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Down-regulation of interferon regulatory factor 4 gene expression in leukemic cells due to hypermethylation of CpG motifs in the promoter region
Although the bcr-abl translocation has been shown to be the causative genetic aberration in chronic myeloid leukemia (CML), there is mounting evidence that the deregulation of other genes, such as the transcription factor interferon regulatory factor 4 (IRF-4), is also implicated in the pathogenesis of CML. Promoter methylation of CpG target sites or direct deletions/insertions of genes are mechanisms of a reversible or permanent silencing of gene expression, respectively. Therefore, we investigated whether IRF-4 promoter methylation or mutation may be involved in the regulation of IRF-4 expression in leukemia cells. Whereas promoter mutations or structural rearrangements could be excluded as a cause of altered IRF-4 expression in hematopoietic cells, the IRF-4 promoter methylation status was found to significantly influence IRF-4 transcription. First, treatment of IRF-4-negative lymphoid, myeloid and monocytic cell lines with the methylation-inhibitor 5-aza-2-deoxycytidine resulted in a time- and concentration-dependent increase of IRF-4 mRNA and protein levels. Second, using a restriction-PCR-assay and bisulfite-sequencing we identified specifically methylated CpG sites in IRF-4-negative but not in IRF-4-positive cells. Third, we clearly determined promoter methylation as a mechanism for IRF-4 down-regulation via reporter gene assays, but did not detect an association of methylational status and mRNA expression of DNA methyltransferases or methyl-CpG-binding proteins. Together, these data suggest CpG site-specific IRF-4 promoter methylation as a putative mechanism of down-regulated IRF-4 expression in leukemia.

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INTRODUCTION
Chronic myeloid leukemia (CML) is a clonal myeloproliferative disorder with a typical three phased course (chronic, accelerated and blastic phase) reflecting the loss of differentiation and malignant progress which inevitably leads to death after the blastic phase (1,2). The hallmark genetic aberration of CML is a reciprocal chromosomal translocation t(9;22) leading to expression of a bcr-abl fusion gene, an aberrant activated tyrosine kinase (2). Treatment with interferon alpha (IFN-alpha) prolongs survival of CML patients and is associated with a complete cytogenetic response in 5-33% of CML patients (1,2). Recently, we described an impaired expression of the interferon regulatory factor 4 (IRF-4) in CML, correlating with poor response to IFN-alpha treatment (3). The cause of the silencing of IRF-4 level remained unclear.
Interferon regulatory factors (IRFs) are a family of transcriptional regulators defined by a characteristic homology in their DNA-binding domain. They play an important role in the regulation of various genes (such as IFNs, interleukins, MHC class I/II), apoptosis and differentiation/maturation (4-6). IRF-4 (ICSAT/Pip/MUM1/LSIRF) is one member with very restricted expression pattern: Predominately B- and activated T-lymphocytes are IRF-4 positive (7-11). In contrast to other IRFs, expression of IRF-4 cannot be induced by IFNs, but by antigen stimulation, crosslinking of T- or B-cell receptors or phorbol-myristate-acetate (10,11). Consistent with the restriction of expression to immunocompetent cells, mice with deletion of IRF-4 failed to develop mature and functionally active B- and T-lymphocytes (12), and the impaired expression of IRF-4 in CML was predominately found in T-cells (3). These data suggest a crucial role for IRF-4 in the function of immune cells.
Methylation of dinucleotide cytosine-guanosine motifs (CpG), especially in CpG islands located in promoter regions, is one of the mechanisms of gene regulation in mammals and a common event of gene silencing in human neoplasias (13,14). As opposed to normal cells, hypermethylation of CpG islands is a frequently observed phenomenon in every cancer type. De novo DNA methylation of genes such as cell cycle, DNA repair, apoptosis and tumor suppressor genes is therefore thought to be involved in tumorigenesis (15-17). Examples for such aberrated genes are MGMT, DAPK, p14ARF, p15INK4b, p16INK4a, BRCA1, CDH13 and APAF-1 (17-19). In CML, methylation is known to regulate expression of the c-abl, the bcr gene and others (20-23), and the extent of methylation in the c-abl promoter has been shown to be associated with advanced disease (24). Hypermethylation due to overexpression of DNA methyltransferases (DNMTs) remains one possible explanation for de novo methylation in tumorigenesis. Recently, DNMTs have been shown to be up-regulated in hematopoietic malignancies (25). Methyl-CpG-binding proteins (MBPs) are thought to inhibit the binding of transcriptional factors to the promoter and are therefore discussed as one mechanism of transcription inhibition by hypermethylation (26).
In this work, we studied mechanisms of IRF-4 gene expression silencing in leukemic cells. We analyzed the IRF-4 promoter region for genetic aberrations and methylational status in IRF-4-positive and -negative hematopoietic cells.

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Cell lines
K-562, Jurkat and U-937 were obtained from the ATCC (American Type Culture Collection, Rockville, USA) and EM-2, LAMA-84, CML-T1, BV-173, SD-1 and RPMI-8226 from the DSMZ (Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH, Braunschweig, Germany). All cell lines, except BV-173, SD-1 and RPMI-8226, were IRF-4-negative.

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Cell culture and stimulation
All cell lines were maintained at 5% CO2 in RPMI 1640 medium with 1% glutamine (Gibco/BRL Eggenstein, Germany) supplemented with 10% fetal calf serum (Gibco/BRL), 1% penicillin/streptomycin (Biochrom, Berlin, Germany). When indicated, cells were treated with 5-aza-2-deoxycytidine (AzadC) or 5-azacytidine (AzaC) (Sigma, Taufkirchen, Germany) for different time periods. Owing to their chemical instability fresh substances were re-added every 24 h.


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Sequencing of the IRF-4 promoter
For analysis of the IRF-4 promoter region for permanent aberrations such as insertions/deletions or mutation, we PCR-amplified two fragments from genomic DNA, which was extracted from depicted cell lines with a commercial kit (Qiagen, Hilde, Germany) as recommended. The primers were 1-forward: 5'-TTGAGATGGAGTCTTGCTCTGT-3', 1-reverse: 5'-CCAGGACCTCAGGAGGCCAGTCA-3'; 2-forward: 5'-AGCGGTGAAACTGAGAGTGCGAGGT-3', 2-reverse: 5'-GCCACATCGCTGCAGTTTAG-3'. The products were cloned with the 'TOPO TA cloning kit' (Invitrogen, Groningen, The Netherlands). After bacterial amplification of the cloned PCR fragments by standard procedures, at least three clones from each sample were sequenced with an automated sequencer (ABI Prism 377, Applied Bio-systems, Foster City, USA) as recommended by the manufacturer.


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Expression analysis
To analyze the IRF-4 transcriptional level, RNA was extracted from cells using the commercial RNAzol-kit (Paesel, Frankfurt, Germany). An aliquot of 1 mug total RNA was used for cDNA synthesis as described previously (27). RNA expression analysis for IRF-4 and the reference gene beta-actin was carried out by semi-quantitative PCR as described previously (3,27). PCR products were verified by automated sequencing. PCR primers and conditions for expression analysis of DNMT or MBP (DNMT1 DNMT3A, DNMT3B, MeCP, MBD1, MBD2 and MBD4) were published elsewhere (28).
For analysis of IRF-4 protein expression, a standard immunoblotting assay was performed as described previously (29). Briefly, protein lysates were generated by incubating 1 x 106 cells in 100 microl RIPA buffer (1% NP-40, 0.5% sodiumdesoxycholate, 0.1% SDS, 100 microg/ml phenylmethylsulfonyl fluoride, 10 microl/ml protease-inhibitory-mix, 1 micromol/ml sodiumorthovanadate in phosphate-buffered saline) for 30 min on ice. After centrifugation, protein concentration of the supernatant was determined by BCA-method (Pierce, Rockford, IL) as recommended. Protein lysates (70-100 microg) were electrophoresed on polyacrylamide gels and transferred to a PVDF-membrane (Immobilon P, 0.45 microm; Millipore, Eschborn, Germany). Membranes were blocked with 2.5% blocking reagent (Boehringer Mannheim, Germany) in TBST buffer (4.44 g/l Tris-HCL, 2.65 g/l TrisOH, 8.07 g/l NaCl, 0.2 g/l KCl and 500 microl/l Tween-20 in H2O) and subsequently incubated with primary antibody as indicated and horseradish peroxidase-conjugated secondary antibody, anti-mouse or anti-goat IgG (DAKO, Hamburg, Germany), respectively. The membranes were then developed with an ECL detection kit (Amersham Pharmacia Biotech, Freiburg, Germany). The primary antibodies were goat anti-IRF-4/ICSAT (M-17) (Santa Cruz Biotechnology, Santa Cruz, CA) and mouse anti-beta-actin (AC-74) (Sigma).

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Methylation-specific restriction-PCR-assay
DNA was extracted with a commercial kit (Qiagen) as recommended. Since the restriction ability of several endonucleases is inhibited by methylation of their target sequence, we used methylation-sensitive enzymes HpaII and HaeII-isochizomer Bsp143II and Bsh1236I (MBI Fermentas, St Leon-Rot, Germany) (20,24). As control the methylation-resistant enzyme MspI and an enzyme with no recognition site in the target promoter, EcoRI, were used. DNA (0.8 microg) was digested by 40 U the respective enzyme for 6 h and, to ensure complete cleavage, additional 20 U for 16 h. Thereafter 100 ng of digested DNA was used to a PCR amplification of two fragments (F1 and F2) spanning part of the IRF-4 promoter (30) (GenBank U52683; see Figure 3A). The sequences of the primers were F1-forward: 5'-TTGAGATGGAGTCTTGCTCTGT-3', F1-reverse: ATCACTTCCAGACTTCAGTTCACCT-3' (341 bp); F2-forward: 5'-AAGGTGAACTGAAGTCTGGAAGTGA-3', F2-reverse: 5'-CCAGGACCTCAGGAGGCCAGTCA-3' (474 bp). The PCR conditions were described elsewhere (3). PCR was performed with an annealing temperature of 62degreesC and 35 cycles. When DNA was methylated at specific sites, the sensitive enzymes were not able to digest the DNA and amplification took place; in case of no methylation, DNA was digested and no product was generated. The PCR products were electrophoresed on a 3% agarose gel, were stained with ethidium bromide and photographed. PCR products were verified by automated sequencing.


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Bisulfite treatment
DNA was extracted as described above. Bisulfite treatment of DNA, leading to conversion of unmethylated cytosine to uracil residues and no change of methylated cytosine residues, was performed as described as follows. Briefly, 1 microg of DNA and 2 microg of poly(dA-dT)(poly(dA-dT) copolymers (Amersham Pharmacia Biotech) were denaturated for 20 min at 42degreesC in 0.3 M NaOH in a volume of 50 microl. Fresh solutions of 30 microl of 10 mM hydrochinon (Sigma) and 530 microl of 3 M sodium bisulfite (pH 5.0; Sigma) were added, the solution was gently mixed, overlayed with mineral oil and incubated in the dark for 12-13 h at 50degreesC. The aqueous phase was recovered using the 'Wizard DNA clean-up system' (Promega, Mannheim, Germany). The purified DNA was subsequently mixed with 1 M NaOH to a final concentration of 0.3 M and incubated for 20 min at 37degreesC to ensure complete desulfonisation. DNA was ethanol precipitated in the presence of 1/10 vol of 3 M sodium acetate, washed with 70% ethanol and resuspended in 50 microl H2O. Subsequent PCR amplification of 4 microl bisulfite-treated DNA was used for cloning of two fragments of the IRF-4 promoter (BS-I and BS-II) into pCR2.1 vector with the 'TOPO TA cloning kit' (Invitrogen) (see Figure 3A). The primers used for PCR amplification of the BS-I and BS-II fragments contain the putative altered sequence of the sense strand due to bisulfite treatment (converted cytosine residues are written in bold letters): BS-I-forward 5'-TATTTGGATTTTTAGGGAGTTTTTTTT-3', BS-I-reverse 5'-ACCCAACTCCCTTAAACTATTAAACT-3' (187 bp); BS-II-forward 5'-AGTTTAATAGTTTAAGGGAGTTGGGT-3', BS-II-reverse 5'-CTCACCCTAAACTCAAAACTAAAAAC-3' (674 bp). After bacterial amplification of the cloned PCR fragments by standard procedures, eight clones from each sample were sequenced with an automated sequencer (ABI Prism 377, Applied Biosystems).


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In vitro methylation and reporter gene assays
The IRF-4 promoter-reporter gene construct was generously provided by J.Hiscott (31). Constructs were methylated in vitro with CpG Methylase (M.Sss I) as recommended by the manufacturer (NE Biolabs) and complete methylation was checked via restriction analysis (Figure 5A). Reporter gene assays using the dual luciferase assay (Promega) were performed similar to previous reports (29). Briefly, 5 nM of the reporter construct and the transfection control construct expressing the renilla luciferase gene were transiently co-expressed via electroporation. The control construct served as an internal reference for transfection efficiency. Forty-eight hours after transfection, luciferase activity was measured with a LB 96 P microlumat (EG&G Berthold, Bad Wildbad, Germany). IRF-4 promoter activation was quantified as a ratio of measured firefly light units (flu) relative to renilla (rlu). Each experiment was carried out at least three times.

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Absence of IRF-4 expression in leukemia cells is not due to promoter alterations
We have previously demonstrated a lack of IRF-4 expression in leukemia patients and specifically in CML T-cells (3). Here, we demonstrate the absence of IRF-4 expression in various hematopoietic cell lines, such as Jurkat, a T-cell leukemia, CML-T1, a bcr-abl-positive T-cell line, K-562, a bcr-abl-positve erythroleukemia, U-937, a monocytic leukemia, EM-2 and LAMA-84, bcr-abl-positve myeloid leukemia, but not in SD-1, a bcr-abl-positive acute lymphoblastic leukemia (pre B-ALL), RPMI-8226, a multiple myeloma and BV-173, a bcr-abl-positive B-cell line (Figures 1A and 5D). After sequencing of the IRF-4 promoter, it could be excluded that absence of IRF-4 expression in any of the above cell lines was due to genetic aberrations. However, 2 bp changes (nucleotide -1081, T-->C and -1068, A-->C) could be detected in both the IRF-4-positive BV-173 and the IRF-4-negative LAMA-84, EM-2 and K-562 (Figure 1B). At position -116 an A-->C substitution was found in EM-2, K-562 and CML-T1, whereas Jurkat, BV-173 and SD-1 exhibited a mixed A/C sequence and U-937, LAMA-84 and RPMI-8226 no substitution at all (Figure 1B). Consequently, these alterations are unlikely to affect IRF-4 expression.

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Increase of IRF-4 expression in hematopoietic cells after demethylating treatment
We next analyzed whether promoter methylation could be responsible for down-regulation of IRF-4 expression. A region including exon1 in the IRF-4 promoter exhibited a large number of CpG-rich sequences (Figure 3A). Several chemical substances such as 5-aza-2-deoxycytidine (AzadC) or 5-azacytidine (AzaC) inhibit de novo and maintenance methylation, and thus can be used to discern promoter methylation (32,33). We used AzadC to generate unmethylated DNA. A 72 h AzadC-treatment resulted in a concentration-dependent activation of IRF-4 transcription in Jurkat and CML-T1 T-cells as well as in U-937, K-562 and EM-2 cell lines (Figure 2A). IRF-4 transcription was induced in a time-dependent manner and was observed as early as 24 h after treatment with AzadC and increased over time until 72 h (Figure 2B). Time and strength of the appearance of IRF-4 transcripts varied among cell lines, i.e. CML-T1 responded strongest to AzadC-treatment (data not shown). In line with this, AzadC-treatment of CML-T1 and LAMA-84 cells also translated in an induction of IRF-4 protein expression (Figure 2C). Accordingly, treatment of the IRF-4-positive cell line BV-173, SD-1 and RPMI-8226 with AzadC had no effect on IRF-4 expression (Figure 2D). There was no difference in the effects of AzaC versus AzadC, as both increased the IRF-4 mRNA level in CML-T1 cells as well (data not shown). This implied that promoter methylation may control IRF-4 expression, but an alternative explanation may be activation of positive transcriptional regulators of IRF-4 by AzadC (or AzaC).

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Methylation-sensitive enzymes do not cut specific sites in the IRF-4 promoter in hematopoietic cells
To further investigate promoter methylation as a regulatory mechanism of IRF-4 gene expression, restriction-PCR-assays were performed (20,24), where only methylated DNA would not be cut enabling subsequent PCR amplification and vice versa. Genomic DNA from leukemic cells Jurkat, CML-T1, U-937, K-562, EM-2 and BV-173 was digested with the methylation-sensitive enzymes HpaII, Bsh1236I and HaeII-isochizomer Bsp143II. EcoRI, which has no recognition site within the IRF-4 promoter, and the methylation-resistant enzyme MspI served as controls. Two separate amplification reactions were performed, generating two fragments, F1 and F2 (Figure 3A). After digestion with HpaII and Bsp143II a sufficient PCR amplification of F1 and F2 was detected in DNA from IRF-4-negative Jurkat, CML-T1, U-937, K-562 and EM-2 cells, suggesting a promoter methylation (and restriction protection) at the respective recognition sites (Figure 3B and C). Notably, in IRF-4-positive SD-1 cells digestion with the methylation-sensitive enzymes completely inhibited amplification of F1 and F2. In IRF-4-positive BV-173 cells a HpaII, but not a Bsh1236I digestion, significantly reduced the amplifiable DNA message of F2 (Figure 3C), whereas amplification of F1 was not affected (Figure 3B). This implied that IRF-4 transcription in SD-1 and BV-173 cells is associated with less promoter methylation (in BV-173 especially at HpaII sites) as compared with the tested IRF-4-negative cells.

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Specific CpG sites in the IRF-4 promoter are methylated in hematopoietic cells
In order to exactly map the methylation sites within the IRF-4 promoter, we treated DNA of Jurkat, CML-T1, U-937, K-562 and EM-2 cells as well as of SD-1, RPMI-8226 and BV-173 control cells with bisulfite, which chemically converts unmethylated cytosine to uracil, whereas it has no effect on methylated cytosine, i.e. in CpG (34). This technique is especially useful for detection of unknown methylation patterns. PCR amplification, cloning and sequencing of the bisulfite-treated DNA showed a specific methylation pattern of the analyzed 62 CpG sites in all cell lines (Figure 4 and Table 1). In general, the methylational status ranged from one cell line with a nearly non-methylated IRF-4 promoter (SD-1, IRF-4-positive) to a completely methylated IRF-4 promoter in CML-T1 (IRF-4-negative). Interestingly, the percentage of CpG methylation in the IRF-4 promoter from IRF-4-positive cells was very low (mean 24%) as compared with IRF-4-negative cells (mean 94%) (Figure 4A and Table 1). A 5'-region (R1) with 13 hypermethylated CpG sites (mean number of methylated clones 5.5 of 8 with 77% methylated CpGs) was found in most cells (except SD-1 and RPMI-8226) and a 3'-region (R3) of 6 hypomethylated CpG sites (mean number of methylated clones 1.7 of 8 with 33% methylated CpGs) was found in most cells (except CML-T1 and U-937) (Figure 4A and Table 1).
Intriguingly, a stretch of 13 CpG sites (#10-22; R2) was detected in between these regions, which were highly methylated in IRF-4-negative (mean number of methylated clones 7.1 of 8 with 89% methylated CpGs) but totally non-methylated in IRF-4-positive cells (Figure 4A and B). Furthermore, three CpG sites at the 5' end (#54, 56, 58) and two CpG motifs at the 3' end (#1, 2) showed this direct correlation between high methylation status and absence of IRF-4 expression. In addition, two CpG sites located in a NFkappaB (#48) and a SP1element (#45) are less methylated in IRF-4-positive than in IRF-4-negative cells (mean number of methylated clones: 1/8 versus 8/8). These results indicate the involvement of CpG methylation in the regulation of IRF-4 expression in leukemic cells.

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In vitro methylation of an IRF-4 promoter-reporter construct decreases its activity
To provide evidence for a direct effect of methylational status on IRF-4 promoter activity we performed reporter gene assays with IRF-4 promoter constructs before and after their in vitro methylation. A complete methylation of these constructs was checked via restriction assays with methylation-sensitive endonucleases (Figure 5A). Intriguingly, methylation of the IRF-4 promoter significantly decreased promoter activity in IRF-4-positive SD-1 cells by 85.0% (Figure 5B). The silencing effect of CpG methylation was not restricted to IRF-4-positive cells, since in vitro methylation led to a 92.9% abrogation of promoter activity in IRF-4-negative Jurkat cells (Figure 5C). In contrast, control methylation of a reporter construct with a different promoter (FasL) as well as an empty vector had no effect on the reporter activity (data not shown). These data proved a direct association between methylation and activity of the IRF-4 promoter.

positive

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negative

decreased

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mRNA expression of DNA methyltransferases and methyl-CpG-binding proteins may not be associated with IRF-4 promoter methylation
Since abundance of DNMT and MBP contribute to promoter regulation via methylation (25,26,28), we studied their mRNA expression to investigate a possible mechanism for the observed methylation differences in the IRF-4 promoter. To this end, we did not detect a significant difference in DNMT (DNMT1, DNMT3A and DNMT3B) or MBP (MBD1, MBD2, MBD4 and MeCP) mRNA expression between IRF-4-positive and -negative cells (Figure 5D). In fact, all analyzed cells had moderate to high mRNA levels of these tested DNMT/MBPs and differences in expression were not correlated with IRF-4 status. These results indicate a distinct cause of the methylation differences in IRF-4-positive and -negative cells rather than changes in the DNMT and MBP mRNA transcription.

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DISCUSSION
Many genetic lesions are known to influence gene expression of tumor suppressor genes. Whereas mutations and deletions or insertions have permanent effects, reversible mechanisms are gene methylation, or expression and activation of transcription factors, respectively. We studied a putative cause for absent IRF-4 expression in leukemia cells and first focused on genetic aberrations of the promoter. We observed no genetic alterations in the IRF-4 promoter, which can account for the lack of IRF-4 expression: The detected base pair changes at position -1081 (T-->C substitution), at position -1068 (A-->C substitution) and at position -116 (A-->C substitution) are unlikely responsible for absent IRF-4-expression since the first two mutations were found both in IRF-4-positive and -negative cells whereas the latter change was not detected consistently in all IRF-4-negative or -positive cells and may thus be a polymorphism. All three substitutions did not change any known putative transcription factor binding sites (30,31) and also do not affect any restriction sites or primer binding sites of the used assays. However, permanent genetic variations in the IRF-4 coding sequence, such as deletions or mutations resulting in stop-codons have not been excluded by sequence analysis. Since IRF-4 expression in cell lines and CML can be induced by demethylation and successful IFN-alpha therapy (3), respectively, the existence of such genetic aberrations seems unlikely.
We then investigated whether the previously described down-regulation of IRF-4 expression in human myeloid leukemias was due to a differential hypermethylation of the promoter, since the presented re-expression due to AzadC-treatment might also be a result of activation of positive transcriptional regulators of IRF-4. Methylation of CpG sites is a common mechanism of silencing genes in leukemia and has also been shown for another IRF, IRF-7 (35) and for PU.1 (36), an interacting partner of IRF-4. To elucidate the relevance of this mechanism for the regulation of IRF-4 expression, various leukemic cells were treated with demethylating agents and promoters were sequenced after bisulfite treatment. We found that IRF-4 expression could indeed be connected to the methylation status of distinct CpG motifs in the IRF-4 promoter. In Figure 4A, those CpG sites are shown (bottom line), whose hypermethylation may account for the absence of IRF-4 expression in the respective cells. One of them (#54) is adjacent to an identified regulatory element (NFkappaB-site), indicating a possible involvement of this site. At two further CpG sites (#48, 45) the methylation status in IRF-4-positive was lower than that of IRF-4-negative cells. These CpG sites are located in an NFkappaB and an SP1 element (31) and thus may also play a role in regulation of IRF-4 expression. It has been shown that NFkappaB elements play an important role in IRF-4 induction as IRF-4 expression depends on binding of the transactivator c-Rel to these elements in the IRF-4 promoter (31,37). Furthermore, methylation of the central CpG in the NFkappaB element inhibits binding of the NFkappaB protein complexes (38), promoting the significance of the observed methylation differences in IRF-4-positive and -negative cells.
Via in vitro methylation and reporter gene assays we could clearly appoint the silencing of the IRF-4 promoter to a methylation effect, which may thus be the mechanism of IRF-4 deregulation in vivo. One possible cause for the aberrant methylation in tumorigenesis is an increased level of DNMTs during the pathogenetic process. In colon, lung and hematologic malignancies, overexpression of DNMT1, a maintenance DNMT, has been detected (39-41). Furthermore, it has been shown that CML cells in the acute phase exhibited elevated levels of the three known DNMTs, while CML cells in chronic phase expressed normal levels of DNMTs if compared with normal bone marrow cells (25). Interestingly, a positive correlation between DNMT1 expression levels and hypermethylation of p15INK4b has been detected in AML (25). In this work, we did not detect significant mRNA expression differences of selected DNMT or MBP, making it an unlikely cause for the observed methylation and thus IRF-4 expression differences in leukemia cells.
The finding that IRF-4 expression is silenced by promoter hypermethylation might represent a mechanism that accounts for the previously observed loss of IRF-4 expression in CML. Indeed, several clinical trials with leukemia patients and patients with myelodysplastic syndromes demonstrated the potential clinical benefit of a treatment with demethylating agents (42-45).
The expression of another IRF, IFN consensus sequence binding protein (ICSBP/IRF-8), is impaired in myeloid leukemias especially CML (27,46,47). But in contrast to IRF-4, the loss of this IRF could not be reverted in ICSBP-negative cell lines (EM-2, CML-T1, K-562 and LAMA-84) by treatment with AzadC (Figure 6) and AzadC has no effect on ICSBP levels in ICSBP-positive U-937 cells (Figure 6). These data suggest a distinct regulatory mechanism for these two IRFs.
IRF-4, similar to many other classical tumor suppressor genes p15INK4b, p16INK4a or p53, may thus be a subject of alterations in the promoter methylation status leading to expression changes, which might contribute to the initiation and/or progression of cancer. Still, the obvious functional diversity of IRF-4 remains remarkable and cannot be fully explained by the IRF-4 promoter methylation status. For example, IRF-4 is primarily known for its oncogenic features. In multiple myeloma (MM) a translocation on chromosome 14q was reported to lead to a fusion gene of immunoglobulin heavy-chain (IgH) and IRF-4 resulting in a subsequent overexpression of IRF-4 (48,49). In addition, abundant IRF-4 expression was found to be a marker for various subsets of lymphomas, such as diffuse large B-cell lymphomas, primary effusion lymphoma, and marginal zone lymphoma, and adult T-cell leukemia (11,31,50-52). This draws a more complex picture of the role of IRF-4. Down-regulation of IRF-4 may promote leukemogenesis in myeloid cell context (3), which was recently confirmed in IRF-4-/- ICSBP-/- double knock-out mice (53), while IRF-4 up-regulation may induce a growth advantage in lymphomas or MM (48).
Taken together, our data suggest that IRF-4 promoter methylation regulates IRF-4 expression, and that aberrant expression of IRF-4 in certain types of leukemia may be a consequence of IRF-4 promoter hypermethylation.

interacting

binding

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binding

NFkappaB element

expression

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positive

negative

negative

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IFN consensus sequence binding protein

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absent

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CpG

binding

silencing

promoter

loss

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impaired

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loss

IFN consensus sequence binding protein

reverted

loss

Down-regulation

responsible

absent

induced

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IFN-alpha

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result

re-expression

account

absence

induction

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binding

overexpression

resulting

overexpression

overexpression

up-regulation

consequence

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change

transcription factor binding sites

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important role

NFkappaB elements

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Foxp3 Represses Retroviral Transcription by Targeting Both NF-kappaB and CREB Pathways
Forkhead box (Fox)/winged-helix transcription factors regulate multiple aspects of immune responsiveness and Foxp3 is recognized as an essential functional marker of regulatory T cells. Herein we describe downstream signaling pathways targeted by Foxp3 that may negatively impact retroviral pathogenesis. Overexpression of Foxp3 in HEK 293T and purified CD4+ T cells resulted in a dose-dependent and time-dependent decrease in basal levels of nuclear factor-kappaB (NF-kappaB) activation. Deletion of the carboxyl-terminal forkhead (FKH) domain, critical for nuclear localization and DNA-binding activity, abrogated the ability of Foxp3 to suppress NF-kappaB activity in HEK 293T cells, but not in Jurkat or primary human CD4+ T cells. We further demonstrate that Foxp3 suppressed the transcription of two human retroviral promoters (HIV-1 and human T cell lymphotropic virus type I [HTLV-I]) utilizing NF-kappaB-dependent and NF-kappaB-independent mechanisms. Examination of the latter identified the cAMP-responsive element binding protein (CREB) pathway as a target of Foxp3. Finally, comparison of the percent Foxp3+CD4+CD25+ T cells to the HTLV-I proviral load in HTLV-I-infected asymptomatic carriers and patients with HTLV-I-associated myelopathy/tropical spastic paraparesis suggested that high Foxp3 expression is associated with low proviral load and absence of disease. These results suggest an expanded role for Foxp3 in regulating NF-kappaB- and CREB-dependent cellular and viral gene expression.

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Synopsis
Over the past several years, mounting evidence has shown that immune tolerance in healthy individuals can be maintained by a population of T lymphocytes known as regulatory T cells (Tregs). As a component of this system, a protein known as Foxp3 has been shown to be absolutely required for the development and function of Tregs. While Foxp3 plays an important role in maintaining immune tolerance by blocking T cell proliferation and production of inflammatory proteins known as cytokines, little is known about the molecular mechanisms that are used by Foxp3 to accomplish these events. The present study expands our understanding of how Foxp3 maintains a check on inappropriate immune responses by demonstrating that Foxp3 can block activation of key inducible proteins such as nuclear factor kappaB (NF-kappaB) and cAMP-responsive element binding protein (CREB). Since NF-kappaB and CREB are integrally involved in controlling cell cycle progression, inflammatory cytokine production, and the replication of numerous viruses at the level of transcription, understanding the mechanisms by which Foxp3 functions to regulate cellular and viral gene expression may aid in the discovery of therapeutic approaches designed to rescue the expression and/or function of Foxp3, which have been found to be deficient in several autoimmune diseases and virus-induced disorders.

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Introduction
Immunological tolerance to self-antigens is the result of the deletion of self-reactive T lymphocytes in the thymus (central tolerance) and suppression of the activation of potentially self-reactive T lymphocytes in the periphery (peripheral tolerance) [1]. Suppression of pathogenic T cell responses is mediated by naturally arising CD4+CD25+ T regulatory cells (Tregs) [2,3]. Deficiencies in Treg development and function have been linked to the severe autoimmune disorder known as immune dysregulation, polyendocrinopathy, enteropathy, X-linked syndrome (IPEX) [4]. In addition, recent studies have provided strong evidence that dysregulation of Treg development and/or function may be a significant factor in the pathogenesis of several autoimmune disorders (e.g., multiple sclerosis [5], myasthenia gravis [6], and type 1 diabetes [7]) and virus-induced immunologic disorders (e.g., human T lymphotropic virus type I [HTLV-I]-associated myelopathy/tropical spastic paraparesis [HAM/TSP], and HIV-induced AIDS [8-10]).
The transcription factor Foxp3 is a 431-amino acid (48-kDa) protein expressed at very high levels in CD4+CD25hi T cells and has previously been shown to be absolutely critical for Treg development and function [11-14]. Foxp3 contains a proline-rich amino-terminal domain reported to function as a nuclear factor of activated T cells (NF-AT) and nuclear factor-kappaB (NF-kappaB) binding domain, a central region containing a zinc finger and leucine zipper potentially important for protein-protein interactions, and a carboxyl-terminal forkhead (FKH) domain required for nuclear localization and DNA-binding activity [14-16]. Functional inactivation of Foxp3 by genetic mutations affecting the Foxp3 coding region, as demonstrated in IPEX, or repression of Foxp3 expression by the HTLV-I-encoded transactivator protein Tax, as recently reported in patients with HAM/TSP, results in loss of regulatory activity in CD4+CD25hi T cells [4,8,17]. Although it is clear that Foxp3 regulates T cell proliferation and cytokine production, very little is known concerning the molecular mechanisms of Foxp3 function.
The first evidence to indicate how Foxp3 promotes the development and function of regulatory T cells came from a report by Ziegler and colleagues [16], which suggested that Foxp3 could inhibit transcriptional activation by physically interacting with forkhead binding sites located immediately adjacent to critical cis-acting NF-AT binding sites found in various cytokine promoters (e.g., IL-2 promoter). That study also demonstrated that Foxp3 could repress activation of a synthetic reporter vector containing an SV40 promoter and three tandem copies of a forkhead binding site. These results provided additional evidence suggesting that Foxp3 transcriptional repression was mediated by binding in a sequence-specific manner to promoters containing forkhead binding sites. A recent study by Bettelli and colleagues [15] further demonstrated that Foxp3 could inhibit NF-AT as well as NF-kappaB activation, although the mechanism of suppression was shown to involve direct protein-protein interactions between NF-AT or NF-kappaB and Foxp3 rather than binding of Foxp3 to promoter elements adjacent to cis-acting NF-AT or NF-kappaB sites. Collectively, these data suggested that Foxp3 may function as a transcriptional repressor, potentially through the formation of both DNA-protein and protein-protein interactions.
In the present study, we expanded upon these observations by defining additional requirements of Foxp3-mediated repression of NF-kappaB activation, and investigated whether Foxp3 could target additional signaling pathways by examining transcriptional activation of NF-kappaB-dependent and NF-kappaB-independent retroviral pathogens. The characterization of the molecular targets of Foxp3 and the mechanism(s) utilized by Foxp3 to support Treg development and function will aid in our understanding of the role Tregs play in the pathogenesis of human autoimmune disease.

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Foxp3 Suppresses NF-kappaB Dependent Transcriptional Activation
To ascertain the molecular mechanisms by which Foxp3 functions to promote the regulatory function of CD4+CD25hi T cells, we first confirmed the function of Foxp3 as a repressor of activation of NF-kappaB, previously implicated as a target of other forkhead/winged-helix family transcription factors (e.g., Foxj1 and Foxo3a) [19,20]. We analyzed the effect of Foxp3 overexpression on NF-kappaB activation in HEK 293T cells in dose-response and time course analyses. Transfection of HEK 293T cells with an NF-kappaB luciferase reporter vector in the presence or absence of increasing concentrations of a Foxp3 expression vector or a control vector (enhanced green fluorescent protein [EGFP]) was performed, and cells were harvested after 24 h to assay for luciferase activity and Foxp3 mRNA expression. Results indicated that as the concentration of Foxp3 transfected into cells increases (from 50 to 2,400 ng), the level of NF-kappaB activation decreases proportionally (Figure 1A). Foxp3 mRNA was also assayed to monitor activity of the Foxp3 expression vector (Figure 1B). Since NF-kappaB activation was partially affected by transfection of high concentrations of the control vector, we determined the fold inhibition of NF-kappaB activation by Foxp3 compared to the control vector at each concentration (Figure 1A). Fold inhibition of NF-kappaB activation was directly proportional to the level of Foxp3 mRNA expression detected by real-time RT-PCR. To determine the level of Foxp3-mediated suppression of NF-kappaB activation over time, HEK 293T cells were transfected with an NF-kappaB luciferase reporter vector and an expression vector encoding Foxp3 or EGFP (control vector) and harvested over 4 d. As shown in Figure 1C, NF-kappaB activation was suppressed by overexpression of Foxp3 at all time points. Extending these results from established, in vitro HEK cell lines to primary human lymphocytes, overexpression of Foxp3 in purified CD4+ T cells from three healthy donors also down-regulated the steady-state level of NF-kappaB activation (Figure 1D). These results recapitulate those from Bettelli and colleagues [15] demonstrating that Foxp3 functions, in part, to block NF-kappaB-dependent transcription in human cell lines as well as in primary human CD4+ T cells.

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The Carboxyl-Terminal FKH Domain Is Not Required for Suppression of NF-kappaB Activation in T Cells
To define the requirements of Foxp3 with respect to inhibition of NF-kappaB-dependent transcription, we utilized a mutant of Foxp3 lacking the FKH domain (Figure 2A) [16], similar to the scurfy mutant Foxp3 of mice, and a mutant Foxp3 protein from a patient with IPEX [4,11,14,17]. Unlike full-length Foxp3, which localizes almost exclusively to the nucleus and can bind in a sequence-specific manner to forkhead binding sites, the DeltaFKH mutant fails to localize to the nucleus and thus cannot interact with promoter elements or nuclear proteins [16]. Therefore, we utilized the DeltaFKH mutant to determine whether nuclear localization (or other function associated with the FKH domain) of Foxp3 was a prerequisite for inhibition of NF-kappaB activation. Although Foxp3 interaction with NF-kappaB presumably takes place in the nucleus, it may also be possible for a cytoplasmic Foxp3 protein to bind to NF-kappaB in the cytoplasm and prevent localization to the nucleus following an activation stimulus. Overexpression of full-length Foxp3, but not of DeltaFKH, was able to suppress activation of a cotransfected NF-kappaB reporter vector in HEK 293T cells (Figure 2B). Both Foxp3 and DeltaFKH were expressed at very high levels following transfection as detected by real-time RT-PCR (unpublished data). These data appear to suggest that the carboxyl-terminal FKH domain is critically important for Foxp3 to down-regulate NF-kappaB-dependent transcription. However, NF-kappaB activation was blocked to a similar extent by both full-length Foxp3 and DeltaFKH in Jurkat T cells (Figure 2C) and primary human CD4+ T cells (Figure 2D). Western blot analysis of NF-kappaB p65 expression demonstrated that Foxp3 and DeltaFKH does not block NF-kappaB activation at the level of p65 protein expression (Figure 2E). These results are very interesting with respect to Foxp3 function, because they suggest that the carboxyl-terminal FKH domain, and possibly nuclear localization, are dispensable for Foxp3 function in T cell populations. Alternative interpretations may include the possibility that the localization of DeltaFKH differ between epithelial cells and T cells. In either case, these results suggest a cell type-specific mechanism of action for this Foxp3 mutant.

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Foxp3 Suppresses HIV-1 Gene Expression in Part through Blocking Activation of NF-kappaB
If Foxp3 functions as a repressor of NF-kappaB-dependent gene expression, then we hypothesized that Foxp3 overexpression could selectively down-regulate transcription from promoters previously shown to be responsive to NF-kappaB. To address this question, we examined the transcriptional activation of the HIV-1 LTR, which contains two tandem cis-acting NF-kappaB binding sites located between positions -102 and -81 with respect to the transcription initiation site [21]. NF-kappaB plays a crucial role in regulating gene expression directed from the HIV-1 LTR in CD4+ T cells [21]. Overexpression of full-length Foxp3, but not DeltaFKH, in HEK 293T cells was able to inhibit basal activation of the HIV-1 LTR (Figure 3A), similar to what was previously demonstrated with the synthetic NF-kappaB reporter vector (Figure 2B). Furthermore, HIV-1 LTR activation was suppressed by full-length Foxp3 and DeltaFKH in Jurkat T cells (Figure 3B). To demonstrate that Foxp3-mediated HIV-1 LTR repression was associated with interactions with NF-kappaB bound to the HIV-1 LTR, we compared basal activation of the HIV-1 LTR or an identical HIV-1 LTR lacking the NF-kappaB sites located between -102 and -81 (HIV-1 Delta-kappaB LTR) (Figure 3C). This mutant HIV-1 LTR construct exhibited reduced levels of transcription compared to the parental HIV-1 LTR in purified healthy donor CD4+ T cells (unpublished data). However, directly comparing the effect of Foxp3 overexpression on the activation of these two viral promoters demonstrated that Foxp3 was more capable of suppressing transcriptional activation of the HIV-1 LTR (Figure 3D) compared to the mutated HIV-1 LTR (Figure 3E). These results suggest that Foxp3 down-regulation of HIV-1 LTR activation was mediated at least in part by cis-acting NF-kappaB binding sites. Residual levels of inhibition of the HIV-1 Delta-kappaB LTR by Foxp3 may be due to NF-AT binding sites located upstream of the NF-kappaB sites within the HIV-1 LTR [22,23].

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The Transactivation Functions of HTLV-I Tax Are Suppressed by Foxp3
Previous studies by Bettelli and colleagues have demonstrated that Foxp3 can repress both the basal levels of NF-kappaB activation as well as tumor necrosis factor-alpha-stimulated NF-kappaB activation [15]. Our next step was to determine whether Foxp3 could also suppress the activation of NF-kappaB caused by a strong viral transactivator protein. HTLV-I encodes a multifunctional transactivator protein, Tax, capable of activating both the NF-kappaB and CREB pathways [24-29]. Since the HTLV-I Tax protein can function at multiple levels in both the cytoplasm and the nucleus to stimulate activation of NF-kappaB [28,29], we hypothesized that overexpression of Foxp3 may interfere with this process. However, since Tax-dependent HTLV-I gene expression is independent of NF-kappaB [18], we also hypothesized that Foxp3 would not affect Tax-dependent activation of the HTLV-I LTR. To test these hypotheses, we overexpressed HTLV-I Tax, full-length Foxp3, and/or DeltaFKH in HEK 293T cells cotransfected with an HTLV-I LTR or NF-kappaB reporter vector. As shown in Figure 4A, HTLV-I Tax strongly up-regulated NF-kappaB-dependent transcriptional activation (~60-fold). Interestingly, overexpression of Foxp3, but not DeltaFKH, suppressed Tax-mediated activation of NF-kappaB-dependent transcription. These observations further suggest that the carboxyl-terminal FKH domain is required for inhibiting activation of NF-kappaB in the presence of Tax in HEK 293T cells, strikingly similar to the requirements of Foxp3 inhibition of basal NF-kappaB activation shown in Figure 2B. Transactivation of the HTLV-I LTR was stimulated about 55-fold by overexpression of Tax (Figure 4B), while transfection of Foxp3 suppressed Tax-dependent HTLV-I LTR activation, although HTLV-I LTR activation in the presence or absence of Tax is independent of NF-kappaB or NF-AT (another transcriptional activator known to interact with Foxp3). Furthermore, overexpression of DeltaFKH also led to suppression of HTLV-I transactivation by Tax to a similar extent as full-length Foxp3 (Figure 4B). The suppressive effects shown in Figure 4A and 4B were not the result of Foxp3 down-regulating the expression of the transfected Tax plasmid as determined by real-time RT-PCR (Figure 4C). These results strongly suggest that Foxp3 interacts with transcriptional regulators in addition to NF-kappaB and NF-AT, and that the carboxyl-terminal FKH domain, and therefore localization to the nucleus, are not required for inhibition of Tax-mediated HTLV-I LTR activation (even in HEK 293T cells).
To determine whether Foxp3 inhibited the transactivation functions of Tax by directly associating with this viral protein, we generated an expression vector in which HTLV-I Tax was fused in-frame to the carboxyl terminus of the Gal4 DNA-binding domain (Gal4-BD). This Gal4-BD-Tax fusion protein activated transcription of a synthetic promoter containing five Gal4 binding sites, while Gal4-BD was insufficient to stimulate transcription by itself (Figure 4D). Transactivation of the Gal4-resposive promoter by Gal4-BD-Tax remained relatively unaffected by overexpression of either EGFP (control), Foxp3, or DeltaFKH, suggesting that Foxp3 does not repress Tax transactivation by directly interfacing with the HTLV-I Tax protein. To confirm that Foxp3 had a direct effect on HTLV-I replication, we transfected HEK 293T cells with a well-characterized HTLV-I infectious molecular clone (termed ACH) [30] in the presence of full-length Foxp3, DeltaFKH, or control vector. ACH has been previously shown to direct the expression of viral antigens, produce infectious virus, and transform CD4+ T cells both in vitro and in vivo [31,32]. After 24 h, the amount of viral antigen expression, in this case Tax mRNA, was detected by a sensitive real-time RT-PCR assay. As illustrated in Figure 5, the level of Tax mRNA synthesized from ACH was down-regulated in the presence of Foxp3 compared to the level produced in the presence of the control vector. DeltaFKH did not have a discernable affect on Tax expression. These data indicate that Foxp3 is capable of repressing the expression of Tax from an infectious HTLV-I molecular clone.

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Increased Foxp3 Protein Expression Is Associated with Low HTLV-I Proviral Load
Since Foxp3 is expressed almost exclusively within CD4+CD25+ T cells, a major viral reservoir for HTLV-I [33], it was important to determine whether there was an association between Foxp3 and HTLV-I replication in infected patients. We therefore quantitated the Foxp3 protein expression in CD4+CD25+ T cells by flow cytometry and the HTLV-I proviral load (a surrogate marker of viral replication) by real-time PCR from eight patients with HAM/TSP and eight asymptomatic carriers (ACs). The data from this analysis is summarized in Table 1. As expected, patients with HAM/TSP exhibited significantly higher proviral loads (indicated as HTLV-I proviral DNA copies/100 cells) (34.68 +/- 23.19) compared to ACs (4.75 +/- 5.47) (p = 0.0008). The percentage of Foxp3+ cells within the CD4+CD25+ T cell population was significantly greater in ACs (43.23 +/- 12.95) than in HAM/TSP patients (18.59 +/- 5.77) (p = 0.0033). These data suggest that high levels of Foxp3 protein expression are associated with reduced HTLV-I replication in vivo. They also support our recent reports that high proviral loads, which have been shown to correlate with high Tax mRNA in HTLV-I-infected patients, are associated with reduced Foxp3 expression [8,34].

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CREB Is a Target for Transcriptional Repression by Foxp3
Although Foxp3 could down-regulate Tax-dependent transactivation of the HTLV-I LTR (Figure 4B) and inhibit Tax expression from an infectious molecular clone (Figure 5), Foxp3 failed to modulate Tax function in the absence of the viral promoter (Figure 4D). These results led us to hypothesize that Foxp3 acts on HTLV-I gene expression by interacting with proteins important for driving HTLV-I LTR activity in vivo. Previous studies have demonstrated that the Tax-responsive elements within the HTLV-I LTR play a crucial role in driving Tax-mediated transactivation of the HTLV-I LTR [18]. The Tax-responsive elements have been shown to resemble CREB binding sites, bind CREB in vitro and in vivo, and facilitate HTLV-I LTR activation both in the presence and in the absence of Tax [35,36]. Ching and colleagues [18] demonstrated that addition of a dominant-negative CREB expression vector resulted in nearly complete inhibition of Tax-mediated activation of the HTLV-I LTR, while blocking NF-kappaB activation by addition of a dominant-negative IKKbeta expression vector had no effect on Tax transactivation of the HTLV-I LTR. Therefore, we hypothesized that Foxp3 may inhibit Tax transactivation of the HTLV-I LTR via disruption of the CREB signaling pathway. To test this possibility, HEK 293T cells were transfected with an HTLV-I LTR or synthetic CREB reporter vector along with a control expression vector (EGFP) or expression vectors encoding Foxp3 or DeltaFKH. As shown in Figure 6A, Foxp3 down-regulated basal activation of the HTLV-I LTR and transcription of a synthetic CREB reporter vector, suggesting that Foxp3 down-regulates HTLV-I LTR activation by targeting the CREB pathway. Deletion of the FKH domain of Foxp3 dampened the suppressive effect of Foxp3, but did not completely abrogate suppression, as is seen with NF-kappaB-responsive promoters in HEK 293T cells. Like NF-kappaB activation, CREB transcriptional activation was also suppressed by expression of Foxp3, and to a similar extent DeltaFKH, in healthy donor CD4+ T cells (Figure 6B). Similarly, Foxp3 and DeltaFKH also repressed basal HTLV-I LTR activation in primary human CD4+ T cells (Figure 6C). To our knowledge, this is the first evidence implicating CREB as a molecular target of Foxp3. As observed with NF-kappaB activation, DeltaFKH was a more potent inhibitor of CREB activation in CD4+ T cells than in HEK 293T cells, further indicating that a cell type-specific mechanism of action may govern the function of this Foxp3 mutant.
To determine whether Foxp3 functioned by directly signaling through CREB, we utilized expression vectors encoding CREB-1 or c-Jun (a member of the activator protein 1 family of transcription factors) fused in-frame to the Gal4-BD (Gal4-BD-CREB-1 and Gal4-BD-c-Jun). As shown in Figure 6D, activation of a Gal4-responsive reporter vector by Gal4-BD-CREB-1 was down-regulated by Foxp3 compared to control vector (EGFP), indicating that Foxp3 functions by directly or indirectly interacting with CREB-1. However, Foxp3 failed to markedly affect transcriptional activation of Gal4-BD-c-Jun (c-Jun has also been demonstrated to bind to the HTLV-I LTR) and Gal4-BD-Tax (see Figure 5). Importantly, the mechanism of Foxp3-mediated inhibition of CREB-dependent transcription was not due to a block in CREB-1 protein expression, as determined by Western blot analysis (Figure 6E). Although these results demonstrate that Foxp3 functions as a co-repressor of CREB activation (in addition to NF-kappaB and NF-AT), we were unable to detect a direct physical interaction between CREB-1 and Foxp3 by coimmunoprecipitation or mammalian two-hybrid analysis (unpublished data). Therefore, our data suggest that Foxp3 may interfere with CREB signaling at an upstream event, such as phosphorylation of CREB or recruitment/function of coactivator proteins CREB-binding protein (CBP)/p300.

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Foxp3 Antagonizes CREB Transcriptional Activation by Disrupting Coactivator Recruitment
Stimulation of CREB-dependent transcription by reagents that activate adenylate cyclase and increase cAMP levels (e.g., forskolin) increase the transactivation potential of CREB through phosphorylation of serine 133 by protein kinase A, which permits binding and recruitment of coactivators CBP/p300 to the promoter [37,38]. Phosphorylation of serine 133 does not, however, affect the DNA-binding activity of CREB in most cases [39-41]. Addition of forskolin to HEK 293T cells stimulated activation of a CREB reporter vector about 65 fold (Figure 7A). Overexpression of Foxp3 was capable of down-regulating forskolin-induced CREB transcriptional activation. The functional interaction between Foxp3 and CREB did not affect the DNA-binding activity of CREB-1, but did show a modest decrease in activating transcription factor 2 (ATF-2) DNA-binding activity in the presence of forskolin as determined by transcription factor ELISA (Figure 7B).
While Foxp3 has been shown to bind to and repress activation of both NF-kappaB and NF-AT, exactly how Foxp3 functions to bring about this affect has not been elucidated. To determine how Foxp3 blocks CREB-dependent transcription, we examined whether Foxp3 was capable of (1) disrupting the recruitment of coactivator proteins and/or (2) preventing phosphorylation of CREB at serine 133 (which is a prerequisite for coactivator recruitment). Since both of these events are required for CREB-dependent gene expression, we hypothesized that Foxp3 may affect CREB activation at both steps. To determine whether Foxp3 can disrupt the function/recruitment of the coactivator protein p300, we introduced a Gal4 reporter vector and a Gal4-BD-CREB-1 expression vector into HEK 293T cells in the absence or presence of Foxp3, p300, and/or control expression vectors (Figure 7C). As expected, p300 overexpression stimulated transcription of the Gal4-BD-CREB-1 fusion protein. Foxp3, again, repressed basal levels of Gal4-BD-CREB-1 activation by more than 2-fold, while effectively neutralizing Gal4-BD-CREB-1 activation in the presence of p300. We next analyzed the effect of Foxp3 on phosphorylation of CREB at serine 133 in forskolin-treated HEK 293T cells by Western blot analysis. Overexpression of Foxp3 failed to reduce the detectable levels of CREB phosphorylation using a phosphospecific antibody for CREB-1 (unpublished data). However, when we attempted to determine whether Foxp3 could physically interact with the coactivator protein p300, we found that p300 immunoprecipitated Foxp3 when both proteins were overexpressed in HEK 293T cells (Figure 7D). Collectively, these results suggest that Foxp3 antagonizes CREB-dependent gene expression by directly interacting with coactivator p300 and interfering with its function and/or recruitment to CREB-responsive promoter sequences.

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Discussion
In the present study, we show that Foxp3 functions as a potent repressor of NF-kappaB- and CREB-dependent transcriptional activation. Furthermore, the carboxyl-terminal FKH domain appears to be dispensable for mediating these effects, at least in T cell populations. This observation may become important in light of recent reports suggesting that Foxp3 expression in thymic epithelial cells was crucial for directing development of T cells in the thymus [42]. Interestingly, the majority of the genetic mutations associated with IPEX, a severe autoimmune disorder caused by functional inactivation of Foxp3, map to the carboxyl-terminal FKH domain or the leucine zipper domain in the central region of the protein. Only one mutation associated with IPEX to date has been mapped to the amino-terminal proline-rich region [43]. It is possible that the FKH domain has a complex tertiary structure that is particularly sensitive to misfolding caused by genetic mutations and that an intact FKH domain is absolutely critical for promoting Foxp3 function in the nucleus, whereas the structure of the amino-terminal proline-rich region may tolerate certain mutations as long as the NF-kappaB/NF-AT binding motif remains unaltered. This motif may also include the zinc finger domain. A logical region that may be targeted by the amino-terminal proline-rich region of Foxp3 is the Rel homology domain found in both NF-kappaB and NF-AT family proteins. A region that may also be important with respect to Foxp3 function is the leucine zipper domain, as demonstrated by the number of mutations associated with IPEX that have been mapped in this region of Foxp3. The role of this domain in Foxp3 function remains uncharacterized, but may play a role in dimer formation as it does in other Foxp family members [44].
Because the pathogenesis of a number of retroviral-induced immunologic disorders such as HIV-1/AIDS and HTLV-I/HAM/TSP have been associated with dysregulation of Foxp3 expression [8,45], we also examined the role of Foxp3 in retroviral gene expression. HIV-1 LTR activation in CD4+ T cells is critically dependent on two tandem NF-kappaB sites located between nucleotide positions -102 and -81 within the HIV-1 enhancer region, whereas HTLV-I LTR activation in the presence or absence of the HTLV-I-encoded transactivator protein Tax is independent of NF-kappaB [18]. To our knowledge for the first time, Foxp3 was shown to have a direct effect on HIV-1 LTR transcription. Deletion of the NF-kappaB sites within the HIV-1 enhancer region reduced the responsiveness of the HIV-1 LTR to Foxp3-mediated suppression. In addition, the FKH domain of Foxp3 was required for this inhibitory effect in HEK 293T cells, but not in Jurkat T cells, similar to Foxp3-mediated suppression of a synthetic NF-kappaB reporter. The direct effect of Foxp3 down-regulating HIV-1 gene expression correlates well with recently reported evidence indicating that higher regulatory activity of CD4+CD25+ T cells from HIV-1-infected patients was associated with lower HIV-1 viral loads in these patients [46].
Foxp3 also affected two well-known functions of HTLV-I Tax: transactivation of the NF-kappaB pathway and, most surprisingly, transactivation of the HTLV-I LTR. Transactivation of the HTLV-I LTR by Tax involves the interaction of ATF/CREB factors with Tax in the nucleus. Binding of Tax enhances ATF/CREB dimerization and promotes assembly of Tax-ATF/CREB complexes onto specific sequences in the viral promoter known as Tax-responsive elements. This series of steps allows Tax to recruit coactivator proteins CBP/p300 to the viral promoter and facilitate a high level of viral gene expression [24-26]. Transactivation of the NF-kappaB pathway by Tax was inhibited by overexpression of full-length Foxp3, but not DeltaFKH, as seen with basal activation of the HIV-1 LTR and a synthetic NF-kappaB reporter in HEK 293T cells. However, Tax-mediated transactivation of the HTLV-I LTR was inhibited by overexpression of both full-length Foxp3 as well as DeltaFKH in both HEK 293T cells and CD4+ T cells. We demonstrated that Foxp3 did not directly affect the functioning of Tax, but rather Foxp3 targeted the transcription factors required for Tax transactivation (i.e., NF-kappaB and a then-unknown cellular factor, which we identified in this study as CREB). The negative effect of Foxp3 on HTLV-I gene expression was confirmed utilizing an HTLV-I infectious molecular clone.
Importantly, we demonstrated that HTLV-I-infected individuals with the highest levels of Foxp3 protein expression within the CD4+CD25+ T cells population exhibited lower proviral loads than did individuals with the lowest levels of Foxp3 protein expression. Previous studies have demonstrated that the HTLV-I proviral load directly correlates with HTLV-I Tax mRNA load, the frequency of immunopathogenic virus-specific CD8+ T cells, and disease severity in patients with HAM/TSP [47]. These results have important implications on the utility of Foxp3 in controlling viral gene expression and thus pathogenesis of HAM/TSP. Therefore, Foxp3 becomes an attractive target for the development of novel therapeutic applications directed at modulating the expression of this important regulatory protein, especially in light of recent observations that the expression of Foxp3 can also be down-regulated by HTLV-I Tax [8].
As the activation of the HTLV-I LTR depends primarily on ATF/CREB proteins (whether in the presence or the absence of Tax), we investigated whether Foxp3 could interact with this additional cellular signaling pathway. While the DNA-binding activity of CREB is, in most cases, constitutive, the transactivation potential of CREB is regulated by the phosphorylation of CREB and recruitment of CBP/p300 [48]. Our data demonstrate that Foxp3 interferes with the latter of these two processes and that the recruitment of the coactivator protein p300, and resulting transcriptional activation are blocked by Foxp3. This may be the result of the physical interaction we detected between Foxp3 and p300. With respect to HTLV-I LTR activity, while full-length Foxp3 inhibited both basal and Tax-dependent transcription by ~50%, DeltaFKH appeared less effective in suppressing basal activation (~25% inhibition) compared to Tax-dependent activation (~50% inhibition). The effect of DeltaFKH on basal activation of the HTLV-I LTR in HEK 293T cells was very similar to that shown for a synthetic CREB reporter, suggesting that the FKH domain of Foxp3 is important at some level. As observed with NF-kappaB activation, the Foxp3 mutant lacking the FKH domain was a stronger inhibitor of CREB activation in CD4+ T cells than in HEK 293T epithelial cells. Therefore, it appears that in CD4+ T cells, the FKH domain is dispensable for the proper functioning of Foxp3 with respect to both NF-kappaB and CREB activation.
In summary, this is, to our knowledge, the first direct evidence implicating a role for the Treg-specific transcription factor Foxp3 in regulating retroviral gene expression. In addition, we identify the CREB pathway as a molecular target of Foxp3. Since CREB has been shown to regulate multiple genes involved in transcription (e.g., JunD, c-Fos, signal transducer of activated T cells 3 [STAT3]), cell cycle (e.g., p15INK4b, cyclin A, cyclin D1), and immune regulation (e.g., IL-2, IL-6, T-cell receptor alpha) (reviewed in [48]), the findings presented in this report broaden the potential range of signaling pathways under the control of the regulatory protein Foxp3. Our evidence stresses the importance of Foxp3 expression and Treg function in the development and maintenance of protective immunity against HIV-1 and HTLV-I. Based on recent findings, Foxp3 may limit HIV-1 and HTLV-I transcription by interfering with activation of NF-kappaB and CREB pathways. However, observing that this inhibitory effect is not absolute, a low level of viral gene expression may persist in CD4+ T cells (in particular regulatory T cells, which are known reservoirs of HIV-1 and HTLV-I) and result in the accumulation of viral proteins that either stimulate NF-kappaB and/or CREB activation or directly inhibit Foxp3 expression or function. The imbalance of NF-kappaB and CREB activation caused by these viral gene products may be a crucial step in the pathogenesis of virus-induced immunological disorders such as AIDS and HAM/TSP. Future studies will be directed at identifying and characterizing cellular proteins that interact with Foxp3 both in the nucleus and cytoplasm, in order to better address how Foxp3 functions to guide the development and function of regulatory T cells in health and disease.

targeted

amino-terminal proline-rich region

dimer formation

interaction

Tax

Binding

Tax

assembly

Tax

recruit

Tax

CBP

recruit

Tax

recruitment

CBP

recruitment

recruitment

interaction

interact

expression

expression

overexpression

overexpression

DeltaFKH

expression

expression

expression

expression

expression

expression

inactivation

lowest

expression

down-regulated

expression

Tax

interferes

recruitment

blocked

recruitment

lacking

FKH domain

inhibit

expression

promotes

Binding

assembly

overexpression

overexpression

DeltaFKH

highest

expression

result

interferes

interaction

result

blocked

interaction

play a role

leucine zipper domain

dimer formation

does

leucine zipper domain

dimer formation

dysregulation

expression

modulating

expression

regulate

JunD

regulate

c-Fos

regulate

signal transducer of activated T cells 3

regulate

regulate

cyclin A

regulate

cyclin D1

regulate

regulate


Input:
Cell culture.
HEK 293T cells were cultured in Dulbecco's modified Eagle medium (Invitrogen, Carlsbad, California, United States). Jurkat T cells and primary human CD4+ T cells were cultured in RPMI-1640 medium (Invitrogen). Media were supplemented with 2 mM L-glutamine, 100 U/ml penicillin, 100 mug/ml streptomycin (Cambrex, East Rutherford, New Jersey, United States), and 10% fetal bovine serum (Atlanta Biologicals, Norcross, Georgia, United States).


Input:
Patients and cell preparation.
PBMCs were prepared by centrifugation over Ficoll-Hypaque gradients (BioWhittaker, Walkersville, Maryland, United States) from eight HAM/TSP patients and eight ACs, and the cells were viably cryopreserved in liquid nitrogen until tested. HAM/TSP was diagnosed according to WHO guidelines [49]. HTLV-I seropositivity was determined by ELISA (Abbott Laboratories, Abbott Park, Illinois, United States), with confirmation by Western blot analysis (Genelabs Technologies, Redwood City, California, United States). Blood samples were obtained after informed consent as part of a clinical protocol reviewed and approved by the NIH institutional review panel.


Input:
Plasmids.
Expression vectors encoding human Foxp3 (pCMV-Foxp3-IRES-EGFP) and human Foxp3 lacking the forkhead (FKH) domain (pCMV-DeltaFKH-IRES-EGFP) were generous gifts from S. Ziegler (Benaroya Research Institute). pEGFP-C2 was provided by I. Lipinski (NIDDK/NIH). pcDNA3 was provided by K. T. Jeang (NIAID/NIH). pCMV4-Tax was a generous gift from W. Greene (University of California San Francisco). pGL4-luc2 and pGL4-TKhRluc2 were purchased from Promega (Madison, Wisconsin, United States). pUC18 was purchased from Stratagene (La Jolla, California, United States). HIV-1 wt LTR and HIV-1 Delta-kappaB LTR luciferase reporter vectors were constructed by cloning the XhoI/HindIII LTR fragments from pHIV-CAT and pDelta-kappaB-HIV-CAT (AIDS Research and Reference Reagent Program, NIAID/NIH) into the multiple cloning site of pGL4-luc2. NF-kappaB, HTLV-I LTR, and CREB luciferase reporter and pCMV-p300-HA expression vectors were generously provided by B. Wigdahl (Drexel University College of Medicine). HTLV-I pACH infectious molecular clone has been described previously [30]. pFR-luc, pFA-CMV, pFA2-CREB-1, and pFA2-c-Jun were purchased from Stratagene. pFA-Tax (encoding a fusion protein consisting of the Gal4 DNA-binding domain fused in-frame to HTLV-I Tax) was constructed by PCR amplification of HTLV-I Tax using pCMV4-Tax as a template and BamHI/BglII-tagged primers. The amplified insert was digested and ligated into the BamHI/BglII sites of pFA-CMV. Plasmid contents were confirmed by DNA sequencing.

Expression

Expression

expression

luciferase

expression

lacking

forkhead (FKH) domain


Input:
Isolation of primary human CD4+ T cells.
CD4+ T cells were isolated from cryopreserved healthy donor PBMCs by negative selection with the CD4+ T Cell Isolation Kit II (Miltenyi Biotech, Bergisch Gladbach, Germany) according to manufacturer's guidelines. Purity of negatively selected CD4+ T cells was consistently higher than 96% as determined by flow cytometry.


Input:
Transient expression and luciferase assays.
HEK 293T cells were plated at a density of 5 x 105 cells/well in six-well culture plates (BD Biosciences, San Diego, California, United States) 1 d prior to transfection with the appropriate plasmid DNA (~2 mug total) using FuGene 6 transfection reagent (Roche, Basel, Switzerland). Jurkat T cells were plated at 1 x 106 cells/well in six-well culture plates the day of transfection with the appropriate plasmid DNA (~2 mug total) using FuGene 6 transfection reagent. Primary human CD4+ T cells (2 x 106) were nucleofected with the specified plasmid DNA (5 mug total) using the Human T Cell Nucleofection Kit (Amaxa, Gaithersburg, Maryland, United States). Forskolin (10 muM; Calbiochem, San Diego, California, United States) was added in some experiments 20 h posttransfection. Cells were harvested 24 h posttransfection and luciferase activity was analyzed using the Dual-luciferase Reporter Assay System (Promega) and a Monolight 2010 luminometer (Analytical Luminescence Laboratory, San Diego, California, United States) according to manufacturer's guidelines. pGL4-TKhRluc2 was used as an internal control to normalize for transfection efficiency. Nucleofected CD4+ T cells were also monitored for transfection efficiency and cell viability 24 h posttransfection as follows. Transfection efficiency was routinely ~30% as determined by flow cytometric analysis of EGFP expression. Cell viability, determined by staining with 7-amino-actinomycin D (7-AAD; BD Biosciences), was routinely ~70%. Both transfection efficiency and cell viability in nucleofected CD4+ T cells was independent of the plasmids used.


Input:
Foxp3 and HTLV-I Tax expression analysis by real-time RT-PCR.
Real-time RT-PCR analysis of Foxp3 and HTLV-I Tax expression was performed as previously described [8,47]. Briefly, total RNA was extracted using the RNeasy Mini Kit (Qiagen, Valencia, California, United States) according to manufacturer's guidelines, and cDNA was synthesized by reverse transcription using TaqMan Gold RT-PCR Kit using random hexamer primers (Applied Biosystems, Foster City, California, United States). Foxp3 and HTLV-I Tax mRNA expression was quantified by real-time PCR using ABI PRISM 7700 Sequence Detection System (Applied Biosystems). The normalized values in each sample were calculated as the relative quantity of Foxp3 or HTLV-I Tax mRNA expression divided by the relative quantity of HPRT mRNA expression. The values were calculated by the following formula: normalized Foxp3 or HTLV-I Tax expression = 2Ct value of HPRT - Ct value of Foxp3 or HTLV-I Tax.

expression

expression

Tax

expression

Tax

expression

expression

expression

Tax

mRNA expression

mRNA expression

Tax

mRNA expression

mRNA expression

Tax

mRNA expression

HPRT


Input:
Real-time PCR.
Real-time PCR analysis of HTLV-I (Tax) proviral load was performed as previously described [47,50]. DNA was extracted from 1 x 106 cells using Puregene DNA Isolation Kit (Gentra, Minneapolis, Minnesota, United States), and 100 ng of the sample DNA solution was analyzed by this system. The HTLV-I proviral DNA load was calculated by the following formula: copy number of HTLV-I (pX) per 100 cells = (copy number of pX)/(copy number of beta-actin/2) x 100.


Input:
Western blot analysis.
HEK 293T cells were plated at a density of 5 x 105 cells/well in six-well culture plates (BD Biosciences) 1 d prior to transfection with the appropriate plasmid DNA (2 mug total) using FuGene 6 transfection reagent (Roche). Cells were harvested 24 h posttransfection for whole-cell lysates in RIPA buffer (50 mM Tris-HCl [pH 7.4], 150 mM NaCl, 1% Igepal (NP-40), 0.5% sodium deoxycholate, 1 mM EDTA, 1 mM DTT, 1 mM PMSF, and 1x Complete Mini Protease Inhibitor [Roche]). Protein concentration was determined by Lowry assay (Bio-Rad, Hercules, California, United States) and colorimetric reactions were read using a VersaMax microplate reader (Molecular Devices, Sunnyvale, California, United States) at an absorbance of 750 nm. Size fractionation was performed on 20 mug of protein/sample by SDS-PAGE, and the protein was transferred to nitrocellulose or PVDF membranes and subjected to immunoblotting using the indicated antibodies. Foxp3 was detected using rabbit anti-human Foxp3 polyclonal antibody (ab4728 or ab10563; Abcam, Cambridge, United Kingdom) and anti-rabbit IgG-HRP secondary antibody (Cell Signaling Technology, Beverly, Massachusetts, United States). NF-kappaB p65 and CREB-1 were detected using rabbit anti-human polyclonal (p65) or monoclonal antibody (CREB-1; 48H2) (Cell Signaling Technology). Beta-actin was detected using a mouse monoclonal antibody (AC-15; Sigma, St. Louis, Missouri, United States). For coimmunoprecipitation analysis, cell lysates were precleared with 30 mul of protein A/G plus-agarose beads (Santa Cruz Biotechnology, Santa Cruz, California, United States) and then incubated with mouse monoclonal anti-HA antibody (6E2; 1:100; Cell Signaling Technology) and 30 mul of protein A/G plus-agarose beads overnight. The immunoprecipitates were washed four times with RIPA buffer, resuspended in SDS sample buffer, and heated at 95 degreesC for 5 min. Proteins were then treated as described for Western blot analysis.


Input:
Transcription factor DNA-binding analysis (TF-ELISA).
CREB-1 and ATF-2 DNA-binding activity was analyzed with the TransFactor Profiling (Inflammation 1) Kit (BD Biosciences) according to the manufacturer's protocol. Nuclear extracts were prepared from HEK 293T cells transfected with a control vector (EGFP) or Foxp3 expression vector (1,000 ng) in the presence or absence of forskolin (10 muM for 4 h) using the TransFactor Extraction Kit (BD Biosciences). Protein concentration was determined using a Biophotometer (Eppendorf, Hamburg, Germany). Nuclear extracts (20 mug) were incubated in preblocked wells containing plate-bound double-stranded oligonucleotides corresponding to an ATF/CREB consensus sequence (emTGACATCAem). Wells were washed, incubated with the appropriate primary antibody, washed, incubated with secondary antibody (HRP-labeled), washed again, and finally developed with TMB substrate. Colorimetric reactions were read using a VersaMax microplate reader (Molecular Devices) at an absorbance of 655 nm.

binding

binding

expression


Input:
Flow cytometric analysis of Foxp3 protein expression.
Cryopreserved PBMCs from HAM/TSP patients or HTLV-I-infected ACs were thawed and washed with FACS buffer (1x PBS, 0.1% NaN3, 5% FBS). Cells (1.5 x 106) were fixed by sequential formaldehyde/methanol fixation as follows. Cells were carefully resuspended in FACS buffer and fixed with 100 mul of reagent A (Fix & Perm kit; Caltag Laboratories, Burlingame, California, United States) at room temperature for 3 min followed by 2 ml of 70% methanol for 5 min at 4 degreesC. Cells were washed twice and permeabilized with 100 mul of reagent B (Fix & Perm kit) and stained for intracellular Foxp3 with mouse anti-human Foxp3 monoclonal antibody (0.5 mug of ab22510; Abcam) or the appropriate isotype control for 30 min. Cells were washed twice and stained with Cy5-conjugated goat anti-mouse immunoglobulin F(ab')2 secondary antibody (Caltag Laboratories) for an additional 30 min. Cells were washed twice and stained for surface CD4 expression with PE-labeled anti-CD4 (BD) and CD25 expression with FITC-labeled anti-CD25 (BD). Cells were washed twice and analyzed on a FACSCalibur (BD). Data analysis was performed using FlowJo (Tree Star, Ashland, Oregon, United States).

expression

expression

expression


Input:
Statistical analyses.
The Mann-Whitney U test was used to compare the data between patients with HAM/TSP and AC.


Input:
EBV Latent Membrane Protein 1 Activates Akt, NFkappaB, and Stat3 in B Cell Lymphomas
Latent membrane protein 1 (LMP1) is the major oncoprotein of Epstein-Barr virus (EBV). In transgenic mice, LMP1 promotes increased lymphoma development by 12 mo of age. This study reveals that lymphoma develops in B-1a lymphocytes, a population that is associated with transformation in older mice. The lymphoma cells have deregulated cell cycle markers, and inhibitors of Akt, NFkappaB, and Stat3 block the enhanced viability of LMP1 transgenic lymphocytes and lymphoma cells in vitro. Lymphoma cells are independent of IL4/Stat6 signaling for survival and proliferation, but have constitutively activated Stat3 signaling. These same targets are also deregulated in wild-type B-1a lymphomas that arise spontaneously through age predisposition. These results suggest that Akt, NFkappaB, and Stat3 pathways may serve as effective targets in the treatment of EBV-associated B cell lymphomas.
Author Summary
Epstein-Barr virus (EBV) is linked to the development of multiple cancers, including post-transplant lymphoma, Hodgkin disease, and nasopharyngeal carcinoma. Latent membrane protein 1 (LMP1) is expressed in many EBV-associated cancers and is responsible for most of the altered cellular growth properties that are induced by EBV infection. This study reveals that LMP1 induces lymphomas in B-1a lymphocytes, a cell type that is susceptible to transformation in aged mice. The lymphomas require Akt, NFkappaB, and Stat3 signaling for enhanced growth and survival. The activation of the Stat3, Akt, and NFkappaB signaling pathways likely underlies the ability of LMP1 to promote malignant transformation.

expressed

Latent membrane protein 1

inhibitors

Akt

inhibitors

Activates

Latent Membrane Protein 1

Akt

Activates

Latent Membrane Protein 1

deregulated

deregulated

deregulated


Input:
Introduction
Epstein-Barr virus (EBV) is a ubiquitous gamma-herpesvirus that infects humans predominantly at an early age with greater than 90% of the adult population infected with EBV [1]. EBV is linked to the development of both B lymphocyte and epithelial cell malignancies, including Burkitt lymphoma, Hodgkin disease (HD), and nasopharyngeal carcinoma (NPC), and cancers linked to immunosuppression, including post-transplant lymphoma and AIDS-associated lymphomas [2,3]. In vitro infection of B lymphocytes with EBV induces permanent growth transformation, and this ability to affect cell growth regulation likely contributes to the development of cancer.
Many of the viral proteins expressed in transformed cells, including the EBV nuclear antigens and latent membrane proteins, have profound effects on cell growth regulation and are required for EBV latent infection and B cell transformation [1]. Latent membrane protein 1 (LMP1) is considered the major oncoprotein of EBV, as it transforms rodent fibroblasts to tumorigenicity in nude mice and is expressed in HD, NPC, and immunosuppression-associated tumors [4-8]. In B lymphocytes, LMP1 mimics CD40 signaling, and both LMP1 and CD40 are essential for EBV-mediated B cell transformation [9-11]. While CD40 interacts with CD40 ligand expressed on activated T cells to induce B cell activation and differentiation, LMP1 acts as a constitutive signal through ligand-independent oligomerization. LMP1 and CD40 interact with the same tumor necrosis factor receptor-associated factors (TRAFs) leading to activation of NFkappaB, c-Jun N terminal kinase (JNK), and p38 MAPK signaling pathways [12-16]. Activation of NFkappaB is required for EBV-induced B cell transformation and its inhibition rapidly results in cell death [17,18]. Recent studies indicate that LMP1 also activates phosphatidylinositol 3 kinase (PI3K)/Akt signaling and that this activation is required for LMP1-mediated transformation of rodent fibroblasts [5,19].
In vitro, primary B cells can be maintained by CD40 ligation in combination with IL4 treatment. In vivo, CD40 signaling is necessary for germinal center (GC) formation such that mice deficient for CD40 or CD40L are unable to form GCs in response to T cell-dependent antigens [20,21]. Both the membrane proximal and distal cytoplasmic regions of CD40 that bind TRAF6 and TRAFs2/3/5, respectively, are necessary for GC formation, but either region is sufficient to induce extrafollicular B cell differentiation and restore low affinity antibody production [22]. Functionally, LMP1 can rescue CD40-deficient mice and restore immunoglobulin (Ig) class switching, most likely because LMP1 recruits similar TRAF molecules, TRAFs 1/2/3/5 and TRAF6, through the C-terminal activation regions 1 and 2 domains, respectively. However, LMP1 is unable to restore affinity maturation and GC formation [23].
Several EBV transforming proteins have been studied in transgenic mouse models, however, only LMP1 induces tumor development when expressed under the control of the Ig heavy chain promoter and enhancer [24-26]. The LMP1 transgenic mice (IgLMP1) express LMP1 in B lymphocytes, and in mice older than 12 mo, lymphoma develops with increased incidence (40%-50%) compared to wild-type control mice (11%), suggesting that LMP1 contributes to tumor development [26]. The LMP1 lymphomas have rearranged Ig genes and have activated Akt, JNK, p38, and NFkappaB, with specific activation of the NFkappaB family member cRel [27].
In this study, the LMP1 transgenic lymphocytes and lymphomas were further characterized and their growth properties in vitro were determined. To obtain pure populations of malignant lymphocytes and to enable more detailed biochemical analyses, examples of primary lymphomas were inoculated and passaged in SCID mice. Interestingly, lymphoma development was restricted to B-1a lymphocytes, a self-replenishing population of cells that are prone to malignancy [28,29]. LMP1 transgenic lymphocytes had increased viability in vitro and viability was increased by the addition of IL4. In contrast, both LMP1-positive and -negative lymphoma cells were independent of IL4 co-stimulation for survival and proliferation in vitro with a complete absence of activated Stat6, the IL4 target. The lymphomas were also distinguished by constitutive activation of Stat3 and deregulation of the Rb cell cycle pathway. Inhibition of the PI3K/Akt, NFkappaB, and Stat3 signaling pathways blocked the enhanced growth of both LMP1 transgenic and malignant lymphocytes, suggesting that these pathways are required for their growth and survival. These appear to be the same targets that are deregulated in wild-type B-1a lymphomas that arise spontaneously through age predisposition. This study reveals that LMP1 promotes malignancy in cells with the inherent ability to proliferate and that the Akt, NFkappaB, and Stat3 signaling pathways are required for its growth stimulatory effects.

interacts

CD40 ligand

oligomerization

interact

interact

ligation

bind

membrane proximal

bind

distal cytoplasmic regions

bind

distal cytoplasmic regions

bind

distal cytoplasmic regions

recruits

TRAFs 1

C-terminal activation regions 1

recruits

C-terminal activation regions 1

recruits

C-terminal activation regions 1

recruits

C-terminal activation regions 1

recruits

2 domains

expressed

Latent membrane protein 1

expressed

CD40 ligand

expressed

express

deficient

deficient

rescue

deficient

deficient

absence

activated

under the control

expressed

activated

Akt

activation

cRel

activated

activation

target

deregulated

Akt

deregulated


Input:
High Levels of LMP1 Expression Correlates with the Development of Lymphoma
LMP1 expression in IgLMP1 mice was directed to B cells under the control of the Ig heavy chain promoter and enhancer. It has previously been shown that in these transgenic mice, LMP1 expression was restricted to B220+ B cells with lymphoma detected in greatly enlarged spleens [23,26]. To investigate whether LMP1 expression contributes to lymphoma development, B cells were purified from splenocytes by positive selection using anti-CD19 MACS magnetic beads, and equivalent amounts of B cells were analyzed by immunoblotting. LMP1 was detectable in LMP1 transgenic B cells, but upon development of lymphoma, LMP1 expression was stronger in 5/7 lymphomas analyzed with concomitant appearance of degradation products (Figure 1A). To determine whether the higher level of LMP1 detected was due to an expansion of malignant lymphocytes, expression of LMP1 in the spleen was further evaluated by immunohistochemical staining. Immunohistochemistry analysis of spleen sections detected LMP1 in the plasma membrane of cells in both the follicular white pulp and circulating lymphocytes in the red pulp (Figure 1B). LMP1 expression was heterogeneous with strong LMP1 staining interspersed amongst a background of cells staining weakly for LMP1. Upon development to lymphoma, LMP1 expression was more abundantly detected with multiple foci of intense LMP1 staining. This demonstrates that the increased LMP1 detected by immunoblotting upon malignant progression reflects an increase in LMP1 expression and an accumulation of cells expressing high levels of LMP1. This correlation between high LMP1 expression and the development of lymphoma suggests that progression to lymphoma results from increased levels of LMP1.

Expression

expression

expression

expression

detectable

expression

expression

detected

expression

expression

expression

expressing

expression

levels

High Levels

Expression

restricted

expression

stronger

expression

higher level

strong

weakly

abundantly

expression

intense

increased

increase

expression

high levels

expressing

high

expression

increased

levels

under the control

expression


Input:
LMP1 Promotes B-1a Lymphomas That Can Escape Allelic Exclusion
To determine if LMP1 signaling affects B cell differentiation and to immunophenotype the lymphomas that arise from LMP1 expression, surface Ig expression of heavy chains (IgM, IgG, IgD) and light chains (kappa, lambda) were analyzed by flow cytometry. Similar numbers of naive (IgM+IgD+IgG-) splenic B cells with a strong bias towards kappa light chain were detected from wild-type or LMP1 transgenic mice, indicating that LMP1 signaling does not affect B cell maturation (unpublished data). Flow cytometry analysis of the SCID-passaged wild-type and LMP1 transgenic lymphomas revealed an IgMhighIgDlow phenotype (Figure 2A), indicative of marginal zone, B-1, or memory B cells. B-1 cells are further separated into CD5+ (B-1a) and CD5- (B-1b) subsets. To differentiate between these cell types, lymphoma cells were further analyzed for the B-1a marker CD5. All (5/5) of the tested LMP1 transgenic lymphomas displayed an IgMhighIgDlowCD5+ phenotype (Figure 2A), an expression pattern that distinguishes B-1a cells. Interestingly, a spontaneous wild-type lymphoma also developed in B-1a cells (Figure 2A). These cells are an interesting population that is self replenishing with an increased likelihood to become malignant in aged mice [30]. Analysis of LMP1 transgenic mice before the development of lymphoma showed similar numbers of splenic B-1a (CD19+CD5+) and B-1b or B-2 populations (CD19+CD5-), indicating that LMP1 does not affect B cell differentiation (Figure 2B).
Due to allelic exclusion, mature B cells that have been exposed to antigen will typically express only one heavy chain isotype (IgG, IgE, or IgA) and either a kappa or lambda light chain. Interestingly, 2/5 LMP1 transgenic lymphomas analyzed (lymphomas 2 and 4) were doubly positive for low levels of both kappa and lambda light chains (Figure 2A). Previous characterization of the LMP1 lymphomas had revealed that the lymphomas were clonal as determined by Ig heavy chain rearrangement [26], and analysis of kappa chain rearrangement (Figure S1) of the samples analyzed in this study confirmed clonality. To further assess light chain expression, the passaged samples were tested by immunoblotting for kappa and lambda light chains (Figure 2C). Interestingly, very low levels of expression of both light chains were detected by flow cytometry. The low levels of expression may reflect a limitation of the total number of light chains that can be expressed on the surface of a B cell. In agreement with the flow cytometry analysis, LMP1 transgenic lymphomas 2 and 4 were also positive for kappa and lambda light chains by immunoblot analysis (Figure 2C), confirming that these lymphomas express both light chains.
A previous study of mice that developed leukemia due to an expansion of self-reactive B-1a cells determined that the B-1a leukemias were also doubly positive for kappa and lambda light chains [31]. These findings indicate that expression of LMP1 in B-1a cells promotes the development of malignancy and can result in the aberrant escape from allelic exclusion.

expression

expression


Input:
LMP1 Promotes B Cell Survival and Proliferation In Vitro
Primary B cell cultures can be maintained through CD40 ligation and supplementation with IL4 [32]. To investigate whether LMP1 affects primary B cell survival and proliferation, splenocytes were cultured in the presence or absence of IL4 and analyzed by MTS as a metabolic marker, by ethidium monoazide (EMA) exclusion for viability, and by 5-bromo-2'-deoxy-uridine (BrdU) incorporation for proliferation. In the MTS assay, as expected, splenocytes from wild-type mice did not survive even with the addition of IL4 due to a lack of CD40 ligation (Figure 3A). In contrast, LMP1 splenocytes had increased metabolism even in the absence of IL4, which was further enhanced upon addition of IL4. Wild-type and LMP1 transgenic lymphoma cells had high levels of MTS activity even in the absence of IL4 (Figure 3A). The LMP1 transgenic lymphoma cells had approximately 4-fold higher MTS activity than the normal transgenic lymphocytes and were at least 2-fold higher than the control lymphoma. As previously published, lymphoma usually develops in mice over 12 mo of age and all mice are sacrificed by 18-20 mo. The ages of the transgenic mice with or without lymphoma ranged between 6 and 20 mo old. There was no correlation between age and MTS activity.
EMA exclusion of CD19+ gated B cells prepared from two wild-type and two LMP1 transgenic mice indicated a 2-fold increase in viability in the LMP1 transgenic lymphocytes compared to wild-type lymphocytes. Two examples of LMP1 transgenic lymphoma cells had greatly increased viability that was not increased by IL4 treatment, indicating that the lymphoma cells are independent of IL4 co-stimulation (Figure 3B). Enhancement in MTS activity was observed in LMP1 transgenic lymphoma cells by the addition of IL4; however, EMA exclusion did not reveal a similar increase. This could reflect a difference for IL4 requirement in the metabolic activity versus the viability of LMP1 transgenic lymphoma cells. Although expression of LMP1 could enhance survival of non-malignant primary lymphocytes, BrdU incorporation revealed that LMP1 expression alone was not sufficient to induce proliferation in culture (unpublished data). Only lymphoma cells had detectable levels of BrdU incorporation detected by flow cytometry (Figure 3C). Interestingly, LMP1 lymphoma cells had significantly higher levels of proliferation in comparison to the spontaneous lymphoma that developed in an LMP1-negative littermate (25% versus 4%). This higher level of proliferation was observed in lymphomas that express both high (Table 1, LMP1-L2 and LMP1-L3) and low (Table 1, LMP1-L5) levels of LMP1, suggesting that even small amounts of LMP1 is sufficient to induce dramatic effects in proliferation. The level of proliferation was not enhanced upon IL4 addition, confirming the IL4 independence observed in the viability studies (Figure 3C; Table1).

ligation

ligation

expression

expression

express

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ligation

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Wild-Type and LMP1 Transgenic Lymphoma Cells Do Not Require IL4 and Stat6 Signaling
To investigate whether IL4 independence was due to endogenous IL4 expression, IL4 transcription was assessed by an Rnase protection assay (RPA). IL4 transcription was detectable with control RNA and faintly in the mouse lymphoma cell line K46mu (Figure 4A). However, IL4 transcription was not detectable in CD19+ MACS-purified B cells from wild-type lymphocytes (unpublished data), LMP1 transgenic lymphocytes, or lymphoma cells, although the GAPDH and L32 controls were effectively protected (Figure 4A). Activated Stat6 (pStat6), a target of the IL4 receptor pathway, was detected in the wild-type and LMP1 transgenic lymphocytes (Figure 4B). In contrast, pStat6 was barely detected in either the wild-type or LMP1 transgenic lymphoma cells. However, the pathway was not disabled, as treatment of the lymphoma cells with IL4 induced Stat6 phosphorylation (Figure 4C).
Although wild-type lymphocytes cannot be maintained in culture with IL4 supplementation alone (Figure 3A), slight enhancement in MTS activity could be detected if the cells were analyzed at an earlier time point, at 1 d (Figure 4D) versus 3 d (Figure 3A) post-harvest. The enhancement of MTS activity induced by IL4 in wild-type lymphocytes could be neutralized by the addition of IL4 antibody (Figure 4D). However, neutralizing antibodies to IL4 did not affect the MTS activity of LMP1 transgenic lymphoma cells (Figure 4E). In summary, the wild-type and LMP1 transgenic lymphoma cells grew independently of IL4 treatment and did not require Stat6 signaling.

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