Biol. Chem. 2014; 395(11): 1265–1274
Review Boet van Riel and Frank Rosenbauer*
Epigenetic control of hematopoiesis: the PU.1 chromatin connection Abstract: Purine-rich box1 (PU.1) is a transcription factor that not only has a key role in the development of most hematopoietic cell lineages but also in the suppression of leukemia. To exert these functions, PU.1 can cross-talk with multiple different proteins by forming complexes in order to activate or repress transcription. Among its protein partners are chromatin remodelers, DNA methyltransferases, and a number of other transcription factors with important roles in hematopoiesis. While a great deal of knowledge has been acquired about PU.1 function over the years, it was the development of novel genome-wide technologies, which boosted our understanding of how PU.1 acts on the chromatin to drive its repertoire of target genes. This review summarizes current knowledge and ideas of molecular mechanisms by which PU.1 controls hematopoiesis and suppresses leukemia. Keywords: chromatin; epigenetics; hematopoiesis; PU.1. DOI 10.1515/hsz-2014-0195 Received May 5, 2014; accepted July 3, 2014; previously published online September 2, 2014
Introduction The transcription factor purine-rich box1 (PU.1) is a key regulator for the development of several hematopoietic lineages. In fact, PU.1 is one of the most intensively studied transcription factors (TFs) in hematopoiesis, and a great deal of knowledge has been produced by now, resulting in more than 1460 publications that can be found in PubMed (as of April 2014: ncbi.nlm.nih.gov/ pubmed). The official gene name for PU.1 is SPI1 in human and Sfpi1 in the mouse. The PU.1-encoding gene was first *Corresponding author: Frank Rosenbauer, Institute of Molecular Tumor Biology, University of Münster, D-48149 Münster, Germany, e-mail: [email protected] Boet van Riel: Institute of Molecular Tumor Biology, University of Münster, D-48149 Münster, Germany
discovered as a target for insertions of the SFFV virus that causes erythroid leukemia (Moreau-Gachelin et al., 1988), followed by the isolation of the 31-kDa protein called PU.1. PU.1 is part of the ETS transcription factor family that consists of 27 different family members in humans (reviewed in Oikawa and Yamada, 2003). This family shares an ETS domain that facilitates direct DNA binding to the core GGAA motif (Wei et al., 2010). PU.1 has next to the ETS domain a winged helix domain that mediates binding to an extended consensus longer than the ETS-binding motif (Pio et al., 1996). Furthermore, PU.1 has an activation domain and a PEST domain involved in protein-protein interactions (Figure 1A). The human SPI1 is located on chromosome 11 and the mouse Sfpi1 on chromosome 2. Strict regulation of the expression of PU.1 is essential for proper hematopoietic development from hematopoietic stem cells (HSCs) into fully differentiated cells. Dysregulation of PU.1 expression in HSCs leads to a depletion of functional HSCs in adult mice (Staber et al., 2013). Also, later in the hematopoietic development, PU.1 expression is important for maturation of macrophages, B cells, early T cell progenitors, and T helper 9 (Th9) cells (Anderson et al., 1998; DeKoter and Singh, 2000; Anderson et al., 2002; Gerlach et al., 2014) (Figure 1B). In progenitor cells, there is also a feedback connection between PU.1 concentrations and the cell cycle that is important for myeloid vs. lymphoid lineage decision (Kueh et al., 2013). Moreover, proper PU.1 expression is essential to avoid the development of myeloid leukemia (Rosenbauer et al., 2004). Of note, downregulation, but not complete abolishment of PU.1 expression, appears to be of advantage for transformation of normal myeloid progenitors into leukemic cells (Steidl et al., 2006; Zhu et al., 2012). Such PU.1 downregulation during leukemia progression can be achieved by several mechanisms, which will be described in more detail later in this review. With the recent revolution in the development of novel genome-wide technologies such as next-generation sequencing methods, we have learned much about the global functions of PU.1 on the genome and are starting to get a glimpse of how PU.1 operates and epigenetically
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1266 B. van Riel and F. Rosenbauer: PU.1 and chromatin
Figure 1 Schematic representation of PU.1, its protein partners, and its role in hematopoiesis. (A) Schematic representation of the PU.1 protein structure with ammo acid numbering. Highlighted domains are the activation domain, PEST domain, and the ETS domain. Binding sites of protein partners are shown. (B) Simplified scheme of mammalian definitive hematopoiesis with cells not expressing PU.1 is depicted in red. Cells expressing PU.1 are depicted in blue: hematopoietic stem cells (HSC), common lymphoid progenitors (CLP), and common myeloid progenitors (CMP). Increase, decrease, or no change in PU.1 expression is symbolized between the cell symbols.
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modifies the chromatin to initiate and maintain expression of its repertoire of target genes. In this review, we will provide an overview of known protein interaction partners by which PU.1 controls the epigenetic state at its genomic target regions and induces the development of myeloid leukemia.
PU.1 interaction with chromatin modifiers PU.1 can either activate or repress the transcription of genes, a notion that is supported by an in vivo study showing that after genetic knockdown of PU.1 in the HSCs, the expression of 225 genes was upregulated and of 97 genes was downregulated (Steidl et al., 2006). This differential control of gene expression is thought to be mediated by the ability of PU.1 to build different complexes with a number of protein partners, thus, either stimulating or inhibiting transcription. One of the first identified proteinbinding partners of PU.1 was the TATA-binding protein (TBP) (Hagemeier et al., 1993). Through TBP, PU.1 is able to recruit the basal transcription machinery to its binding sites at promoters and enhancers as a requirement for transcription (Kihara-Negishi et al., 2001). Indeed, many PU.1 target genes lack TATA box sequences and, thus, are unable to bind TBP directly (reviewed in Sandra Clarke and Siamon Gordon, 1998). Subsequent studies have shown that PU.1 can form complexes with a number of proteins that can directly modify chromatin or DNA. For example, PU.1 can bind the histone acetyltransferases CREB-binding protein (CBP) and p300 to activate transcription (Yamamoto et al., 1999; Bai et al., 2005). Acetylation of histone tails is thought to open the chromatin and allow other proteins to bind to the DNA and activate transcription (Sterner and Berger, 2000). Furthermore, PU.1 can also bind a histone deacetylase complex consisting of histone deacetylase 1 (HDAC1) and mammalian Sin3a (mSIN3a) (Kihara-Negishi et al., 2001). At least in reporter assays using the c-Myc promoter, HDAC1-mSIN3a functions as a repressor of PU.1 transactivation capacity (Kihara-Negishi et al., 2001). Later on, it was reported that together with HDAC1-mSIN3a, PU.1 can also bind the methyl CpG-binding protein 2 (MeCP2) to repress genes (Suzuki et al., 2006; Imoto et al., 2010). A possible link between methylated DNA sequences and PU.1 has also been suggested by the observation that the PU.1 protein is able to form a complex with the de novo DNA methyltransferases Dnmt3a and Dnmt3b (Suzuki et al., 2006). These interactions were discovered by
co-transfection experiments in 293T cells and are believed to site-specifically methylate DNA in order to repress transcription of selected genomic PU.1 target sites such as the tumor-suppressor gene p16 (INK4A) promoter (Suzuki et al., 2006). Importantly, the PU.1-DNMT3b complex has also been observed to occur endogenously during monocyte to ostoclast differentiation (de la Rica et al., 2013). However, genome-wide data have yet failed to identify a significant overlay between PU.1 occupancy and methylated DNA sequences (Pham et al., 2013), thus, challenging the concept that PU.1 can recruit the DNA methylation machinery to repress gene expression. Hence, further functional experiments are needed to solve this obvious discrepancy. As a limitation to the interpretation of the relevance of many of these PU.1 protein complexes, most of them have been identified (and perhaps exclusively observed) using in vitro co-transfection experiments in non- hematopoietic 293T or 3T3 cells. Thus, it remains unclear at the moment if at all, or in what hematopoietic cell types, these complexes exist endogenously. Nevertheless, these protein complex data do at least provide a hint for a model in which PU.1 can operate as a platform for directing the recruitment of different chromatin-modifying proteins, with either gene-repressive or -activating functions, to its target genes (Figures 1B and 2). Importantly, such a model is instrumental for explaining how one single transcription factor is capable of controlling expression of different repertoires of target genes in a dynamic and cell-type specific pattern.
PU.1 cross-talk with other lineagespecific transcription factors PU.1 can cross-talk with a number of other lineage-specific TFs, most of which are known as important regulators of blood cell development. In early hematopoietic differentiation stages such as HSCs and multipotent progenitors, CCAAT-enhancer-binding proteins (C/EBPs) and Runt-related transcription factor 1 (Runx1/AML1) play important roles in concert with PU.1. For example, several essential myeloid lineage growth factor receptor genes, such as those encoding the M-CSFR and GM-GSFRα, are regulated by a PU.1 collaboration with C/EBPα (Hohaus et al., 1995; Li et al., 2005). Runx1 is a critical factor for the development of the definitive HSC pool and also has a role in the specification of some hematopoietic lineages (Wang et al., 1996; van Riel et al., 2012). Similar to the PU.1-C/EBPα complex, PU.1 also collaborates with Runx1
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1268 B. van Riel and F. Rosenbauer: PU.1 and chromatin
Figure 2 Model for chromatin preparation for PU.1 binding to lower-affinity binding sites. For PU.1 to bind DNA, the chromatin has to be opened by other TFs. In the HSC, Runx1 opens PU.1 binding sites. Immunoresponsive TFs, like STAT6 and NF-κB, open latent enhancers for PU.1 binding, and in myeloid development, C/EBP needs to open additional enhancers in the Sfpi1 locus. This leads to PU.1 binding and recruitment of additional TFs and co-factors to regulate transcription. The figure also illustrates three examples of low-affinity PU.1 DNA consensus binding site in humans (Pham et al., 2013).
to activate transcription of the Csf1r gene encoding the M-CSF receptor (Li et al., 2005). Moreover, expression of the PU.1-encoding gene (Sfpi1), itself, is activated by the PU.1-Runx1 protein interaction, via their collaborative binding to adjacent consensus sites within a critical Sfpi1 enhancer, the -14-kb upstream regulatory element (-14 kb URE) (Huang et al., 2008). In myeloid cells, steadily increasing PU.1 expression is a prerequisite for guiding early progenitors through their complete differentiation program all the way toward reaching the mature state (Leddin et al., 2011). In agreement, it was shown that high expression levels of PU.1, together with C/EBPα or β, can convert other lineages to macrophages (Laiosa et al., 2006, Feng et al., 2008). During the developmental cascade, PU.1 appears to dynamically collaborate with changing protein-binding partners to orchestrate the step-wise process of differentiation. For example, as stated above, its collaboration with C/EBPα drives the expression of a number of genes, which are important for early myeloid differentiation (Hohaus et al., 1995). In agreement, using genetic studies in mice,
C/EBPα was found to be required for the transition from the common myeloid progenitor (CMP) to the granulocyte macrophage progenitor (GMP) differentiation step, but is no longer required thereafter (Traver et al., 2001; Zhang et al., 2004). A model seems likely, in which PU.1 transiently interacts with C/EBPα to guide the early myeloid developmental process. Once this differentiation step has past, the PU.1-to-C/EBPα interaction is probably dispensable for further myeloid differentiation steps. C/EBPβ was also shown to bind to PU.1 and to form a transactivating complex (Nagulapalli et al., 1995). C/EBPβ is a factor important for later myeloid differentiation steps (reviewed in Huber et al., 2012), thus suggesting a role for the PU.1-C/ EBPβ complex during terminal myeloid maturation. Concurrently, a likely scenario is that PU.1 actively switches between different C/EBP family members as binding partners to drive progenitors along myeloid differentiation. A repressive complex that is known to be important in granulocyte development is the one between PU.1 and growth factor independant-1 (GFI-1) (Dahl et al., 2007). GFI-1 is a transcriptional repressor, which is already important in HSCs (Lancrin et al., 2012). Interestingly, in myeloid cells of gfi-1-/- mice, PU.1 target genes are upregulated (Dahl et al., 2007). In agreement, GFI-1 can repress macrophage-specific PU.1 target genes, but has little to no effect on PU.1 target genes expressed in granulocytes (Dahl et al., 2007). This bias by GFI-1 is argued to be caused by differences in PU.1 concentrations needed for activation of macrophage and granulocytic genes (Dahl et al., 2007). Another salient example for the interaction of PU.1 with other lineage-specific TFs is its complex with GATAbinding factor 1 (Gata1) (Zhang et al., 1999). Gata1 is expressed in erythroid and megakaryocytic lineages (for reviews see Cantor and Orkin, 2002 and Goldfarb, 2007). It is the key driver of erythroid lineage differentiation, and mice in which Gata1 is lacking are devoid of erythroid progenitors (Cantor and Orkin, 2002). Moreover, Gata1 is erythroid lineage instructive because its ectopic expression in GMPs causes, in part, reprogramming to erythroid cells (Iwasaki et al., 2003). In myeloid progenitors, PU.1 binds to Gata1 and physically blocks Gata1 binding activity to the DNA (Zhang et al., 2000). PU.1 expression is downregulated at the erythroblast stage of definitive erythropoiesis (Hromas et al., 1993), and failure of its downregulation causes a block in erythroid differentiation and subsequent development of erythroid leukemia (Galson et al., 1993). Therefore, one important prerequisite for erythroid differentiation is the capacity of Gata1 to inhibit binding of co-activators to PU.1 and to thereby block its transcriptional activation activity (Zhang et al., 1999). This block includes transcription of the Sfpi1 gene,
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itself, and thus shuts down PU.1 expression. Thus, the antagonistic interplay between PU.1 and Gata1 appears to be important for the selection of bipotential progenitors to differentiate into either myeloid or erythroid-megakaryocytic progeny. Within the lymphoid compartment, PU.1 is expressed in common lymphoid progenitors (CLPs), B cells, and the earliest T-cell progenitors in the thymus (Nutt et al., 2005). Importantly and similar to the erythroid lineage, PU.1 expression is required in later T-cell progenitors to become repressed (Anderson et al., 2002), stays repressed in Th2 cells and CD8 T cells, but is reexpressed in Th9 cells (Gerlach et al., 2014). To achieve PU.1 repression in early T cells, the activity of PU.1 is antagonized by Notch, which is essential to push thymocytes into further maturation (Del Real and Rothenberg, 2013). Moreover, T-cell-specific repression of PU.1 expression is thought to be facilitated by binding of Runx1 to a silencer element located between the Sfpi1 promoter and its upstream enhancer elements (Zarnegar et al., 2010). In summary, PU.1 exerts its functions in concert with a variety of other lineage-specific TFs (Figures 1B and 2). These collaborative and antagonistic interactions appear to be an important prerequisite for the dynamic orchestration of lineage fate choice and differentiation outcomes of hematopoietic progenitors.
Chromatin binding of PU.1 In recent years, as a result of the ability to map genomewide binding patterns of TFs by chromatin immunoprecipitation-sequencing (ChIP-seq), PU.1 has been found to operate as a key factor for initiation and maintaining transcriptionally permissive chromatin (Ostuni et al., 2013; Staber et al., 2013). Multiple ChIP-seq data sets have been generated to identify PU.1 chromatin occupancy patterns in a number of different hematopoietic cell types, including monocytes and macrophages (Heinz et al., 2010; Pham et al., 2012) in the presence or absence of immune stimulation (Ostuni et al., 2013), B cells (Heinz et al., 2010) and a HCS-like cell line (Wilson et al., 2010). These data showed that PU.1 occupies a great number of different genomic locations at promoters as well as intragenic and intergenic regions. The sequence of the PU.1 binding site can differ significantly throughout the genome. More than 2500 potential 12-mer consensus motifs with variable frequencies for PU.1 binding have been identified in monocytes and macrophages (Pham et al., 2013). Furthermore, the same study showed that only 1% of all potential PU.1
binding motifs within the genome are actually occupied by PU.1 (Pham et al., 2013). How does the selection of binding sites of TFs in the genome occur? The DNA binding consensus sequences, which in vitro were shown to have the highest affinity for PU.1 binding, are in vivo in a large part located within gene deserts, and lack any obvious binding motifs of other known TF nearby (Pham et al., 2013). Also, these isolated high-affinity PU.1 binding sites are often located in unconserved regions, and the biological significance of these sites is currently unclear. The functional binding sites of PU.1 in promoters and enhancers have an overall lower binding affinity for PU.1 and are principally always flanked by binding sites of one or more additional TFs (Pham et al., 2013). The most likely scenario for functional binding site selection seems, therefore, to be a combination of ‘low’ affinity with adjacent sites to allow combinatorial binding of additional TFs. Collaborative TF binding with PU.1 is either important to stabilize PU.1 DNA occupancy, or to enhance the transactivating function of PU.1, or most likely both. Also, in many cases, it will be the occupancy of all collaborative factors, which is necessary to create and maintain open chromatin (Heinz et al., 2010, 2013). The relative expression level of PU.1 in progenitor cells has been shown to act as a driving event of lineage choice selection during hematopoietic differentiation (DeKoter and Singh, 2000; Leddin et al., 2011). PU.1 protein concentrations appear to have an impact on the occurrence of PU.1 binding to functional DNA elements after comparing occupancy between hematopoietic progenitor cells and monocytes (Pham et al., 2013). With the increase in protein concentration, the number of binding sites increases, and more low-affinity sites are bound (Pham et al., 2013). Another important question is how PU.1 penetrates the chromatin at its different binding sites. Most likely, PU.1 competes with nucleosomes at high-affinity sites, which is indicated by its binding to sites that show high affinity in vitro but are not hypersensitive for DNase I in vivo and are most likely devoid of collaborative binding events (Pham et al., 2013). In contrast, lower-affinity binding sites may not be accessed by PU.1 alone and are frequently flanked by additional TF binding sites when bound. This assumption is supported by the finding that the chromatin at several PU.1 binding sites have to be opened or primed earlier in development by global chromatin changes mediated by factors like Runx1 (Hoogenkamp et al., 2009) and that PU.1 is unable to rescue the Runx1 knock-out phenotype (Lancrin et al., 2012). It appears that PU.1 needs a pre-prepared global ‘chromatin context’ in the cell for functional binding and
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1270 B. van Riel and F. Rosenbauer: PU.1 and chromatin subsequent transactivation of important target genes. Indeed, a recent report described a set of enhancers that was inactive in unstimulated macrophages but became active after immune stimulation (Ostuni et al., 2013). PU.1 was shown to bind to these so-called ‘latent’ enhancers only after immune stimulation, and PU.1 binding was preceded and dependent on binding by immune responsive TFs such as SMAD6 and NF-κB. Another example is that high expression of PU.1 in macrophages requires two autoregulatory loops at its Sfpi1 gene, built via the -14-kb and -12-kb enhancers (Leddin et al., 2011). However, before PU.1 is able to bind to the macrophagespecific -12-kb enhancer, C/EBPα has to bind to the -14-kb enhancer to subsequently ‘open’ the chromatin at the -12-kb enhancer so that PU.1 can enter and bind (Leddin et al., 2011). It is likely that similar to the situation at its own gene locus, the 1% of the potential binding sites that are indeed occupied by PU.1 in the human genome is globally selected by being located in active DNA regions that had been ‘opened’ for PU.1 to enter chromatin. A model of PU.1 occupancy at lower-affinity binding sites is shown in Figure 2. As yet, we have a very poor understanding, if at all, of the impact of posttranslational modifications at the PU.1 peptide on the in vivo function and DNA binding site selection of PU.1. Several posttranslational PU.1 modifications have been described. By transient transfection studies, phosphorylation of serine 148 and 142 has been shown to influence PU.1 transactivation capacity (Lodie et al., 1997; Wang et al., 2003). Interestingly, acetylation of PU.1 has an influence on the protein complex selection of PU.1 in vitro (Lodie et al., 1997; Kihara-Negishi et al., 2005). Mutation of the lysine stretch within the C-terminal end of the ETS domain of PU.1 into arginines, which cannot be acetylated anymore, had little effect on the binding potential of PU.1 to CBP, but reduced binding to mSIN3a using GST pull down assays. Therefore, it is likely that posttranslational modifications can, at least in part, influence the ability of PU.1 to direct the differentiation choice by affecting the nature of the protein complexes.
PU.1 and the three-dimensional chromatin structure Distal regulatory gene elements such as enhancers, insulators, and silencers must often cross-talk with proximal promoters over long genomic distances (for reviews on this subject see Pennacchio et al., 2013; Chetverina
et al., 2014). Several models have been put forward to explain how this cross-talk may occur, one of which is DNA looping. Indeed, owing to novel technological developments such as chromosome conformation capturing (3C) and related approaches, there is now overwhelming evidence that DNA looping operates as a major gene regulatory mechanism (Carter et al., 2002; Dekker et al., 2002; Tolhuis et al., 2002). However, formation of such three-dimensional chromatin structures must be tightly controlled to participate in the dynamic regulation of gene expression. Indeed, several lineage-specific hemato poietic TFs, such as PU.1 and Runx1, have been shown to facilitate DNA looping (Levantini et al., 2011; Schonheit et al., 2013; Staber et al., 2013). PU.1 binds to its -14-kb URE enhancer to autoregulate expression of its own Spfi1 gene (Li et al., 2001). This enhancer was found to loop to the proximal Sfpi1 promoter (Ebralidze et al., 2008), a process, which is impaired by deletion of the PU.1 binding motifs with the -14-kb URE (Staber et al., 2013). Interestingly, dependency of this looping structure between the -14-kb URE and the proximal promoter required PU.1 binding exclusively in HSCs but not in macrophages (Staber et al., 2013). A likely explanation for this cell type-associated dynamics lies in an additional Sfpi1 enhancer element (-12 kb enhancer) in the vicinity of the -14-kb URE, which is active in macrophages but not in HSCs (Leddin et al., 2011). This enhancer might preserve the DNA loop even while PU.1 binding to the -14-kb URE is abolished. Another example of a regulatory gene element of which chromatin looping is dependent on PU.1 is the -50-kb enhancer in the Irf8 locus (Schonheit et al., 2013). Here, knock-down of PU.1 decreases looping frequency between this enhancer and the proximal Irf8 promoter. An important question that these single gene studies did not yet answer is whether on a more global scale, myeloid enhancers, in general, depend on PU.1 binding to facilitate looping with their respective genes. However, this appears to be likely, because studies in B cells have shown that PU.1 occupancy is associated with enhancers and genomic regions involved in DNA looping (Heinz et al., 2010; Lin et al., 2012). Another important question is on the molecular mechanism by which PU.1 controls DNA looping. The most likely answer to this question, again, may be found in the choice of protein partners, which may require PU.1 occupancy at the chromatin to bind to DNA and subsequently facilitate chromosomal looping. The use of genome-wide variants of the 3C technology (Carter et al., 2002; Tolhuis et al., 2002) in myeloid cells will probably have the power to provide answers to these important questions.
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PU.1, chromatin, and myeloid leukemia It has been known for several years that decreased PU.1 expression levels can directly lead to the development of AML in mice (Rosenbauer et al., 2004). Of note, however, only very few cases with mutations in SPI1 have been reported in human AML (Mueller et al., 2003). However, PU.1 has been shown to be frequently downregulated by leukemia-driving oncogenes in AML patients, suggesting that similar to mice, PU.1 can act as a tumor suppressor in man (Vangala et al., 2003; Mueller et al., 2006). Although one study in mice has shown that complete absence of PU.1 can lead to leukemia (Metcalf et al., 2006), in humans, complete loss of PU.1 function has not yet been reported, suggesting that its graded reduction is supportive for cancer development. This is similar to the situation of C/EBPα mutations in AML, which lead to reduced but not completely abolished function (Pabst and Mueller, 2009). An explanation why residual PU.1 expression may be required for AML development has recently been illustrated by the genome-wide finding that the PML-RARα oncogene requires PU.1 for its chromatin entry and control of target genes (Wang et al., 2010). Thus, it appears that a fine-tuned reduction in PU.1 expression may provide the optimal concentration for blocking normal myeloid differentiation, on the one hand, and on the other hand, still allows oncogenes, such as PML/RARα, to regulate their target genes. Interestingly, in patients with complex karyotype AML, a single nucleotide polymorphism (SNP) in the SPI1 URE enhancer, which disrupts binding of the chromatin organizer special AT-rich sequence-binding protein-1 (SATb1) occurs with higher frequency than in healthy cells, suggesting a role for this SNP in leukemia predisposition (Steidl et al., 2007). Indeed, functional investigation of this SNP has shown that it leads to reduced levels of PU.1 (Steidl et al., 2007). Intriguingly, SATb1 is involved in the control of DNA looping factor in early T cells (Yasui et al., 2002), and it therefore seems likely, but not yet proven, that the SNP within the PU.1 enhancer might impair its DNA looping to the proximal promoter. In four out of 120 AML patients, sequence variations in a NF-κB binding site in the URE have been identified, and it was shown that this aberration resulted in impaired PU.1 expression (Bonadies et al., 2010). Importantly, NF-κB has recently been revealed to coincide with de novo PU.1 binding to many immune-responsive enhancers in macrophages (Ostuni et al., 2013). Hence, it is tempting to speculate that also in leukemia cells, failed NF-kB binding
to the URE may reduce chromatin entry of PU.1 to autoregulate its own expression.
Conclusion PU.1 is a central regulator of hematopoiesis, and intensive research over almost three decades has provided ample understanding of its pattern of expression and gene regulation, its role in differentiation, its function as a tumor suppressor in myeloid leukemia, its genomewide nature of chromatin occupancy and binding site selection, and its many protein interaction partners. So, a valid question is why is continuing research on PU.1 still needed? The main reason is that in spite of the existing wealth of knowledge, many important questions still remain. For example, how do the different PU.1 expression levels drive specific hematopoietic differentiation outcomes at the chromatin level? Are there different PU.1 concentration-dependent protein-protein interactions, and/or do the different PU.1 concentrations lead to a selection of different binding sites in the genome? Also, why does PU.1 bind to some of its genomic consensus motifs but not to others, and how is this selection regulated? An example highlighting this specific question is the lack of PU.1 occupancy at ‘latent’ enhancers prior to, but not after, immune stimulation of macrophages. This leads to the question why is PU.1 able to prepare enhancers at some genomic sites but not at others? Another interesting unresolved topic is on the molecular nature of the PU.1-dependent chromatin looping structures. For example, how does PU.1 affect the spatial organization of the chromatin? Often, addressing novel questions requires the utilization of newly emerging technologies. Importantly, it is a major advantage of our great knowledge on PU.1 that exists today to enable us to put new findings into the proper molecular and cellular perspectives. Thus, PU.1 does not only serve as an important factor for the regulation of chromatin but also continues to serve as a role model in research on transcription factors. Acknowledgments: We thank Michael Rehli (Regensburg, Germany) for valuable suggestions on the manuscript. The literature regarding PU.1 is overwhelming, and we wish to apologize to those whose work we were unable to cite. This work was supported by the Deutsche Forschungsgemeinschaft Research Unit 1336 and Cells-in-Motion Cluster of Excellence (EXC 1003 – CiM), University of Münster, Germany.
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1272 B. van Riel and F. Rosenbauer: PU.1 and chromatin
References Anderson, K.L., Smith, K.A., Conners, K., McKercher, S.R., Maki, R.A., and Torbett, B.E. (1998). Myeloid development is selectively disrupted in PU.1 null mice. Blood 91, 3702–3710. Anderson, M.K., Weiss, A.H., Hernandez-Hoyos, G., Dionne, C.J., and Rothenberg, E.V. (2002). Constitutive expression of PU.1 in fetal hematopoietic progenitors blocks T cell development at the pro-T cell stage. Immunity 16, 285–296. Bai, Y., Srinivasan, L., Perkins, L., and Atchison, M.L. (2005). Protein acetylation regulates both PU.1 transactivation and Ig κ 3′ enhancer activity. J. Immunol. 175, 5160–5169. Bonadies, N., Neururer, C., Steege, A., Vallabhapurapu, S., Pabst, T., and Mueller, B.U. (2010). PU.1 is regulated by NF-κB through a novel binding site in a 17 kb upstream enhancer element. Oncogene 29, 1062–1072. Cantor, A.B. and Orkin, S.H. (2002). Transcriptional regulation of erythropoiesis: an affair involving multiple partners. Oncogene 21, 3368–3376. Carter, D., Chakalova, L., Osborne, C.S., Dai, Y.F., and Fraser, P. (2002). Long-range chromatin regulatory interactions in vivo. Nat. Genet. 32, 623–626. Chetverina, D., Aoki, T., Erokhin, M., Georgiev, P., and Schedl, P. (2014). Making connections: insulators organize eukaryotic chromosomes into independent cis-regulatory networks. Bioessays 36, 163–172. Clarke S. and Gordon S. (1998). Myeloid-specific gene expression. J. Leukoc. Biol. 63, 153–168. Dahl, R., Iyer, S.R., Owens, K.S., Cuylear, D.D., and Simon, M.C. (2007). The transcriptional repressor GFI-1 antagonizes PU.1 activity through protein-protein interaction. J. Biol. Chem. 282, 6473–6483. de la Rica, L., Rodriguez-Ubreva, J., Garcia, M., Islam, A.B., Urquiza, J.M., Hernando, H., Christensen, J., Helin, K., Gomez-Vaquero, C., and Ballestar, E. (2013). PU.1 target genes undergo Tet2coupled demethylation and DNMT3b-mediated methylation in monocyte-to-osteoclast differentiation. Genome Biol. 14, R99. Dekker, J., Rippe, K., Dekker, M., and Kleckner, N. (2002). Capturing chromosome conformation. Science 295, 1306–1311. DeKoter, R.P. and Singh, H. (2000). Regulation of B lymphocyte and macrophage development by graded expression of PU.1. Science 288, 1439–1441. Del Real, M.M. and Rothenberg, E.V. (2013). Architecture of a lymphomyeloid developmental switch controlled by PU.1, Notch and Gata3. Development 140, 1207–1219. Ebralidze, A.K., Guibal, F.C., Steidl, U., Zhang, P., Lee, S., Bartholdy, B., Jorda, M.A., Petkova, V., Rosenbauer, F., Huang, G., et al. (2008). PU.1 expression is modulated by the balance of functional sense and antisense RNAs regulated by a shared cisregulatory element. Genes Dev. 22, 2085–2092. Feng, R., Desbordes, S.C., Xie, H., Tillo, E.S., Pixley, F., Stanley, E.R., and Graf, T. (2008). PU.1 and C/EBPα/β convert fibroblasts into macrophage-like cells. Proc. Natl. Acad. Sci. USA 105, 6057–6062. Galson, D.L., Hensold, J.O., Bishop, T.R., Schalling, M., D’Andrea, A.D., Jones, C., Auron, P.E., and Housman, D.E. (1993). Mouse β-globin DNA-binding protein B1 is identical to a proto-oncogene, the transcription factor Spi-1/PU.1, and is restricted in expression to hematopoietic cells and the testis. Mol. Cell Biol. 13, 2929–2941.
Gerlach, K., Hwang, Y., Nikolaev, A., Atreya, R., Dornhoff, H., Steiner, S., Lehr, H.A., Wirtz, S., Vieth, M., Waisman, A., et al. (2014). T9 cells that express the transcription factor PU.1 drive T cell-mediated colitis via IL-9 receptor signaling in intestinal epithelial cells. Nat. Immunol. 15, 676–686. Goldfarb, A.N. (2007). Transcriptional control of megakaryocyte development. Oncogene 26, 6795–6802. Hagemeier, C., Bannister, A.J., Cook, A., and Kouzarides, T. (1993). The activation domain of transcription factor PU.1 binds the retinoblastoma (RB) protein and the transcription factor TFIID in vitro: RB shows sequence similarity to TFIID and TFIIB. Proc. Natl. Acad. Sci. USA 90, 1580–1584. Heinz, S., Benner, C., Spann, N., Bertolino, E., Lin, Y.C., Laslo, P., Cheng, J.X., Murre, C., Singh, H., and Glass, C.K. (2010). Simple combinations of lineage-determining transcription factors prime cis-regulatory elements required for macrophage and B cell identities. Mol. Cell 38, 576–589. Heinz, S., Romanoski, C.E., Benner, C., Allison, K.A., Kaikkonen, M.U., Orozco, L.D., and Glass, C.K. (2013). Effect of natural genetic variation on enhancer selection and function. Nature 503, 487–492. Hohaus, S., Petrovick, M.S., Voso, M.T., Sun, Z., Zhang, D.E., and Tenen, D.G. (1995). PU.1 (Spi-1) and C/EBPα regulate expression of the granulocyte-macrophage colonystimulating factor receptor α gene. Mol. Cell Biol. 15, 5830–5845. Hoogenkamp, M., Lichtinger, M., Krysinska, H., Lancrin, C., Clarke, D., Williamson, A., Mazzarella, L., Ingram, R., Jorgensen, H., Fisher, A., et al. (2009). Early chromatin unfolding by RUNX1: a molecular explanation for differential requirements during specification versus maintenance of the hematopoietic gene expression program. Blood 114, 299–309. Hromas, R., Orazi, A., Neiman, R.S., Maki, R., Van, B.C., Moore, J., and Klemsz, M. (1993). Hematopoietic lineage- and stagerestricted expression of the ETS oncogene family member PU.1. Blood 82, 2998–3004. Huang, G., Zhang, P., Hirai, H., Elf, S., Yan, X., Chen, Z., Koschmieder, S., Okuno, Y., Dayaram, T., Growney, J.D., et al. (2008). PU.1 is a major downstream target of AML1 (RUNX1) in adult mouse hematopoiesis. Nat. Genet. 40, 51–60. Huber, R., Pietsch, D., Panterodt, T., and Brand, K. (2012). Regulation of C/EBPβ and resulting functions in cells of the monocytic lineage. Cell Signal. 24, 1287–1296. Imoto, A., Okada, M., Okazaki, T., Kitasato, H., Harigae, H., and Takahashi, S. (2010). Metallothionein-1 isoforms and vimentin are direct PU.1 downstream target genes in leukemia cells. J. Biol. Chem. 285, 10300–10309. Iwasaki, H., Mizuno, S., Wells, R.A., Cantor, A.B., Watanabe, S., and Akashi, K. (2003). GATA-1 converts lymphoid and myelomonocytic progenitors into the megakaryocyte/erythrocyte lineages. Immunity. 19, 451–462. Kihara-Negishi, F., Yamamoto, H., Suzuki, M., Yamada, T., Sakurai, T., Tamura, T., and Oikawa, T. (2001). In vivo complex formation of PU.1 with HDAC1 associated with PU.1-mediated transcriptional repression. Oncogene 20, 6039–6047. Kihara-Negishi, F., Suzuki, M., Yamada, T., Sakurai, T., and Oikawa, T. (2005). Impaired repressor activity and biological functions of PU.1 in MEL cells induced by mutations in the acetylation motifs within the ETS domain. Biochem. Biophys. Res. Commun. 335, 477–484.
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B. van Riel and F. Rosenbauer: PU.1 and chromatin 1273 Kueh, H.Y., Champhekar, A., Nutt, S.L., Elowitz, M.B., and Rothenberg, E.V. (2013). Positive feedback between PU.1 and the cell cycle controls myeloid differentiation. Science 341, 670–673. Laiosa, C.V., Stadtfeld, M., Xie, H., de Andres-Aguayo, L., and Graf, T. (2006). Reprogramming of committed T cell progenitors to macrophages and dendritic cells by C/EBPα and PU.1 transcription factors. Immunity 25, 731–744. Lancrin, C., Mazan, M., Stefanska, M., Patel, R., Lichtinger, M., Costa, G., Vargel, O., Wilson, N.K., Moroy, T., Bonifer, C., et al. (2012). GFI1 and GFI1B control the loss of endothelial identity of hemogenic endothelium during hematopoietic commitment. Blood 120, 314–322. Leddin, M., Perrod, C., Hoogenkamp, M., Ghani, S., Assi, S., Heinz, S., Wilson, N.K., Follows, G., Schonheit, J., Vockentanz, L., et al. (2011). Two distinct auto-regulatory loops operate at the PU.1 locus in B cells and myeloid cells. Blood 117, 2827–2838. Levantini, E., Lee, S., Radomska, H.S., Hetherington, C.J., BerichJorda, M., Amabile, G., Zhang, P., Gonzalez, D.A., Zhang, J., Basseres, D.S., et al. (2011). RUNX1 regulates the CD34 gene in haematopoietic stem cells by mediating interactions with a distal regulatory element. EMBO J. 30, 4059–4070. Li, Y., Okuno, Y., Zhang, P., Radomska, H.S., Chen, H., Iwasaki, H., Akashi, K., Klemsz, M.J., McKercher, S.R., Maki, R.A., et al. (2001). Regulation of the PU.1 gene by distal elements. Blood 98, 2958–2965. Li, X., Vradii, D., Gutierrez, S., Lian, J.B., van Wijnen, A.J., Stein, J.L., Stein, G.S., and Javed, A. (2005). Subnuclear targeting of Runx1 is required for synergistic activation of the myeloid specific M-CSF receptor promoter by PU.1. J. Cell Biochem. 96, 795–809. Lin, Y.C., Benner, C., Mansson, R., Heinz, S., Miyazaki, K., Miyazaki, M., Chandra, V., Bossen, C., Glass, C.K., and Murre, C. (2012). Global changes in the nuclear positioning of genes and intraand interdomain genomic interactions that orchestrate B cell fate. Nat. Immunol. 13, 1196–1204. Lodie, T.A., Savedra, R., Jr., Golenbock, D.T., Van Beveren, C.P., Maki, R.A., and Fenton, M.J. (1997). Stimulation of macrophages by lipopolysaccharide alters the phosphorylation state, conformation, and function of PU.1 via activation of casein kinase II. J. Immunol. 158, 1848–1856. Metcalf, D., Dakic, A., Mifsud, S., Di, R.L., Wu, L., and Nutt, S. (2006). Inactivation of PU.1 in adult mice leads to the development of myeloid leukemia. Proc. Natl. Acad. Sci. USA 103, 1486–1491. Moreau-Gachelin, F., Tavitian, A., and Tambourin, P. (1988). Spi-1 is a putative oncogene in virally induced murine erythroleukaemias. Nature 331, 277–280. Mueller, B.U., Pabst, T., Osato, M., Asou, N., Johansen, L.M., Minden, M.D., Behre, G., Hiddemann, W., Ito, Y., and Tenen, D.G. (2003). Heterozygous PU.1 mutations are associated with acute myeloid leukemia. Blood 101, 2074. Mueller, B.U., Pabst, T., Fos, J., Petkovic, V., Fey, M.F., Asou, N., Buergi, U., and Tenen, D.G. (2006). ATRA resolves the differentiation block in t (15;17) acute myeloid leukemia by restoring PU.1 expression. Blood 107, 3330–3338. Nagulapalli, S., Pongubala, J.M., and Atchison, M.L. (1995). Multiple proteins physically interact with PU.1. Transcriptional synergy with NF-IL6 β (C/EBPδ, CRP3). J. Immunol. 155, 4330–4338. Nutt, S.L., Metcalf, D., D’Amico, A., Polli, M., and Wu, L. (2005). Dynamic regulation of PU.1 expression in multipotent hematopoietic progenitors. J. Exp. Med. 201, 221–231.
Oikawa, T. and Yamada, T. (2003). Molecular biology of the Ets family of transcription factors. Gene 303, 11–34. Ostuni, R., Piccolo, V., Barozzi, I., Polletti, S., Termanini, A., Bonifacio, S., Curina, A., Prosperini, E., Ghisletti, S., and Natoli, G. (2013). Latent enhancers activated by stimulation in differentiated cells. Cell 152, 157–171. Pabst, T. and Mueller, B.U. (2009). Complexity of CEBPA dysregulation in human acute myeloid leukemia. Clin. Cancer Res. 15, 5303–5307. Pennacchio, L.A., Bickmore, W., Dean, A., Nobrega, M.A., and Bejerano, G. (2013). Enhancers: five essential questions. Nat. Rev. Genet. 14, 288–295. Pham, T.H., Benner, C., Lichtinger, M., Schwarzfischer, L., Hu, Y., Andreesen, R., Chen, W., and Rehli, M. (2012). Dynamic epigenetic enhancer signatures reveal key transcription factors associated with monocytic differentiation states. Blood 119, e161–e171. Pham, T.H., Minderjahn, J., Schmidl, C., Hoffmeister, H., Schmidhofer, S., Chen, W., Langst, G., Benner, C., and Rehli, M. (2013). Mechanisms of in vivo binding site selection of the hematopoietic master transcription factor PU.1. Nucleic Acids Res. 41, 6391–6402. Pio, F., Kodandapani, R., Ni, C.Z., Shepard, W., Klemsz, M., McKercher, S.R., Maki, R.A., and Ely, K.R. (1996). New insights on DNA recognition by ets proteins from the crystal structure of the PU.1 ETS domain-DNA complex. J. Biol. Chem. 271, 23329–23337. Rosenbauer, F., Wagner, K., Kutok, J.L., Iwasaki, H., Le Beau, M.M., Okuno, Y., Akashi, K., Fiering, S., and Tenen, D.G. (2004). Acute myeloid leukemia induced by graded reduction of a lineagespecific transcription factor, PU.1. Nat. Genet. 36, 624–630. Schonheit, J., Kuhl, C., Gebhardt, M.L., Klett, F.F., Riemke, P., Scheller, M., Huang, G., Naumann, R., Leutz, A., Stocking, C., et al. (2013). PU.1 level-directed chromatin structure remodeling at the Irf8 gene drives dendritic cell commitment. Cell Rep. 3, 1617–1628. Staber, P.B., Zhang, P., Ye, M., Welner, R.S., Nombela-Arrieta, C., Bach, C., Kerenyi, M., Bartholdy, B.A., Zhang, H., Berich-Jorda, M., et al. (2013). Sustained PU.1 levels balance cell-cycle regulators to prevent exhaustion of adult hematopoietic stem cells. Mol. Cell 49, 934–946. Steidl, U., Rosenbauer, F., Verhaak, R.G., Gu, X., Ebralidze, A., Otu, H.H., Klippel, S., Steidl, C., Bruns, I., Costa, D.B., et al. (2006). Essential role of Jun family transcription factors in PU.1 knockdown-induced leukemic stem cells. Nat. Genet. 38, 1269–1277. Steidl, U., Steidl, C., Ebralidze, A., Chapuy, B., Han, H.J., Will, B., Rosenbauer, F., Becker, A., Wagner, K., Koschmieder, S., et al. (2007). A distal single nucleotide polymorphism alters longrange regulation of the PU.1 gene in acute myeloid leukemia. J. Clin. Invest. 117, 2611–2620. Sterner, D.E. and Berger, S.L. (2000). Acetylation of histones and transcription-related factors. Microbiol. Mol. Biol. Rev. 64, 435–459. Suzuki, M., Yamada, T., Kihara-Negishi, F., Sakurai, T., Hara, E., Tenen, D.G., Hozumi, N., and Oikawa, T. (2006). Site-specific DNA methylation by a complex of PU.1 and Dnmt3a/b. Oncogene 25, 2477–2488. Tolhuis, B., Palstra, R.J., Splinter, E., Grosveld, F., and de, L.W. (2002). Looping and interaction between hypersensitive sites in the active β-globin locus. Mol. Cell 10, 1453–1465.
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1274 B. van Riel and F. Rosenbauer: PU.1 and chromatin Traver, D., Miyamoto, T., Christensen, J., Iwasaki-Arai, J., Akashi, K., and Weissman, I.L. (2001). Fetal liver myelopoiesis occurs through distinct, prospectively isolatable progenitor subsets. Blood 98, 627–635. van Riel, B., Pakozdi, T., Brouwer, R., Monteiro, R., Tuladhar, K., Franke, V., Bryne, J.C., Jorna, R., Rijkers, E.J., van, I.W., et al. (2012). A novel complex, RUNX1-MYEF2, represses hematopoietic genes in erythroid cells. Mol. Cell Biol. 32, 3814–3822. Vangala, R.K., Heiss-Neumann, M.S., Rangatia, J.S., Singh, S.M., Schoch, C., Tenen, D.G., Hiddemann, W., and Behre, G. (2003). The myeloid master regulator transcription factor PU.1 is inactivated by AML1-ETO in t (8;21) myeloid leukemia. Blood 101, 270–277. Wang, Q., Stacy, T., Binder, M., Marin-Padilla, M., Sharpe, A.H., and Speck, N.A. (1996). Disruption of the Cbfa2 gene causes necrosis and hemorrhaging in the central nervous system and blocks definitive hematopoiesis. Proc. Natl. Acad. Sci. USA 93, 3444–3449. Wang, J.M., Lai, M.Z., and Yang-Yen, H.F. (2003). Interleukin-3 stimulation of mcl-1 gene transcription involves activation of the PU.1 transcription factor through a p38 mitogen-activated protein kinase-dependent pathway. Mol. Cell Biol. 23, 1896–1909. Wang, K., Wang, P., Shi, J., Zhu, X., He, M., Jia, X., Yang, X., Qiu, F., Jin, W., Qian, M., et al. (2010). PML/RARα targets promoter regions containing PU.1 consensus and RARE half sites in acute promyelocytic leukemia. Cancer Cell 17, 186–197. Wei, G.H., Badis, G., Berger, M.F., Kivioja, T., Palin, K., Enge, M., Bonke, M., Jolma, A., Varjosalo, M., Gehrke, A.R., et al. (2010). Genome-wide analysis of ETS-family DNA-binding in vitro and in vivo. EMBO J. 29, 2147–2160. Wilson, N.K., Foster, S.D., Wang, X., Knezevic, K., Schutte, J., Kaimakis, P., Chilarska, P.M., Kinston, S., Ouwehand, W.H., Dzierzak,
E., et al. (2010). Combinatorial transcriptional control in blood stem/progenitor cells: genome-wide analysis of ten major transcriptional regulators. Cell Stem Cell 7, 532–544. Yamamoto, H., Kihara-Negishi, F., Yamada, T., Hashimoto, Y., and Oikawa, T. (1999). Physical and functional interactions between the transcription factor PU.1 and the coactivator CBP. Oncogene 18, 1495–1501. Yasui, D., Miyano, M., Cai, S., Varga-Weisz, P., and Kohwi-Shigematsu, T. (2002). SATB1 targets chromatin remodelling to regulate genes over long distances. Nature 419, 641–645. Zarnegar, M.A., Chen, J., and Rothenberg, E.V. (2010). Cell-type-specific activation and repression of PU.1 by a complex of discrete, functionally specialized cis-regulatory elements. Mol. Cell Biol. 30, 4922–4939. Zhang, P., Behre, G., Pan, J., Iwama, A., Wara-Aswapati, N., Radomska, H.S., Auron, P.E., Tenen, D.G., and Sun, Z. (1999). Negative cross-talk between hematopoietic regulators: GATA proteins repress PU.1. Proc. Natl. Acad. Sci. USA 96, 8705–8710. Zhang, P., Zhang, X., Iwama, A., Yu, C., Smith, K.A., Mueller, B.U., Narravula, S., Torbett, B.E., Orkin, S.H., and Tenen, D.G. (2000). PU.1 inhibits GATA-1 function and erythroid differentiation by blocking GATA-1 DNA binding. Blood 96, 2641–2648. Zhang, P., Iwasaki-Arai, J., Iwasaki, H., Fenyus, M.L., Dayaram, T., Owens, B.M., Shigematsu, H., Levantini, E., Huettner, C.S., Lekstrom-Himes, J.A., et al. (2004). Enhancement of hematopoietic stem cell repopulating capacity and self-renewal in the absence of the transcription factor C/EBPα. Immunity 21, 853–863. Zhu, X., Zhang, H., Qian, M., Zhao, X., Yang, W., Wang, P., Zhang, J., and Wang, K. (2012). The significance of low PU.1 expression in patients with acute promyelocytic leukemia. J. Hematol. Oncol. 5, 22.
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Review Boet van Riel and Frank Rosenbauer*
Epigenetic control of hematopoiesis: the PU.1 chromatin connection Abstract: Purine-rich box1 (PU.1) is a transcription factor that not only has a key role in the development of most hematopoietic cell lineages but also in the suppression of leukemia. To exert these functions, PU.1 can cross-talk with multiple different proteins by forming complexes in order to activate or repress transcription. Among its protein partners are chromatin remodelers, DNA methyltransferases, and a number of other transcription factors with important roles in hematopoiesis. While a great deal of knowledge has been acquired about PU.1 function over the years, it was the development of novel genome-wide technologies, which boosted our understanding of how PU.1 acts on the chromatin to drive its repertoire of target genes. This review summarizes current knowledge and ideas of molecular mechanisms by which PU.1 controls hematopoiesis and suppresses leukemia. Keywords: chromatin; epigenetics; hematopoiesis; PU.1. DOI 10.1515/hsz-2014-0195 Received May 5, 2014; accepted July 3, 2014; previously published online September 2, 2014
Introduction The transcription factor purine-rich box1 (PU.1) is a key regulator for the development of several hematopoietic lineages. In fact, PU.1 is one of the most intensively studied transcription factors (TFs) in hematopoiesis, and a great deal of knowledge has been produced by now, resulting in more than 1460 publications that can be found in PubMed (as of April 2014: ncbi.nlm.nih.gov/ pubmed). The official gene name for PU.1 is SPI1 in human and Sfpi1 in the mouse. The PU.1-encoding gene was first *Corresponding author: Frank Rosenbauer, Institute of Molecular Tumor Biology, University of Münster, D-48149 Münster, Germany, e-mail: [email protected] Boet van Riel: Institute of Molecular Tumor Biology, University of Münster, D-48149 Münster, Germany
discovered as a target for insertions of the SFFV virus that causes erythroid leukemia (Moreau-Gachelin et al., 1988), followed by the isolation of the 31-kDa protein called PU.1. PU.1 is part of the ETS transcription factor family that consists of 27 different family members in humans (reviewed in Oikawa and Yamada, 2003). This family shares an ETS domain that facilitates direct DNA binding to the core GGAA motif (Wei et al., 2010). PU.1 has next to the ETS domain a winged helix domain that mediates binding to an extended consensus longer than the ETS-binding motif (Pio et al., 1996). Furthermore, PU.1 has an activation domain and a PEST domain involved in protein-protein interactions (Figure 1A). The human SPI1 is located on chromosome 11 and the mouse Sfpi1 on chromosome 2. Strict regulation of the expression of PU.1 is essential for proper hematopoietic development from hematopoietic stem cells (HSCs) into fully differentiated cells. Dysregulation of PU.1 expression in HSCs leads to a depletion of functional HSCs in adult mice (Staber et al., 2013). Also, later in the hematopoietic development, PU.1 expression is important for maturation of macrophages, B cells, early T cell progenitors, and T helper 9 (Th9) cells (Anderson et al., 1998; DeKoter and Singh, 2000; Anderson et al., 2002; Gerlach et al., 2014) (Figure 1B). In progenitor cells, there is also a feedback connection between PU.1 concentrations and the cell cycle that is important for myeloid vs. lymphoid lineage decision (Kueh et al., 2013). Moreover, proper PU.1 expression is essential to avoid the development of myeloid leukemia (Rosenbauer et al., 2004). Of note, downregulation, but not complete abolishment of PU.1 expression, appears to be of advantage for transformation of normal myeloid progenitors into leukemic cells (Steidl et al., 2006; Zhu et al., 2012). Such PU.1 downregulation during leukemia progression can be achieved by several mechanisms, which will be described in more detail later in this review. With the recent revolution in the development of novel genome-wide technologies such as next-generation sequencing methods, we have learned much about the global functions of PU.1 on the genome and are starting to get a glimpse of how PU.1 operates and epigenetically
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1266 B. van Riel and F. Rosenbauer: PU.1 and chromatin
Figure 1 Schematic representation of PU.1, its protein partners, and its role in hematopoiesis. (A) Schematic representation of the PU.1 protein structure with ammo acid numbering. Highlighted domains are the activation domain, PEST domain, and the ETS domain. Binding sites of protein partners are shown. (B) Simplified scheme of mammalian definitive hematopoiesis with cells not expressing PU.1 is depicted in red. Cells expressing PU.1 are depicted in blue: hematopoietic stem cells (HSC), common lymphoid progenitors (CLP), and common myeloid progenitors (CMP). Increase, decrease, or no change in PU.1 expression is symbolized between the cell symbols.
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B. van Riel and F. Rosenbauer: PU.1 and chromatin 1267
modifies the chromatin to initiate and maintain expression of its repertoire of target genes. In this review, we will provide an overview of known protein interaction partners by which PU.1 controls the epigenetic state at its genomic target regions and induces the development of myeloid leukemia.
PU.1 interaction with chromatin modifiers PU.1 can either activate or repress the transcription of genes, a notion that is supported by an in vivo study showing that after genetic knockdown of PU.1 in the HSCs, the expression of 225 genes was upregulated and of 97 genes was downregulated (Steidl et al., 2006). This differential control of gene expression is thought to be mediated by the ability of PU.1 to build different complexes with a number of protein partners, thus, either stimulating or inhibiting transcription. One of the first identified proteinbinding partners of PU.1 was the TATA-binding protein (TBP) (Hagemeier et al., 1993). Through TBP, PU.1 is able to recruit the basal transcription machinery to its binding sites at promoters and enhancers as a requirement for transcription (Kihara-Negishi et al., 2001). Indeed, many PU.1 target genes lack TATA box sequences and, thus, are unable to bind TBP directly (reviewed in Sandra Clarke and Siamon Gordon, 1998). Subsequent studies have shown that PU.1 can form complexes with a number of proteins that can directly modify chromatin or DNA. For example, PU.1 can bind the histone acetyltransferases CREB-binding protein (CBP) and p300 to activate transcription (Yamamoto et al., 1999; Bai et al., 2005). Acetylation of histone tails is thought to open the chromatin and allow other proteins to bind to the DNA and activate transcription (Sterner and Berger, 2000). Furthermore, PU.1 can also bind a histone deacetylase complex consisting of histone deacetylase 1 (HDAC1) and mammalian Sin3a (mSIN3a) (Kihara-Negishi et al., 2001). At least in reporter assays using the c-Myc promoter, HDAC1-mSIN3a functions as a repressor of PU.1 transactivation capacity (Kihara-Negishi et al., 2001). Later on, it was reported that together with HDAC1-mSIN3a, PU.1 can also bind the methyl CpG-binding protein 2 (MeCP2) to repress genes (Suzuki et al., 2006; Imoto et al., 2010). A possible link between methylated DNA sequences and PU.1 has also been suggested by the observation that the PU.1 protein is able to form a complex with the de novo DNA methyltransferases Dnmt3a and Dnmt3b (Suzuki et al., 2006). These interactions were discovered by
co-transfection experiments in 293T cells and are believed to site-specifically methylate DNA in order to repress transcription of selected genomic PU.1 target sites such as the tumor-suppressor gene p16 (INK4A) promoter (Suzuki et al., 2006). Importantly, the PU.1-DNMT3b complex has also been observed to occur endogenously during monocyte to ostoclast differentiation (de la Rica et al., 2013). However, genome-wide data have yet failed to identify a significant overlay between PU.1 occupancy and methylated DNA sequences (Pham et al., 2013), thus, challenging the concept that PU.1 can recruit the DNA methylation machinery to repress gene expression. Hence, further functional experiments are needed to solve this obvious discrepancy. As a limitation to the interpretation of the relevance of many of these PU.1 protein complexes, most of them have been identified (and perhaps exclusively observed) using in vitro co-transfection experiments in non- hematopoietic 293T or 3T3 cells. Thus, it remains unclear at the moment if at all, or in what hematopoietic cell types, these complexes exist endogenously. Nevertheless, these protein complex data do at least provide a hint for a model in which PU.1 can operate as a platform for directing the recruitment of different chromatin-modifying proteins, with either gene-repressive or -activating functions, to its target genes (Figures 1B and 2). Importantly, such a model is instrumental for explaining how one single transcription factor is capable of controlling expression of different repertoires of target genes in a dynamic and cell-type specific pattern.
PU.1 cross-talk with other lineagespecific transcription factors PU.1 can cross-talk with a number of other lineage-specific TFs, most of which are known as important regulators of blood cell development. In early hematopoietic differentiation stages such as HSCs and multipotent progenitors, CCAAT-enhancer-binding proteins (C/EBPs) and Runt-related transcription factor 1 (Runx1/AML1) play important roles in concert with PU.1. For example, several essential myeloid lineage growth factor receptor genes, such as those encoding the M-CSFR and GM-GSFRα, are regulated by a PU.1 collaboration with C/EBPα (Hohaus et al., 1995; Li et al., 2005). Runx1 is a critical factor for the development of the definitive HSC pool and also has a role in the specification of some hematopoietic lineages (Wang et al., 1996; van Riel et al., 2012). Similar to the PU.1-C/EBPα complex, PU.1 also collaborates with Runx1
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1268 B. van Riel and F. Rosenbauer: PU.1 and chromatin
Figure 2 Model for chromatin preparation for PU.1 binding to lower-affinity binding sites. For PU.1 to bind DNA, the chromatin has to be opened by other TFs. In the HSC, Runx1 opens PU.1 binding sites. Immunoresponsive TFs, like STAT6 and NF-κB, open latent enhancers for PU.1 binding, and in myeloid development, C/EBP needs to open additional enhancers in the Sfpi1 locus. This leads to PU.1 binding and recruitment of additional TFs and co-factors to regulate transcription. The figure also illustrates three examples of low-affinity PU.1 DNA consensus binding site in humans (Pham et al., 2013).
to activate transcription of the Csf1r gene encoding the M-CSF receptor (Li et al., 2005). Moreover, expression of the PU.1-encoding gene (Sfpi1), itself, is activated by the PU.1-Runx1 protein interaction, via their collaborative binding to adjacent consensus sites within a critical Sfpi1 enhancer, the -14-kb upstream regulatory element (-14 kb URE) (Huang et al., 2008). In myeloid cells, steadily increasing PU.1 expression is a prerequisite for guiding early progenitors through their complete differentiation program all the way toward reaching the mature state (Leddin et al., 2011). In agreement, it was shown that high expression levels of PU.1, together with C/EBPα or β, can convert other lineages to macrophages (Laiosa et al., 2006, Feng et al., 2008). During the developmental cascade, PU.1 appears to dynamically collaborate with changing protein-binding partners to orchestrate the step-wise process of differentiation. For example, as stated above, its collaboration with C/EBPα drives the expression of a number of genes, which are important for early myeloid differentiation (Hohaus et al., 1995). In agreement, using genetic studies in mice,
C/EBPα was found to be required for the transition from the common myeloid progenitor (CMP) to the granulocyte macrophage progenitor (GMP) differentiation step, but is no longer required thereafter (Traver et al., 2001; Zhang et al., 2004). A model seems likely, in which PU.1 transiently interacts with C/EBPα to guide the early myeloid developmental process. Once this differentiation step has past, the PU.1-to-C/EBPα interaction is probably dispensable for further myeloid differentiation steps. C/EBPβ was also shown to bind to PU.1 and to form a transactivating complex (Nagulapalli et al., 1995). C/EBPβ is a factor important for later myeloid differentiation steps (reviewed in Huber et al., 2012), thus suggesting a role for the PU.1-C/ EBPβ complex during terminal myeloid maturation. Concurrently, a likely scenario is that PU.1 actively switches between different C/EBP family members as binding partners to drive progenitors along myeloid differentiation. A repressive complex that is known to be important in granulocyte development is the one between PU.1 and growth factor independant-1 (GFI-1) (Dahl et al., 2007). GFI-1 is a transcriptional repressor, which is already important in HSCs (Lancrin et al., 2012). Interestingly, in myeloid cells of gfi-1-/- mice, PU.1 target genes are upregulated (Dahl et al., 2007). In agreement, GFI-1 can repress macrophage-specific PU.1 target genes, but has little to no effect on PU.1 target genes expressed in granulocytes (Dahl et al., 2007). This bias by GFI-1 is argued to be caused by differences in PU.1 concentrations needed for activation of macrophage and granulocytic genes (Dahl et al., 2007). Another salient example for the interaction of PU.1 with other lineage-specific TFs is its complex with GATAbinding factor 1 (Gata1) (Zhang et al., 1999). Gata1 is expressed in erythroid and megakaryocytic lineages (for reviews see Cantor and Orkin, 2002 and Goldfarb, 2007). It is the key driver of erythroid lineage differentiation, and mice in which Gata1 is lacking are devoid of erythroid progenitors (Cantor and Orkin, 2002). Moreover, Gata1 is erythroid lineage instructive because its ectopic expression in GMPs causes, in part, reprogramming to erythroid cells (Iwasaki et al., 2003). In myeloid progenitors, PU.1 binds to Gata1 and physically blocks Gata1 binding activity to the DNA (Zhang et al., 2000). PU.1 expression is downregulated at the erythroblast stage of definitive erythropoiesis (Hromas et al., 1993), and failure of its downregulation causes a block in erythroid differentiation and subsequent development of erythroid leukemia (Galson et al., 1993). Therefore, one important prerequisite for erythroid differentiation is the capacity of Gata1 to inhibit binding of co-activators to PU.1 and to thereby block its transcriptional activation activity (Zhang et al., 1999). This block includes transcription of the Sfpi1 gene,
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B. van Riel and F. Rosenbauer: PU.1 and chromatin 1269
itself, and thus shuts down PU.1 expression. Thus, the antagonistic interplay between PU.1 and Gata1 appears to be important for the selection of bipotential progenitors to differentiate into either myeloid or erythroid-megakaryocytic progeny. Within the lymphoid compartment, PU.1 is expressed in common lymphoid progenitors (CLPs), B cells, and the earliest T-cell progenitors in the thymus (Nutt et al., 2005). Importantly and similar to the erythroid lineage, PU.1 expression is required in later T-cell progenitors to become repressed (Anderson et al., 2002), stays repressed in Th2 cells and CD8 T cells, but is reexpressed in Th9 cells (Gerlach et al., 2014). To achieve PU.1 repression in early T cells, the activity of PU.1 is antagonized by Notch, which is essential to push thymocytes into further maturation (Del Real and Rothenberg, 2013). Moreover, T-cell-specific repression of PU.1 expression is thought to be facilitated by binding of Runx1 to a silencer element located between the Sfpi1 promoter and its upstream enhancer elements (Zarnegar et al., 2010). In summary, PU.1 exerts its functions in concert with a variety of other lineage-specific TFs (Figures 1B and 2). These collaborative and antagonistic interactions appear to be an important prerequisite for the dynamic orchestration of lineage fate choice and differentiation outcomes of hematopoietic progenitors.
Chromatin binding of PU.1 In recent years, as a result of the ability to map genomewide binding patterns of TFs by chromatin immunoprecipitation-sequencing (ChIP-seq), PU.1 has been found to operate as a key factor for initiation and maintaining transcriptionally permissive chromatin (Ostuni et al., 2013; Staber et al., 2013). Multiple ChIP-seq data sets have been generated to identify PU.1 chromatin occupancy patterns in a number of different hematopoietic cell types, including monocytes and macrophages (Heinz et al., 2010; Pham et al., 2012) in the presence or absence of immune stimulation (Ostuni et al., 2013), B cells (Heinz et al., 2010) and a HCS-like cell line (Wilson et al., 2010). These data showed that PU.1 occupies a great number of different genomic locations at promoters as well as intragenic and intergenic regions. The sequence of the PU.1 binding site can differ significantly throughout the genome. More than 2500 potential 12-mer consensus motifs with variable frequencies for PU.1 binding have been identified in monocytes and macrophages (Pham et al., 2013). Furthermore, the same study showed that only 1% of all potential PU.1
binding motifs within the genome are actually occupied by PU.1 (Pham et al., 2013). How does the selection of binding sites of TFs in the genome occur? The DNA binding consensus sequences, which in vitro were shown to have the highest affinity for PU.1 binding, are in vivo in a large part located within gene deserts, and lack any obvious binding motifs of other known TF nearby (Pham et al., 2013). Also, these isolated high-affinity PU.1 binding sites are often located in unconserved regions, and the biological significance of these sites is currently unclear. The functional binding sites of PU.1 in promoters and enhancers have an overall lower binding affinity for PU.1 and are principally always flanked by binding sites of one or more additional TFs (Pham et al., 2013). The most likely scenario for functional binding site selection seems, therefore, to be a combination of ‘low’ affinity with adjacent sites to allow combinatorial binding of additional TFs. Collaborative TF binding with PU.1 is either important to stabilize PU.1 DNA occupancy, or to enhance the transactivating function of PU.1, or most likely both. Also, in many cases, it will be the occupancy of all collaborative factors, which is necessary to create and maintain open chromatin (Heinz et al., 2010, 2013). The relative expression level of PU.1 in progenitor cells has been shown to act as a driving event of lineage choice selection during hematopoietic differentiation (DeKoter and Singh, 2000; Leddin et al., 2011). PU.1 protein concentrations appear to have an impact on the occurrence of PU.1 binding to functional DNA elements after comparing occupancy between hematopoietic progenitor cells and monocytes (Pham et al., 2013). With the increase in protein concentration, the number of binding sites increases, and more low-affinity sites are bound (Pham et al., 2013). Another important question is how PU.1 penetrates the chromatin at its different binding sites. Most likely, PU.1 competes with nucleosomes at high-affinity sites, which is indicated by its binding to sites that show high affinity in vitro but are not hypersensitive for DNase I in vivo and are most likely devoid of collaborative binding events (Pham et al., 2013). In contrast, lower-affinity binding sites may not be accessed by PU.1 alone and are frequently flanked by additional TF binding sites when bound. This assumption is supported by the finding that the chromatin at several PU.1 binding sites have to be opened or primed earlier in development by global chromatin changes mediated by factors like Runx1 (Hoogenkamp et al., 2009) and that PU.1 is unable to rescue the Runx1 knock-out phenotype (Lancrin et al., 2012). It appears that PU.1 needs a pre-prepared global ‘chromatin context’ in the cell for functional binding and
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1270 B. van Riel and F. Rosenbauer: PU.1 and chromatin subsequent transactivation of important target genes. Indeed, a recent report described a set of enhancers that was inactive in unstimulated macrophages but became active after immune stimulation (Ostuni et al., 2013). PU.1 was shown to bind to these so-called ‘latent’ enhancers only after immune stimulation, and PU.1 binding was preceded and dependent on binding by immune responsive TFs such as SMAD6 and NF-κB. Another example is that high expression of PU.1 in macrophages requires two autoregulatory loops at its Sfpi1 gene, built via the -14-kb and -12-kb enhancers (Leddin et al., 2011). However, before PU.1 is able to bind to the macrophagespecific -12-kb enhancer, C/EBPα has to bind to the -14-kb enhancer to subsequently ‘open’ the chromatin at the -12-kb enhancer so that PU.1 can enter and bind (Leddin et al., 2011). It is likely that similar to the situation at its own gene locus, the 1% of the potential binding sites that are indeed occupied by PU.1 in the human genome is globally selected by being located in active DNA regions that had been ‘opened’ for PU.1 to enter chromatin. A model of PU.1 occupancy at lower-affinity binding sites is shown in Figure 2. As yet, we have a very poor understanding, if at all, of the impact of posttranslational modifications at the PU.1 peptide on the in vivo function and DNA binding site selection of PU.1. Several posttranslational PU.1 modifications have been described. By transient transfection studies, phosphorylation of serine 148 and 142 has been shown to influence PU.1 transactivation capacity (Lodie et al., 1997; Wang et al., 2003). Interestingly, acetylation of PU.1 has an influence on the protein complex selection of PU.1 in vitro (Lodie et al., 1997; Kihara-Negishi et al., 2005). Mutation of the lysine stretch within the C-terminal end of the ETS domain of PU.1 into arginines, which cannot be acetylated anymore, had little effect on the binding potential of PU.1 to CBP, but reduced binding to mSIN3a using GST pull down assays. Therefore, it is likely that posttranslational modifications can, at least in part, influence the ability of PU.1 to direct the differentiation choice by affecting the nature of the protein complexes.
PU.1 and the three-dimensional chromatin structure Distal regulatory gene elements such as enhancers, insulators, and silencers must often cross-talk with proximal promoters over long genomic distances (for reviews on this subject see Pennacchio et al., 2013; Chetverina
et al., 2014). Several models have been put forward to explain how this cross-talk may occur, one of which is DNA looping. Indeed, owing to novel technological developments such as chromosome conformation capturing (3C) and related approaches, there is now overwhelming evidence that DNA looping operates as a major gene regulatory mechanism (Carter et al., 2002; Dekker et al., 2002; Tolhuis et al., 2002). However, formation of such three-dimensional chromatin structures must be tightly controlled to participate in the dynamic regulation of gene expression. Indeed, several lineage-specific hemato poietic TFs, such as PU.1 and Runx1, have been shown to facilitate DNA looping (Levantini et al., 2011; Schonheit et al., 2013; Staber et al., 2013). PU.1 binds to its -14-kb URE enhancer to autoregulate expression of its own Spfi1 gene (Li et al., 2001). This enhancer was found to loop to the proximal Sfpi1 promoter (Ebralidze et al., 2008), a process, which is impaired by deletion of the PU.1 binding motifs with the -14-kb URE (Staber et al., 2013). Interestingly, dependency of this looping structure between the -14-kb URE and the proximal promoter required PU.1 binding exclusively in HSCs but not in macrophages (Staber et al., 2013). A likely explanation for this cell type-associated dynamics lies in an additional Sfpi1 enhancer element (-12 kb enhancer) in the vicinity of the -14-kb URE, which is active in macrophages but not in HSCs (Leddin et al., 2011). This enhancer might preserve the DNA loop even while PU.1 binding to the -14-kb URE is abolished. Another example of a regulatory gene element of which chromatin looping is dependent on PU.1 is the -50-kb enhancer in the Irf8 locus (Schonheit et al., 2013). Here, knock-down of PU.1 decreases looping frequency between this enhancer and the proximal Irf8 promoter. An important question that these single gene studies did not yet answer is whether on a more global scale, myeloid enhancers, in general, depend on PU.1 binding to facilitate looping with their respective genes. However, this appears to be likely, because studies in B cells have shown that PU.1 occupancy is associated with enhancers and genomic regions involved in DNA looping (Heinz et al., 2010; Lin et al., 2012). Another important question is on the molecular mechanism by which PU.1 controls DNA looping. The most likely answer to this question, again, may be found in the choice of protein partners, which may require PU.1 occupancy at the chromatin to bind to DNA and subsequently facilitate chromosomal looping. The use of genome-wide variants of the 3C technology (Carter et al., 2002; Tolhuis et al., 2002) in myeloid cells will probably have the power to provide answers to these important questions.
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B. van Riel and F. Rosenbauer: PU.1 and chromatin 1271
PU.1, chromatin, and myeloid leukemia It has been known for several years that decreased PU.1 expression levels can directly lead to the development of AML in mice (Rosenbauer et al., 2004). Of note, however, only very few cases with mutations in SPI1 have been reported in human AML (Mueller et al., 2003). However, PU.1 has been shown to be frequently downregulated by leukemia-driving oncogenes in AML patients, suggesting that similar to mice, PU.1 can act as a tumor suppressor in man (Vangala et al., 2003; Mueller et al., 2006). Although one study in mice has shown that complete absence of PU.1 can lead to leukemia (Metcalf et al., 2006), in humans, complete loss of PU.1 function has not yet been reported, suggesting that its graded reduction is supportive for cancer development. This is similar to the situation of C/EBPα mutations in AML, which lead to reduced but not completely abolished function (Pabst and Mueller, 2009). An explanation why residual PU.1 expression may be required for AML development has recently been illustrated by the genome-wide finding that the PML-RARα oncogene requires PU.1 for its chromatin entry and control of target genes (Wang et al., 2010). Thus, it appears that a fine-tuned reduction in PU.1 expression may provide the optimal concentration for blocking normal myeloid differentiation, on the one hand, and on the other hand, still allows oncogenes, such as PML/RARα, to regulate their target genes. Interestingly, in patients with complex karyotype AML, a single nucleotide polymorphism (SNP) in the SPI1 URE enhancer, which disrupts binding of the chromatin organizer special AT-rich sequence-binding protein-1 (SATb1) occurs with higher frequency than in healthy cells, suggesting a role for this SNP in leukemia predisposition (Steidl et al., 2007). Indeed, functional investigation of this SNP has shown that it leads to reduced levels of PU.1 (Steidl et al., 2007). Intriguingly, SATb1 is involved in the control of DNA looping factor in early T cells (Yasui et al., 2002), and it therefore seems likely, but not yet proven, that the SNP within the PU.1 enhancer might impair its DNA looping to the proximal promoter. In four out of 120 AML patients, sequence variations in a NF-κB binding site in the URE have been identified, and it was shown that this aberration resulted in impaired PU.1 expression (Bonadies et al., 2010). Importantly, NF-κB has recently been revealed to coincide with de novo PU.1 binding to many immune-responsive enhancers in macrophages (Ostuni et al., 2013). Hence, it is tempting to speculate that also in leukemia cells, failed NF-kB binding
to the URE may reduce chromatin entry of PU.1 to autoregulate its own expression.
Conclusion PU.1 is a central regulator of hematopoiesis, and intensive research over almost three decades has provided ample understanding of its pattern of expression and gene regulation, its role in differentiation, its function as a tumor suppressor in myeloid leukemia, its genomewide nature of chromatin occupancy and binding site selection, and its many protein interaction partners. So, a valid question is why is continuing research on PU.1 still needed? The main reason is that in spite of the existing wealth of knowledge, many important questions still remain. For example, how do the different PU.1 expression levels drive specific hematopoietic differentiation outcomes at the chromatin level? Are there different PU.1 concentration-dependent protein-protein interactions, and/or do the different PU.1 concentrations lead to a selection of different binding sites in the genome? Also, why does PU.1 bind to some of its genomic consensus motifs but not to others, and how is this selection regulated? An example highlighting this specific question is the lack of PU.1 occupancy at ‘latent’ enhancers prior to, but not after, immune stimulation of macrophages. This leads to the question why is PU.1 able to prepare enhancers at some genomic sites but not at others? Another interesting unresolved topic is on the molecular nature of the PU.1-dependent chromatin looping structures. For example, how does PU.1 affect the spatial organization of the chromatin? Often, addressing novel questions requires the utilization of newly emerging technologies. Importantly, it is a major advantage of our great knowledge on PU.1 that exists today to enable us to put new findings into the proper molecular and cellular perspectives. Thus, PU.1 does not only serve as an important factor for the regulation of chromatin but also continues to serve as a role model in research on transcription factors. Acknowledgments: We thank Michael Rehli (Regensburg, Germany) for valuable suggestions on the manuscript. The literature regarding PU.1 is overwhelming, and we wish to apologize to those whose work we were unable to cite. This work was supported by the Deutsche Forschungsgemeinschaft Research Unit 1336 and Cells-in-Motion Cluster of Excellence (EXC 1003 – CiM), University of Münster, Germany.
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1272 B. van Riel and F. Rosenbauer: PU.1 and chromatin
References Anderson, K.L., Smith, K.A., Conners, K., McKercher, S.R., Maki, R.A., and Torbett, B.E. (1998). Myeloid development is selectively disrupted in PU.1 null mice. Blood 91, 3702–3710. Anderson, M.K., Weiss, A.H., Hernandez-Hoyos, G., Dionne, C.J., and Rothenberg, E.V. (2002). Constitutive expression of PU.1 in fetal hematopoietic progenitors blocks T cell development at the pro-T cell stage. Immunity 16, 285–296. Bai, Y., Srinivasan, L., Perkins, L., and Atchison, M.L. (2005). Protein acetylation regulates both PU.1 transactivation and Ig κ 3′ enhancer activity. J. Immunol. 175, 5160–5169. Bonadies, N., Neururer, C., Steege, A., Vallabhapurapu, S., Pabst, T., and Mueller, B.U. (2010). PU.1 is regulated by NF-κB through a novel binding site in a 17 kb upstream enhancer element. Oncogene 29, 1062–1072. Cantor, A.B. and Orkin, S.H. (2002). Transcriptional regulation of erythropoiesis: an affair involving multiple partners. Oncogene 21, 3368–3376. Carter, D., Chakalova, L., Osborne, C.S., Dai, Y.F., and Fraser, P. (2002). Long-range chromatin regulatory interactions in vivo. Nat. Genet. 32, 623–626. Chetverina, D., Aoki, T., Erokhin, M., Georgiev, P., and Schedl, P. (2014). Making connections: insulators organize eukaryotic chromosomes into independent cis-regulatory networks. Bioessays 36, 163–172. Clarke S. and Gordon S. (1998). Myeloid-specific gene expression. J. Leukoc. Biol. 63, 153–168. Dahl, R., Iyer, S.R., Owens, K.S., Cuylear, D.D., and Simon, M.C. (2007). The transcriptional repressor GFI-1 antagonizes PU.1 activity through protein-protein interaction. J. Biol. Chem. 282, 6473–6483. de la Rica, L., Rodriguez-Ubreva, J., Garcia, M., Islam, A.B., Urquiza, J.M., Hernando, H., Christensen, J., Helin, K., Gomez-Vaquero, C., and Ballestar, E. (2013). PU.1 target genes undergo Tet2coupled demethylation and DNMT3b-mediated methylation in monocyte-to-osteoclast differentiation. Genome Biol. 14, R99. Dekker, J., Rippe, K., Dekker, M., and Kleckner, N. (2002). Capturing chromosome conformation. Science 295, 1306–1311. DeKoter, R.P. and Singh, H. (2000). Regulation of B lymphocyte and macrophage development by graded expression of PU.1. Science 288, 1439–1441. Del Real, M.M. and Rothenberg, E.V. (2013). Architecture of a lymphomyeloid developmental switch controlled by PU.1, Notch and Gata3. Development 140, 1207–1219. Ebralidze, A.K., Guibal, F.C., Steidl, U., Zhang, P., Lee, S., Bartholdy, B., Jorda, M.A., Petkova, V., Rosenbauer, F., Huang, G., et al. (2008). PU.1 expression is modulated by the balance of functional sense and antisense RNAs regulated by a shared cisregulatory element. Genes Dev. 22, 2085–2092. Feng, R., Desbordes, S.C., Xie, H., Tillo, E.S., Pixley, F., Stanley, E.R., and Graf, T. (2008). PU.1 and C/EBPα/β convert fibroblasts into macrophage-like cells. Proc. Natl. Acad. Sci. USA 105, 6057–6062. Galson, D.L., Hensold, J.O., Bishop, T.R., Schalling, M., D’Andrea, A.D., Jones, C., Auron, P.E., and Housman, D.E. (1993). Mouse β-globin DNA-binding protein B1 is identical to a proto-oncogene, the transcription factor Spi-1/PU.1, and is restricted in expression to hematopoietic cells and the testis. Mol. Cell Biol. 13, 2929–2941.
Gerlach, K., Hwang, Y., Nikolaev, A., Atreya, R., Dornhoff, H., Steiner, S., Lehr, H.A., Wirtz, S., Vieth, M., Waisman, A., et al. (2014). T9 cells that express the transcription factor PU.1 drive T cell-mediated colitis via IL-9 receptor signaling in intestinal epithelial cells. Nat. Immunol. 15, 676–686. Goldfarb, A.N. (2007). Transcriptional control of megakaryocyte development. Oncogene 26, 6795–6802. Hagemeier, C., Bannister, A.J., Cook, A., and Kouzarides, T. (1993). The activation domain of transcription factor PU.1 binds the retinoblastoma (RB) protein and the transcription factor TFIID in vitro: RB shows sequence similarity to TFIID and TFIIB. Proc. Natl. Acad. Sci. USA 90, 1580–1584. Heinz, S., Benner, C., Spann, N., Bertolino, E., Lin, Y.C., Laslo, P., Cheng, J.X., Murre, C., Singh, H., and Glass, C.K. (2010). Simple combinations of lineage-determining transcription factors prime cis-regulatory elements required for macrophage and B cell identities. Mol. Cell 38, 576–589. Heinz, S., Romanoski, C.E., Benner, C., Allison, K.A., Kaikkonen, M.U., Orozco, L.D., and Glass, C.K. (2013). Effect of natural genetic variation on enhancer selection and function. Nature 503, 487–492. Hohaus, S., Petrovick, M.S., Voso, M.T., Sun, Z., Zhang, D.E., and Tenen, D.G. (1995). PU.1 (Spi-1) and C/EBPα regulate expression of the granulocyte-macrophage colonystimulating factor receptor α gene. Mol. Cell Biol. 15, 5830–5845. Hoogenkamp, M., Lichtinger, M., Krysinska, H., Lancrin, C., Clarke, D., Williamson, A., Mazzarella, L., Ingram, R., Jorgensen, H., Fisher, A., et al. (2009). Early chromatin unfolding by RUNX1: a molecular explanation for differential requirements during specification versus maintenance of the hematopoietic gene expression program. Blood 114, 299–309. Hromas, R., Orazi, A., Neiman, R.S., Maki, R., Van, B.C., Moore, J., and Klemsz, M. (1993). Hematopoietic lineage- and stagerestricted expression of the ETS oncogene family member PU.1. Blood 82, 2998–3004. Huang, G., Zhang, P., Hirai, H., Elf, S., Yan, X., Chen, Z., Koschmieder, S., Okuno, Y., Dayaram, T., Growney, J.D., et al. (2008). PU.1 is a major downstream target of AML1 (RUNX1) in adult mouse hematopoiesis. Nat. Genet. 40, 51–60. Huber, R., Pietsch, D., Panterodt, T., and Brand, K. (2012). Regulation of C/EBPβ and resulting functions in cells of the monocytic lineage. Cell Signal. 24, 1287–1296. Imoto, A., Okada, M., Okazaki, T., Kitasato, H., Harigae, H., and Takahashi, S. (2010). Metallothionein-1 isoforms and vimentin are direct PU.1 downstream target genes in leukemia cells. J. Biol. Chem. 285, 10300–10309. Iwasaki, H., Mizuno, S., Wells, R.A., Cantor, A.B., Watanabe, S., and Akashi, K. (2003). GATA-1 converts lymphoid and myelomonocytic progenitors into the megakaryocyte/erythrocyte lineages. Immunity. 19, 451–462. Kihara-Negishi, F., Yamamoto, H., Suzuki, M., Yamada, T., Sakurai, T., Tamura, T., and Oikawa, T. (2001). In vivo complex formation of PU.1 with HDAC1 associated with PU.1-mediated transcriptional repression. Oncogene 20, 6039–6047. Kihara-Negishi, F., Suzuki, M., Yamada, T., Sakurai, T., and Oikawa, T. (2005). Impaired repressor activity and biological functions of PU.1 in MEL cells induced by mutations in the acetylation motifs within the ETS domain. Biochem. Biophys. Res. Commun. 335, 477–484.
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B. van Riel and F. Rosenbauer: PU.1 and chromatin 1273 Kueh, H.Y., Champhekar, A., Nutt, S.L., Elowitz, M.B., and Rothenberg, E.V. (2013). Positive feedback between PU.1 and the cell cycle controls myeloid differentiation. Science 341, 670–673. Laiosa, C.V., Stadtfeld, M., Xie, H., de Andres-Aguayo, L., and Graf, T. (2006). Reprogramming of committed T cell progenitors to macrophages and dendritic cells by C/EBPα and PU.1 transcription factors. Immunity 25, 731–744. Lancrin, C., Mazan, M., Stefanska, M., Patel, R., Lichtinger, M., Costa, G., Vargel, O., Wilson, N.K., Moroy, T., Bonifer, C., et al. (2012). GFI1 and GFI1B control the loss of endothelial identity of hemogenic endothelium during hematopoietic commitment. Blood 120, 314–322. Leddin, M., Perrod, C., Hoogenkamp, M., Ghani, S., Assi, S., Heinz, S., Wilson, N.K., Follows, G., Schonheit, J., Vockentanz, L., et al. (2011). Two distinct auto-regulatory loops operate at the PU.1 locus in B cells and myeloid cells. Blood 117, 2827–2838. Levantini, E., Lee, S., Radomska, H.S., Hetherington, C.J., BerichJorda, M., Amabile, G., Zhang, P., Gonzalez, D.A., Zhang, J., Basseres, D.S., et al. (2011). RUNX1 regulates the CD34 gene in haematopoietic stem cells by mediating interactions with a distal regulatory element. EMBO J. 30, 4059–4070. Li, Y., Okuno, Y., Zhang, P., Radomska, H.S., Chen, H., Iwasaki, H., Akashi, K., Klemsz, M.J., McKercher, S.R., Maki, R.A., et al. (2001). Regulation of the PU.1 gene by distal elements. Blood 98, 2958–2965. Li, X., Vradii, D., Gutierrez, S., Lian, J.B., van Wijnen, A.J., Stein, J.L., Stein, G.S., and Javed, A. (2005). Subnuclear targeting of Runx1 is required for synergistic activation of the myeloid specific M-CSF receptor promoter by PU.1. J. Cell Biochem. 96, 795–809. Lin, Y.C., Benner, C., Mansson, R., Heinz, S., Miyazaki, K., Miyazaki, M., Chandra, V., Bossen, C., Glass, C.K., and Murre, C. (2012). Global changes in the nuclear positioning of genes and intraand interdomain genomic interactions that orchestrate B cell fate. Nat. Immunol. 13, 1196–1204. Lodie, T.A., Savedra, R., Jr., Golenbock, D.T., Van Beveren, C.P., Maki, R.A., and Fenton, M.J. (1997). Stimulation of macrophages by lipopolysaccharide alters the phosphorylation state, conformation, and function of PU.1 via activation of casein kinase II. J. Immunol. 158, 1848–1856. Metcalf, D., Dakic, A., Mifsud, S., Di, R.L., Wu, L., and Nutt, S. (2006). Inactivation of PU.1 in adult mice leads to the development of myeloid leukemia. Proc. Natl. Acad. Sci. USA 103, 1486–1491. Moreau-Gachelin, F., Tavitian, A., and Tambourin, P. (1988). Spi-1 is a putative oncogene in virally induced murine erythroleukaemias. Nature 331, 277–280. Mueller, B.U., Pabst, T., Osato, M., Asou, N., Johansen, L.M., Minden, M.D., Behre, G., Hiddemann, W., Ito, Y., and Tenen, D.G. (2003). Heterozygous PU.1 mutations are associated with acute myeloid leukemia. Blood 101, 2074. Mueller, B.U., Pabst, T., Fos, J., Petkovic, V., Fey, M.F., Asou, N., Buergi, U., and Tenen, D.G. (2006). ATRA resolves the differentiation block in t (15;17) acute myeloid leukemia by restoring PU.1 expression. Blood 107, 3330–3338. Nagulapalli, S., Pongubala, J.M., and Atchison, M.L. (1995). Multiple proteins physically interact with PU.1. Transcriptional synergy with NF-IL6 β (C/EBPδ, CRP3). J. Immunol. 155, 4330–4338. Nutt, S.L., Metcalf, D., D’Amico, A., Polli, M., and Wu, L. (2005). Dynamic regulation of PU.1 expression in multipotent hematopoietic progenitors. J. Exp. Med. 201, 221–231.
Oikawa, T. and Yamada, T. (2003). Molecular biology of the Ets family of transcription factors. Gene 303, 11–34. Ostuni, R., Piccolo, V., Barozzi, I., Polletti, S., Termanini, A., Bonifacio, S., Curina, A., Prosperini, E., Ghisletti, S., and Natoli, G. (2013). Latent enhancers activated by stimulation in differentiated cells. Cell 152, 157–171. Pabst, T. and Mueller, B.U. (2009). Complexity of CEBPA dysregulation in human acute myeloid leukemia. Clin. Cancer Res. 15, 5303–5307. Pennacchio, L.A., Bickmore, W., Dean, A., Nobrega, M.A., and Bejerano, G. (2013). Enhancers: five essential questions. Nat. Rev. Genet. 14, 288–295. Pham, T.H., Benner, C., Lichtinger, M., Schwarzfischer, L., Hu, Y., Andreesen, R., Chen, W., and Rehli, M. (2012). Dynamic epigenetic enhancer signatures reveal key transcription factors associated with monocytic differentiation states. Blood 119, e161–e171. Pham, T.H., Minderjahn, J., Schmidl, C., Hoffmeister, H., Schmidhofer, S., Chen, W., Langst, G., Benner, C., and Rehli, M. (2013). Mechanisms of in vivo binding site selection of the hematopoietic master transcription factor PU.1. Nucleic Acids Res. 41, 6391–6402. Pio, F., Kodandapani, R., Ni, C.Z., Shepard, W., Klemsz, M., McKercher, S.R., Maki, R.A., and Ely, K.R. (1996). New insights on DNA recognition by ets proteins from the crystal structure of the PU.1 ETS domain-DNA complex. J. Biol. Chem. 271, 23329–23337. Rosenbauer, F., Wagner, K., Kutok, J.L., Iwasaki, H., Le Beau, M.M., Okuno, Y., Akashi, K., Fiering, S., and Tenen, D.G. (2004). Acute myeloid leukemia induced by graded reduction of a lineagespecific transcription factor, PU.1. Nat. Genet. 36, 624–630. Schonheit, J., Kuhl, C., Gebhardt, M.L., Klett, F.F., Riemke, P., Scheller, M., Huang, G., Naumann, R., Leutz, A., Stocking, C., et al. (2013). PU.1 level-directed chromatin structure remodeling at the Irf8 gene drives dendritic cell commitment. Cell Rep. 3, 1617–1628. Staber, P.B., Zhang, P., Ye, M., Welner, R.S., Nombela-Arrieta, C., Bach, C., Kerenyi, M., Bartholdy, B.A., Zhang, H., Berich-Jorda, M., et al. (2013). Sustained PU.1 levels balance cell-cycle regulators to prevent exhaustion of adult hematopoietic stem cells. Mol. Cell 49, 934–946. Steidl, U., Rosenbauer, F., Verhaak, R.G., Gu, X., Ebralidze, A., Otu, H.H., Klippel, S., Steidl, C., Bruns, I., Costa, D.B., et al. (2006). Essential role of Jun family transcription factors in PU.1 knockdown-induced leukemic stem cells. Nat. Genet. 38, 1269–1277. Steidl, U., Steidl, C., Ebralidze, A., Chapuy, B., Han, H.J., Will, B., Rosenbauer, F., Becker, A., Wagner, K., Koschmieder, S., et al. (2007). A distal single nucleotide polymorphism alters longrange regulation of the PU.1 gene in acute myeloid leukemia. J. Clin. Invest. 117, 2611–2620. Sterner, D.E. and Berger, S.L. (2000). Acetylation of histones and transcription-related factors. Microbiol. Mol. Biol. Rev. 64, 435–459. Suzuki, M., Yamada, T., Kihara-Negishi, F., Sakurai, T., Hara, E., Tenen, D.G., Hozumi, N., and Oikawa, T. (2006). Site-specific DNA methylation by a complex of PU.1 and Dnmt3a/b. Oncogene 25, 2477–2488. Tolhuis, B., Palstra, R.J., Splinter, E., Grosveld, F., and de, L.W. (2002). Looping and interaction between hypersensitive sites in the active β-globin locus. Mol. Cell 10, 1453–1465.
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1274 B. van Riel and F. Rosenbauer: PU.1 and chromatin Traver, D., Miyamoto, T., Christensen, J., Iwasaki-Arai, J., Akashi, K., and Weissman, I.L. (2001). Fetal liver myelopoiesis occurs through distinct, prospectively isolatable progenitor subsets. Blood 98, 627–635. van Riel, B., Pakozdi, T., Brouwer, R., Monteiro, R., Tuladhar, K., Franke, V., Bryne, J.C., Jorna, R., Rijkers, E.J., van, I.W., et al. (2012). A novel complex, RUNX1-MYEF2, represses hematopoietic genes in erythroid cells. Mol. Cell Biol. 32, 3814–3822. Vangala, R.K., Heiss-Neumann, M.S., Rangatia, J.S., Singh, S.M., Schoch, C., Tenen, D.G., Hiddemann, W., and Behre, G. (2003). The myeloid master regulator transcription factor PU.1 is inactivated by AML1-ETO in t (8;21) myeloid leukemia. Blood 101, 270–277. Wang, Q., Stacy, T., Binder, M., Marin-Padilla, M., Sharpe, A.H., and Speck, N.A. (1996). Disruption of the Cbfa2 gene causes necrosis and hemorrhaging in the central nervous system and blocks definitive hematopoiesis. Proc. Natl. Acad. Sci. USA 93, 3444–3449. Wang, J.M., Lai, M.Z., and Yang-Yen, H.F. (2003). Interleukin-3 stimulation of mcl-1 gene transcription involves activation of the PU.1 transcription factor through a p38 mitogen-activated protein kinase-dependent pathway. Mol. Cell Biol. 23, 1896–1909. Wang, K., Wang, P., Shi, J., Zhu, X., He, M., Jia, X., Yang, X., Qiu, F., Jin, W., Qian, M., et al. (2010). PML/RARα targets promoter regions containing PU.1 consensus and RARE half sites in acute promyelocytic leukemia. Cancer Cell 17, 186–197. Wei, G.H., Badis, G., Berger, M.F., Kivioja, T., Palin, K., Enge, M., Bonke, M., Jolma, A., Varjosalo, M., Gehrke, A.R., et al. (2010). Genome-wide analysis of ETS-family DNA-binding in vitro and in vivo. EMBO J. 29, 2147–2160. Wilson, N.K., Foster, S.D., Wang, X., Knezevic, K., Schutte, J., Kaimakis, P., Chilarska, P.M., Kinston, S., Ouwehand, W.H., Dzierzak,
E., et al. (2010). Combinatorial transcriptional control in blood stem/progenitor cells: genome-wide analysis of ten major transcriptional regulators. Cell Stem Cell 7, 532–544. Yamamoto, H., Kihara-Negishi, F., Yamada, T., Hashimoto, Y., and Oikawa, T. (1999). Physical and functional interactions between the transcription factor PU.1 and the coactivator CBP. Oncogene 18, 1495–1501. Yasui, D., Miyano, M., Cai, S., Varga-Weisz, P., and Kohwi-Shigematsu, T. (2002). SATB1 targets chromatin remodelling to regulate genes over long distances. Nature 419, 641–645. Zarnegar, M.A., Chen, J., and Rothenberg, E.V. (2010). Cell-type-specific activation and repression of PU.1 by a complex of discrete, functionally specialized cis-regulatory elements. Mol. Cell Biol. 30, 4922–4939. Zhang, P., Behre, G., Pan, J., Iwama, A., Wara-Aswapati, N., Radomska, H.S., Auron, P.E., Tenen, D.G., and Sun, Z. (1999). Negative cross-talk between hematopoietic regulators: GATA proteins repress PU.1. Proc. Natl. Acad. Sci. USA 96, 8705–8710. Zhang, P., Zhang, X., Iwama, A., Yu, C., Smith, K.A., Mueller, B.U., Narravula, S., Torbett, B.E., Orkin, S.H., and Tenen, D.G. (2000). PU.1 inhibits GATA-1 function and erythroid differentiation by blocking GATA-1 DNA binding. Blood 96, 2641–2648. Zhang, P., Iwasaki-Arai, J., Iwasaki, H., Fenyus, M.L., Dayaram, T., Owens, B.M., Shigematsu, H., Levantini, E., Huettner, C.S., Lekstrom-Himes, J.A., et al. (2004). Enhancement of hematopoietic stem cell repopulating capacity and self-renewal in the absence of the transcription factor C/EBPα. Immunity 21, 853–863. Zhu, X., Zhang, H., Qian, M., Zhao, X., Yang, W., Wang, P., Zhang, J., and Wang, K. (2012). The significance of low PU.1 expression in patients with acute promyelocytic leukemia. J. Hematol. Oncol. 5, 22.
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