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Cancer Research 67, 1461-1471, February 15, 2007. doi: 10.1158/0008-5472.CAN-06-2615
© 2007 American Association for Cancer Research
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Molecular Biology, Pathobiology, and Genetics

Activation of TLX3 and NKX2-5 in t(5;14)(q35;q32) T-Cell Acute Lymphoblastic Leukemia by Remote 3'-BCL11B Enhancers and Coregulation by PU.1 and HMGA1

Stefan Nagel1, Michaela Scherr2, Alexander Kel3, Klaus Hornischer3, Gregory E. Crawford4, Maren Kaufmann1, Corinna Meyer1, Hans G. Drexler1 and Roderick A.F. MacLeod1

1 Deutsche Sammlung von Mikroorganismen und Zellkulturen (German Collection of Microorganisms and Cell Cultures), Department of Cell Cultures, Braunschweig, Germany; 2 Department of Hematology, Hemostasis and Oncology, Hanover Medical School, Hanover, Germany; 3 BIOBASE GmbH, Wolfenbüttel, Germany; and 4 Duke University, Institute for Genome Sciences and Policy, Durham, North Carolina

Requests for reprints: Roderick A.F. MacLeod or Stefan Nagel, Deutsche Sammlung von Mikroorganismen und Zellkulturen, Department of Cell Cultures, Inhoffenstrasse 7B, 38124 Braunschweig, Germany. Phone: 49-531-261-6167; Fax: 49-531-261-6150; E-mail: rml{at}dsmz.de or sna{at}dsmz.de.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In T-cell acute lymphoblastic leukemia, alternative t(5;14)(q35;q32.2) forms effect dysregulation of either TLX3 or NKX2-5 homeobox genes at 5q35 by juxtaposition with 14q32.2 breakpoints dispersed across the BCL11B downstream genomic desert. Leukemic gene dysregulation by t(5;14) was investigated by DNA inhibitory treatments with 26-mer double-stranded DNA oligonucleotides directed against candidate enhancers at, or near, orphan T-cell DNase I hypersensitive sites located between 3'-BCL11B and VRK1. NKX2-5 down-regulation in t(5;14) PEER cells was almost entirely restricted to DNA inhibitory treatment targeting enhancers within the distal breakpoint cluster region and was dose and sequence dependent, whereas enhancers near 3'-BCL11B regulated that gene only. Chromatin immunoprecipitation assays showed that the four most effectual NKX2-5 ectopic enhancers were hyperacetylated. These enhancers clustered ~1 Mbp downstream of BCL11B, within a region displaying multiple regulatory stigmata, including a TCRA enhancer motif, deep sequence conservation, and tight nuclear matrix attachment relaxed by trichostatin A treatment. Intriguingly, although TLX3/NKX2-5 promoter/exon 1 regions were hypoacetylated, their expression was trichostatin A sensitive, implying extrinsic regulation by factor(s) under acetylation control. Knockdown of PU.1, known to be trichostatin A responsive and which potentially binds TLX3/NKX2-5 promoters, effected down-regulation of both homeobox genes. Moreover, genomic analysis showed preferential enrichment near ectopic enhancers of binding sites for the PU.1 cofactor HMGA1, the knockdown of which also inhibited NKX2-5. We suggest that HMGA1 and PU.1 coregulate ectopic homeobox gene expression in t(5;14) T-cell acute lymphoblastic leukemia by interactions mediated at the nuclear matrix. Our data document homeobox gene dysregulation by a novel regulatory region at 3'-BCL11B responsive to histone deacetylase inhibition and highlight a novel class of potential therapeutic target amid noncoding DNA. [Cancer Res 2007;67(4):1461–71]


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
BCL11B (also known as CTIP2/Rit1) is a Krüppel family zinc finger gene located at 14q32, which is expressed during thymocyte maturation from CD4–/CD8– to CD4+/CD8+ stages (1). BCL11B may act either as a tumor-suppressor gene, first detected via inactivating mutations or deletions in murine radiation-induced lymphomas (2) and more recently in T-cell acute lymphoblastic leukemia (T-cell ALL; ref. 3); or, as an oncogenic activator, first detected by its juxtaposition with the TLX3 (also known as HOX11L2) homeobox gene at 5q35 via the recurrent t(5;14)(q35;q32) rearrangement also in T-cell ALL (4–6). A neighboring related homeobox gene, NKX2-5 (also known as CSX), is similarly activated in T-cell ALL by a variant t(5;14)(q35;q32) or by t(5;14)(q35;q11.2) involving the T-cell receptor D locus at 14q11.2 (7, 8). Recently, interleukin-2 has been identified as a transcriptional target of BCL11B in both developing and transformed T cells (9). The ectopic expression of Nirenberg-Kim family homeobox genes (e.g., TLX3) has been recently shown to drive proliferation of myeloid progenitor cells in a mouse model (10). Nevertheless, the mechanism underlying leukemic deregulation by t(5;14) remains obscure, mainly because 14q32.2 breakpoints are widely scattered over the ~1 Mbp downstream breakpoint cluster region. Analogies with T-cell receptor loci and the nature of the "genomic desert" covering 3'-BCL11B support the existence of underlying interactions between TLX3/NKX2-5 and remote transcriptional enhancers.

Chromosomal mechanisms governing remote transcriptional regulation have lately attracted increasing interest. In particular, conserved noncoding elements/regions/sequences identified near metazoan developmental genes show coordinate enhancer activity when tested in zebrafish embryos (11). Physical juxtaposition between regulators and their targets has now been shown (12). These interactions are believed to occur at chromatin loops tethered to nuclear scaffold matrix attachment regions (13–15). DNaseI hypersensitive sites denote "active" chromatin and are believed to heighten the accessibility of nucleosome-depleted regulatory DNA regions and facilitate loading of transcription factor binding sites. Chromatin relaxation is also associated with histone hyperacetylation (16) and is antagonized by histone deacetylases. A global survey of regulatory regions in CD4+ T cells uncovered >5,000 DNaseI hypersensitive sites mostly bracketing active genes (17). A notable exception included an orphan DNaseI hypersensitive site cluster lying inside the BCL11B gene desert amid a region subsequently shown to host an acetylation island in T cells (18).

To investigate the role of 3'-BCL11B in leukemic cis-activation, we adapted DNA inhibitory treatments by using matching oligonucleotides directed against specific noncoding sequences based on previous work on the IgH enhancer (19, 20). Regulatory potential was further analyzed via nuclear matrix attachment in nuclear halo preparations, and by characterizing protein acetylation associated with both genes and noncoding regions. Binding site matrices for transcription factors PU.1 and HMGA1, identified in the promoter regions of TLX3 and near regulatory DNaseI hypersensitive sites, respectively, by using the TRANSFAC database (21) were targeted for inhibition by small interfering RNA (siRNA) treatment, and their effects on leukemic homeobox gene expression in t(5;14) cells were measured.

Our data support the existence of a remote regulatory DNaseI hypersensitive site cluster associated with leukemic homeobox gene activation in t(5;14) T-cell ALL. TRANSFAC data show this putative regulatory enhancer region to be enriched in HMGA1 binding sites juxtaposed by partner PU.1 sites at promoter regions of both TLX3 and NKX2-5. We propose that leukemic activation overlays HMGA1-PU.1 interactions mediated cytogenetically by transcription factor binding site juxtaposition, highlighting the role of noncoding DNA in neoplasia and as a potential therapeutic target therein.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cells and culture. Detailed descriptions of cell lines and culture methods are given by Drexler (22). Molecular karyotypes of CCRF-CEM, HPB-ALL, and PEER, all established from pediatric T-cell ALL, have been published elsewhere (6, 7). Antisense treatment was done as described previously (23). Briefly, 5 µmol/L 21-mer oligonucleotide phosphorothioate, modified at positions 1-2/20-21 (MWG, Ebersberg, Germany) and targeting the start codon, was added to the culture medium at days 0 and 1. Cells were harvested on day 3 for gene expression analysis.

Cytogenetic analysis. Harvesting, slide preparation, trypsin G-banding, and fluorescence in situ hybridization (FISH) are detailed elsewhere (24). Bacterial artificial chromosome (BAC) clones were obtained from BAC/PAC Resources (Oakland, CA), RZPD (Berlin, Germany), The Sanger Institute (Cambridge, United Kingdom), and Drs. Olivier Bernard (Hopital Necker, Paris, France) and Reiner Siebert (Institut für Genetik, Kiel, Germany). Fosmid clones were obtained from The Sanger Institute. Clone DNA was prepared using Princeton Big-PAC kits (EMP, Berlin, Germany) and labeled with Spectrum Red-dUTP or Spectrum Green-dUTP (Vysis, Bergisch Gladbach, Germany), or Cy3-dUTP [GE Healthcare (formerly Amersham), Munich, Germany], by nick translation (Invitrogen, Karlsruhe, Germany). FISH images were captured and analyzed using Smart Capture 2 software using a cooled Cohu charge coupled device camera (Applied Imaging, Newcastle, United Kingdom) configured to an Axioskop 2 microscope (Zeiss, Göttingen, Germany).

Preparation of DNA fibers. FISH was done on DNA fibers following the method described by Mejia et al. (25). Briefly, 10 µL cell suspension adjusted to 1 x 106 in PBS was placed immediately under the frosted part of a cleaned microscope slide preheated to 50°C and dried. The slide was then placed near vertically, and DNA fibers were extruded by rinsing with 150 µL lysis buffer applied dropwise (0.05 mol/L NaOH in 28.6% ethanol); fixed by a second rinse using 200 µL methanol; and thereafter dehydrated in ethanol (70%, 90%, and 100%, each for 2 min), air-dried, and processed for FISH.

Nuclear matrix attachment. Nuclear halos were prepared as previously described (26, 27). After FISH analysis, the nuclear locations of ≥30 nuclear signals from each of 20 BACs were recorded and subsequently classed as follows: predominantly matrix (score 2), perimatrix or both matrix and halo (score 1), and predominantly halo (score 0). Nuclear matrix attachment scores for each clone were taken as the means of these values. For histone deacetylase inhibition, cells were treated with 10 µmol/L trichostatin A (Upstate, Lake Placid, NY) dissolved in ethanol.

Chromatin immunoprecipitation assay. Chromatin immunoprecipitation (ChIP) was done as recommended using the ChIP Assay kit (Upstate), whereby 106 cells were sonicated for 30 seconds, producing fragments ranging 100 to 1,000 bp, and precipitated with 10 µg of the anti–acetyl-histone H3 antibody. For PCR analysis, 400 ng of DNA (ChIP or complete genomic DNA) were used as templates (Supplementary Table S1).

PCR. Total RNA was isolated 6 or 24 h after trichostatin A treatment, either 24 h after electroporation with the individual oligonucleotides, or 4 to 5 days after lentiviral transduction. RNA extraction, cDNA synthesis, and PCR were done as described previously (7). Real-time quantitative-PCR (RQ-PCR) was done as described previously (28). Oligonucleotide sequences are listed in Supplementary Table S1.

Oligonucleotide inhibitory treatments. Transient inhibitory treatments were modified from that described by Cutrona et al. (20), substituting double-stranded 26-mer double-stranded oligonucleotide (DSO) phosphorothioate modified at positions 1-2/25-26 (purchased from MWG) for peptide nucleic acid oligonucleotides. In 10 of 16 cases, DSO sequences (Table 1 ) were uniquely located at, or near (≤1 kbp), orphan T-cell DNaseI hypersensitive sites (17), whereas the remainder lay either offset by ~1 to 10 kbp (DSO.12-14), or remote by >10 kbp (DSO.3/5), from DNaseI hypersensitive sites. Target sequences were chosen using TRANSFAC databanks listing paired transcription factor binding sites displaced by <10 bp (the so-called composite elements; ref. 21). The selection process is shown in Supplementary Fig. S1A for DSO.6. DSOs were produced by pooling equimolar amounts of complementary single-stranded oligonucleotides dissolved in water, heating to 95°C for 5 min and incubating at room temperature for 30 min. For electroporation, 2 µg pEGFP-N1 reporter (Clontech, Heidelberg, Germany) or DSO (20 µmol/L) were used to treat 106 cells in culture medium using a EPI-2500 impulse generator (Fischer, Heidelberg, Germany) for 10 ms at 350 V. Cells were subsequently transferred into 1 mL fresh culture medium and incubated overnight.


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  		Table 1. DSO sequences, efficacy, and distances from DNaseI hypersensitive sites and transcription factor binding matrices

 
Chemically synthesized siRNA directed against HMGA1. Expression of HMGA1 was knocked down using validated siRNA obtained from Qiagen (Hilden, Germany). As control, a synthetic siRNA directed against an irrelevant sequence was used (BioSpring, Frankfurt, Germany). For treatments, 1 µL of a 20 µmol/L stock solution of siRNA was electroporated into PEER cells as described above.

Lentiviral-mediated RNA interference directed against PU.1. DNA oligonucleotides corresponding to the siRNA nucleotide targeting sequence 720 to 738 bp of the human PU.1 gene (Genbank accession no. NM_003120; ref. 29) were chemically synthesized with a 5' BglII overhang and a 3' SalI restriction site for cloning (Biospring, Frankfurt, Germany). The numbering of the first nucleotide of the short hairpin RNA (shRNA) refers to the ATG start codon. The shRNA oligonucleotide sequences were as follows:

FP-PU.1-720: 5'-GATCCCCGAAGCTCACCTACCAGTTCTTCAAGAGAGAACTGGTAGGTGAGCTTCTTTTTTGGAAG-3'
RP-PU.1-720: 5'-TCGACTTCCAAAAAAGAAGCTCACCTACCAGTTCTCTCTTGAAGAACTGGTAGGTGAGCTTCGGG-3'.

The noncomplementary 9-nt loop sequences are underlined, and the sense oligonucleotide harbors six T's as a polymerase III transcription termination signal. The oligonucleotides were annealed and inserted 3' of the H1-RNA promoter into the BglII/SalI–digested pBlueScript-derived pH1-plasmid to generate pH1-PU.1-720 as described by Scherr et al. (30). The sequence was confirmed by sequencing analysis. Two plasmids, pH1-shRNA controls 1 and 2, directed against irrelevant sequences, were used as controls (30).

pdc-SR was used to generate lentiviral transgenic plasmids containing H1-siRNA expression cassettes located in the U3 region of the 3' long terminal repeat. To generate the lentiviral plasmid pdcH1-siRNA-SR, the pH1-PU.1-720 plasmid was digested with SmaI and HincII, and the resulting DNA fragments (360 nt) were blunt-end ligated into the SnaBI site of the pdc-SR to generate pdcH1-PU.1-SR plasmid.

VSV.G pseudotyped lentiviral particles were generated by calcium phosphate cotransfection of 293T cells, and viral supernatants were concentrated and titered as previously described, whereas recombinant lentiviral supernatants were used to transduce 106 PEER cells with a multiplicity of infection of ~2 as described previously (31).

Mapping DNaseI hypersensitive sites. DNaseI hypersensitive sites were mapped by methods previously described (17, 32). Briefly, intact nuclei from human CD4+ T cells were treated with small amounts of DNaseI (Roche, Indianapolis, IN), and blunt-ended fragments were ligated to a biotinylated linker. Approximately 230,000 sequences were generated from the linked ends using massively parallel signature sequencing.5 Because this level of sequencing depth only identified 20% of all DNaseI hypersensitive sites, we have recently sequenced more than 3 million tags using the 454 Life Sciences GS-20 sequencer (Life Sciences).6 Multiple sequences from the DNase-treated library that map within a 250 base window have been shown to be highly accurate at identifying valid DNaseI hypersensitive sites.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Cytogenetic analysis for further delineation of BCL11B breakpoint cluster region. The consensus karyotype of the DND-41 pediatric T-cell ALL cell line used to demarcate the BCL11B breakpoint cluster region is 86-98<4n>XXYY,–9,–15,+20. FISH analysis using a 14q32 BAC contig revealed paired (i.e., pretetraploidization) cryptic ins(5;14)(q35.2;q32.2q32.2) rearrangements undetectable by G-banding (Fig. 1A, right inset ), involving a two-component microinsertion comprising ~500 kbp of the 3'-BCL11B region at 14q32.2 extending from within clones 61o1 to 1127d7, significantly lacking the interstitial region covered by clones 1140p17 and 1082a3 (Fig. 1A). The results of fiber FISH showed that the insertion point of 3'-BCL11B material at 5q35 in DND-41 cells directly juxtaposes TLX3 in cells (Fig. 1A). This insertion is presumably responsible for activating TLX3 in DND-41 cells (33) by introducing ectopic (3'-BCL11B) regulatory enhancers and thus resembles that reported in a T-cell ALL patient (34). After fine-mapping using fosmid clones (Fig. 1B), the 14q32 breakpoint in PEER, first mapped using BAC clones (7), was revised (telomerically) to 97.81 Mbp, ~900 kbp downstream of BCL11B, and that of CCRF-CEM to 98.43 Mbp in close agreement with the previous BAC data (7), whereas 3'-BCL11B itself is located at 98.71 Mbp (Fig. 1C). The configuration of ins(5;14) in DND-41 implies a target telomeric of BAC 74h1 within that part of the breakpoint cluster region covered by the insertion. Furthermore, remaining instances of t(5;14) cell lines CCRF-CEM, HPB-ALL, and PEER all involved microinsertions, ins(14;5), raising a question mark over clinical instances where such complex rearrangements may be underascertained (6, 7, 10). To reconcile insertion and breakpoint data, the critical target(s) of t(5;14) rearrangements should lie within the genomic desert between clone 74h1 and BCL11B itself probably within the consensus region as defined by the centromeric portion of the DND-41 insertion.


Figure 1
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  		Figure 1. Cytogenetic analysis and the BCL11B breakpoint cluster region. A and B, cytogenetic analysis of DND-41 and PEER cells. FISH preparations were counterstained with 4',6-diamidino-2-phenylindole (DAPI). A, cryptic formation of ins(5;14)(q35;q32) in DND-41. Right insets, normal G-banding, involving the discontinuous insertion of material from 3'-BCL11B at 5q35. Note the absence of flanking clones 15e14 (green) and 2348n10 (purple) from the insertion (detailed in C). Inset (A, left, bottom), fiber FISH confirming juxtaposition of TLX3 containing clone 117l6 (green) with clone 61o1 (red). B, fine mapping of 14q32 breakpoint in PEER cells to lie between fosmid clones G248P8-5772g10 (orange) and G248P8-9081b5 (red). Positions of these clones and the breakpoint in PEER (C). Note the insertion of material from fosmid G248P8-229B6 (green) at the 14q32 breakpoint. C, available cytogenetic data (breakpoints and insertions) on the far downstream region of BCL11B from t(5;14) T-cell ALL cell lines (large arrows) and patients (small arrows) from Su et al. (33). Coordinates are given in Mbp based on http://genome.ucsc.edu/index.html (National Center for Biotechnology Information Build 36.1). Positions of DSO.1-16 (downward pointing triangles) were shaded according to specificity in DNA inhibitory treatment–mediated down-regulation of NKX2-5 (black), BCL11B (gray), or neither (white). Bottom, DNaseI hypersensitive sites (DNAseI-HS; rockets) targeted by DSO.6-9 in the 97.72-.78 Mbp region, together with comparison data (HSS sites) described by Su et al. (33). Black rockets, activity, whether against NKX2-5 (DSO.6-9) or in heterologous promoter assays (HSS-3/4). Note the absence of promoter activity at HSS-1/2/5 (white rockets). NB, coincident active DNaseI hypersensitive site in both studies. Note also the TCRA enhancer–like motif (AP-1, LEF-1, ETS-1, AML1, and TATA box) at DSO.6/7. Acetylation islands (18) and duplicated noncoding sequences recently posited to host insulators (34) are also shown.

 
Analysis of nuclear matrix attachment. Nuclear matrix attachment, which reflects transcriptional recruitment of coding and regulatory regions (13–15), possibly related to nuclear territories, was assayed cytologically from 3'-BCL11B to VRK1 in PEER cells (Fig. 2A and B ). Illustrating the correlation between nuclear matrix attachment and gene activity in PEER cells, clones for NOTCH1 (obligate expression in T cells), PAX5 (silenced in T cells), and highly repetitive centromeric {alpha}-satellite DNA from chromosome 17 displayed progressively looser nuclear matrix attachment scored 2, 1, and 0, respectively (Fig. 2A, bottom left). In PEER cells, the highest nuclear matrix attachment was observed close to BCL11B expression of which is T-cell specific, followed by a central plateau (97.3–97.9 Mbp), straddling putative distal regulatory regions DSO.3 and DSO.6-9 (Fig. 2A and B). In SC-1 B-cell lymphoma cells in which BCL11B and BC043585 are silent, neither the BCL11B peak nor the adjacent plateau were discernible (data not shown)—findings consistent with tissue-specific variations in nuclear matrix attachment (Fig. 2A, top). Trichostatin A treatment for 24 h reduced nuclear matrix attachment uniformly across the distal breakpoint cluster region, including both plateau and neighboring telomeric valley regions (Fig. 2A, bottom right, B). Interestingly, this valley shared coordinates with noninserted/deleted material in ins(5;14) in DND-41 cells (Fig. 1A and C) and in a t(5;14) patient reported by Su et al. (33). In overextracted preparations (8 min), differences in nuclear matrix attachment between ins(14;5) responsible for leukemic NKX2-5 activation, which exhibited high nuclear matrix attachment, forming a tight structure reminiscent of "active chromatin hubs" (35), were evident, whereas nonleukemogenic der(5) and der(14) partner chromosomes looped outside the nuclear matrix (Fig. 2A, bottom middle).


Figure 2
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  		Figure 2. Nuclear matrix attachment in PEER cells. The three-color FISH images in (A) depict nuclear matrix preparations from PEER nuclei and hybridized with contrast-labeled BAC clones. Top panels, nuclear matrix attachment along the noncoding 3'-BCL11B region: Note the low nuclear matrix attachment of clones 14l13 and 354a24 (yellow) derived from the region immediately centromeric of the BCL11B breakpoint cluster region, contrasting with clones covering the neighboring leukemic regulatory region, 61o1, 3181d5, and 74h1 (red), which exhibited high nuclear matrix attachment. On the other hand, clones from the intervening region closer to BCL11B, 1057p17, and 3104h21 (green) showed intermediate nuclear matrix attachment. In overextracted preparations (8 min), it was possible to distinguish both the correlation of nuclear matrix attachment with gene activity, as shown by the progressive margination of NOTCH1 (red), PAX5 (green), respectively, silent and active in PEER cells, and centromeric-17 (cen 17, yellow, bottom left); and between ins(14;5) responsible for leukemic NKX2-5 activation that exhibited high nuclear matrix attachment, forming a tight structure reminiscent of active chromatin hubs, whereas der(5) and der(14) "silent" partner chromosomes looped outward from the matrix (bottom middle). Trichostatin A (TSA) treatment–affected histone deacetylase inhibition, which induces silencing of NKX2-5 transcription in PEER cells, reduced matrix attachment, as shown by clones 15e14 (red) or 3181d5 (green; bottom right). Preparations were counterstained with DAPI and reversed to yield gray background staining. B, nuclear matrix attachment data (points) over the entire 2.3 Mbp region between VRK1 and BCL11B; bars, SE. Note central plateau of high nuclear matrix attachment including putative enhancer regions (DSO.3, DSO.6-9) and EST gene (BC043585), flanking lowered nuclear matrix attachment zones, and elevated nuclear matrix attachment close to transcribed genes VRK1 and BCL11B. Note the uniform reduction in nuclear matrix attachment along the region tested after trichostatin A treatment to inhibit histone deacetylase.

 
Inhibitory treatments of candidate enhancer regions. To characterize the gene regulatory potential of noncoding regions identified near DNaseI hypersensitive sites, these were targeted using DNA inhibitory treatments based on reports describing the use of short inhibitory peptide nucleic acid probes matching IgH regulatory sequences, thus inhibiting oncogene expression driven by IgH enhancers (19, 20). The efficiency of DNA transfer by electroporation was checked by a reporter construct (pEGFP) in several B-cell lines with t(8;14)(q24;32)/t(14;18)(q32;q21) and in T-cell lines with t(5;14)(q35;q32), among which 380 and PEER cells, respectively, displayed the best survival (both >80%) and transfection efficiencies (~60% and ~80%, respectively). The 380 B-cell lymphoma cell line carries a t(14;18)(q32;q21), which juxtaposes BCL2 with the IgH-Eµ enhancer (22). Preliminary investigations showed that inhibition of BCL2 and NKX2-5 transcription by complementary oligonucleotides was (a) restricted to DSOs by both reverse transcription-PCR (RT-PCR; Supplementary Fig. S1B) and RQ-PCR (Supplementary Table S2); (b) abolished by point mutations, confirming sequence specificity (Supplementary Table S2); and (c) concentration dependent (Supplementary Fig. S1C). DNA inhibitory treatments directed against regulatory regions was validated against the IgH-Eµ enhancer sequence driving BCL2 transcription in 380 B-cell lymphoma cells (Supplementary Fig. S1D), analogous to that used by Cutrona et al. (20).

Enhancer inhibition was assayed by measuring BCL11B and NKX2-5 expression in PEER cells in which these loci are juxtaposed via a ~100 kbp genomic insertion of NKX2-5 ~1.1 Mbp downstream of BCL11B (Fig. 1B and C; ref. 7). Assayed by RT-PCR, DSO.3/6-10/12-14 inhibited expression of NKX2-5 (Fig. 3A ), whereas DSO.15/16 inhibited BCL11B only (Fig. 3B). Control RT-PCR studies showed that inhibition depended on both double-strandedness and sequence fidelity (Supplementary Fig. S1B). Quantification of NKX2-5 down-regulation by RQ-PCR yielded broadly similar results as well as confirming the dependence of inhibition on double strandedness and sequence fidelity (Fig. 3C; Supplementary Table S2). Gene expression analysis of NKX2-5 and BCL11B distinguished three types of DSO target site in PEER cells: Those that affected down-regulation of NKX2-5 (by DSO.3, DSO.6-10, DSO.12-14), of BCL11B (by DSO.15/16), or of neither (by DSO.1/2, DSO.4/5, DSO.11; Fig. 3C). Interestingly, principal components of the TCRA enhancer comprising binding sites for activator protein-1, high-mobility group domain containing protein LEF-1, ETS-1, AML1, and a TATA box were identified within the BAC sequence 61o1 at 6,895 bp adjoining the breakpoint in PEER cells. Expression of BC043585 and VRK1 were unaffected by any DSO tested, including DSO.4 located within intron 4 of BC043585 (Fig. 3D), and DSO.1/2 at 3'-VRK1. Of 10 DNaseI hypersensitive sites closely targeted (sites <1 kbp), seven were inhibitory (five of NKX2-5 and two of BCL11B), whereas two of the three nonefficacious sites (DSO.1/2) lay most distant outside the breakpoint cluster region and close to VRK1. Neither non–DNaseI hypersensitive sites DSO.4/5 proved inhibitory of NKX2-5 by RT-PCR or RQ-PCR. Although the immediate proximity of DSO to DNaseI hypersensitive sites did not dictate efficacy, the association of the latter with targeting specificity was nonrandom: eight of nine NKX2-5 inhibitory sites lay inside the breakpoint cluster region, of which the four most efficacious (DSO.6-9) clustered immediately centromeric of the 14q32 breakpoint in PEER cells. On the other hand, both BCL11B-inhibitory sites (DSO.15/16), although inside the breakpoint cluster region, lay most distant from the breakpoint and nearest to BCL11B itself (Fig. 1C). One outlying inhibitory DSO site (DSO.3) located >500 kbp centromeric of the PEER breakpoint lay within the high nuclear matrix attachment region (Fig. 2B). In summary, the most remarkable feature was the cluster of efficacious sites (DSO.6-9) located near the insertion point of NKX2-5 in PEER cells.


Figure 3
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  		Figure 3. DNA inhibitory treatment by complementary DSOs. A and B, inhibition of NKX2-5 or BCL11B expression in PEER cells electroporated with DSO complementary to noncoding sequences at 3'-BCL11B. Note the complete inhibition of NKX2-5 alone by DSO.3, DSO.6-10, and DSO.12/13 (A), and of BCL11B alone by DSO.15/16 and partially by DSO.14 (B). C, RQ-PCR data (detailed in Supplementary Table S1). Note peak inhibition by DSO.6-9 clustered near 14q32 breakpoint in PEER cells. Note also abrogation of DSO inhibition by use of either single-stranded (SSO.6a or SSO.6b) or mutated oligonucleotides. D, ineffectiveness of DNA inhibitory treatment using various DSO on transcription of BC043585 (top) or VRK1 (bottom), highlighting the specificity of the NKX2-5 inhibition.

 
Multiple DNaseI hypersensitive sites. Multiple DNaseI hypersensitive sites (clustered within 500 bp to minimize noise) are taken to serve as proxy enhancers (32). Multiple DNaseI hypersensitive sites coordinates derived by deep sequencing inside the BCL11B genomic desert are given in Supplementary Table S3 and the intersite distances shown in Supplementary Fig. S2. DNaseI hypersensitive site densities fluctuate along 3'-BCL11B, usually peaking near gene loci, notably BCL11B itself, but only weakly near VRK1, if at all (Supplementary Fig. S2). Density peaks often coincided with the largest DNaseI hypersensitive sites, although these sometimes occurred within troughs. A similar pattern occurred near acetylation islands. The second highest DNaseI hypersensitive site peak coincided with the DSO.6-9 region, which is devoid of known genes. Interestingly, this peak included the largest concentration of multiple DNaseI hypersensitive sites (Supplementary Fig. S2).

Histone acetylation analyses. In higher organisms, gene activity is profoundly associated with histone acetylation in promoter/enhancer regions (36). We did ChIP analysis to assay histone-H3 acetylation of promoter/exon 1 regions of genes BC043585, BCL11B, NKX2-5, and VRK1, together with DSO target regions in both PEER (NKX2-5 expressing) and DND-41 cells (TLX3 expressing). We found both BCL11B and BC043585 to be highly acetylated in PEER and moderately in DND-41 cells, whereas VRK1 was moderately acetylated in both cell lines. In contrast, NKX2-5 and TLX3 displayed negligible promoter/exon 1 acetylation in PEER and DND-41 cells (Fig. 4A ). Among DNA inhibitory treatment target regions, the DSO.6-9 cluster juxtaposing the NKX2-5 insertion and TLX3 recipient breakpoints were strongly acetylated in both PEER and DND-41, whereas more distant DSO targets were weakly acetylated despite their efficacy (Fig. 4B). Thus, the acetylation islands of T-cell ALL cells resemble those of nonmalignant T-cells (described in ref. 18) shown in Fig. 1C.


Figure 4
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  		Figure 4. Acetylation and leukemic expression of TLX3 and NKX2-5. Equal amounts of ChIP or genomic control DNA, respectively, was used for PCR analysis of promoter/exon 1 regions of genes VRK1, BC043585, NKX2-5, TLX3, and BCL11B (A), and noncoding regions hosting DSOs (B). Comparison of PCR product intensities reveals acetylation of VRK1, BC043585, and BCL11B, contrasting with NKX2-5 or TLX3, which remained unacetylated. Note acetylation of DSO.6-9, present within a known T-cell acetylation island (18) in both DND-41 and PEER, whereas other DSOs remained inconsistently acetylated, weakly acetylated, or nonacetylated. Effects of trichostatin A treatment on the expression of TLX3 in DND-41 assayed by RT-PCR (C) and of NKX2-5 expression in PEER assayed by RQ-PCR (D). TLX3, BCL11B, and GATA3 were down-regulated by trichostatin A treatment of DND-41 cells in contrast to unchanged levels of UBF, VRK1, MYB, and NOTCH1. Similarly, NKX2-5 exhibited trichostatin A–sensitive expression in PEER cells.

 
Histone deacetylase inhibitors reactivate genes silenced by histone deacetylation. Expression of MYB, NOTCH1, UBF, and VRK1 (ubiquitously expressed in T cells) were unmodulated by trichostatin A treatment, whereas TLX3, BCL11B, and GATA3 (subject to developmental regulation in T cells) were down-regulated (Fig. 4C). Similarly, in PEER cells, trichostatin A treatment affected down-regulation of NKX2-5 by 62% (Fig. 4D; Supplementary Table S2). The trichostatin A sensitivities of both TLX3 and NKX2-5 considered together with the absence of promoter/exon-1 acetylation, revealed by ChIP assay (Fig. 4A), implies that leukemic activation of both homeobox genes by t(5;14) is subject to indirect regulation by histone deacetylase. Obvious intermediates are transcription factors such as PU.1, down-regulated by trichostatin A in multiple hematopoietic lineages (37), whereas both GATA3 and PU.1 are repressed by direct and promoter acetylation, respectively (38).

PU.1 and HMGA1 knockdown. Inspection of upstream promoter regions of both TLX3 and NKX2-5 revealed GAGGAA sequences corresponding to consensus PU.1 binding sites 100 and 366 bp upstream of their respective transcription start points. TRANSFAC analysis done at DSO sites (±30 bp) to uncover transcription factor binding sites matrices showed the greatest degree of enrichment for HMGA1 (P < 0.005) at efficacious sites, whereas PU.1 sites were distributed randomly (Table 1). To investigate TLX3/NKX2-5 regulation by transcription factors subject to acetylation control, we treated DND-41 cells with antisense oligonucleotides directed against GATA3 and PU.1 and subsequently monitored expression of TLX3. These experiments showed that antisense PU.1, but not antisense GATA3 treatment, inhibited transcription of TLX3 in DND-41 cells (Fig. 5A ). Prompted by these results, lentivirally mediated RNA interferences directed against PU.1 or chemically synthesized siRNA against HMGA1 were done in PEER cells using RQ-PCR of NKX2-5 as end point. These experiments showed that knockdown of PU.1 (by 50%) or HMGA1 (by 70%) effected down-regulation of NKX2-5 by 80% and 70%, respectively (Fig. 5B; Supplementary Table S2). Taken together, we concluded that PU.1 and HMGA1 are both necessary to sustain TLX3/NKX2-5 transcription in t(5;14) cells.


Figure 5
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  		Figure 5. Effects of PU.1 and HMGA1 knockdown on TLX3 and NKX2-5. A, effects of antisense treatment of DND-41 cells using oligonucleotides directed against GATA3 or PU.1 compared with control oligonucleotide (AS-C). RT-PCR analysis shows down-regulation of TLX3 after as-PU.1 treatment of DND-41 cells, indicating the dependence of TLX3 on PU.1. B, quantitative RT-PCR of PU.1, HMGA1, and NKX2-5 after RNA interference–mediated inhibition. Knockdown of PU.1 was done by lentiviral-mediated transfer of shRNA vectors directed against PU.1. PU.1 expression was reduced by 40% and correlated with 30% reduction of NKX2-5. Inhibition of HMGA1 was done by electroporation of synthetic siRNA. Compared with controls, expression of both HMGA1 and NKX2-5 was reduced by 70%, respectively. Data indicate regulation of NKX2-5 expression by PU.1 and HMGA1.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
We investigated leukemic activation of Nirenberg-Kim family homeobox genes NKX2-5 and TLX3 involved in alternative t(5;14) translocations by carrying out DNA inhibitory treatments directed against candidate enhancers within the 3'-BCL11B breakpoint cluster region, initially focusing on T-cell single DNaseI hypersensitive sites identified therein by global screening (17). It remains to be determined whether DNA inhibitory treatment acts directly on DNA substrates by, for example, blocking transcription factor access, or by sequestering activatory transcription factors themselves. Inhibition was largely restricted to targets located within the breakpoint cluster region that hosts 10 of 11 regulatory sites, whereas only one of five outliers proved regulatory. Regulatory sequences in PEER cells sometimes lay close to DNaseI hypersensitive sites, although absolute coincidence was not a requirement. Sites were dedicated to either NKX2-5 or BCL11B, their orientation apparently dictated by proximity. Maximal inhibition of NKX2-5 in PEER cells was affected by treatments with DSO.6-9, which cluster within a ~10 kbp region located ~1 Mbp downstream of BCL11B and which, in PEER cells, immediately juxtapose NKX2-5. In addition to DNaseI hypersensitive sites, this region displays a diverse array of known and likely cis-regulatory stigmata, including elevated nuclear matrix attachment, hyperacetylation colocalization with an orphan T-cell acetylation island (18), abyssal cross-species homology related to "five-way regulatory potential" (39), a proximate TCRA enhancer–like motif (AP-1, LEF-1, ETS-1, AML1, and TATA box), and placement at the centromeric breakpoint cluster region boundary near the NKX2-5 insertion in PEER cells. A recent study has also mapped DNaseI hypersensitive sites in primary T-cell ALL (33), a pair of which (HSS-3/4), respectively, match DSO.6/7 and DSO.8/9 described herein (Fig. 1C). Remarkably, of the five DNaseI hypersensitive sites tested by the authors, heterologous promoter activity was solely manifested by HSS-3/4, consistent with our finding that DSO.6-9 exerted the greatest effects on NKX2-5 expression. Taken together, these data provide compelling evidence for a distal regulatory enhancer complex ~1 Mbp downstream of BCL11B—rivaling sonic hedgehog in remoteness from its putative coordinate target gene (40).

Remoteness, together with the failure of DSO.6-9 to inhibit BCL11B expression, raise the question of whether BCL11B is indeed the normal subordinate of this enhancer. The clinical homogeneity of t(5;14) cases irrespective of breakpoint, together with the failure of flanking DSO to inhibit BC043585, and indeed the absence of any convincing alternative, leaves BCL11B by far the most likely target. Furthermore, in PEER cells that exceptionally (among T cells) express very low levels of BCL11B (7), the distal regulatory region containing DSO.6-9 is cut off from BCL11B by the intervening ~100 kbp insertion containing NKX2-5, consistent with regulatory deprivation. Unlike the deeply conserved region surrounding DSO.6-9, locally expressed transcripts, including BC043585, are primate specific, belying any regulatory connection with deeply conserved sequences also represented in fugu and zebrafish. However, because 3'-BCL11B breakpoints are variable, the role of additional enhancers in the region remains to be addressed in detail. Further experiments are planned to assess the regulatory repertoire of the DSO.6-9 and other candidate enhancers within 3'-BCL11B and the potential of the former as a locus control region, inter alia, contributing to the maintenance of local hyperacetylation.

To direct spatiotemporal transcriptional regulation, developmental genes may require boundary elements (insulators). In this regard, DSOs effectual against NKX2-5 in PEER cells are separated from DSO.15/16 inhibitory of BCL11B only by flanking multiple noncoding elements/regions/sequences duplicated 3' of the homologous BCL11A. This type of homology hallmarks transcriptional insulators (35). The need to exclude transcriptional insulators would help explain the unprecedentedly widespread yet enigmatic formation of BCL11B rearrangements via microinsertions (6, 7, 34), or complex deletion-associated inversions (3), to the possible exclusion of conventional translocations, at least in cell lines where the most detailed studies have been documented hitherto.

In addition to confirming the existence in malignant T cells of an acetylation island recently described in their normal human counterparts (18), but absent or displaced in mice (41), ChIP assays revealed that indigenous loci at 14q32.2 (VRK1, BC043585, and BCL11B) were promoter/exon 1 acetylated. This finding contrasts with immigrant TLX3 and NKX2-5, which were, nevertheless, both sensitive to histone deacetylase inhibition by trichostatin A and knockdown of PU.1 or HMGA1. To resolve these apparently contradictory findings, given the provision of PU.1 binding site matrices at both TLX3 and NKX2-5 promoters, we propose that the sensitivities of the latter t(5;14) targets to histone deacetylase inhibition may be mediated indirectly by PU.1, the expression of which is trichostatin A sensitive (37). All four acetylated DSO sites (DSO.6-9) target DNaseI hypersensitive sites clustered inside an orphan acetylation island recently described in primary resting and activated T cells (18), and imply the retention and possible exploitation of an epigenetic alteration by malignant T cells. Although PU.1 GAGGAA consensus binding site sequences occur at 5'-TLX3 (at –100 bp), NKX2-5 (at –366 bp), and BCL11B (at –61 bp), the TRANSFAC database shows no preferential enrichment of PU.1 matrices near effectual DSO, in contrast to its regulatory consort HMGA1 for which matrices occur at all DSO inhibitory of NKX2-5 with the sole omission of DSO.15. These findings serve to encourage wider investigation of the implied relationship between the respective expression levels of PU.1/HMGA1 and Nirenberg-Kim family homeobox genes in cases of T-cell ALL with t(5;14). PU.1 regulates IgH developmental expression via enhancer binding (42) and interacts with HMGA1 by increasing its transcriptional binding activity (43). HMGA1 also modifies DNA structure at the IFNß enhancer to consolidate the enhanceosome (44, 45), whereas HMGA1 acetylation by CBP effects its disruption (46). HMGA1 functions as a transcriptional coactivator of the IgH-µ enhancer in B cells through indirect association with DNA, being but loosely bound to enhancer DNA (47).

Our data showing that PU.1 and HMGA1 are both required to sustain leukemic homeobox gene transcription in T cells, and that der(14) breakpoints near effectual DSO sites are flanked by PU.1 and HMGA1matrices, suggest a possible nucleochromosomal mechanism underpinning the transient regulatory association of these transcription factors as invoked in B cells. A recent study has shown enrichment of binding sites for ETS family transcription factors (including PU.1) near DNaseI hypersensitive sites implicated in long-distance regulation of tissue-specific genes in endothelial cells (48). The selectively tight nuclear matrix attachment displayed at 14q32, reminiscent of active chromosomal hubs, and its alleviation by trichostatin A treatments inhibitory of TLX3/NKX2-5, support a model whereby PU.1/HMGAl interactions are mediated nucleochromosomally, possibly via tethering to SATB1, which is known to act as a substrate of HMGA1 (49).

In T cells, SATB1 occupies chromosomal scaffolds to facilitate tissue-specific nuclear matrix attachment and histone modification, thereby linking chromatin modification and looping to transcription (50). Our findings that the centromeric boundary of the 3'-BCL11B breakpoint cluster region coincides with a steep decline in nuclear matrix attachment, while the DSO.6-9 region occupies an elevated nuclear matrix attachment plateau, also support a role for nuclear geography in leukemogenic activation. The reasons underlying deep cross-species conservation, as present around DSO.6-9, remain obscure but may reflect combined structural and functional constraints, such as the need to effect transient interactions between appropriate transcription factors. In the present case, this might involve interactions between HMGA1 bound near enhancers, and PU.1 bound to promoters of homeobox genes subject to a variety of acetylation controls: both direct, by opening DNaseI hypersensitive sites, and indirect, mediated via HMGA1/PU.1. According to such a model, a putative t(5;14) enhanceosome might interact with distal DNaseI hypersensitive sites convened at the nuclear matrix embodying "active chromatin hubs."

To summarize, we developed a method (DNA inhibitory treatment) for the targeting of specific noncoding DNA sequences that permitted identification of remote leukemic enhancers at 3'-BCL11B in t(5;14) cells. Interestingly, the most effectual enhancers lay within a DNaseI hypersensitive site cluster located ~0.9 to 1.0 Mbp downstream of BCL11B and displayed multiple regulatory stigmata. Ectopic NKX2-5 and TLX3 expression were inhibited by knockdown of either HMGA1 or PU.1, highlighting their complicity in leukemic homeobox deregulation in t(5;14). As well as providing novel paradigms for oncogene dysregulation, t(5;14) T-cell ALL is a major clinical entity in which the identification of potential therapeutic targets amid noncoding DNA sensitive to DNA inhibitory treatments and histone deacetylase inhibition merits wider investigation.


   Acknowledgments
 
Grant support: José Carreras Leukemia Research Fund grants (R.A.F. MacLeod and S. Nagel).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Sandra Goetze and Silke Winkelman for help in establishing nuclear matrix assays, Drs. Olivier Bernard and Reiner Siebert for supplying clones, Greg Elgar for access to unpublished data, SriLaxmi Kalavalapalli and Sebastian Becker for help with data analysis, and Professor Jürgen Bode for critically reading the manuscript.


   Footnotes
 
Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

5 http://research.nhgri.nih.gov/DNaseHS/May2005/ Back

6 G.E. Crawford, unpublished observations. Back

Received 7/14/06. Revised 11/ 2/06. Accepted 12/ 8/06.


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 Introduction
 Materials and Methods
 Results
 Discussion
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