Multiple PU.1 sites cooperate in the regulation ofp40^phox transcription during granulocytic differentiation of myeloid cells

NADPH oxidase is the major inducible source of superoxide and superoxide-derived reactive oxygen species in phagocytes and is a key enzyme in host defense against microbial infection.1-3 Genetic deficiency of this enzyme results in the inherited disorder chronic granulomatous disease (CGD), which is characterized by impaired phagocyte microbicidal activity and a clinical syndrome of recurrent life-threatening infections.1-3 The reactive oxygen species generated by this enzyme are also toxic to host cells and may account for much of the tissue injury that occurs during inflammation.4-6Therefore, the expression and activity of the phagocyte NADPH oxidase components must be tightly regulated.

The fully assembled active NADPH oxidase complex is composed of membrane-bound and cytosolic components.7,8 The membrane-associated core component of the oxidase is a heterodimeric b-type cytochrome comprised of light (p22^phox) and heavy (gp91^phox) chain polypeptides. Cytosolic components include p67^phox, p47^phox, p40^phox, and the small GTPase protein, Rac2. X-linked CGD, the most prevalent form of the disorder, is a result of mutations in the gp91^phox gene, whereas autosomal recessive CGD is associated with mutations of thep47^phox, p67^phox, orp22^phox genes.1-3 

Numerous studies have demonstrated essential roles for gp91^phox, p22^phox, p67^phox, p47^phox, and Rac2 in phagocyte NADPH oxidase function.9-20Reconstitution of the oxidase by transfection into cell lines deficient in the oxidase proteins or recombination of the components in vitro shows that efficient superoxide generation by the NADPH oxidase complex is absolutely dependent on these components. The role of p40^phox, on the other hand, is less clearly defined because it does not appear to be absolutely necessary for the core enzymatic function, namely the univalent reduction of oxygen. Nevertheless, several lines of evidence suggest that p40^phox is directly involved in the normal physiologic function of the oxidase. Thus, p40^phox forms a trimolecular complex with p47^phox and p67^phoxin the cytosol of resting phagocytes and on activation translocates to the membrane with these components.21-23 Moreover, p40^phox can bind p67^phox and p47^phox, and the strong binding of p40^phox to p67^phox can be disrupted by the activated guanosine triphosphate-bound form of Rac, a key intermediate in oxidase activation.23-31 The SH3 domain of p40^phox competes for the C-terminal, proline-rich domain in p47^phox, which also interacts with the C-terminal SH3 domain of p67^phox and thereby down-regulates NADPH oxidase activity.24,25,29,30 In contrast, other studies suggest that p40^phox actually participates in the activation of NADPH oxidase by increasing the affinity of p47^phox for the flavocytochrome.32More recently, it has been reported that the 57-kd actin-binding protein coronin associates with the p40^phox–p67^phox–p47^phoxcytosolic complex through the C-terminal domain of p40^phox and that it colocalizes with the phox proteins in the periphery of the phagocytic vacuole during phagocytosis and activation of the oxidase.33Finally, recent studies have shown that p40^phox is hyperphosphorylated during NADPH oxidase activation and dephosphorylated during enzyme inactivation.34 35 In summary, these studies suggest that p40^phox acts as a regulator of oxidase activity and as an adapter that links oxidase to the cytoskeletal elements that participate in cell activation.

Tissue distribution studies have demonstrated that, similar to the gp91^phox, p47^phox, and p67^phox components of NADPH oxidase, p40^phox is preferentially expressed in cells of myeloid lineage.36,37 Transcription of many myeloid-specific genes is regulated by PU.1, a member of theets family of transcription factors.38Gene-targeting studies have demonstrated that PU.1 is essential for the normal embryonic development of hematopoietic cells of myeloid, B-cell, and T-cell lineages.39,40 We have previously characterized the promoter of the p47^phox gene and demonstrated that PU.1 is essential for p47^phoxtranscription in myeloid cells.41 PU.1 appears to regulate transcription by binding to a single cis element located in the proximal region of the p47^phox gene. In this report, we describe our studies on the transcriptional regulation of the p40^phox gene in the HL-60 myeloid cell line. Our data show that p40^phoxtranscription is regulated by 3 PU.1 binding sites located within the proximal region of the p40^phox promoter and that the PU.1 transcription factor binds to each site, in vitro and in vivo. The central role of PU.1 in the regulation ofp40^phox expression supports previous observations that this phox protein is mainly restricted to cells of myeloid lineage and is of particular importance in phagocyte function.

RPMI 1640 was obtained from Life Technologies (Gaithersburg, MD). Restriction enzymes, T4 polynucleotide kinase, RNasin, and pGL3-Basic luciferase reporter vector and the dual luciferase assay kit were from Promega (Madison, WI). γ-[³²P]Adenosine triphosphate (ATP), 6000 Ci/mmol (370 MBq/mol), was obtained from DuPont-NEN (Boston, MA). The TOPO-TA Cloning Kit, with the pCRII vector for cloning products of the polymerase chain reaction (PCR), was obtained from Invitrogen (San Diego, CA). Oligonucleotides were synthesized at the Advanced DNA Technology Unit (University of Texas Health Science Center at San Antonio). The Sequenase DNA sequencing kit was obtained from United States Biochemical (Cleveland, OH). PU.1 antibody was purchased from Santa Cruz Biotechnology (Santa Cruz, CA).

The p40^phox 5′-flanking region was cloned using the PromoterFinder Kit (Clontech, Palo Alto, CA) according to the manufacturer's protocol. PCR was performed using the human genomic libraries provided as templates to amplify the desired sequences. The forward primer was complementary to the adaptor ligated to the genomic DNA fragments contained in each library. The reverse primer (5′-CCAGGGAGCAGGTGGAGAGTCTCGC-3′, Figure 2A) was complementary to base pair (bp) +142 to +117 of thep40^phox gene.22 36Amplified products were analyzed on a 1.2% agarose gel and then subjected to a second round of PCR with nested primers. The nested reverse primer (5′-AGGCTGAGTTCACCTCTCACTTCC-3′) was complementary to bp +105 to +81 of the gene. Final PCR products were cloned directly into the pCRII vector, and their identities were confirmed by sequencing. Nucleotide sequencing was carried out in both directions using the dideoxy termination procedure and sequence-specific oligonucleotide primers.

Reporter vectors were constructed in the pGL3-Basic luciferase vector. Promoter regions were amplified using human genomic DNA as template. The reverse primer was similar to the nested reverse primer used above, but a site for cleavage by XhoI was added (5′-CACTCGAGAGGCTGAGTTCACCTCTC-3′). The forward primers corresponded to the upstream sequences of desired promoter regions, with a KpnI cleavage site added at the 5′ end. PCR products were digested with KpnI and XhoI and were cloned into the pGL3-Basic reporter vector. Inserts of the constructs generated all extend downstream to +104 relative to the transcription start site of the p40^phoxgene.36 Mutations of the PU.1 binding sites were generated by site-directed mutagenesis (QuikChange kit; Stratagene, La Jolla, CA) using the primers illustrated in Figure 2B. Constructs were confirmed by restriction mapping and sequencing.

As previously described,41 the human myeloid cell lines HL-60, THP-1, U937, and PLB985 and the human B-cell line Raji were maintained in RPMI 1640 supplemented with 10% fetal bovine serum, 10 mM HEPES, penicillin, and streptomycin. Transfection was carried out by electroporation as we have described.41 Briefly, approximately 10⁷ cells were resuspended in 0.5 mL complete medium containing 20 μg luciferase reporter constructs and 0.2 μg of a Renilla luciferase vector (pRL-null; Promega) as a transfection efficiency control. In Figure 1, transfection was also carried out with equimolar amounts (7 pmol) of each construct, plus an unrelated plasmid, pUC19, to provide an equal amount of total DNA. For cotransfection assays, 10 μg each reporter construct and PU.1 expression plasmid (a generous gift from Dr M. Klemsz, Indiana University, Indianapolis) were used. Electroporation was carried out at 960 μF and 250 V for HL-60 cells or 320 V for Raji cells. At 48 hours the cells were washed 3 times in phosphate-buffered saline (pH 7.4), lysed in 100 μL 1× reporter lysis buffer (Promega), and centrifuged at 400g for 5 minutes at ambient temperature, and 20-μL aliquots of the supernatants were tested for reporter gene activity using the dual luciferase assay system (Promega) and a Turner Designs TD–20/20 luminometer. The human cervical carcinoma epithelial cell line HeLa was grown in Dulbecco modified Eagle medium supplemented with 10% fetal bovine serum and transfected by Lipofectamine Plus reagent (Gibco) according to the manufacturer's protocol.

Mouse PU.1 cDNA42 was subcloned into pBluescript SK and was translated in vitro using T3 RNA polymerase and the TnT-coupled reticulocyte lysate system (Promega). Synthesized [³⁵S]-methionine–labeled PU.1 was analyzed by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) and fluorography. A predominant band measuring approximately 38 kd was observed, consistent with the molecular mass previously reported.43 The control un-programmed sample (ie, no cDNA) produced no corresponding band.

Cells were disrupted by nitrogen cavitation using a technique originally described for neutrophils,44 and nuclear extracts were prepared as we have previously reported in detail.41 Extracts were collected and stored in aliquots at −70°C. The protein concentration was determined using the Bradford reagent (Bio-Rad, Hercules, CA).

Complementary DNA oligonucleotides were annealed by heating in 1× NET at 95°C for 5 minutes and cooling at ambient temperature. Probes were then labeled with [γ-³²P]ATP and T4 polynucleotide kinase. For gel shift assays, nuclear extract (6 μg) was incubated for 20 minutes at ambient temperature with 5 × 10⁴ cpm of the labeled DNA probe in 20 μL binding buffer containing 10 mM Tris-HCl (pH 7.6), 50 mM NaCl, 1 mM EDTA, 1 mM dithiothreitol, 5% glycerol, 1 μg/μL bovine serum albumin, and 2 μg poly-d(I-C). For supershift assays, 2 μL specific antibody was added, and the reaction was continued for 15 minutes. Samples were loaded on 5% nondenaturing polyacrylamide gels, and electrophoresis was carried out at 200 V in 25 mM Tris (pH 8.5) with 190 mM glycerol and 1 mM EDTA. Competition assays were carried out in the same manner, except that the above reaction mixture was preincubated with competitor DNA for 10 minutes at 4°C before the labeled probe was added. Signals were quantitated with a PhosphorImager and ImageQuant software (Molecular Dynamics). Relative binding avidity of various DNA probes for PU.1 were determined by comparisons of band intensities observed in the presence of varied amounts of probes or a series of dilutions of competing oligonucleotides.

Chromatin immunoprecipitation (ChIP) analysis was carried out as described by Boyd et al.45 Briefly, formaldehyde was added directly to the cell culture to a final concentration of 1% (wt/vol), and the cells were incubated at 23°C for 10 minutes. Glycine was added to stop the reaction, and the cells were collected, washed, and allowed to swell on ice for 10 minutes in 5 mM PIPES (pH 8.0) containing 85 mM KCl, 0.5% NP-40, 0.5 mM phenylmethylsulfonyl fluoride, and 100 ng/mL leupeptin and aprotinin. Pelleted nuclei were collected and incubated on ice for 10 minutes in 50 mM Tris HCl (pH 8.1), 1% SDS, 10 mM EDTA, and the protease inhibitors, sonicated on ice to break the chromatin DNA, and precleared with Staph A cells (protein A–positive Staphylococcus aureus cells; Roche Molecular Biochemicals, Indianapolis, IN). Precleared chromatin from 2 × 10⁷ cells was incubated with 1.5 μg affinity-purified rabbit polyclonal antibody to PU.1 (anti-PU.1 sc-352X; Santa Cruz Biotechnology) or without antibody, and it was rotated at 4°C for 12 to 16 hours. Immunoprecipitation was carried out with Staph A cells. The supernatant from the reaction lacking primary antibody was saved as total input of chromatin and was processed in the same way as the eluted immunoprecipitates, starting with the cross-link reversal step. Cross-linking was reversed by incubation at 65°C for 5 hours in the presence of 300 mM NaCl and 0.01% RNase A. Samples were then precipitated, resusupended, and treated with proteinase K, followed by extraction with phenol-chloroform-isoamyl alcohol and precipitation with sodium acetate and ethanol plus tRNA and glycogen as carriers. Pellets were collected by microcentrifugation, resuspended in 30 μL H₂O, and analyzed by PCR. Total input samples were resuspended in 100 μL H₂O and then diluted 1:100 before PCR. PCR reactions were carried out in a total volume of 25 μL containing 2-3 μL immunoprecipitate or diluted total input as the template and using recombinant Taq DNA polymerase (Life Technologies). After 35 cycles of amplification, PCR products were separated on 1.5% agarose containing ethidium bromide. PCR primers (p40-PU.1 forward, 5′-GCGCCAAGGACTGACATC-3′ and reverse, 5′-GAGCAGGTGGTGCGTCTC-3′) were designed to amplify a 299-bp region of the p40^phox promoter that contains the 3 PU.1 sites.

Total cellular RNA was isolated using the Ultraspec reagent (Biotecx Laboratories, Houston, TX) and dissolved in nuclease-free water. Samples of RNA (10 μg) were separated by electrophoresis in 1% agarose under denaturing conditions (formaldehyde), transferred to Nytran Plus membrane (Schleicher & Schuell, Keene, NH), UV cross-linked, and probed with p40^phoxcDNA labeled using the High Prime DNA labeling kit, (Roche Molecular Biochemicals) and [α-³²P]-dCTP (DuPont-NEN). Following autoradiography at −70°C, the membrane was stripped and probed with³²P-labeled cDNA for human acidic ribosomal phosphoprotein 36B4 as a loading control.

Previous studies have shown that humanp40^phox mRNA is expressed almost exclusively in hematopoietic cells that exhibit NADPH oxidase activity.36,37 To identify the sites of promoter activity in the p40^phox gene, regions of the 5′-flanking sequence were prepared by PCR, cloned into the luciferase reporter vector pGL3-Basic, and assayed by transient transfection and expression in the promyelocytic HL-60 cell line. The highest level of activity (approximately 37-fold increase over pGL3-Basic vector control) was observed with the construct pGL3-p40-106 (Figure1), which contained the first 106 nucleotides upstream of the p40^phoxtranscription start site.36 Constructs extending further upstream exhibited progressively lower luciferase activity, suggesting that the minimal p40^phox promoter lies within the first 106 nucleotides of the 5′-flanking sequence of thep40^phox gene and that negative regulatory elements may be present in upstream regions. Other myeloid cell lines tested (THP-1, U937, and PLB985) showed similar results (data not shown).

Inspection of the p40^phox promoter for potential regulatory elements identified several core-binding sequences (GAGGAA) for the PU.1 transcription factor. Applying criteria for optimal flanking nucleotides that we identified previously,46 3 of these PU.1 sites (PU.1a, −95 to −100; PU.1b, −65 to −60; and PU.1c, +80 to +85; Figure2A) were considered to have a high probability of binding PU.1. To determine whether these sites bound PU.1, electrophoretic mobility shift assay (EMSA) was carried out using³²P-labeled double-stranded oligonucleotides corresponding to these regions (Figure 2B) and nuclear extracts of HL-60 cells. Several DNA–protein complexes were formed between the PU.1 probes and HL-60 nuclear extracts (Figure 2C). Effective competition with excess unlabeled wild-type, but not mutated, DNA probes demonstrated that these bands were specific. The addition of specific antibody to PU.1 resulted in a dramatic decrease in band intensity and in the appearance of new supershifted bands, providing clear evidence that the complexes contained PU.1 protein. Comparison with previous studies suggested that the major bands of slower mobility contained intact PU.1 protein, whereas the faster-migrating bands were probably formed by PU.1 degradation products.46 

To investigate the role of these 3p40^phox PU.1 binding sites on promoter activity, a series of mutation and deletion promoter–luciferase constructs were prepared in which the 3 PU.1 sites were deleted or mutated individually or in combination. These constructs were tested as before by transient expression in HL-60 cells. Mutation of PU.1a, PU.1b, or PU.1c in the −1599 bpp40^phox promoter construct reduced activity by 82%, 63%, and 18%, respectively (Figure3A). Similar results were observed with the deletion series (Figure 3B). The p40^phoxpromoter activity in the −1599 bp construct was nearly abolished in the HL-60 cells when all 3 binding sites were mutated, indicating the essential requirement for PU.1 in this cell type (Figure 3B). Because PU.1 and p40^phox are also expressed in B lymphocytes, we investigated the roles of the PU.1 sites on p40^phox promoter activity in this cell type using the Raji human B-cell line. The PU.1 sites in thep40^phox promoter bound to PU.1 protein in Raji nuclear extracts (Figure 4A). Furthermore, the promoter was active in these cells, though at levels lower than those observed in the HL-60 cells (Figure 4B). More important, however, the promoter function was dependent on the presence of intact PU.1a and PU.1b sites (Figure 4B; PU.1c not tested). To investigate further the requirement for the PU.1 sites inp40^phox promoter activity, the mutated constructs were also analyzed in the nonmyeloid HeLa cell line, which does not express PU.1. Although the activity of the wild type −1599 bpp40^phox promoter construct was high in HeLa (Figure 4C), this activity was not affected by mutation of the PU.1 sites. The relatively high basal activity seen in HeLa cells with these constructs was surprising and might have been mediated by transcription factors present in these cells that bind the −1599p40^phox promoter construct at sites distinct from the PU.1 elements. These sites might normally be repressed in nonmyeloid tissues in vivo through elements in the gene not included in the −1599 p40^phoxconstruct. Such factors would be absent, functionally inactive, or PU.1-dependent in HL-60 and Raji cells because in both cases, mutation of the PU.1 sites was sufficient to eliminate most promoter activity.

Our previous studies indicated a correlation between the transactivation activity of PU.1 sites from a number of promoters and their affinity for the PU.1 protein.46 Thus, we hypothesized that the different levels of transactivation activity of the PU.1 sites in the p40^phox promoter might be attributed in part to their relative PU.1 binding avidity. To investigate this possibility, the binding affinities of the 3 sites for PU.1 present in HL-60 nuclear extracts or in vitro synthesized PU.1 were estimated by 2 methods. First, increasing amounts of each labeledp40^phox PU.1 oligonucleotide probe were incubated with a fixed amount of PU.1 protein (Figure5A). Second, the ability of unlabeledp40^phox PU.1 oligonucleotide probes to compete for binding with the well-characterizedp47^phox PU.1 site41 46 was determined (Figure 5B). Semiquantitative analysis of these binding studies suggested that the binding avidity of the PU.1a site is 2- to 4-fold greater than that of the PU.1b site (compare lane 2 with lanes 8 and 9 in Figure 5A and lane 2 with lane 6 in Figure 5B). The binding avidity of PU.1b is, in turn, 3- to 8-fold greater than that of the PU.1c site (compare lane 7 with lane 15 in Figure 5A and lane 5 with lane 9 in Figure 5B). Combining these EMSA results with the data obtained by deletion and mutation transfection studies indicated a direct relationship between the avidity with which these sites bind PU.1 and their capacity to dictate reporter gene transcription (PU.1a is greater than PU.1b is greater than PU.1c). However, the decrease in promoter activity on deletion of the PU.1c site was 20% when the other 2 PU.1 binding sites were intact, but it was 36% when the PU.1a site was deleted and 45% when the PU.1a and PU.1b sites were deleted (data not shown), suggesting that compensation may take place when one or another of the PU.1 sites is inactivated.

To determine whether PU.1 protein binds thep40^phox promoter at the identified PU.1 sites in vivo, we performed ChIP analysis on HL-60 cells using a specific polyclonal antibody to PU.1. Following immunoprecipitation and reversal of cross-linking, the DNA was purified and used as a template for PCR amplification using primers that encompass the region of thep40^phox promoter containing the 3 PU.1 binding sites. As a negative control, we carried out immunoprecipitation with antibody to c-Jun, consensus binding sites for which are absent from this region of thep40^phox promoter. Thep40^phox primers produced an amplicon of predicted size from the total input DNA and the anti–PU.1-precipitated DNA, but not from anti–c-Jun–precipitated DNA (Figure6). Furthermore, PCR primers for an unrelated gene, CCR5, generated products only from the total input DNA, confirming the specificity of the assay. The CCR5primers were designed to amplify a region of the CCR5gene that does not contain PU.1 or c-Jun binding sites. These data indicate that endogenous PU.1 protein in HL-60 cells binds thep40^phox promoter in the region containing the PU.1 sites and thus may promote transcription of the gene in vivo.

To provide further evidence that PU.1 is critical forp40^phox promoter activity, thep40^phox luciferase construct pGL3-p40-106 or pGL-3-Basic control were transfected into HL-60 cells with or without cotransfection of either the PU.1 expression plasmid pJ6-mPU.1 or the dominant-negative mutant plasmid pJ6-NN, in which the transactivation domain is deleted. Overexpression of PU.1 increased the activity of the p40^phox promoter by approximately 2.5-fold (Figure 7), whereas expression of the dominant-negative mutant PU.1 decreased activity by approximately 50%, presumably because of competition with endogenous PU.1. That the transactivation was dependent on the binding of PU.1 to the putative PU.1 sites in the proximal portion of thep40^phox promoter was confirmed in control experiments using the pGL3-p40-106mt construct, in which all 3 PU.1 sites were mutated. No effect of wild-type or mutant PU.1 was observed with this mutant reporter construct (Figure 7).

HL-60 cells can be induced to differentiate toward a granulocytic phenotype by treatment with dimethyl sulfoxide (DMSO).47Components of the phagocyte NADPH oxidase system are induced by DMSO, and enzymatic activity can be detected within 1 to 2 days of treatment48-50 Althoughp40^phox is not an essential component of the functional NADPH oxidase, it is expressed predominantly in phagocytic cells, complexed with the essential oxidase components p47^phox and p67^phox, and, in all probability, has a role in enzyme activation and regulation.29 32 To investigate the influence of granulocytic differentiation on p40^phoxgene expression, we first examined the induction ofp40^phox mRNA during DMSO treatment. Total RNA was isolated from DMSO-treated HL-60 cells at daily intervals, and Northern blot analysis was carried out using a cDNA probe specific for human p40^phox. Levels of p40^phox mRNA increased approximately 8-fold within 1 day of treatment (Figure8A). Subsequently, transcript levels decreased but were maintained at approximately 4-fold over controls for 2 to 3 days. To determine whether increased gene transcription contributed to this observed increase inp40^phox gene expression, HL-60 cells were first transfected with the pGL-3–p40-106 reporter gene construct, then treated with DMSO for 38 hours, and luciferase reporter activity was measured. DMSO increased reporter gene activity by 3- to 4-fold over the untreated control (Figure 8B). However, when we transfected the construct in which all 3 PU.1 sites were mutated, only a small increase in luciferase activity was seen following DMSO treatment. This suggests that the increase in transcriptional activity induced by DMSO in the wild-type promoter was mediated through increased PU.1 activity. These data were supported by EMSA and immunoblot analysis because nuclear extracts from HL-60 cells taken 38 hours after DMSO treatment showed increased PU.1 binding activity (Figure9A) and increased PU.1 protein using specific anti-PU.1 antibodies (Figure 9B).

PU.1, the most divergent member of the ets family of transcription factors, is important in the early development of multiple hematopoietic progenitors. Targeted disruption of the PU.1 locus results in multilineage defects that affect B and T lymphocytes, monocytes, osteoclasts, alveolar macrophages, and neutrophils.39,40 The PU.1 transcription factor is expressed specifically in hematopoietic tissues, particularly in cells of granulocytic, monocytic, and B lymphoid lineages.42,51-53 These patterns of expression are shared with the components of the phagocyte NADPH oxidase.1-3Moreover, previous studies have shown that the components of the phagocytic respiratory burst oxidase are transcriptionally regulated by PU.1.41,54-57 For example, expression of the large subunit of the membrane-bound cytochrome b₅₅₈, gp91^phox is controlled by PU.1, along with other transcription factors.54-57 Of particular note, the PU.1-binding site is critical for function of the promoter of thegp91^phox (CYBB) gene, and point mutations in this site lead to the CGD clinical phenotype.55,56 In the case of the cytosolic oxidase component p47^phox, we have shown that its myeloid-specific expression is regulated primarily through a single essential cis element that binds PU.1 with high avidity.41 In addition, we and others have reported that p67^phox transcription is regulated by a complex array of transcription factors that includes PU.1.57-59Adding to the body of evidence for a key role of PU.1 in regulation of NADPH oxidase expression are our current data that the cytosolic oxidase component p40^phox is also regulated by PU.1. We have shown that a unique combination of 3 PU.1-binding elements, 2 in the proximal promoter and one in the 5′ UTR of the gene, comprise the major regulatory components ofp40^phox gene transcription in the HL-60 myeloid and Raji B-cell lines. Moreover, we have demonstrated that the 3 PU.1-binding sites also mediate increased expression of thep40^phox gene observed during DMSO-induced granulocytic differentiation of HL-60 cells.

Recent studies indicate that regulation by multiple PU.1 promoter elements may not be unusual for myeloid-restricted genes. Two functional PU.1 sites have been identified in the promoter of the myeloid-specific CD18 gene60 and, interestingly, in the promoter of the PU.1 gene itself.61The PU.1-binding sites in the PU.1 promoter thus appear to act as a pathway for autoregulation, a potentially important concept for lineage development of myeloid cells. A recent report has indicated that a low level of expression of PU.1 in hematopoietic progenitors is associated with the development of B cells, whereas a high level of expression blocks B-cell development and promotes macrophage differentiation.62 Nevertheless, the specific functional implications of the presence of multiple functional PU.1 sites in a promoter are yet to be determined. It is possible that in some genes they could act in a synergistic or cooperative manner, allowing a relatively strong transcriptional response in the presence of low activity of the PU.1 transcription factor, a situation that might be found in the early stages of myeloid development. This may well be the case with the p40^phox gene. Transcripts for p40^phox can be readily identified in undifferentiated myeloid cell lines by Northern blot analysis,36 whereas the mRNA forp47^phox, p67^phox, andgp91^phox is generally more difficult to detect.36,50,63 64 

We showed previously that the avidity of binding of PU.1 by a particular cis element correlates with its capacity to dictate reporter gene transcription.46 The results of our analyses of the 3 PU.1 sites in thep40^phox promoter support this earlier observation. The affinity of binding of PU.1 to thep40^phox PU.1 oligonucleotide probes in the gel mobility shift assay and the transacting activity mediated by each site in the transfection studies were of the same rank order (PU.1a is greater than PU.1b is greater than PU.1c). The relatively high activity of the PU.1a site is probably caused by the presence of optimal (AT-rich) sequences flanking the 5′ end of the core GAGGAA sequence.46 Interestingly, these sequences are not found in the weakly acting PU.1c site. The proximity of the 3 PU.1-binding elements to each other and their different activities suggest that together they produce a synergistic activation ofp40^phox gene transcription. However, our deletion–mutation studies indicated that the relative contribution to promoter activity of each of the 3 PU.1 sites was not reduced when the other sites were functionally lost. Therefore, at least in resting HL-60 cells, it appears more likely that each PU.1 site acts independently of the others.

Transfection of the −1599 bp p40^phoxpromoter construct into the nonmyeloid HeLa cell line resulted in moderate levels of reporter gene activity. Consistent with the absence of PU.1 protein in these cells, this activity was not affected by mutation of the PU.1 sites in the promoter construct. These data suggest that nonmyeloid-specific factors are capable of binding and transactivating the −1599 bp p40^phoxpromoter construct in locations other than the PU.1 sites we have described here. However, if present in myeloid cells and B lymphocytes, these factors must be dependent on PU.1 binding because mutation of the PU.1 sites alone markedly reduced promoter activity in these cells. Given that the expression of p40^phox is restricted to myeloid cells and B lymphocytes, transactivation of the gene by these putative factors in nonmyeloid tissues must be inhibited in some way. However, a previous report suggests that some myeloid-specific promoter–reporter constructs may show activity in HeLa cells, as opposed to other nonmyeloid cell types.65 

There is increasing evidence that PU.1 can act in association with accessory factors to transactivate genes. Early studies showed an essential functional requirement for the factor NF-EM5 (PIP) to act in conjunction with PU.1 to transactivate the immunoglobulin kappa gene.66 The recruitment of NF-EM5 to the PU.1–DNA complex was dependent on the activation of PU.1 by phosphorylation of serine 148, possibly by casein kinase II. In the immunoglobulin kappa gene, nonphosphorylated PU.1 was able to bind its cognate site but could not stimulate promoter activity. It is possible that the binding of PU.1 to the p40^phox gene recruits such accessory factors into a ternary complex that in turn dictates promoter activity. Our data do not directly support such a model because the pattern of complexes formed on EMSA with thep40^phox PU.1 probes was similar for HL-60 nuclear extracts and in vitro–synthesized PU.1 protein. However, the recruitment of accessory factors may require additional nucleotide sequences that extend beyond those used for the probes.

The induction of granulocytic differentiation in HL-60 cells with DMSO led to an increase in the net cellular levels ofp40^phox mRNA. DMSO treatment also produced an increase in the total cellular levels of PU.1, particularly in the slower-migrating form of PU.1, which most likely corresponds to a hyper-phosphorylated form of the protein.67,68 These data are consistent with earlier studies using enriched human CD34⁺ progenitor cells in which the induction of myeloid differentiation with granulocyte macrophage–colony-stimulating factor resulted in an increase in PU.1 mRNA and protein.69 70 The increase in PU.1 protein in the HL-60 cells was accompanied by an increase in the levels of PU.1 binding to DNA and PU.1-mediatedp40^phox promoter activity. Taken together with our functional mutation studies, these data suggest that the increased p40^phox gene transcription observed during maturation and differentiation is mediated primarily by the 3 identified PU.1 sites. This does not preclude the contribution of additional accessory or coactivator factors, perhaps recruited through the activation and phosphorylation of PU.1. However, these putative factors would have to function through the PU.1 elements because mutation of these sites reduced the promoter activity in the HL-60 cells to near-background levels.

We thank Dr Michael J. Klemsz for providing PU.1 expression plasmids and Dr Long Wang for assistance with graphics and statistical analyses.