most of the studies on the PU.1 gene report that the PU.1 protein is expressed tissue-specifically in hematopoietic cells but is not expressed outside the blood cell system (39, 40). However, it was recently reported that the PU.1 gene is found in microglial cell lines and brain microglial cells (41) where it may be associated with the immunological functions of opioid receptors in the brain. Since initial studies of opioid receptor expression in the CNS evidence has accumulated which suggests that there is a neuroimmune circuit involving opioid pathways. For example, radiolabeled agonist and antagonist binding analyses show that opioid receptors are also expressed in cells of the immune system (42–44). Moreover, µ, {kappa}, and {delta} opioid receptors have been cloned by RT-PCR from mRNA isolated from lymphocytes and macrophages and are identical to receptors in the CNS (8, 45, 46). However, the detailed mechanism of transcriptional regulation of MOR involving transcription factors in immune cells has been poorly understood so far. Thus, we used nuclear extracts from an immune cell line, macrophage-like RAW264.7 cells, to determine in vitro physical interactions between the endogenous PU.1 protein from the RAW264.7 cells and the negative cis-acting element of the MOR distal promoter by EMSA using the SL-1 probe. A single major band was identified by competition with self- and mutate competitors (Fig. 2A) and by supershift assay using PU.1-specific antibodies (Fig. 2C). Competition either with a SL-mPU oligonucleotide or an mPU.1 oligonucleotide, which were both mutated at the PU.1 sites, had no effect on the major band (lanes 4 and 6 of Fig. 2B, respectively). Competition either with the self-competitor or the PU.1 consensus competitor (39) was able to abolish the major band (lanes 3 and 5 of Fig. 2B, respectively), indicating that the PU.1 protein interacts with the cis-acting element in a sequence-specific manner. To further confirm the PU.1 interactions, a supershift assay was carried out. Two separate PU.1 antibodies, PU.1 (T-21, C-terminal epitope) and PU.1 (D-19, N-terminal epitope), were able to supershift the major band, but the major band was not supershifted with either preimmune serum as a negative control or neuron-restrictive silencer factor antibody as a nonspecific antibody control (Fig. 2C), indicating that the major band consists of a complex between PU.1 protein from the immune cells RAW264.7 and the cis-acting element. These results demonstrate that PU.1 binds to the mouse MOR promoter, and it is possible that this factor may play a role in the regulation of MOR promoter activity in immune cells.

PU.1 Protein Regulates MOR Transcription through the PU.1 Binding Motif of MOR Distal Promoter—To determine how the PU.1 protein regulates transcription of the mouse MOR gene, the full-length PU.1 DNA in the mammalian expression vector pJ6 was used for further functional analysis. This PU.1 DNA was co-transfected with the mouse distal promoter linked to the luciferase reporter gene into neuronal NMB cells endogenously expressing MOR. When the full-length PU.1 construct was co-transfected into NMB cell line, the PU.1 protein repressed MOR distal promoter activity (pL1.3K/687) in a dose-dependent manner (Fig. 3A). The similar repressive function of the overexpressed PU.1 was also obtained from promoter reporter analysis in macrophage-like RAW264.7 cells (data not shown).



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  		FIG. 3.
The PU.1 binding motif is important for the regulation of the MOR distal promoter by PU.1. A, neuronal NMB cells endogenously expressing MOR gene were co-transfected with the indicated amounts of PU.1 construct in expression vector pJ6 and 0.5 µg of luciferase reporter MOR promoter construct pL1.3K/687 in each well of the 6-well dishes. The relative luciferase activity was expressed as n-fold relative to the reporter activity with vector alone transfected, which was assigned an activity value of 1.0 and normalized by {beta}-galactosidase activity as described under "Experimental Procedures." The data shown are the means of three independent experiments with at least two different plasmid preparations. Error bars indicate the range of standard errors. B, transcriptional analysis was performed similar as above in the transient transfection and luciferase assays. The reporter constructs of pL1.3K/687, pL1.3K/728, and various mutant pL1.3K/687 were co-transfected in the absence (instead, the same amount of vector added) or presence of PU.1-expressing plasmid in the MOR-expressing NMB cells (PU.1-negative cells). The X mark in the filled oval indicates the mutated corresponding site including PU.1 binding site or its flanking sequences. LUC and DP represent luciferase reporter gene and MOR distal promoter, respectively. Transfection efficiencies were normalized as described above. The activities of the luciferase reporter were expressed as n-fold relative to the activity of each corresponding luciferase reporter with vector alone transfected, which was assigned an activity value of 1.0.

 
To further confirm the role of the PU.1 binding motif and PU.1 function, PU.1 construct was co-transfected with various MOR promoter constructs containing mutations in different regions of the negative cis-acting element into the MOR-expressing NMB cells (PU.1-negative cells) (Fig. 3B). The PU.1 protein could not repress promoter activities of the construct pL1.3K/728 (deleted PU.1 binding motif) and Linker 6 (mutated PU.1 binding motif), whereas the wild-type construct pL1.3K/687 and construct Linker 3 (mutated 5' region of the negative cis-acting element) resulted in a significant decrease of MOR promoter activity by PU.1. However, overexpression of PU.1 still repressed about 30% of the MOR promoter activity in Linker 7, which was mutated at the 3'-flanking region of PU.1 binding motif, indicating that the flanking sequence of the PU.1 binding motif may interfere with the action of PU.1 protein that was also reported in other promoters for the function of PU.1 (47, 48). All the above functional results further confirm that the PU.1 protein regulates MOR transcription through the PU.1 binding motif of the MOR distal promoter.

PU.1 and HDAC1 Associate with the MOR Distal Promoter in Vivo in Mouse Macrophage-like RAW264.7 Cells—In this article we have presented evidence for the physical interaction between PU.1 and the cis-acting element of the MOR promoter and for PU.1-mediated regulation of MOR gene transcription through its binding motif in the MOR promoter. Next, we worked to confirm the in vivo interaction between PU.1 and the cis-acting element of the MOR promoter by ChIP with a PU.1 antibody (Fig. 4A). Murine RAW264.7 cells were cross-linked with formaldehyde for the ChIP assay. Chromatin co-precipitated with anti-PU.1 antibody was amplified by PCR using primers including the PU.1 binding site spanning 245 bp for the MOR promoter. As shown in Fig. 4A, ChIP PCR product was detected with the PU.1 antibody in RAW264.7 cells but not with either the NF-{kappa}B (p65) antibody as a nonspecific control or preimmune serum. This suggests that endogenous PU.1 from the RAW264.7 cells specifically binds to the MOR distal promoter. This is consistent with the in vitro results of our EMSA and supershift assays in Fig. 2. Additional ChIP PCR using the same anti-PU.1 antibody was performed with primers spanning 142 bp of the MOR promoter including the same PU.1 binding site but shorter than the above 245-bp promoter region. The expected ChIP PCR product (142 bp) was detected, confirming the first ChIP (245-bp band) assay (data not shown). As a negative control ChIP experiment, we used the MOR-positive C6 glioma cells, which have no detectable PU.1 expression. There was no pulled-down ChIP PCR product observed using anti-PU.1 antibody in rat C6 glioma cells, which is consistent with the result that no PU.1 band was detected in the C6 cells by our EMSA (data not shown).



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  		FIG. 4.
ChIP using PU.1 antibody and HDAC inhibitor TSA treatment. A, the murine macrophage-like cell line RAW264.7 (PU.1-positive) cells were used for a ChIP assay, which could demonstrate PU.1 protein binding to MOR promoter DNA in vivo. Anti-PU.1 and control antibodies were used in a ChIP assay as described under "Experimental Procedures," and the sequence flanking the PU.1 binding site on the MOR gene was examined by ChIP PCR reaction. Lane 1 shows input, lanes 2 and 5 show anti-NF-{kappa}B (P65) as a nonspecific antibody control and preimmune serum as a negative control to precipitate lysates, respectively. Lanes 3 and 4 show the results of positive reactions using anti-PU.1 (T-21) and anti-HDAC1 antibodies, respectively. The ChIP PCR product was the same size as expected and confirmed the integrity by sequencing. B, HDAC inhibitor activates transcription from the MOR distal promoter and inhibits PU.1-mediated repression of the promoter activity. The pL1.3K/687 plasmid was transiently co-transfected either with vector pJ6 or PU.1 plasmid into NMB cells, and 24 h after the co-transfection with the given amount of either TSA or 5-Aza-dC were treated to the corresponding transfected cells. Luciferase activities were analyzed after 24 h treatment of HDAC inhibitor TSA and methylation inhibitor 5-Aza-dC. Data were normalized by protein concentration and expressed as n-fold activation of the luciferase activity of non-treated vector control, which is arbitrarily defined as 1.0. Error bars indicate the range of S.E. In addition, the fold increase in promoter activity by TSA compared with corresponding non-treated control is shown on the top of the error bars.

 
Regarding the functional mechanism of PU.1 on transcriptional repression, it was recently reported that PU.1 recruits HDAC1 to the c-Myc and several other promoters for altering chromatin structure (49). Thus, we used an anti-HDAC1 antibody in place of the anti-PU.1 antibody in the same ChIP assay. The addition of anti-HDAC1 antibody also resulted in an identical band corresponding to 245-bp MOR promoter region containing the PU.1 binding site as detected by the anti-PU.1 antibody (Fig. 4A). This suggests an interaction between PU.1 and HDAC1, resulting in transcriptional repression. No other transcription factors that can interact with HDAC1 have been reported in the 245-bp MOR promoter region other than that for PU.1. Computerized searching for transcription factor binding sites using TRANSFAC (www.cbil.upenn.edu/tess (50)) also revealed no other known factor sites that can interact with HDAC1 adjacent to the PU.1 binding site.

The Treatment of TSA as a Specific Inhibitor of Histone Deacetylase (HDAC) Activity Abolishes PU.1-mediated Repression of the MOR Promoter Activity—PU.1 was originally identified as an activator in the transcriptional regulation of various genes (39, 51). There is accumulating evidence that PU.1 also functions as a transcriptional repressor (49, 52, 53). As described earlier, it was reported that PU.1 interacts with HDAC1 to tighten the chromatin structure and prevent access of other transcriptional activators into the corresponding gene promoter regions, thus leading to transcriptional repression (49).

To investigate whether PU.1-mediated repression of transcription of the MOR gene functions through an interaction with HDACs in addition to ChIP experiment, reporter gene analysis of MOR promoter activity was carried out in NMB cells treated with or without the HDAC inhibitor, TSA. Altering chromatin structure by blocking HDAC activity with specific inhibitors such as TSA can result in an up-regulation of gene expression (54). As shown in Fig. 4B, HDAC inhibitor TSA increased MOR promoter activity about 4.3-fold in NMB cells (co-transfection of vector plus pL1.3K/687 as vector control) and abolished PU.1-mediated repression (PU.1 plus pL1.3K/687). An increase in MOR promoter activity in the absence of PU.1 (vector control) by TSA suggests that other transcription factors interacting with HDACs may also be involved in NMB cells that do not express PU.1.

In addition, methylation inhibitor, 5-Aza-dC, was used. It was reported that direct association between PU.1 and MeCP2, a methyl CpG-binding protein, could recruit mSin3A-HDAC complex for PU.1-mediated transcriptional repression in murine erythroleukemia (MEL) cells (54). In that study the MeCP2 repressed PU.1-mediated activation of transcriptional activity containing PU.1 binding sites, and this down-regulation was recovered in the presence of histone deacetylase inhibitor, TSA. However, in our study, the treatment with 5-Aza-dC as a methylation inhibitor did not influence MOR promoter activities of both vector control and PU.1 co-transfection (Fig. 4B). This result suggests that MeCP2, previously known to act as a co-repressor of PU.1, does not facilitate complex formation with mSin3A and HDACs in NMB cells. Collectively, the results from ChIP using HDAC1 antibody (Fig. 4A) and TSA treatment experiments (Fig. 4B) suggest that PU.1 represses MOR promoter activity through an interaction with HDAC1, indicating the alteration of chromatin structure.

The Endogenous MOR Gene Regulation of PU.1 Protein Was Analyzed by Using PU.1 Knock-out Mice and a siRNA Strategy—We utilized PU.1 knock-out mice to provide in vivo evidence of PU.1 regulation of the MOR gene. PU.1-negative mice were generated using the same procedure as the previous report (32). The mouse brain samples were obtained from the mid- and hindbrain of the animals. MOR gene expression at the mRNA level was analyzed by RT-PCR (Fig. 5A). In homozygous PU.1 knock-out mice (PU.1–/–), expression of the MOR gene was significantly increased 3–8-fold compared with normal mice (PU.1+/). The difference of MOR expression level between two normal mice (lane 1 and lane 2, Fig. 5A) may be the result of normal variability of heterozygous PU.1 mice, or due to a difference between PU.1+/– and +/+ genotypes, since the genotypes remain to be confirmed. The control {beta}-actin gene was expressed at the same level from all four mouse samples (normal and mutant mice), and there were no detectable bands from the samples without reverse transcriptase during cDNA synthesis (–RT, lanes 6–9), indicating specific MOR expression by RT-PCR. Quantitative data in Fig. 5B were analyzed by using ImageQuant 5.2 (Amersham Biosciences) software after {beta}-actin normalization.



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  		FIG. 5.
Analysis of MOR gene regulation of PU.1 protein in vivo using PU.1 knock-out mice. A, PU.1 negative mice were generated from the same procedure as the previous report (32). The mice brain samples were obtained from the mid- and hindbrain of the animals. RT-PCR was performed to analyze the MOR gene expression using cDNAs synthesized from the brain samples of the animals. Homozygous PU.1-negative mice are represented as PU.1–/–. Normal mice controls are represented as PU.1+/. Lanes 1–4 included reverse transcriptase (+RT) in cDNA synthesis as normal reaction, whereas lanes 6–9 did not include the RT in cDNA synthesis as negative control reactions (–RT). The RT-PCR for the {beta}-actin gene (33) was carried out in a separate tube as an internal control. Lane 5 shows a 100-bp DNA marker (Invitrogen). The PCR products for MOR and the {beta}-actin gene were the same size as expected and confirmed the integrity by sequencing. B, quantitative data for lanes 1–4 of A were analyzed by using ImageQuant 5.2 (Amersham Biosciences) software after {beta}-actin normalization. Bars depict the sum of signal intensity in the same size area and S.D. between experiments. The average signal from normal mice (PU.1 +/) was set at 1.

 
siRNA strategy (34) was employed to silence the endogenous PU.1 expression in RAW264.7 cells since there may be complicating factors in gene knock-out mice studies, making it difficult to assign a direct role for a target gene. PU.1 and scrambled siRNAs were generated according to Donze's procedure (34) and the T7 RiboMAX Express RNAi system (Promega). The concentration of siRNAs was optimized to 2.5 µg in each transfection for RAW264.7 cells. In Fig. 6A, transfection of PU.1 siRNA resulted in a complete silencing of the PU.1 gene at the protein level. Transfection of scrambled siRNA as a nonspecific sequence control had no effect on PU.1 expression compared with the no siRNA transfection control, indicating high efficiency of the siRNA strategy. Expression of the target MOR was dramatically increased with PU.1 siRNA transfection both at the protein and mRNA levels (Fig. 6, A and B, respectively), compared with scrambled siRNA and no transfection, showing in vivo evidence of the PU.1 function on endogenous MOR gene regulation. Fig. 6C is a quantitative analysis of two independent RT-PCR experiments measuring changes in MOR mRNA levels. Data were normalized to {beta}-actin, and the graph was generated by using ImageQuant 5.2 (Amersham Biosciences) software.



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  		FIG. 6.
Analysis of MOR gene regulation by PU.1 protein in vivo using a siRNA strategy. A, results of siRNA experiments in RAW264.7 cells are shown here by Western blotting. Lanes 1 and 2 show transfection cells without siRNA and with 2.5 µg of scrambled (scb) siRNA, respectively. Lane 3 shows transfection cells with 2.5 µg of PU.1 siRNA. The design and synthesis of siRNA were carried out as Donze's procedure (34) and the T7 RiboMAX Express RNAi system (Promega), as described under "Experimental Procedures." The upper Western blot shows PU.1 expression with anti-PU.1 (T-21) antibody (Santa Cruz Biotechnology), and the lower Western blot shows MOR expression with anti-MOR (generated from our laboratory) antibody using ECFTM substrate (Amersham Biosciences). B, a parallel experiment was conducted using total RNAs from the above samples to analyze changes in the mRNA level of MOR after siRNA treatment. RT-PCR was performed using the same PCR conditions as in "PU.1 Knock-out Mice and RT-PCR Experiments" under "Experimental Procedures" of PU.1 knock-out mice. Total RNAs were isolated by TRI Reagent (Molecular Research Center, Inc.). C, a quantitative analysis of two independent experiments at the MOR mRNA level of B is shown in here after {beta}-actin normalization; the graph was generated by using ImageQuant 5.2 (Amersham Biosciences) software. Bars depict the sum of signal intensity in same size area and the S.D. between experiments. The signal from non-transfected cells was set at 1.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Precise transcriptional regulation of opioid receptor genes in the brain is crucial for normal neuropharmacological function. Several classes of nuclear proteins are intricately involved in controlling expression of these genes (55). The µ opioid receptor (MOR) gene, which is regulated by opioid and non-opioid drugs, is under tight inhibitory control operating at the level of the 5'-distal promoter regulatory sequence, which has consensus PU.1 binding sites (25). This negative cis-acting element of the MOR distal promoter exerts a strong repressor function on both the MOR distal promoter and the heterologous, SV40 early promoter (25). We report here that a nuclear factor, PU.1, which binds to this negative regulatory element, represses the MOR promoter in transiently transfected MOR-positive neuronal (NMB) and immune (RAW264.7) cells.

In this study we identified a transcription factor PU.1 as a transcriptional regulator of the mouse MOR gene. This regulatory activity was mediated by a PU.1 binding site in the 34-bp cis-acting element, located downstream of the MOR distal promoter. Nuclear extracts from mouse brain can specifically bind to this 34-bp cis-acting element by EMSA. We were able to isolate a PU.1 protein from mouse brain by the yeast one-hybrid system using the cis-acting sequence as bait. Deletion or mutation of this PU.1 binding motif abolished PU.1-mediated repression of mouse MOR distal promoter activity in transient transfection assays. Nuclear extracts from RAW264.7 cells could specifically bind to this PU.1 binding motif, and the protein was confirmed to be PU.1 by EMSA and supershift assay. When the PU.1 gene is disrupted, as in PU.1 knock-out mice, and by using siRNA for PU.1 gene in immune (RAW264.7) cells, the endogenous target MOR gene is significantly increased. This increase is possibly the result of modification of the chromatin structure, as suggested by the observed increase in MOR promoter activity in NMB cells co-expressing PU.1 and the MOR promoter reporter construct after treatment with TSA. Thus, the identified PU.1 binding motif plays an important role in the regulation of MOR distal promoter through interaction with the PU.1 protein in cells that express this protein. We also found a consensus PU.1 binding site in the proximal promoter of the mouse MOR gene, but further experiments suggested that this site appears to be functionally inactive (data not shown).

The MOR distal promoter is known to be 20-fold less active than the proximal promoter in mouse brain (23). In immune cells it is not known whether the distal promoter is so much less active than the proximal promoter as in mouse brain. In the present study, disruption of the PU.1 gene results in an up-regulation of MOR from distal promoter in immune cells. This up-regulation may overwhelm the MOR expression from the proximal promoter since the distal promoter is physically located upstream of the proximal promoter of MOR gene. However, this speculation requires further investigation. The current results from the PU.1 knock-out mice study (Fig. 5) may not be conclusive in this regard since there may be complicating factors in gene knock-out mice studies, making it difficult to assign a direct role for a target gene.

PU.1 is a member of the ets family of transcription factors (39, 56, 57) and was originally identified as a protein that binds to a purine-rich sequence (PU box, 5'-GAGGAA-3') in the mouse major histocompatibility complex class II gene I-A{beta} (39). PU.1 is found mainly in cells of the hematopoietic system (39). Recent studies show that PU.1 plays an important role in the development of hematopoietic cell lineages, since disruption of the PU.1 gene in mice is lethal to animals that lack B cells and macrophages (32, 58). PU.1 has been implicated in regulation of the expression of many myeloid (granulocytes, monocytes, and macrophages) and B cell genes (i.e. immunoglobulin genes (51, 59), macrophage colony-stimulating factor receptor (CSF) (60), granulocyte-macrophage CSF receptor {alpha} (61), granulocyte CSF receptor (62), and mannose receptor (63)). It may also serve self-regulatory functions (64). Many of these genes play important roles in the growth, development, and function of the hematopoietic system. In addition, cloning of the PU.1 protein from the adult mouse brain cDNA library in this present study indicates the PU.1 gene is also present in the adult mouse brain, which is consistent with the recent report that PU.1 is expressed in microglial cells (the brains resident macrophage) of rat brain and microglial cell lines (41). This finding suggests a possible immunological role for PU.1 in the brain. Microglia respond rapidly to brain damage by proliferating, changing morphology, and producing a number of cytokines (65). PU.1 is known to bind to Goosecoid, which reduces cell proliferation and enhances neurite outgrowth in response to NGF in PC12 cells (66). Co-expression of the N-terminal portion of PU.1 and Goosecoid repressed neurite outgrowth and rescued the proliferation of PC12 cultures, indicating a potential role of PU.1 in neuronal development. As a result of the initial studies of opioid receptor expression in the CNS, evidence has accumulated that suggests a neuroimmune circuit involving opioid pathways. Strong evidence for the direct modulation of the immune system by opioids is well documented (9, 67). µ opioids have been shown to alter the release of cytokines important for both host defense and the inflammatory response. Pharmacological and molecular biological studies show that opioid receptors are expressed on cells of the immune system (6, 42, 68) such as in lymphocytes and macrophages and are identical to receptors in the CNS (8, 45, 46). Moreover, MOR-deficient mice show increased proliferation of granulocyte-macrophage, erythroid, and multipotential progenitor cells (69), indicating an important link between the opioid system and hematopoietic development.

PU.1 interacts with several proteins, such as CREB-binding protein (CBP/p300) (70), interferon regulatory factor 4 (IRF4), and interferon consensus sequence-binding protein, to function as a transcriptional activator or repressor. The activation domain of the N-terminal region of PU.1, including the glutamine-rich region, has been known to interact with CBP (70), a shared co-activator for several transcription factors including CREB, AP-1, c-Myb, nuclear hormone receptors, STATs, NF-{kappa}B, E2F-1, and p53 (70, 71). CBP/p300 possesses intrinsic histone acetyltransferase activities, acetylating histones, and nonhistone nuclear proteins (71). The C-terminal region of PU.1 has recently been reported to interact with HDAC1 and mSin3A, where it functions as a repressor (49). Acetylation of histones is associated with a relaxed chromatin configuration, which is thought to facilitate transcription factor access to DNA. Gene activation is generally associated with hyperacetylated histones, and repressed genes are usually found in hypoacetylated chromatin (72). Our data in Fig. 4 has proven, in part, that the repressive function of PU.1 in neuronal cells and immune cells acts through interaction with HDACs.

IRF4 (also called Pip) and interferon consensus sequence-binding proteins, belonging to the family of IRF proteins, can also form specific complexes with PU.1 and can activate or repress transcription via binding to PU.1/IRF composite sequences depending on protein-interaction partners and/or context of DNA binding (53). These IRF proteins bind to interferon-stimulated response elements (ISRE) or IFN-{gamma}-activated sequences (GAS) within interferon-responsive genes (73). IRF4 is recruited to its binding site on DNA by PU.1 phosphorylated at Ser-148 of its PEST region (74). Interaction between the interferon consensus sequence-binding protein (IRF8) and TEL (another Ets member), which is triggered by IFN-{gamma}, recruits the histone deacetylase HDAC3 to the interferon-responsive element (75). A typical ISRE motif follows the consensus 5'-RYAAAGTGAAANC-3' and appears to be a tandem repeat with dyad symmetry (76). This consensus sequence represents a double repeat of the monomeric sequence 5'-GAAAN(T/C)-3'. A permutation of ISRE motif is the hybrid PU.1/ISRE motif that follows the consensus 5'-RRRGAAGTGAAANY-3'. The 5' PU.1 and 3'ISRE core motifs are indicated by underlined bold letters. A similar PU.1/ISRE motif is found in the region from nucleotides –701 to –688 (5'-GAGGAACTAAAGGG-3') of the 34-bp cis-acting element of the MOR promoter. An additional putative ISRE sequence (5'-taAAAGaGAAAAT-3' in the opposite orientation; mismatches are indicated by small letters) is located at nucleotide –761 to –749 upstream of the 34-bp cis-acting element. Even if the latter putative ISRE sequence can bind IRF proteins, the physical binding distance between these two sites (ISRE and PU.1) is much greater than between known PU.1/IRF motifs, which are two bases apart as shown above. However, the existence of Sox binding sites (29) in between these two sites may explain how PU.1 interacts with IRF proteins. Sox proteins are supposed to bend the target DNA to allow interactions between multiple transcription factors. Sox proteins belong to the high mobility group (HMG) proteins, and HMG I/Y, another HMG protein, has been shown to interact with PU.1. Binding of the Sox protein to its consensus binding site on the MOR promoter may bring the PU.1 and IRF binding sites close enough for a functional interaction.

Another possible mechanism of PU.1 regulation in the MOR gene could involve cytokine IL-4 and IFN-{gamma}, based on the recent report that the MOR was induced at the transcriptional level by IL-4 through Stat6 interaction on the IL-4 response element of human and rat MOR promoters in human primary blood cells, immune cell lines, dendritic cells, and neuronal cells (77). This IL-4 response element is located 23 bases upstream of the 34-bp cis-acting element including the PU.1 binding site. The cytokines IL-4 and IFN-{gamma} exert biologically antagonistic effects that, in part, reflect opposing influences on gene transcription. IFN-{gamma} directly suppresses IL-4 gene expression. IFN regulatory factor-1 (IRF1) and IRF2 induced by IFN-{gamma} bind to the distinct IL-4 promoter sites and function as transcriptional repressors (78). Additionally, IL-4 inhibited the IFN-{gamma}-inducible promoter of IRF1 by activating Stat6 in a cell line expressing low endogenous Stat6. The Stat1-dependent IFN-{gamma} activation sequence element of the IRF1 promoter is a target for Stat6-mediated inhibition despite apparently normal Stat1 DNA binding (79), and this repression is attenuated by increasing the amount of IFN-{gamma}. IFN-{gamma}-dependent transcription of the IRF1 reporter gene is suppressed by IL-4, but IL-4 alone has no trans-activating function (80). Morphine and the endogenous opioid {beta}-endorphin suppressed production of IFN-{gamma} by cultured human peripheral blood mononuclear cells (81) in vivo (82), and these opioid-mediated effects were abolished in MOR-deficient mice (82), clearly showing that opioid-induced immunosuppression is mediated by the MOR. A significant enhancement of IFN-{gamma} release was observed in lipopolysaccharide-stimulated whole blood cultures obtained from patients undergoing surgery after induction of anesthesia (83). Morphine has been shown to up-regulate its own receptor (µ opioid receptor) in human and monkey lymphocytes (17). In addition to the interactions of ets member PU.1 with HDAC1 and mSin3A, the direct interaction between the other ets member, TEL, and IRF proteins on the IFN-stimulated response element (ISRE) recruits HDAC3 in macrophages in an IFN-{gamma}-dependent manner, causing increased repression of IFN-{gamma}-mediated reporter activity through the ISRE (75). With all of this previous information we propose that PU.1 may suppress IL-4-mediated activation of the MOR gene through IRFs. Thus, PU.1 may play an important role in the transcriptional regulation of the MOR gene depending on interacting factor context and be involved in known regulatory mechanisms of PU.1 such as interactions with IRF proteins and HDACs.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Research Grants DA00564, DA01583, DA11806, and K05-DA70554 and by the A&F Stark Fund of the Minnesota Medical Foundation. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ To whom correspondence should be addressed: Dept. of Pharmacology, University of Minnesota Medical School, 6-120 Jackson Hall, 321 Church St. S.E., Minneapolis, MN 55455. Tel.: 612-626-6539; Fax: 612-625-8408; E-mail: hwang025{at}umn.edu.

1 The abbreviations used are: CNS, central nervous system; MOR, µ (mu)-opioid receptor; RT-PCR, reverse transcriptase-polymerase chain reaction; EMSA, electrophoretic mobility shift assay; HDAC, histone deacetylase; CREB, cAMP-response element-binding protein; CBP/p300, CREB-binding protein; IRF4, interferon regulatory factor 4; IFN-{gamma}, interferon {gamma}; IL-4, interleukin-4; Sox, Sry like HMG box; STAT, signal transducers and activators of transcription; TSA, trichostatin A; 5-Aza-dC, 5-aza-2'-deoxycytidine; ChIP, chromatin immunoprecipitation; siRNA, small interfering RNA-based; ISRE, interferon-stimulated response element. Back


   ACKNOWLEDGMENTS
 
We thank Drs. Ping-Yee Law, Li-Na Wei, Sabita Roy, Ursula D'Souza, and Santosh Talreja for helpful suggestions and manuscript review. We also thank Dr. M. J. Klemsz for kindly providing both the PU.1 expression plasmids and the vector pJ6 plasmid.



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 DISCUSSION
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