Methods for modulating ATRX-dependent gene repression (2024)

This application is a § 371 National Stage Application of PCT/US2015/058338, filed Oct. 30, 2015, which claims the benefit of U.S. Provisional Patent Application Ser. No. 62/072,962, filed on Oct. 30, 2014. The entire contents of the foregoing are hereby incorporated by reference.

This invention was made with Government support under Grant No. RO1-GM090278 awarded by the National Institutes of Health. The Government has certain rights in the invention.

This invention relates to methods for modulation of the activity of alpha thalassemia/mental retardation syndrome X-linked (ATRX), e.g., modulation of DNA-ATRX or RNA-ATRX interactions, and methods for identifying and using compounds that modulate DNA-ATRX or RNA-ATRX interactions, as well as the compounds themselves.

Transcriptome analyses have suggested that, although only 1-2% of the mammalian genome is protein-coding, 70-90% is transcriptionally active (Carninci et al., Science 309, 1559-1563, 2005; Kapranov et al., Science 316, 1484-148, 2007; Mercer et al., Nat Rev Genet 10, 155-159, 2009). Ranging from 100 nt to >100 kb, these transcripts are largely unknown in function, may originate within or between genes, and may be conserved and developmentally regulated (Kapranov et al., 2007, supra; Guttman et al., 2009). Methods for targeting these transcripts allow for modulation of gene expression.

The present invention is based, at least in part, on the discovery that ATRX protein is a required specificity determinant for locus-specific PRC2 targeting and effects on gene expression. Thus, methods and compounds targeting the ATRX-RNA interaction can be used to modulate gene expression.

In one aspect, the invention provides methods for increasing expression of a selected gene listed in Tables 1 or 2 or 3 or 4 or 5, in a cell, the method comprising contacting the cell with a nucleic acid triplex-forming oligonucleotide (TFO) that binds specifically to an ATRX localization sequence or binding site associated with the selected gene. In some embodiments, the selected gene is DMD, and the TFO targets a sequence within SEQ ID NOs.7867-7900. In some embodiments, the selected gene is XIST, and the TFO targets a sequence within SEQ ID NOs.7957, 7959, 7961, or 7962.

In another aspect, the invention provides methods for treating a subject who has duch*enne muscular dystrophy. The methods include administering to the subject a therapeutically effective amount of a TFO that targets a sequence within SEQ ID NOs. 7867-7900.

In a further aspect, the invention provides methods for treating a subject who has Rett Syndrome. The methods include administering to the subject a therapeutically effective amount of a TFO that targets a sequence within SEQ ID NOs. 7957, 7959, 7961, or 7962.

In some embodiments of the methods described herein, the TFO comprises one or more of DNA, RNA, PNA, HNA, MNA, ANA, LNA, CAN, INA, CeNA, TNA, (2′-NH)-TNA, (3′-NH)-TNA, alpha-L-Ribo-LNA, alpha-L-Xylo-LNA, beta-D-Ribo-LNA, beta-D-Xylo-LNA, [3.2.1]-LNA, Bicyclo-DNA, 6-Amino-Bicyclo-DNA, 5-epi-Bicyclo-DNA, alpha-Bicyclo-DNA, Tricyclo-DNA, Bicyclo[4.3.0]-DNA, Bicyclo[3.2.1]-DNA, Bicyclo[4.3.0]amide-DNA, beta-D-Ribopyranosyl-NA, alpha-L-Lyxopyranosyl-NA, 2′-R-RNA, 2′-OR-RNA, 2′-AE-RNA, alpha-L-RNA, and beta-D-RNA.

In some embodiments of the methods described herein, the TFO includes one or more modifications described herein.

In another aspect, the invention provides methods for preparing a library of nuclear ribonucleic acids (nRNAs) that specifically bind ATRX. Preferably, the methods include (a) contacting a sample containing nRNAs, e.g. at least 10⁴, 10⁵, or 10⁶ different nRNAs, with (i) ATRX protein and (ii) a ATRX binding agent, under conditions sufficient to form complexes between the nRNA, ATRX protein and the ATRX binding agent, and

(b) isolating the complexes.

In some embodiments, the methods further include (c) synthesizing cDNA complementary to the nRNA, and (d) selecting cDNAs that (i) have RPKM above a desired threshold or (ii) are enriched compared to a control library, or both (i) and (ii).

In a further aspect, the invention provides methods for preparing a plurality of cDNAs complementary to a pool of nuclear ribonucleic acids (nRNAs). Preferably, the methods include providing a sample comprising nuclear ribonucleic acids, e.g., a sample comprising nuclear lysate, e.g., comprising nRNAs bound to nuclear proteins; contacting the sample with an agent, e.g., an antibody, that binds specifically to ATRX protein, under conditions sufficient to form complexes between the agent and ATRX proteins, e.g., such that the nRNAs remain bound to the ATRX proteins; isolating the complexes; synthesizing DNA complementary to the nRNAs to provide an initial population of cDNAs; optionally PCR-amplifying the cDNAs using strand-specific primers; purifying the initial population of cDNAs to obtain a purified population of cDNAs that are at least about 20 nucleotides (nt) in length, e.g., at least 25, 50, 100, 150 or 200 nt in length; sequencing at least part of substantially all of the purified population of cDNAs; comparing the high-confidence sequences to a reference genome, and selecting those sequences that have a high degree of identity to sequences in the reference genome, e.g., at least 95%, 98%, or 99% identity, or that have fewer than 10, 5, 2, or 1 mismatches; and selecting those cDNAs that have (i) reads per kilobase per million reads (RPKM) above a desired threshold, and (ii) are enriched as compared to a control library (e.g., a protein-null library or library made from an IgG pulldown done in parallel); thereby preparing the library of cDNAs.

In some embodiments, the methods further include a step of crosslinking the nRNAs bound to nuclear proteins, e.g., using methods known in the art, including chemical or other crosslinkers, e.g., ultraviolet irradiation.

In some embodiments of the methods described herein, the agent is an antibody and isolating the complexes comprises immunoprecipitating the complexes.

In some embodiments of the methods described herein, the cDNAs are synthesized using strand-specific adaptors.

In some embodiments, the methods described herein include sequencing substantially all of the cDNAs.

In a further aspect, the invention provides libraries of cDNAs complementary to a pool of nuclear ribonucleic acids (nRNAs) prepared by a method described herein. In some embodiments, each of the cDNAs is linked to an individually addressable bead or area on a substrate.

In a further aspect, the invention provides methods for identifying compounds that disrupts binding of one or more nuclear RNAs (nRNAs), e.g., long non-coding RNAs (lncRNAs) or coding mRNAs, to ATRX protein. Preferably, the methods include providing a sample comprising an nRNA and ATRX, wherein the nRNA can bind to the ATRX and form nRNA-ATRX complexes; contacting the sample with a test compound; and detecting the formation of nRNA-ATRX complexes in the presence and the absence of the test compound, wherein a decrease in formation of nRNA-ATRX complexes in the presence of the test compound as compared to formation of nRNA-ATRX complexes in the absence of the test compound indicates that the test compound disrupts binding of the lncRNA to ATRX. In some embodiments, the interacting RNA is and lncRNA; in some embodiments, the nRNA is a coding origin (mRNA). Herein, nRNA includes nuclear lncRNA and mRNA.

In some embodiments of the methods described herein, the sample is a cell-free sample. In some embodiments, the sample comprises a cell expressing the nRNA and ATRX. In some embodiments, the sample is from a mammalian cell, e.g., a human cell or a non-human animal cell, e.g., a non-human primate, cow, pig, sheep, horse, cat, dog, or other domestic or agricultural animal.

In some embodiments of the methods described herein, the ATRX, the nRNA, or both, is labeled.

In some embodiments, the test compound is a nucleic acid, e.g., an antagomir, mixmer, or gapmer of LNA.

In some embodiments, the methods described herein further include isolating nRNA-ATRX complexes from the sample, and optionally isolating unbound ATRX from the sample, e.g., by contacting the sample with an anti-ATRX antibody, and isolating nRNA-ATRX-antibody complexes and unbound ATRX.

In some embodiments, the methods further include selecting a compound that disrupts binding of the nRNA to ATRX; contacting a tumor cell with the compound; measuring proliferation, survival, or invasiveness of the tumor cell in the presence and absence of the compound; and identifying as a candidate therapeutic compound a compound that inhibits proliferation, affects survival, e.g., induces or promotes cell death, or reduces or delays metastasis, of the tumor cell.

In some embodiments, the methods further include administering the candidate compound to an animal model of cancer, and detecting an effect of the compound on cancer in the animal model, e.g., an effect on tumor size or metastasis.

In a further aspect, the invention provides methods for identifying an RNA target for the treatment of cancer, the method comprising: (a) comparing (i) a library of nRNAs that specifically bind ATRX prepared from a normal cell with (ii) a library of nRNAs that specifically bind ATRX prepared from a cancerous cell, wherein the to normal cell and cancerous cell are of the same tissue type; and (b) identifying an nRNA that is differentially expressed between the libraries of (a)(i) and (a)(ii) as an RNA target for treatment of cancer.

As used herein, “ATRX” refers to transcriptional repressor protein ATRX, the human hom*olog of which has an mRNA sequence as set forth in the GenBank database at Accession No. NM_000489.4, and a protein sequence as set forth at Accession No. NP_000480.3.

Unless otherwise defined, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this invention belongs. Methods and materials are described herein for use in the present invention; other, suitable methods and materials known in the art can also be used. The materials, methods, and examples are illustrative only and not intended to be limiting. All publications, patent applications, patents, sequences, database entries, and other references mentioned herein are incorporated by reference in their entirety. In case of conflict, the present specification, including definitions, will control.

Other features and advantages of the invention will be apparent from the following detailed description and figures, and from the claims.

Middle: Western blot showing ATRX depletion but constant EZH2 levels in shATRX-1 and shATRTX-2 female MEFs.

Right: H3K27me3 staining and Xist RNA FISH show no change in the intensity or foci number after transient ATRX KD.

(J,K) Immunostaining of ATRX, EZH2 (J), and H3K27me3 (K), and Xist RNA/DNA FISH in indicated cells lines after 24 h dox induction. % with shown pattern and sample size (n) are indicated.

FIGS. 3A-I: ATRX binds RNA and stimulates RNA binding to PRC2.

(A,B) RIP+UV crosslinking in X+P transgenic MEFs after dox-induction for 24 h (A) or WT MEF (B) Primers spanning Xist exons 1-3 and U1 snRNAs were used for qPCR. 1% input was processed in parallel. S.E. from 3 independent experiments. P values calculated using Student t-test.

(C) One possible structure of Repeat A (Maenner et al., 2010).

(D) Coomassie stain of purified full-length FLAG-ATRX-HA (left), C-terminal SNF2/helicase (middle), and N-terminal ADD (right) domains.

(E) RNA EMSA with ATRX at indicated concentrations and 0.2 nM of each probe. B, bound; U, unbound probe.

(F) Left: Binding Isotherms of ATRX for indicated RNAs. S.E. from 3 independent experiments shown. Right: Table summarizing K_d's and R² values. >>200 nM indicates K_d's out of the assay range. N/A, not applicable.

(G) EMSA of ATRX with different RNA, dsDNA, or ssDNA probes.

(H) Left: Binding Isotherms of ATRX for indicated DNAs. S.E. for 3 independent experiments shown for each point. Right: Table summarizing K_d's and R² values. >>200 nM indicates K_d's out of the assay range. N/A, not applicable.

(I) RNA and DNA EMSAs using ATRX truncation mutants, with summary of results.

FIGS. 4A-F: A ternary complex of ATRX, PRC2, and RNA, with stimulation of PRC2 binding in an ATP-dependent manner.

(A) FLAG-ATRX-HA binding to PRC2±ATP in the absence of RNA. HA-immunoprecipitated material was probed with α-EZH2 antibodies for Western.

(B) Tandem IP to detect ternary complex formation between ATRX, Repeat A RNA, and PRC2.

(C) Binding reaction to test for simultaneous association of ATRX with RNA and DNA. RNA and DNA can be discerned by their different electrophoretic mobilities.

(D) Top: Inverted image of RNA gel showing Repeat A RNA recovered after EZH2 IP±ATRX and ±ATP. Bottom: Western blot showing protein levels in input and recovered after IP. Representative results from 3 independent experiments shown.

(E) Photo crosslinking of PRC2 and Repeat A RNA in the presence of ATRX with ATP or AMP-PNP. Representative results from 5 biological replicates shown.

(F) Filter binding of ATRX binding to Repeat A RNA (left) or DNA (right)±ATP and +AMP-PNP. ATRX concentrations are indicated and S.E. from 3 independent experiments shown. *, p<0.05.

FIGS. 6A-G: Global roles of ATRX in regulating PRC2 localization and function

(A) RefSeq genes (21,677) divided into equal quartiles (Q1-Q4) based on ATRX coverage. Average ATRX coverages and number of EZH2 target genes (peaks) in each quartile are shown.

(B) Dot plots showing Log 2 densities of EZH2 and H3K27me3 in ATRX+vs. ATRX KD MEFs for each quartile. ****, P<<0.0001, as calculated by the Student t test.

(C) Scatterplot of EZH2 and H3K27me3 densities in Log 2 scale over all genes (grey dots) and Q1 PRC2 target genes (purple dots) in ATRX+ or ATRX KD MEFs. The difference between ATRX+ and ATRX KD cells of Q1 PRC2 target genes is highly significant (P<2.2e-16, Student t test) for both epitopes.

(D) Probability density function for EZH2 and H3K27me3 based on coverages over the Q1-PRC2 target genes in the indicated MEF samples.

(E) Metagene profiles of EZH2 (top) and H3K27me3 (bottom) coverage. The metagenes are scaled 0 to 1 from genic start (TSS, x=0) to end (TTS, x=1).

(F) Distribution of EZH2 and H3K27me3 peaks in indicated MEF samples.

Peaks are categorized as TSS (+/−3 Kb), genic, or intergenic. Total number of peaks called for each dataset is indicated next to each chart.

(G) ChIP-seq tracks showing EZH2 and H3K27me3 densities in ATRX+ and ATRX-KD MEFs. Black bars, significantly enriched segments. H3K4me3 (pink) and RNA

Seq (blue) tracks are as published (Yang et al., 2010; Yildirim et al., 2012).

FIGS. 7A-E: Loss of EZH2 from target genes as a result of ATRX knockdown results in activation of these genes.

(A, B) Left panel: ChIP-seq tracks of the Hoxd cluster (A) or individual genes (B) showing EZH2 and H3K27me3 patterns in ATRX+ and ATRX-KD MEFs. Black bars, statistically significant enriched segments. Replicates are labeled Rep1 and Rep2. H3K4me3 (pink) and RNA Seq (blue) tracks were previously published (Yang et al., 2010; Yildirim et al., 2012).

(C) qRT-PCR analysis of expression levels before and after ATRX KD. Averages and S.E. of 3 independent experiments shown with Student t-test P values.

(D) EZH2 and H3K27me3 coverages over TSS before and after ATRX KD.

(E) Model: ATRX-dependent targeting of PRC2 on a genome-wide scale. ATRX mediates targeting either by directly binding DNA, or via regulatory RNAs such as those in the PRC2 interactome.

FIGS. 8A-B: FLAG-macroH2A pulldown and mass spectrometry of 293 cells identifiy interacting proteins. FIG. 8 relates to FIG. 1.

(A) Colloidal blue staining of FLAG immunoprecipitates from a control (293F) and 293F FLAG macroH2A expressing cell line run on a 4-20% SDS gradient gel (top). Samples are indicated on top of the panels. Peptides recovered and validated as high confidence hits for each protein are shown to the right. Histone peptides were too numerous to be labeled. Bands labeled with asterisks correspond to the following: Black asterisk, one ATRX peptide at a lower than expected molecular weight and red asterisk, macroH2A peptides.

(B) The validated spectra for ATRX is shown at the bottom.

FIGS. 10A-F: Biochemical analysis of ATRX-RNA and PRC2-RNA interactions. FIGS. 10A-F relates to FIG. 3.

(A) In vitro transcribed Repeat A and MBP RNAs were run on a polyacrylamide UREA gel and gel purified to obtain the full length RNA for gel shift assays.

(B) Left: RNA EMSA PRC2 at indicated concentrations and 0.2 nM probe of species indicated. Locations of bound (B) and unbound (U) RNA probe are marked. Right: Table summarizing K_d's and R² values of PRC2 bound to different RNA species. >>1000 for PRC2 indicate K_d's that were out of the assay range. N/D, not determined. N/A, not applicable.

(C) RNA EMSA with ATRX at indicated concentrations and 0.2 nM probe of species indicated. Locations of bound (B) and unbound (U) RNA probe are marked.

(D) Double filter binding assay using increasing concentrations of ATRX (0, 2.5, 5 and 10 nM) with either Repeat A RNA or MBP RNA. Upper panel represents RNA bound to ATRX and the lower panel free or unbound RNA. Experiment was done in triplicate to show reproducibility across wells.

(E) ATRX binding to ³²P-labeled Repeat A (I-IV) RNA in the presence of increasing concentrations of Repeat A DNA competitor.

(F) ATRX binding to ³²P-labeled Repeat A (I-IV) dsDNA in the presence of increasing concentrations of Repeat A (I-IV) RNA competitor.

FIGS. 11A-D: ATRX densities across the X chromosome and chromosome 13 in WT and ATRX KD MEFs. FIGS. 11A-D relate to FIG. 5.

(A) ATRX densities with 50-kb bin sizes across the entire X chromosome. The highest density of ATRX on the X chromosome is observed around the X inactivation center (Xic) that is marked in red.

(B) ATRX densities with 50-kb bin sizes across 50 kb of the Xic, centered on Xist. The genomic co-ordinates are indicated on the x axis.

(C) ATRX ChIP-seq tracks showing M. musculus (mus), M. castaneus (cas), and composite (comp; sum of both allelic and non-alleleically aligned reads) across a Chr13 locus in day 0 and 7 mESC and MEFs. Positions of peaks/enriched segments (grey) are shown under comp tracks.

(D) ChIP-seq tracks showing Mus musculus (MUS, red), Mus castaneus (CAS, red) and total allelic and non-allelically assigned (COMP, black) ATRX reads across the Xist locus in WT (top) and ATRX KD (bottom) MEFs. The ATRX-KD samples show significant depletion of signal along the Xist gene.

FIGS. 12A-B: Correlation between EZH2 and H3K27me3 coverages. FIGS. 12A-B relates to FIG. 6.

(A) In ATRX+ cells, correlation between two EZH2 ChIP-seq replicates and between EZH2 and H3K27me3 ChIP-seq profiles. Pearson's correlation coefficient (r) is shown.

(B) In ATRX-deficient cells, correlation between two EZH2 ChIP-seq replicates and between EZH2 and H3K27me3 ChIP-seq profiles. Pearson's correlation coefficient (r) is shown.

FIGS. 13A-D: EZH2 and H3K27me3 distributions in ChIP-seq replicate experiments and their effect on genic loci upon ATRX KD. FIGS. 13A-D relate to FIG. 7.

(A,B) ChIP-seq tracks of the Hoxd cluster (Replicate 2) or Hoxc cluster (replicates 1 and 2) showing EZH2 and H3K27me3 patterns in ATRX+ and ATRX-KD MEFs. Statistically significant enriched segments are shown below each track as black bars. Replicates are labeled Rep1 and Rep2. H3K4me3 (pink) and RNA-seq (blue) tracks were previously published (Yang et al., 2010; Yildirim et al., 2012).

(C) qRT-PCR analysis of expression levels before and after ATRX KD. Expression levels were consistently higher upon ATRX knockdown, though the differences were modest (˜2-fold) and bordered statistical significance for Hox genes, with P-values ranging from 0.01-0.1, possibly because Hox genes are tissue-specific and are repressed by multiple mechanisms (i.e., ATRX and PRC2 are not the only regulator). Averages and S.E. of 3 independent experiments with Student t-test P values are shown.

(D) EZH2 and K27me3 TSS coverage tables for individual genes in ChIP-seq Rep2.

FIG. 14: Behavior of Polycomb target genes without ATRX coverage (Q4 quartile). FIG. 14 relates to FIGS. 6 and 7.

ChIP-seq track of two Polycomb target genes, Onecut1 and Klh131, without ATRX coverage (Q4 quartile). ATRX-deficiency minimally affected EZH2 localization. Statistically significant enriched segments are shown below each track as black bars. H3K4me3 (pink) and RNA-seq (blue) tracks were previously published (Yang et al., 2010; Yildirim et al., 2012).

Table 1: ATRX binding sites (ChIP-Seq peaks) from Mus musculus mouse embryonic fibroblasts. The columns correspond to c1: Chromosome number. c2: Read start position of ATRX binding site. c3: read end position of binding site. c4: coding strand of the nearest gene. c5: nearest gene name. ‘int’ signifies an intergenic binding site, or a binding site not located within any annotated gene. All coordinates in mm9

Table 2: Conserved human genomic regions that correspond to ATRX ChIP-Seq peaks from Mus musculus embryonic fibroblasts. Coordinates of ATRX Chip seq peaks that are aligned to the mouse mm9 genome are converted or ‘lifted over’ to the human Hg19 genome. The columns correspond to c1: Chromosome number. c2: Read start position of ATRX binding site. c3: read end position of binding site. c4: coding strand of the nearest gene. c5: nearest gene name. ‘int’ signifies an intergenic binding site, or a binding site not located within any annotated gene.

Table 4: PRC2 target genes in the Q1 quartile.

Table 5: List of 100 genes with highest ATRX coverage in MEFs, with coverage values shown.

Table 6: Primer sequences and Antibodies used in Examples 1-6.

Polycomb repressive complex 2 (PRC2) (Dupont and Gribnau, 2013; Lee, 2012; Lee and Bartolomei, 2013; Starmer and Magnuson, 2009; Wutz, 2011) is a histone methyltransferase complex that trimethylates histone H3 at lysine 27 (H3K27me3) and establishes repressive chromatin (Muller and Verrijzer, 2009; Simon and Kingston, 2013). Because PRC2 controls both normal development and the pathogenesis of disease, PRC2 has become a high-priority drug target (Helie and Dhanak, 2013). Apart from the Xi, PRC2 binds thousands of specific sites in the mammalian genome. Still not fully understood is how PRC2 is targeted when the core subunits are not sequence-specific DNA binding proteins. PRC2 preferentially occupies CpG-rich regions and is aided in recruitment by substoichiometric association with the Jumonji protein, JARID2, and the Zinc-finger protein, AEPB2 (Cifuentes-Rojas et al., 2014; da Rocha et al., 2014; Kaneko et al., 2014; Simon and Kingston, 2013). However, other specificity determinants must exist in vivo, given that JARID2 and AEPB2 are nonspecific DNA-binding proteins and cannot by themselves impart specificity to PRC2 localization.

The example of RepA/Xist RNA demonstrates that cis-regulatory RNAs can serve as locus-specific recruiting factors (Zhao et al., 2008). Because such transcripts are unique in the genome, their cis-action enables targeting of chromatin complexes to a singular location (Lee, 2012). During XCI, PRC2 is first targeted to the Xic by RepA via the Repeat A motif Xist RNA then co-transcriptionally binds PRC2 via Repeat A and loads in cis onto a nucleation center before spreading outwardly to envelop the Xi (Jeon and Lee, 2011). PRC2 is now known to have a large RNA interactome, with membership exceeding 9,000 transcripts (Kaneko et al., 2013; Kanhere et al., 2010; Khalil et al., 2009; Zhao et al., 2010). In vitro, PRC2 can bind RNA with a range of affinities (Cifuentes-Rojas et al., 2014; Davidovich et al., 2013). The large RNA interactome raises the question of how PRC2 discriminates between RNA species in the physiological context. Here, we investigate this question by carrying out an unbiased screen for novel specificity determinants. We identify the chromatin remodeler, ATRX.

Although in vitro analysis shows that PRC2 is intrinsically capable of distinguishing between cognate and nonspecific transcripts (Cifuentes-Rojas et al., 2014), the thousands of possible RNA partners in vivo (Zhao et al., 2010) argue that additional specificity determinants must exist in the physiological context in order for PRC2 to be targeted in a locus-specific manner. Our proteomics screen has identified ATRX as an essential discrimination factor for PRC2 localization and function on a global scale. During XCI, ATRX interacts with RepA/Xist RNA and promotes loading of PRC2 (FIG. 5L). ATRX is an avid but also specific RNA-binding protein, with an impressive K_d of ˜5 nM for Repeat A RNA. ATRX is also an avid DNA-binding protein (Kd ˜15 nM). ATRX's interaction with Repeat A DNA leaves it poised, enabling ATRX to bind RepA/Xist RNAs in a co-transcriptional manner. Augmentation of ATRX's effect on Repeat A-PRC2 interactions by ATP suggests a role, potentially as an RNA helicase, to remodel Repeat A RNA to a permissive structure. This activity may parallel that of the MLE helicase for the MSL-roX dosage compensation complex of the fruitfly (Ilik et al., 2013; Maenner et al., 2013). In the case of XCI, absence of ATRX action at both Repeat A RNA and DNA precludes PRC2 targeting and spread along the Xi in cis.

ATRX's specificity function extends beyond XCI. Without ATRX, PRC2 cannot discriminate true targets from nonspecific sequences. Loss of ATRX causes global shifts in PRC2 localization and function, with PRC2 redistributing en masse to ectopic sites in intergenic space and to non-canonical sites within coding genes. At ectopic loci, PRC2 seems to lack proper context and cofactors to carry out H3K27 methylation in a specific manner. While fewer enriched H3K27me3 peaks are observed, overall H3K27me3 levels are unchanged. Loss of ATRX-dependent PRC2 function results in upregulation of Polycomb target genes (FIG. 7, 13A-D). We do not know whether ATRX action at autosomal genes depends on noncoding transcripts, such as RepA/Xist in the case of XCI. Because ATRX can bind DNA directly (FIG. 3H), ATRX-dependent targeting of PRC2 at autosomal loci may be mediated by either RNA or DNA (FIG. 7E). The large RNA interactome of PRC2 may also play a role. Finally the presence of EZH2 peaks in Q3 and Q4 genes (FIG. 6A) suggests the presence of an ATRX-independent mechanism for PRC2 recruitment as well: For example, at the Q4 Polycomb targets, Onecut1 and Klh131, loss of ATRX did not affect EZH2 targeting (FIG. 14).

ATRX's role in PRC2 targeting and function may help elucidate the pathogenesis of the X-linked alpha-thalassemia mental retardation (ATR-X) syndrome (Gibbons et al., 2008), a disease associated with aberrant chromatin at telomeres and pericentric heterochromatin (Clynes et al., 2013). Mutations in ATRX and its interacting partner, DAXX, are also frequently mutated in tumors that rely on hom*ologous recombination instead of telomerase activation to circumvent telomere shortening. Our present study suggests that, in addition to abnormalities in constitutive heterochromatin caused by abnormal H3K9 methylation, ATRX-associated diseases may also be ascribed to dysregulated PRC2 function in facultative heterochromatin. ATRX-mediated targeting of PRC2 thus provides a new framework for understanding Polycomb biology and human disease.

Methods of Producing ATRX-Binding Nuclear RNAs

Described herein are methods for producing libraries of nRNAs, e.g., nuclear IncRNAs and coding mRNAs, that bind to ATRX. In some embodiments, the methods include the of RIP-SEQ, e.g., as shown in FIG. 1A of WO2012/065143, with ATRX substituted for a-EZH2; one of skill in the art will appreciate that other techniques can be substituted for those shown, including the use of PAR-CLIP or HITS-CLIP, e.g., as described in Konig et al., Nature Reviews Genetics 13:77-83 (2012).

In some embodiments, the methods include providing a sample comprising nuclear ribonucleic acids (nRNAs) bound to ATRX; and contacting the sample with an agent, e.g., an antibody, that binds specifically to ATRX, under conditions and for a time sufficient to form complexes between the agent and the protein; isolating the complexes; synthesizing DNA complementary to the nRNAs to provide an initial population of cDNAs; PCR-amplifying, if necessary, using strand-specific primers; purifying the initial population of cDNAs to obtain a purified population of cDNAs that are at least 20 nucleotides (nt) in length; high-throughput sequencing the purified population of cDNAs. hom*opolymer reads are filtered, and reads matching the mitochondrial genome and ribosomal RNAs are excluded from all subsequent analyses. Reads that align to a reference genome with ≤1 mismatch are retained, excluding hom*opolymers, reads that align to the mitochondrial genome, and ribosomal RNAs. High probability ATRX-interacting transcripts are then called based on two criteria: (1) that the candidate transcript has a minimum read density in RPKM terms (number of reads per kilobase per million reads); (2) that the candidate transcript is enriched in the wildtype library versus a suitable control library (such as a protein-null library, a library made from an IgG pulldown done in parallel, or a minus-crosslink (−UV) control library done in parallel).

In general, to construct native RIP-seq libraries, cell nuclei are prepared, treated with DNAse, and incubated with antibodies directed against a chromatin-associated factor of interest, along with a control IgG reaction in parallel. RNA-protein complexes are then immunoprecipitated with agarose beads, magnetic beads, or any other platform in solution or on a solid matrix (e.g., columns, microfluidic devices). RNAs are extracted using standard techniques. To capture all RNAs (not just polyA RNAs) and to preserve strand information, asymmetric primers are used to generate cDNA from the RNA template, in which the first adaptor (adaptor 1) to make the first strand cDNA contains a random multimer sequence (such as random hexamers) at the 3′ end. A reverse transcriptase is used to create the first strand. A distinct second adaptor (adaptor2) is used to create the second strand. One example is as follows: If Superscript II is used, it will add non-template CCC 3′ overhangs, which can then be used to hybridize to a second adaptor containing GGG at the 3′ end, which anneal to the non-template CCC overhangs. Other methods of creating second strands may be substituted. PCR using adaptor1- and adaptor2-specific primer pairs is then the performed to amplify the cDNAs and the products sequenced via standard methods of high throughput sequencing. Prior to sequencing, a size-selection step can be incorporated (if desired) in which RNAs or cDNAs of desired sizes are excised after separation by gel electrophoresis (e.g., on a Nu-Sieve agarose gel or in an acrylamide gel) or other methods of purification, such as in a microfluidic device or in standard biochemical columns.

To construct libraries from UV-crosslinked samples, a number of techniques are currently available, including CLIP, iCLIP, PAR-CLIP, and HITS-CLIP (e.g., as described in Konig et al., Nature Reviews Genetics 13:77-83 (2012)), with appropriate modifications made based on the epitope of interest. As one example of a crosslinking protocol, cells are trypsinized and resuspended in PBS. Cells for +UV experiments are crosslinked with 256 nm UV (or 365 nm for PAR-CLIP) in a 15-cm dish at 250 mJ/cm² using the Stratalinker 1800 (Stratagene). ±UV cell pellets are resuspended in 1-2 mL Buffer A (10 mM HEPES pH 7.9, 1.5 mM MgCl₂, 10 mM KCl, 0.5 mM PMSF) and incubated on ice for 30 min with frequent vortexing. Nuclei are pelleted at 2500×g for 15 min, washed in PBS, resuspended in 500 mL Buffer C (20 mM HEPES pH 7.5, 420 mM NaCl, 15% glycerol, 1.5 mM MgCl₂, 0.5 mM PMSF, protease and RNase inhibitors) and incubated at 4° C. for 30 min with rotation. Nuclear lysates are diluted with one volume of 20 mM HEPES pH 7.5 and treated with 40 U TURBO DNase at 37° C. for 30 min to liberate chromatin-associated CTCF-RNA complexes. After quenching the DNase with 10 mM EDTA, 5% is removed and saved for RNA-seq, while the remainder is added with sarkosyl to 0.5% and the RNA fragmented by sonication with Diagenode Bioruptor XL twice for 20 min each (with 30 s on, 30 s off cycles). Cell debris is pelleted at 16,000×g for 10 min, the lysate is diluted again with 1 volume of 20 mM HEPES and divided into three aliquots. 15 uL of Anti-FLAG M2 Magnetic Beads (Sigma-Aldrich A2220) is added to each aliquot and incubated at 4° C. overnight with rotation. Beads from all aliquots is recombined, washed 3× with high salt Wash Buffer I (20 mM HEPES pH 7.5, 250 mM NaCl, 0.1% SDS, 0.5% sodium deoxycholate, 0.5% Nonident-P40, protease and RNase inhibitors), once with 1× TURBO DNase buffer, then treated with 100 U/mL TURBO DNase at 37° C. for 30 min. Beads are further washed 2× with Wash Buffer I supplemented with 10 mM EDTA, then 2× with low salt Wash Buffer II (50 mM Tris-HCl pH 7.5, 1% NP40, 0.5% sodium deoxycholate, 50 mM NaCl, 10 mM EDTA), and 1× with PNK buffer (50 mM Tris-HCl pH 7.5, 10 mM MgCl₂, 0.5% NP40, 5 mM DTT). CLIP-tags on beads are radiolabelled with [γ-³²P]ATP using T4 polynucleotide kinase (New England Biolabs) for 20 min at 37° C., and washed 4× with PNK buffer. Beads are resuspended in SDS-PAGE loading buffer at heated for 5 min at 70° C., run on 8% Bis-Tris SDS-PAGE in MOPS buffer (50 mM MOPS, 50 mM Tris, 0.1% SDS, 1 mM EDTA) at 120 V, transferred to nitrocellulose membrane, and exposed to film for autoradiography or used for immunoblot with 1:3000 αFLAG antibodies (Sigma-Aldrich F1804).

Membrane fragments containing CLIP signal, as confirmed by immunoblot, and corresponding positions on control lanes are excised, and RNA is eluted by incubation in prewarmed proteinase K buffer (100 mM Tris-HCl pH 7.5, 50 mM NaCl, 10 mM EDTA, 0.5% SDS, 4 mg/mL proteinase K) for 20 min at 37° C., then incubation for an additional 20 min in proteinase K buffer supplemented with 7 M urea, followed by TRIzol extraction and ethanol precipitation. RNA size and quality are verified using RNA 6000 Pico chips on the Agilent Bioanalyzer. CLIP-seq library is then constructed from CLIP RNA using the NEBNext Small RNA Library Prep set (New England Biolabs E7330), size-selected and cleaned up of primer/adaptor-dimers using Agencourt AMPure XP beads (Beckman Coulter A63880), verified with DNA High Sensitivity chips on the Agilent Bioanalyzer, quantitated using KAPA Biosystems library quantification kit (KK4844), and sequenced with the Illumina HiSeq 2000 system with 50 cycles paired end reads.

ATRX-Binding nRNAs and nRNA Libraries

The present invention includes libraries of nRNAs produced by methods described herein. In some embodiments, the libraries are in solution, or are lyophilized. In some embodiments, the libraries are bound to a substrate, e.g., wherein each member of the library is bound to an individually addressable member, e.g., an individual area on an array (e.g., a microarray), or a bead.

nRNAs may be functionally conserved without being highly conserved at the level of overall nucleotide identity. For example, mouse Xist shows only 76% overall nucleotide identity with human XIST using sliding 21-bp windows, or an overall sequence identity of only 60%. However, within specific functional domains, such as Repeat A, the degree of conservation can be >70% between different mammalian species. The crucial motif in Repeat A is the secondary structures formed by the repeat. For ATRX-Xist interactions, the crucial motif appears to be Repeat A. Other nRNAs interacting with ATRX may therefore be similarly low in overall conservation but still have conservation in secondary structure within specific domains of the RNA, and thereby demonstrate functional conservation with respect to recruitment of ATRX.

Calculations of hom*ology or sequence identity between sequences (the terms are used interchangeably herein) are performed as follows.

To determine the percent identity of two nucleic acid sequences, the sequences are aligned for optimal comparison purposes (e.g., gaps can be introduced in one or both of a first and a second amino acid or nucleic acid sequence for optimal alignment and non-hom*ologous sequences can be disregarded for comparison purposes). The length of a reference sequence aligned for comparison purposes is at least 80% of the length of the reference sequence, and in some embodiments is at least 90% or 100%. The nucleotides at corresponding amino acid positions or nucleotide positions are then compared. When a position in the first sequence is occupied by the same nucleotide as the corresponding position in the second sequence, then the molecules are identical at that position (as used herein nucleic acid “identity” is equivalent to nucleic acid “hom*ology”). The percent identity between the two sequences is a function of the number of identical positions shared by the sequences, taking into account the number of gaps, and the length of each gap, which need to be introduced for optimal alignment of the two sequences.

For purposes of the present invention, the comparison of sequences and determination of percent identity between two sequences can be accomplished using a Blossum 62 scoring matrix with a gap penalty of 12, a gap extend penalty of 4, and a frameshift gap penalty of 5.

There are several potential uses for the nRNAs described herein in the ATRX transcriptome: The RNAs themselves, or antagomirs and small molecules designed against them, can be utilized to modulate expression (either up or down) of ATRX target genes. In addition, the nRNAs can be used in methods of detecting or identifying cancerous cells, as described herein.

Modulating ATRX Binding to Genomic DNA

As described herein, ATRX localizes to genomic DNA in a sequence-specific manner. To inhibit this localization, and thus disrupt the ATRX-dependent loading of PRC2 and increase expression of nearby genes, oligonucleotides are used that bind to genomic DNA at or near (e.g., within 100, 50, or 25) nucleotides of an ATRX localization site, identified as an ATRX ChIP-Seq peak in Tables 1 and 2. Table 1 lists genes corresponding to the ATRX ChIP-Seq peaks from Mus musculus mouse embryonic fibroblasts, and Table 2 provides the Human genomic regions determined by LiftOver analysis to correspond to the ATRX ChIP-Seq peaks from Mus musculus mouse embryonic fibroblasts. Each table provides the SEQ ID NO: of the peak(s) (i.e., the ATRX localization site(s)) that correspond to each of the listed genes.

In some embodiments, the oligonucleotides are triplex-forming oligonucleotides (TFOs). TFOs are defined as triplex-forming oligonucleotides which bind as third strands to duplex DNA in a sequence specific manner. Triplex-forming oligonucleotides may be comprised of any possible combination of nucleotides and modified nucleotides. Modified nucleotides may contain chemical modifications of the heterocyclic base, sugar moiety or phosphate moiety. TFOs, and methods of making them, are known in the art; see, e.g., Frank-Kamenetskii and Mirkin, Annual Review of Biochemistry, 64:65-95 (1995); Vasquez and Glazer, Quarterly Reviews of Biophysics, 35(01):89-107 (2002); US PGPub Nos. 20070219122; US20110130557; and US20090216003.

In general, the TFO is a single-stranded nucleic acid molecule between 5 and 100 nucleotides in length, preferably between 7 and 40 nucleotides in length, e.g., 10 to 20 or 20 to 30 nucleotides in length. In some embodiments, the base composition is hom*opurine or hom*opyrimidine, polypurine or polypyrimidine. The oligonucleotides can be generated using known DNA synthesis procedures.

The nucleotide sequence of the oligonucleotides is selected based on a target sequence of an ATRX localization sequence as provided herein; in addition, the sequence can be determined based on physical constraints imposed by the need to achieve binding of the oligonucleotide within the major groove of the target region, and preferably have a low dissociation constant (Kd) for the oligonucleotide/target sequence. The oligonucleotides should have a base composition that is conducive to triple-helix formation and can be generated based on known structural motifs for third strand binding. The most stable complexes are formed on polypurine:polypyrimidine elements, which are relatively abundant in mammalian genomes. Triplex formation by TFOs can occur with the third strand oriented either parallel or anti-parallel to the purine strand of the duplex. In the anti-parallel, purine motif, the triplets are G.G:C and A.A:T, whereas in the parallel pyrimidine motif, the canonical triplets are C⁺.G:C and T.A:T. The triplex structures are stabilized by two Hoogsteen hydrogen bonds between the bases in the TFO strand and the purine strand in the duplex. See U.S. Pat. No. 5,422,251 for additional information on base compositions for third strand binding oligonucleotides.

The TFOs can include one or more modifications, e.g., backbone modifications such as incorporation of the flexible basestacking monomers (Bulge insertions of (R)-1-O-[4-(1-pyrenylethynyl)phenylmethyl]glycerol into the middle of hom*opyrimidine oligodeoxynucleotides (twisted intercalating nucleic acids, TINA)) as described in US PGPub No 20090216003; intercalating nucleic acid monomers as described in WO2006125447A2; intercalator (R)-1-O-[4-(1-pyrenylethynyl)benzyl]-glycerol (see, e.g., Filichev et al., J. Am. Chem. Soc. 127:14849 (2005); Filichev et al., Eur. J. Org. Chem. 17:3960-3968 (2006); Globisch et al., Helv. Chim. Acta, 91:805 (2008)); 2-phenyl or 2-naphth-1-yl-phenanthroimidazole intercalators as described in US20110130557.

In addition or in alternative, modifications can be made to the nucleobases (see, e.g., Roig and Asseline, J. Am. Chem. Soc. 2003, 125, 4416; Hildbrand et al., J. Am. Chem. Soc. 1997, 119, 5499; and Xodo et al., Nucleic Acids Res. 1991, 19, 5625); to the sugar (sugar moiety modifications include, but are not limited to, 2′-O-aminoetoxy, 2′-O-amonioethyl (2′-OAE), 2′-O-methoxy, 2′-O-methyl, 2-guanidoethyl (2′-OGE), 2′-O,4′-C-methylene (LNA), 2′-O-(methoxyethyl) (2′-OME) and 2′-O—(N-(methyl)acetamido) (2′-OMA). 2′-O-aminoethyl sugar moiety substitutions are especially preferred; see, e.g., Carlomagno et al., J. Am. Chem. Soc. 2001, 123, 7364; Cuenoud et al., Angew. Chem. Int. Ed. 1998, 37, 1288; Wengel, Acc. Chem. Res. 1999, 32, 301; Obika et al., Tetrahedron Let. 2000, 41, 8923; Sun et al., Biochemistry, 2004, 43, 4160; Basye et al., Nucleic Acids Res. 2001, 29, 4873); and/or to the phosphate backbone (see, e.g., Michel, et al., Chem Bio Chem. 2005, 6, 1254; Ehrenmann et al., Nucl. Nucl. Nucleic Acids. 2001, 20, 797; Michel et al., J. Biomol. Struct. Dyn. 2003, 21, 435; Tereshko et al., J. Am. Chem. Soc. 1998, 120, 269; Escude et al., Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 4365; Gryaznov et al., Proc. Natl. Acad. Sci. U.S.A. 1995, 92, 5798; Gryaznov and Chen, J. Am. Chem. Soc. 1994, 116, 3143; and Chur et al., Nucleic Acids Res. 1993, 21, 5179). Chemical modifications of heterocyclic bases or heterocyclic base analogs can be used to increase the binding affinity of a nucleotide or its stability in a triplex. Chemically-modified heterocyclic bases include, but are not limited to, inosine, 5-(1-propynyl) uracil (pU), 5-(1-propynyl) cytosine (pC), 5-methylcytosine, 8-oxo-adenine, pseudocytosine, pseudoisocytosine, 5 and 2-amino-5-(2′-deoxy-beta-D-ribofuranosyl)pyridine (2-aminopyridine), and various pyrrolo- and pyrazolopyrimidine derivatives. Substitution of 5-methylcytosine or pseudoisocytosine for cytosine in triplex-forming molecules such as TFOs and PNAs helps to stabilize triplex formation at neutral and/or physiological pH, especially in triplex-forming molecules with isolated cytosines. See, e.g., US20110268810.

For example, each nucleotide monomer can be selected from the group consisting of DNA, RNA, HNA, MNA, ANA, LNA, CAN, INA, CeNA, TNA, (2′-NH)-TNA, (3′-NH)-TNA, alpha-L-Ribo-LNA, alpha-L-Xylo-LNA, beta-D-Ribo-LNA, beta-D-Xylo-LNA, [3.2.1]-LNA, Bicyclo-DNA, 6-Amino-Bicyclo-DNA, 5-epi-Bicyclo-DNA, alpha-Bicyclo-DNA, Tricyclo-DNA, Bicyclo[4.3.0]-DNA, Bicyclo[3.2.1]-DNA, Bicyclo[4.3.0]amide-DNA, beta-D-Ribopyranosyl-NA, alpha-L-Lyxopyranosyl-NA, 2′-R-RNA, 2′-OR-RNA, 2′-AE-RNA, alpha-L-RNA, and beta-D-RNA, and combinations and modifications thereof; in some embodiments, some or all of the TFO is a peptide nucleic acid (PNA), in which the phosphate backbone of oligonucleotides is replaced in part or in its entirety by repeating N-(2-aminoethyl)-glycine units, and the phosphodiester bonds are replaced by peptide bonds. In addition, the TFO can include one or more of the modifications described in WO2012/065143.

In some embodiments, the TFO includes a “tail” or “tail clamp” added to the Watson-Crick binding portion that binds to target strand outside the triple helix and reduces the requirement for a polypurine:polypyrimidine stretch, increasing the number of potential target sites. Tail clamps added to PNAs (referred to as tcPNAs) have been described by Kaihatsu, et al., Biochemistry, 42(47):13996-4003 (2003); Bentin, et al., Biochemistry, 42(47):13987-95 (2003) Rogers, et al., Proc. Natl. Acad. Sci. USA., 99(26):16695-700 (2002)), and are known to bind to DNA more efficiently due to low dissociation constants. The addition of the tail also increases binding specificity and binding stringency of the triplex-forming molecules to the target duplex.

In some embodiments, the TFOs are modified with, or administered with, amidoanthraquinones as described in Fox et al., Proc. Natl. Acad. Sci. USA 92:7887-7891 (1995).

Methods of Treatment

Oligos, e.g., TFOs that target the sequences of ATRX binding sites associated with disease-related genes can also be used to treat subjects. For example, the DMD gene is a causal factor in duch*enne muscular dystrophy; administration of an oligo, e.g., a TFO that targets an ATRX localization site associated with the DMD gene can be used to treat subjects who have duch*enne muscular dystrophy. In addition, administration of an oligo, e.g., a TFO, that targets an ATRX localization site associated with the XIST gene can be used to treat subjects who have Rett Syndrome. One of skill in the art would be able to identify other disease-related genes from among those listed in Tables 1, 2, 3, 4, or 5. In particular, an oligo, e.g., a TFO, that targets an ATRX localization site associated with a human disease-related gene as set forth in Table 2 or 3 can be used to treat a human having the disease to which the gene is related; in some embodiments, the oligos, e.g., TFOs, are used to reactivate a normal gene in a heterozygous individual, i.e., an individual with one normal copy and one affected copy of the gene. The oligo, e.g., TFO, can be administered in a pharmaceutical composition or formulation as known in the art, e.g., as described herein. Subjects having a genetic disease, e.g., a disease related to a gene listed in Table 2 or 3, can be identified using methods known in the art. Table 3 includes additional examples of diseases and the ATRX localization sites associated with the disease-related gene from Table 2. Disease identifications follow OMIM nomenclature.

Pharmaceutical Compositions

The methods described herein can include the administration of pharmaceutical compositions and formulations comprising oligonucleotides (e.g., TFOs) designed to target an ATRX localization site sequence.

In some embodiments, the compositions are formulated with a pharmaceutically acceptable carrier. The pharmaceutical compositions and formulations can be administered parenterally, topically, orally or by local administration, such as by aerosol or transdermally. The pharmaceutical compositions can be formulated in any way and can be administered in a variety of unit dosage forms depending upon the condition or disease and the degree of illness, the general medical condition of each patient, the resulting preferred method of administration and the like. Details on techniques for formulation and administration of pharmaceuticals are well described in the scientific and patent literature, see, e.g., Remington: The Science and Practice of Pharmacy, 21st ed., 2005.

The nucleic acids can be administered alone or as a component of a pharmaceutical formulation (composition). The compounds may be formulated for administration, in any convenient way for use in human or veterinary medicine. Wetting agents, emulsifiers and lubricants, such as sodium lauryl sulfate and magnesium stearate, as well as coloring agents, release agents, coating agents, sweetening, flavoring and perfuming agents, preservatives and antioxidants can also be present in the compositions.

Formulations of the compositions of the invention include those suitable for intradermal, inhalation, oral/nasal, topical, parenteral, rectal, and/or intravagin*l administration. The formulations may conveniently be presented in unit dosage form and may be prepared by any methods well known in the art of pharmacy. The amount of active ingredient (e.g., nucleic acid sequences of this invention) which can be combined with a carrier material to produce a single dosage form will vary depending upon the host being treated, the particular mode of administration, e.g., intradermal or inhalation. The amount of active ingredient which can be combined with a carrier material to produce a single dosage form will generally be that amount of the compound which produces a therapeutic effect, e.g., an antigen specific T cell or humoral response.

Pharmaceutical formulations of this invention can be prepared according to any method known to the art for the manufacture of pharmaceuticals. Such drugs can contain sweetening agents, flavoring agents, coloring agents and preserving agents. A formulation can be admixtured with nontoxic pharmaceutically acceptable excipients which are suitable for manufacture. Formulations may comprise one or more diluents, emulsifiers, preservatives, buffers, excipients, etc. and may be provided in such forms as liquids, powders, emulsions, lyophilized powders, sprays, creams, lotions, controlled release formulations, tablets, pills, gels, on patches, in implants, etc.

Pharmaceutical formulations for oral administration can be formulated using pharmaceutically acceptable carriers well known in the art in appropriate and suitable dosages. Such carriers enable the pharmaceuticals to be formulated in unit dosage forms as tablets, pills, powder, dragees, capsules, liquids, lozenges, gels, syrups, slurries, suspensions, etc., suitable for ingestion by the patient. Pharmaceutical preparations for oral use can be formulated as a solid excipient, optionally grinding a resulting mixture, and processing the mixture of granules, after adding suitable additional compounds, if desired, to obtain tablets or dragee cores. Suitable solid excipients are carbohydrate or protein fillers include, e.g., sugars, including lactose, sucrose, mannitol, or sorbitol; starch from corn, wheat, rice, potato, or other plants; cellulose such as methyl cellulose, hydroxypropylmethyl-cellulose, or sodium carboxy-methylcellulose; and gums including arabic and tragacanth; and proteins, e.g., gelatin and collagen. Disintegrating or solubilizing agents may be added, such as the cross-linked polyvinyl pyrrolidone, agar, alginic acid, or a salt thereof, such as sodium alginate. Push-fit capsules can contain active agents mixed with a filler or binders such as lactose or starches, lubricants such as talc or magnesium stearate, and, optionally, stabilizers. In soft capsules, the active agents can be dissolved or suspended in suitable liquids, such as fatty oils, liquid paraffin, or liquid polyethylene glycol with or without stabilizers.

Aqueous suspensions can contain an active agent (e.g., nucleic acid sequences of the invention) in admixture with excipients suitable for the manufacture of aqueous suspensions, e.g., for aqueous intradermal injections. Such excipients include a suspending agent, such as sodium carboxymethylcellulose, methylcellulose, hydroxypropylmethylcellulose, sodium alginate, polyvinylpyrrolidone, gum tragacanth and gum acacia, and dispersing or wetting agents such as a naturally occurring phosphatide (e.g., lecithin), a condensation product of an alkylene oxide with a fatty acid (e.g., polyoxyethylene stearate), a condensation product of ethylene oxide with a long chain aliphatic alcohol (e.g., heptadecaethylene oxycetanol), a condensation product of ethylene oxide with a partial ester derived from a fatty acid and a hexitol (e.g., polyoxyethylene sorbitol mono-oleate), or a condensation product of ethylene oxide with a partial ester derived from fatty acid and a hexitol anhydride (e.g., polyoxyethylene sorbitan mono-oleate). The aqueous suspension can also contain one or more preservatives such as ethyl or n-propyl p-hydroxybenzoate, one or more coloring agents, one or more flavoring agents and one or more sweetening agents, such as sucrose, aspartame or saccharin. Formulations can be adjusted for osmolarity.

In some embodiments, oil-based pharmaceuticals are used for administration of nucleic acid sequences of the invention. Oil-based suspensions can be formulated by suspending an active agent in a vegetable oil, such as arachis oil, olive oil, sesame oil or coconut oil, or in a mineral oil such as liquid paraffin; or a mixture of these. See e.g., U.S. Pat. No. 5,716,928 describing using essential oils or essential oil components for increasing bioavailability and reducing inter- and intra-individual variability of orally administered hydrophobic pharmaceutical compounds (see also U.S. Pat. No. 5,858,401). The oil suspensions can contain a thickening agent, such as beeswax, hard paraffin or cetyl alcohol. Sweetening agents can be added to provide a palatable oral preparation, such as glycerol, sorbitol or sucrose. These formulations can be preserved by the addition of an antioxidant such as ascorbic acid. As an example of an injectable oil vehicle, see Minto (1997) J. Pharmacol. Exp. Ther. 281:93-102.

Pharmaceutical formulations can also be in the form of oil-in-water emulsions. The oily phase can be a vegetable oil or a mineral oil, described above, or a mixture of these. Suitable emulsifying agents include naturally-occurring gums, such as gum acacia and gum tragacanth, naturally occurring phosphatides, such as soybean lecithin, esters or partial esters derived from fatty acids and hexitol anhydrides, such as sorbitan mono-oleate, and condensation products of these partial esters with ethylene oxide, such as polyoxyethylene sorbitan mono-oleate. The emulsion can also contain sweetening agents and flavoring agents, as in the formulation of syrups and elixirs. Such formulations can also contain a demulcent, a preservative, or a coloring agent. In alternative embodiments, these injectable oil-in-water emulsions of the invention comprise a paraffin oil, a sorbitan monooleate, an ethoxylated sorbitan monooleate and/or an ethoxylated sorbitan trioleate.

The pharmaceutical compounds can also be administered by in intranasal, intraocular and intravagin*l routes including suppositories, insufflation, powders and aerosol formulations (for examples of steroid inhalants, see e.g., Rohatagi (1995) J. Clin. Pharmacol. 35:1187-1193; Tjwa (1995) Ann. Allergy Asthma Immunol. 75:107-111). Suppositories formulations can be prepared by mixing the drug with a suitable non-irritating excipient which is solid at ordinary temperatures but liquid at body temperatures and will therefore melt in the body to release the drug. Such materials are cocoa butter and polyethylene glycols.

In some embodiments, the pharmaceutical compounds can be delivered transdermally, by a topical route, formulated as applicator sticks, solutions, suspensions, emulsions, gels, creams, ointments, pastes, jellies, paints, powders, and aerosols.

In some embodiments, the pharmaceutical compounds can also be delivered as microspheres for slow release in the body. For example, microspheres can be administered via intradermal injection of drug which slowly release subcutaneously; see Rao (1995) J. Biomater Sci. Polym. Ed. 7:623-645; as biodegradable and injectable gel formulations, see, e.g., Gao (1995) Pharm. Res. 12:857-863 (1995); or, as microspheres for oral administration, see, e.g., Eyles (1997) J. Pharm. Pharmacol. 49:669-674.

In some embodiments, the pharmaceutical compounds can be parenterally administered, such as by intravenous (IV) administration or administration into a body cavity or lumen of an organ. These formulations can comprise a solution of active agent dissolved in a pharmaceutically acceptable carrier. Acceptable vehicles and solvents that can be employed are water and Ringer's solution, an isotonic sodium chloride. In addition, sterile fixed oils can be employed as a solvent or suspending medium. For this purpose any bland fixed oil can be employed including synthetic mono- or diglycerides. In addition, fatty acids such as oleic acid can likewise be used in the preparation of injectables. These solutions are sterile and generally free of undesirable matter. These formulations may be sterilized by conventional, well known sterilization techniques. The formulations may contain pharmaceutically acceptable auxiliary substances as required to approximate physiological conditions such as pH adjusting and buffering agents, toxicity adjusting agents, e.g., sodium acetate, sodium chloride, potassium chloride, calcium chloride, sodium lactate and the like. The concentration of active agent in these formulations can vary widely, and will be selected primarily based on fluid volumes, viscosities, body weight, and the like, in accordance with the particular mode of administration selected and the patient's needs. For IV administration, the formulation can be a sterile injectable preparation, such as a sterile injectable aqueous or oleaginous suspension. This suspension can be formulated using those suitable dispersing or wetting agents and suspending agents. The sterile injectable preparation can also be a suspension in a nontoxic parenterally-acceptable diluent or solvent, such as a solution of 1,3-butanediol. The administration can be by bolus or continuous infusion (e.g., substantially uninterrupted introduction into a blood vessel for a specified period of time).

In some embodiments, the pharmaceutical compounds and formulations can be lyophilized. Stable lyophilized formulations comprising a nucleic acid can be made by lyophilizing a solution comprising a pharmaceutical of the invention and a bulking agent, e.g., mannitol, trehalose, raffinose, and sucrose or mixtures thereof. A process for preparing a stable lyophilized formulation can include lyophilizing a solution about 2.5 mg/mL protein, about 15 mg/mL sucrose, about 19 mg/mL NaCl, and a sodium citrate buffer having a pH greater than 5.5 but less than 6.5. See, e.g., U.S. 20040028670.

The compositions and formulations can be delivered by the use of liposomes. By using liposomes, particularly where the liposome surface carries ligands specific for target cells, or are otherwise preferentially directed to a specific organ, one can focus the delivery of the active agent into target cells in vivo. See, e.g., U.S. Pat. Nos. 6,063,400; 6,007,839; Al-Muhammed (1996) J. Microencapsul. 13:293-306; Chonn (1995) Curr. Opin. Biotechnol. 6:698-708; Ostro (1989) Am. J. Hosp. Pharm. 46:1576-1587. As used in the present invention, the term “liposome” means a vesicle composed of amphiphilic lipids arranged in a bilayer or bilayers. Liposomes are unilamellar or multilamellar vesicles that have a membrane formed from a lipophilic material and an aqueous interior that contains the composition to be delivered. Cationic liposomes are positively charged liposomes that are believed to interact with negatively charged DNA molecules to form a stable complex. Liposomes that are pH-sensitive or negatively-charged are believed to entrap DNA rather than complex with it. Both cationic and noncationic liposomes have been used to deliver DNA to cells.

Liposomes can also include “sterically stabilized” liposomes, i.e., liposomes comprising one or more specialized lipids. When incorporated into liposomes, these specialized lipids result in liposomes with enhanced circulation lifetimes relative to liposomes lacking such specialized lipids. Examples of sterically stabilized liposomes are those in which part of the vesicle-forming lipid portion of the liposome comprises one or more glycolipids or is derivatized with one or more hydrophilic polymers, such as a polyethylene glycol (PEG) moiety. Liposomes and their uses are further described in U.S. Pat. No. 6,287,860.

The formulations of the invention can be administered for prophylactic and/or therapeutic treatments. In some embodiments, for therapeutic applications, compositions are administered to a subject who is need of reduced triglyceride levels, or who is at risk of or has a disorder described herein, in an amount sufficient to cure, alleviate or partially arrest the clinical manifestations of the disorder or its complications; this can be called a therapeutically effective amount. For example, in some embodiments, pharmaceutical compositions of the invention are administered in an amount sufficient to decrease serum levels of triglycerides in the subject.

The amount of pharmaceutical composition adequate to accomplish this is a therapeutically effective dose. The dosage schedule and amounts effective for this use, i.e., the dosing regimen, will depend upon a variety of factors, including the stage of the disease or condition, the severity of the disease or condition, the general state of the patient's health, the patient's physical status, age and the like, and can be determined using methods known in the art. In calculating the dosage regimen for a patient, the mode of administration also is taken into consideration.

The dosage regimen also takes into consideration pharmaco*kinetics parameters well known in the art, i.e., the active agents' rate of absorption, bioavailability, metabolism, clearance, and the like (see, e.g., Hidalgo-Aragones (1996) J. Steroid Biochem. Mol. Biol. 58:611-617; Groning (1996) Pharmazie 51:337-341; Fotherby (1996) Contraception 54:59-69; Johnson (1995) J. Pharm. Sci. 84:1144-1146; Rohatagi (1995) Pharmazie 50:610-613; Brophy (1983) Eur. J. Clin. Pharmacol. 24:103-108; Remington: The Science and Practice of Pharmacy, 21st ed., 2005). The state of the art allows the clinician to determine the dosage regimen for each individual patient, active agent and disease or condition treated. Guidelines provided for similar compositions used as pharmaceuticals can be used as guidance to determine the dosage regiment, i.e., dose schedule and dosage levels, administered practicing the methods of the invention are correct and appropriate.

Single or multiple administrations of formulations can be given depending on for example: the dosage and frequency as required and tolerated by the patient, the degree and amount of therapeutic effect generated after each administration (e.g., effect on tumor size or growth), and the like. The formulations should provide a sufficient quantity of active agent to effectively treat, prevent or ameliorate conditions, diseases or symptoms.

In alternative embodiments, pharmaceutical formulations for oral administration are in a daily amount of between about 1 to 100 or more mg per kilogram of body weight per day. Lower dosages can be used, in contrast to administration orally, into the blood stream, into a body cavity or into a lumen of an organ. Substantially higher dosages can be used in topical or oral administration or administering by powders, spray or inhalation. Actual methods for preparing parenterally or non-parenterally administrable formulations will be known or apparent to those skilled in the art and are described in more detail in such publications as Remington: The Science and Practice of Pharmacy, 21st ed., 2005.

Various studies have reported successful mammalian dosing using complementary nucleic acid sequences. For example, Esau C., et al., (2006) Cell Metabolism, 3(2):87-98 reported dosing of normal mice with intraperitoneal doses of miR-122 antisense oligonucleotide ranging from 12.5 to 75 mg/kg twice weekly for 4 weeks. The mice appeared healthy and normal at the end of treatment, with no loss of body weight or reduced food intake. Plasma transaminase levels were in the normal range (AST 3/4 45, ALT 3/4 35) for all doses with the exception of the 75 mg/kg dose of miR-122 ASO, which showed a very mild increase in ALT and AST levels. They concluded that 50 mg/kg was an effective, non-toxic dose. Another study by Kriitzfeldt J., et al., (2005) Nature 438, 685-689, injected anatgomirs to silence miR-122 in mice using a total dose of 80, 160 or 240 mg per kg body weight. The highest dose resulted in a complete loss of miR-122 signal. In yet another study, locked nucleic acid molecules (“LNA molecules”) were successfully applied in primates to silence miR-122. Elmen J., et al., (2008) Nature 452, 896-899, report that efficient silencing of miR-122 was achieved in primates by three doses of 10 mg kg-1 LNA-antimiR, leading to a long-lasting and reversible decrease in total plasma cholesterol without any evidence for LNA-associated toxicities or histopathological changes in the study animals.

Methods of Screening

Included herein are methods for screening test compounds, e.g., polypeptides or peptides, polynucleotides or oligonucleotides, inorganic or organic large or small molecule test compounds, to identify agents useful in the treatment of cancer. As used herein, “small molecules” refers to small organic or inorganic molecules of molecular weight below about 3,000 Daltons. In general, small molecules useful for the invention have a molecular weight of less than 3,000 Daltons (Da). The small molecules can be, e.g., from at least about 100 Da to about 3,000 Da (e.g., between about 100 to about 3,000 Da, about 100 to about 2500 Da, about 100 to about 2,000 Da, about 100 to about 1,750 Da, about 100 to about 1,500 Da, about 100 to about 1,250 Da, about 100 to about 1,000 Da, about 100 to about 750 Da, about 100 to about 500 Da, about 200 to about 1500, about 500 to about 1000, about 300 to about 1000 Da, or about 100 to about 250 Da).

The test compounds can be, e.g., natural products or members of a combinatorial chemistry library. A set of diverse molecules should be used to cover a variety of functions such as charge, aromaticity, hydrogen bonding, flexibility, size, length of side chain, hydrophobicity, and rigidity. Combinatorial techniques suitable for synthesizing small molecules are known in the art, e.g., as exemplified by Obrecht and Villalgordo, Solid-Supported Combinatorial and Parallel Synthesis of Small-Molecular-Weight Compound Libraries, Pergamon-Elsevier Science Limited (1998), and include those such as the “split and pool” or “parallel” synthesis techniques, solid-phase and solution-phase techniques, and encoding techniques (see, for example, Czarnik, Curr. Opin. Chem. Bio. 1:60-6 (1997)). In addition, a number of small molecule libraries are commercially available. A number of suitable small molecule test compounds are listed in U.S. Pat. No. 6,503,713, incorporated herein by reference in its entirety.

In some embodiments, the test compounds are nucleic acids, e.g., one or more nucleic acids that have identity to all or a portion of the ATRX-binding RNA or DNA, or a set of randomly generated oligos. The oligos can be LNAs, and can be antagomirs, mixmers, or gapmers.

Libraries screened using the methods of the present invention can comprise a variety of types of test compounds. A given library can comprise a set of structurally related or unrelated test compounds. In some embodiments, the test compounds are peptide or peptidomimetic molecules. In some embodiments, the test compounds are nucleic acids.

In some embodiments, the test compounds and libraries thereof can be obtained by systematically altering the structure of a first test compound, e.g., a first test compound that is structurally similar to a known natural binding partner of the target polypeptide, or a first small molecule identified as capable of binding the target polypeptide, e.g., using methods known in the art or the methods described herein, and correlating that structure to a resulting biological activity, e.g., a structure-activity relationship study. As one of skill in the art will appreciate, there are a variety of standard methods for creating such a structure-activity relationship. Thus, in some instances, the work may be largely empirical, and in others, the three-dimensional structure of an endogenous polypeptide or portion thereof can be used as a starting point for the rational design of a small molecule compound or compounds. For example, in one embodiment, a general library of small molecules is screened, e.g., using the methods described herein.

In some embodiments, a test compound is applied to a test sample, e.g., a cancer cell, and one or more effects of the test compound is evaluated. In a cultured cancer cell for example, the ability of the test compound to inhibit proliferation or affect survival, e.g., to induce or promote cell death, is evaluated.

In some embodiments, the test sample is, or is derived from (e.g., a sample taken from) a tumor, e.g., a primary or cultured tumor cell.

Methods for evaluating each of these effects are known in the art. For example, assays of proliferation or cell survival/viability are well known in the art.

A test compound that has been screened by a method described herein and determined to inhibit proliferation or affect survival, e.g., induce or promote cell death, can be considered a candidate compound. A candidate compound that has been screened, e.g., in an in vivo model of a disorder, e.g., a xenograft model, and determined to have a desirable effect on the disorder, e.g., on growth or metastasis of a tumor, can be considered a candidate therapeutic agent. Candidate therapeutic agents, once screened in a clinical setting, are therapeutic agents. Candidate compounds, candidate therapeutic agents, and therapeutic agents can be optionally optimized and/or derivatized, and formulated with physiologically acceptable excipients to form pharmaceutical compositions.

Thus, test compounds identified as “hits” (e.g., test compounds that inhibit proliferation or affect survival, e.g., induce or promote cell death) in a first screen can be selected and systematically altered, e.g., using rational design, to optimize binding affinity, avidity, specificity, or other parameter. Such optimization can also be screened for using the methods described herein. Thus, in one embodiment, the invention includes screening a first library of compounds using a method known in the art and/or described herein, identifying one or more hits in that library, subjecting those hits to systematic structural alteration to create a second library of compounds structurally related to the hit, and screening the second library using the methods described herein.

Test compounds identified as hits can be considered candidate therapeutic compounds, useful in treating cancer. A variety of techniques useful for determining the structures of “hits” can be used in the methods described herein, e.g., NMR, mass spectrometry, gas chromatography equipped with electron capture detectors, fluorescence and absorption spectroscopy. Thus, the invention also includes compounds identified as “hits” by the methods described herein, and methods for their administration and use in the treatment, prevention, or delay of development or progression of a disorder described herein.

Test compounds identified as candidate therapeutic compounds can be further screened by administration to an animal model of a tumor, e.g., a xenograft model, as known in the art. The animal can be monitored for a change in the disorder, e.g., for an improvement in a parameter of the disorder, e.g., a parameter related to clinical outcome. In some embodiments, the parameter is tumor size, and an improvement would be a reduction or stabilization of tumor size, or a reduction in growth rate; in some embodiments, the parameter is invasiveness, and an improvement would be a reduction or delay in metastasis.

The invention is further described in the following examples, which do not limit the scope of the invention described in the claims.

Experimental Procedures

The following materials and methods were used in Examples 1-6.

Cell Lines:

Clonal Xist knockout MEFs, Tsix^TST/+ ESC, and X+P and X-RA male MEF lines were described previously (Jeon and Lee, 2011; Ogawa et al., 2008; Zhang et al., 2007). Mouse embryonic fibroblasts (MEFs) were immortalized (SV40T) and selected for clonal cell lines with Xi of mus origin. The Tsix^TST allele ensures exclusive inactivation of the mus X-chromosome in ESC.

RNA FISH, DNA FISH and Immunostaining:

RNA FISH, DNA FISH or sequential RNA-DNA FISH and immunostaining were described previously (Zhang et al., 2007).

Immunoprecipitation and Mass Spectrometry:

Nuclear extract from 293F FLAG macroH2A cell lines was prepared and dialyzed into BC100 buffer (20 mM Tris Cl pH 7.6, 2 mM EDTA, 100 mM KCl, 10% glycerol and 0.2 mM PMSF), incubated with M2 agarose beads and washed 3× with BC300. Flag peptide eluted complexes were run SDS-PAGE gels and colloidal blue stained. MacroH2A IP specific bands were sent for mass spectrometry. Proteins were digested and extracted peptides (Wilker et al., 2007) loaded on a precolumn and separated by reverse phase HPLC (Agilent) over a 75 min gradient before nanoelectrospray using Orbitrap XL mass spectrometer (Thermo). Raw mass spectral data files were processed as described (Johnson et al., 2012). Mascot peptide identifications were verified manually with the assistance of CAMV (Curran et al., 2013).

qRT-PCR, ChIP, and UV-RNA IP:

RNA isolation and RT-PCR were performed according to manufacturer's protocols (Invitrogen). ChIP and UV crosslinked RNA immunoprecipitation were performed as described previously (Jeon and Lee, 2011). Primers and antibodies are listed in Table 6.

Protein Expression and Purification:

Full length ATRX containing an N term FLAG tag and C term HA tag was cloned into a pfastbac vector (invitrogen). Sf9 cells were infected with virus for 60 h and protein was purified using Flag M2 agarose. The helicase domain of ATRX with an N term FLAG was cloned into the pdest10 vector (invitrogen). The ADD domain of ATRX was cloned into pet101 vector and purified via a C term His tag using Ni-NTA resin (Qiagen) according to manufacturer's instructions.

RNA and DNA EMSA and Double Filter Binding Assays:

EMSAs and filter binding assays were performed as described previously (Cifuentes-Rojas et al., 2014).

In Vitro Protein-RNA Interaction Assays: 2 μl g ATRX, 2 μg PRC2 and 1 μg Repeat A RNA were combined in buffer containing 50 mM Tris-HCl pH 8.0, 100 mM NaCl, 1.5 mM MgCl₂, 10 μg/ml BSA, 0.05% NP40, 1 mM DTT, 20 U RNaseOUT (Invitrogen™), 50 ng/μl yeast tRNA (Ambion® Cat # AM7119) and 1 mM ATP in a 20 μl reaction and incubated at 37° C. for 30 min. Reactions were added to Protein G coupled a-EZH2 at 4° C. for 2 h and washed with 1× binding buffer. Beads were split in 2: for RNA and protein analyses. RNA was eluted with formamide-urea buffer, loaded on a 6% Urea-PAGE and SYBR Gold stained. Proteins were eluted by boiling beads in SDS loading buffer.

For ternary complex IPs, 3 μg ATRX, PRC2 and Repeat A RNA were incubated at 30° C. in an RNA-EMSA binding reaction (see above) and precipitated using α-HA beads. Beads were washed with 1× binding buffer and complexes eluted with 1 μg/μl HA peptide. HA eluate was incubated with a-EZH2-protein G dynabeads. After washing, beads were split into 2 parts: for RNA and protein western blot. RNA was eluted with 0.2M glycine pH 2.0, neutralized with Tris.Cl pH 8.0 and end labeled in a PNK reaction for visualization.

200 ng of 5′ Biotin tagged Repeat A dsDNA was immobilized to Streptavidin Myone dynabeads. 25 nM ATRX and 35 nM RNA were combined in an EMSA reaction at 30° C. for 20 min, added to DNA and incubated for 20 min at 30° C. Beads were washed (as above) and nucleic acids were end labeled and resolved on an 18% Urea-PAGE gel.

Photocrosslinking:

In vitro transcription reactions were done as previously described (Hernandez et al., 2008) for 16 hrs at 37° C. with addition of 1 nM 4-Thio-UTP and ³²Pα-GTP and ³²Pα-CTP at 0.6 μM. Binding reactions were performed as described previously (Cifuentes-Rojas et al., 2014), irradiated for 12 min with 365-nm UV and digested with 0.1 ng RNase A, 1 U RNase T1 and 1 U RNase V1 for 20 min. SDS loading buffer was added and samples run on a 3-8% Tris-acetate SDS gradient gel.

ChIP-Seq Read Mapping:

ChIP-seq samples were sequenced using Illumina HiSeq2000 resulting in 25-50 million 50 bp paired-end reads per sample. Using ASAP tool (bioinformatics.babraham.ac.uk/projects/ASAP/), reads were aligned allele-specifically to CAST/EiJ and 129S1/SvJm genomes constructed using high quality polymorphisms to the C57/B16 reference genome (mm9). 25% of reads mapped uniquely and allele-specifically and 45% equally well to both genomes. Composite coverage tracks were generated using alignment against mm9 with ≤2 alternative mappings allowed. 80% of reads were aligned, with 10-15% of them discarded due to multiple alternate mappings. Input-normalized read densities were computed using SPP (Kharchenko et al., 2008) after PCR duplicate removal.

Peak Calling:

Regions of ATRX enrichment were determined based on tag counts in a 1 Kb window sliding with a 200 bp step. Statistical significance of enrichment of ChIP vs input was estimated using negative binomial distribution, with mean based on input tag count in input and size parameter (s) selected based on manual inspection of resulting peak calls. Regions of enrichment were defined by merging adjacent significantly enriched windows separated by ≤1 Kb.

Coverage Over Genomic Elements:

Coverage over TSS-proximal regions, gene bodies, or non-overlapping genome-wide 1 Kb bins was calculated from input-normalized SPP read densities. Actively expressed genes were identified using the cutoff of FPKM≥1.0 based on RNA-Seq data from (Yang et al., 2010). Input-normalized coverage over 1 Kb windows for Xist CHART and EZH2 ChIP-Seq was computed based on data from (Pinter et al., 2012; Simon et al., 2013). Significance of change in EZH2 and H3K27me3 densities produced by ATRX KD was assessed by t-test in separate quartiles of unique RefSeq genes ranked by WT ATRX density over gene body.

Because the macroH2A (mH2A) histone variant is enriched within gene-dense bands of the Xi together with Xist RNA and PRC2 (Chadwick and Willard, 2004; Costanzi and Pehrson, 1998), we performed an unbiased proteomics screen using mH2A as bait in an affinity purification. We expressed FLAG-tagged mH2A in 293, a human fibroblast cell line, and carried out FLAG immunoprecipitation followed by mass spectrometry (IP-MS). We observed several known interactors of mH2A, including PARP1 and linker histone H1 (FIG. 1A, left and middle panels; FIG. 8A)(Nusinow et al., 2007), as well as MECP2 and HP1. In addition, IP-MS revealed the 280-kD ATRX protein (FIG. 1A, 8B). We validated all interacting proteins by Western blot after FLAG-mH2A IP (FIG. 1A, right panel, and data not shown).

ATRX caught our attention, as it was shown to be enriched on the Xi by immunofluorescence (Baumann and De La Fuente, 2009) and has an ATPase and helicase domain (Clynes et al., 2013; Ratnakumar and Bernstein, 2013). ATRX is known as an unusual SNF2-like member of the SWI/SNF family of chromatin remodelers, as it appears to have only weak remodeling activity and does not affect nucleosome phasing in vitro. It does, however, have ATPase activity that is mildly stimulated by naked DNA and mononucleosomes (Tang et al., 2004; Xue et al., 2003), and a translocase activity that displaces a third strand of a DNA triplex (Mitson et al., 2011; Xue et al., 2003). The atypical N-terminal PHD finger domain of ATRX (a.k.a., ATRX-Dnmt3-Dnmt3L [ADD] domain) binds to unmodified histone H3-lysine 4 and to di- or tri-methylated H3-lysine 9 (Dhayalan et al., 2011; Eustermann et al., 2011; Iwase et al., 2011), in keeping with ATRX's role in maintaining pericentric heterochromatin and telomeres (Goldberg et al., 2010; Lewis et al., 2010). Mutations in ATRX cause X-linked mental retardation and alpha-thalassemia in humans, with ˜80% of mutations mapping to the PHD finger and helicase domains (Gibbons et al., 2008). Although ATRX appears enriched on the Xi and plays a role in peri-implantation development (Garrick et al., 2006), whether ATRX plays a role in XCI is unknown.

To investigate, we generated clones of mouse embryonic fibroblasts (MEF) with stable ATRX knockdown (KD) (shATRX-1, -2) and >90% depletion of ATRX protein (FIG. 1B). We examined localization of EZH2, the PRC2 subunit that catalyzes the trimethylation of histone H3 at lysine-27 (H3K27me3). Interestingly, immunofluorescence showed that, while ≈=60% of WT cells showed prominent EZH2 foci on the Xi, only 16-20% of ATRX KD cells retained EZH2 (FIG. 1B). Moreover, H3K27me3 on the X-chromosome was either absent or markedly reduced in shATRX cells. At the same time, RNA fluorescent in situ hybridization (FISH) showed that Xist RNA localized poorly, with quantitative RT-PCR showing that total Xist RNA levels were reduced 40-60% (FIG. 1C). Transient ATRX KDs in MEFs (48 h) showed that EZH2 localization was similarly compromised (FIG. 1D). In transient knockdowns, there was no obvious disturbance to Xist localization or H3K27me3, consistent with the short-term stability of Xist RNA and H3K27me3. Similar results were obtained using two independent ATRX shRNAs, whereas scrambled shRNA controls (Scr) resulted in no changes to Xist, EZH2, or H3K27me3 localization. Xist levels and localization were affected only in the long term, consistent with the notion that H3K27me3 at the Xist promoter targeted by RepA facilitates Xist upregulation (Sun et al., 2006; Zhao et al., 2008) and that the H3K27me3 mark is depleted only after several rounds of DNA replication. These findings hinted at a role of ATRX in recruiting and spreading PRC2 to form a “cloud” on the Xi.

To examine whether there were sex-specific phenotypes of ATRX deficiency, we used a mouse embryonic stem cell (ESC) model to recapitulate random XCI during ex vivo cell differentiation. We generated male and female ESC clones with stable ATRX KD (FIG. 2A,B; representative clones shown) and compared their behavior to shScr control clones in time-course analyses from differentiation days 0-10. While no differences were evident from days 0-4, sex-specific differences emerged between days 4-8 (FIG. 2C,D). shATRX female cells showed defective embryoid body (EB) outgrowth, whereas shScr control and male shATRX clones grew robustly (FIG. 2C). Immunostaining showed appropriate downregulation of pluripotency markers, such as NANOG, between days 0-8 in all clones (FIG. 9A, B), indicating that the defect was not caused by failed entry into the differentiation pathway per se. The Xist regulator, Tsix, was also appropriately downregulated in shATRX and shScr female ESC between days 0-10 (FIG. 2D). These results indicated a developmental defect downstream of the differentiation signal. In spite of normal Tsix downregulation, however, Xist RNA was not appropriately upregulated in shATRX cells. Furthermore, EZH2 targeting was severely compromised, as only 9% of shATRX cells (n=184) showed EZH2 enrichment (FIG. 2E) and only 17% of cells showed H3K27me3 enrichment (FIG. 2F,G) on the Xi, even though H3K27me3 levels were unperturbed globally (FIG. 2G). Therefore, ATRX depletion resulted in female-specific outgrowth defects related to an inability to initiate XCI.

To probe further, we circumvented a dependency of Xist expression on ATRX and used an inducible transgenic system in which Xist RNA is upregulated by addition of doxycycline (dox)(Jeon and Lee, 2011). In this “X+P” system, Xist is carried on an autosomal transgene in male cells and is upregulated >20-fold upon induction, accompanied by recruitment of PRC2 along the autosome in cis. In stable shATRX X+P clones (FIG. 2H; representative clone #4 shown), ATRX KD did not affect total cellular EZH2 levels (FIG. 2H) and dox-induction led to an appropriate increase of Xist expression (FIG. 2I), indicating successful bypass of shATRX's effect on Xist expression. Immuno-RNA FISH showed that EZH2 and H3K27me3 enrichment were observed on the Xist-coated chromosome in 66% and 74% of ATRX+ transgenic cells, respectively, confirming the sufficiency of ectopic Xist expression to recruit and spread EZH2 in cis (FIG. 2J,K). However, on ATRX KD, EZH2 and H3K27me3 recruitment decreased to 34% and 42%, respectively. Thus, ATRX is required for Xist RNA-mediated recruitment of PRC2.

To ask if Xist and ATRX directly interact in vivo, we performed UV-crosslink RNA immunoprecipitation (UV-RIP; FIG. 3A). As expected, in ATRX+ transgenic system (X+P), EZH2 pulled down Xist RNA under UV crosslinking conditions, consistent with a direct interaction between EZH2 and Xist RNA (Zhao et al., 2008). Intriguingly, ATRX also pulled down Xist RNA. This association was detectable only when cells were subjected to UV-crosslinking, suggesting that Xist is directly bound to ATRX. Neither EZH2 nor ATRX pulled down the nonspecific RNA control, U1 RNA. Significantly, when ATRX was knocked down (X+P shATRX), the ability of EZH2 to interact with Xist RNA was compromised (P=0.02; FIG. 3A). Similarly, EZH2-Xist RNA interactions were disrupted in wildtype (WT) female MEF cells when ATRX was depleted (P=0.007; FIG. 3B). These data demonstrate that ATRX-Xist interaction facilitates binding of PRC2 to Xist RNA in vivo.

To understand mechanism, we characterized ATRX-RNA interactions in vitro. PRC2 binds RepA/Xist via Repeat A (Zhao et al., 2008), a 435-nt motif consisting of 8.5 repeats of a 28-nucleotide sequence, with the potential to form several structures, including two long stem-loop structures, I-II and III-IV (FIG. 3C) (Duszczyk et al., 2011; Maenner et al., 2010; Wutz et al., 2002; Zhao et al., 2008). To ask how ATRX facilitates PRC2 interaction with Repeat A, we first performed RNA electromobility shift assays (EMSA) using purified full-length ATRX (FIG. 3D, left panels, 10B). ATRX robustly shifted Repeat A in a concentration-dependent manner (FIG. 3E), with ˜100% probe binding at <100 nM ATRX protein (FIG. 10C). To determine which region of Repeat A is sufficient for ATRX binding, we split Repeat A into I-II versus III-IV (FIG. 10A) and found that binding still occurred, albeit at lower levels. We calculated dissociation constants (K_d) from binding isotherms plotted from densitometric analysis of double-filter binding assays (FIG. 10D). Binding curves were fitted using a nonlinear regression model, with high R² values showing excellent fit of data points to the curve (FIG. 3F). ATRX strongly bound Repeat A I-IV, with a K_d of 5.41±0.40 nM. Subfragments I-II and III-IV retained binding with K_d's of 9.96±0.89 nM and 17.90±3.35 nM, respectively. To test the specificity of ATRX-RNA interactions, we used the first 300 nt of the Maltose Binding Protein (MBP) mRNA of Escherichia coli and the P4-P6 ribozyme of Tetrahymena, two non-cognate RNAs. Neither bound ATRX to an appreciable extent (K_d>>200 nM). We conclude that ATRX has high affinity and specificity for Repeat A RNA.

Interestingly, ATRX could also bind Repeat A double-stranded DNA (dsDNA; K_d=15.7)(FIG. 3G,H). However, it could not bind well to the single-stranded DNA (ssDNA), nor MBP and mouse Hotair dsDNAs (FIG. 3G,H; K_d>>200 nM). Therefore, ATRX has high affinity for both Repeat A RNA and dsDNA, and belongs to an emerging class of “bivalent” chromatin factors capable of binding both RNA and DNA [e.g., CTCF (Sun et al., 2013) and YY1 (Jeon and Lee, 2011)]. Competition analysis showed that 10-1000× molar excess of cold Repeat A dsDNA could not titrate away ATRX from Repeat A I-IV RNA probe (FIG. 10E) and, reciprocally, cold Repeat A I-IV RNA could not titrate ATRX away from the dsDNA probe (FIG. 10F), suggesting that ATRX interacts with RNA and dsDNA via distinct domains. To delineate binding domains, we generated subfragments of ATRX (FIG. 3D,I). The helicase domain could bind Repeat A RNA at 10-25 nM, but not so well as full-length ATRX. On the other hand, it bound dsDNA poorly, if at all (FIG. 3I). Gel shifts with the ADD domain showed binding to RNA at 100 nM, but not to DNA at the same concentration.

Because ATRX and PRC2 both bind Repeat A RNA, we asked whether ATRX and PRC2 could make direct contact to create an ATRX-RNA-PRC2 ternary complex. Using purified proteins, we observed that they do (FIG. 4A). This finding is consistent with a yeast two-hybrid screen in which EZH2 was identified as a directly interacting partner for ATRX (Cardoso et al., 1998), but which has not been verified by in vivo analyses. Notably, PRC2 components were not identified by our IP-MS analysis, though ATRX was a clear interacting partner (FIG. 1A). This apparent discrepancy may be due to substoichiometric ATRX-PRC2 associations in vivo and to mH2A being used as a bait in the proteomics screen. To confirm the ternary complex in vitro, we performed tandem IP, first pulling down ATRX via the HA tag, then immunoprecipitating PRC2 with an EZH2 antibody. Indeed, we observed end-labelled Repeat A RNA in the tandem IP (FIG. 4B), indicating that ATRX, PRC2, and Repeat A RNA formed a ternary complex.

Given pairwise interactions between ATRX-RNA and ATRX-DNA (FIG. 3E-I), we also explored the possibility of an ATRX-RNA-DNA ternary complex. We immobilized biotinylated Repeat A dsDNA on streptavidin beads and tested its ability to bind the ATRX-Repeat A RNA complex. We reasoned that, if the ATRX-RNA complex could engage dsDNA, RNA would be visualized after elution from the column. Under these conditions, RNA was minimally present, if at all, in the eluate (FIG. 4C). Thus, if an RNA:ATRX:dsDNA ternary complex could form, it may be too transient to detect in vitro. We conclude that ATRX, PRC2, and Repeat A RNA forms a stable ternary complex, but that ATRX likely does not form a stable ternary complex with RNA and dsDNA in vitro.

With ATRX possessing an ATPase domain, we examined how interaction dynamics might be changed by ATP. We assessed PRC2-RNA binding in the presence or absence of ATRX and/or ATP by immunoprecipitating EZH2 to quantitate the amount of RNA bound (FIG. 4D). Interestingly, ATRX's stimulatory effect was modestly increased (2-fold) in the presence of ATP, raising the possibility that ATRX may act by remodeling Repeat A to a configuration that promotes PRC2 binding. To test this idea, we carried out an RNAse protection assay with photocrosslinking. We incubated PRC2 with thiolated radiolabeled Repeat A RNA in the presence of ATRX and ATP or the non-hydrolysable analogue, AMP-PNP. The resulting complexes were subjected to UV crosslinking and treated with RNase A, T1, and V1. In principle, protein-bound radiolabeled RNA would be protected from digestion and detected in an SDS-PAGE autoradiograph. In five biological replicates, the presence of ATRX and ATP consistently resulted in greater protection of Repeat A RNA from RNAses, as visualized by the increased RNA signal bound to EZH2/SUZ12 (FIG. 4E), two known RNA-binding subunits of PRC2 (Cifuentes-Rojas et al., 2014). Together, these experiments show that ATRX stimulates binding of PRC2 to Repeat A RNA in a manner that is enhanced by ATP.

Intriguingly, ATRX bound RNA and DNA less well in the presence of ATP (FIG. 4F). However, when AMP-PNP was used, binding was not affected. This observation suggests that the nucleic acids are released by ATRX upon ATP hydrolysis. That is, ATRX may release RNA (and DNA) once the nucleic acid is reconfigured for PRC2 binding. The transient nature of the interactions may explain why a dsDNA-ATRX-RNA ternary complex was difficult to detect (FIG. 4C). The to ATRX:RNA:PRC2 ternary complex may be easier to detect (FIG. 4B) because it is held in place by both PRC2-RNA and PRC2-ATRX interactions. Combined, our data demonstrate that ATRX directly promotes binding of PRC2 to RepA/Xist RNA in vivo and in vitro.

We investigated genomic binding patterns for ATRX by performing chromatin immunoprecipitation with deep sequencing (ChIP-seq). To distinguish between Xa and Xi, we used genetically marked hybrid cell lines in which the Xi is invariably of Mus musculus (mus) origin and the Xa of Mus castaneus (cas) origin (Ogawa et al., 2008; Pinter et al., 2012). Analysis of two biological replicates revealed that, in MEFs and ESC, ATRX was broadly distributed along all chromosomes (Chr)—including ChrX and a representative autosome, Chr13—with preferential enrichment in gene-dense regions (FIG. 5A). Allele-specific analysis of ChIP-seq coverage density plots for ATRX, EZH2, and H3K27me3 (Pinter et al., 2012) and CHART-seq analysis of Xist RNA (Simon et al., 2013) revealed that the four epitopes were concentrated in the same regions of the Xi in female MEFs (FIG. 5B), supporting the idea that ATRX, PRC2, and Xist RNA are functionally linked. Strong ATRX binding occurred at ˜4987 sites, with 4% in promoter regions (transcription start site [TSS]±3 kb), 24% in coding gene bodies, and the rest in intergenic space (including enhancers, unannotated lncRNA transcription units)(FIG. 5C). This distribution is consistent with previous ChIP-seq analysis in male ESCs (Law et al., 2010).

Among thousands of ATRX sites on ChrX, one locus stood out (FIG. 5D-E, 11A-B). Xist was among the top genic hits in female MEFs (FIG. 5F) and was covered by ATRX along its entire length, contrasting with the restricted peaks characteristic of other loci (FIG. 11C). Allele-specific ChIP-seq of shATRX MEFs showed loss of enriched signals at Xist (FIG. 11D), confirming the specificity of the ATRX ChIP-seq analysis. ATRX enrichment at Xist was developmentally specific, being absent in pre-XCI ESC and present in post-XCI MEF. Furthermore, enrichment was observed only on the Xi (mus, FIG. 5E). However, ATRX coverage across Xi alleles was not enriched relative to the genomic average (FIG. 5G, representative Chr13 shown; KS test, P=1). Nor was ATRX coverage different on average between two classes of Xi genes—those expressed on Xa versus those that are silent (FIG. 5H; KS test, P=0.8031), indicating that ATRX was no more enriched on X-genes that are expressed in MEFs. Furthermore, ATRX coverage in genes that escape XCI was not significantly different between Xa and Xi (FIG. 5I, purple dots; KS test, P=0.75).

ChIP-qPCR confirmed that, in WT female MEFs, ATRX was significantly enriched (relative to IgG pulldowns) within the Xist gene regardless of what positions were queried (FIG. 5J). By contrast, Jarid1c—an X-linked gene that escapes XCI did not show significant ATRX enrichment. Enrichment at all Xist positions, including the nucleation center (uRF) was lost upon ATRX KD (FIG. 5J). ChIP-qPCR also verified the allele-specific nature of ATRX localization. The MEF line, Xa^WT Xi^2lox (clone 111.8), carried two loxP sites flanking a 16 kb Xist sequence on the Xi, whereas Xa^WT Xi^ΔXist (clone 111.20) carried a 16 kb conditional deletion of Xist on the Xi (Zhang et al., 2007). ATRX was significantly enriched at Xist only in the Xa^WT Xi^2lox, the cell line bearing an intact Xist gene on the Xi (FIG. 5J). Given that ATRX-RNA interactions are mediated by Repeat A RNA (FIG. 1-4), we asked whether the localization of ATRX to the chromatin hotspot depended on Repeat A DNA. In the Xist transgene model, ChIP-qPCR showed that, while deleting Repeat A (X-RA) did not affect spreading of Xist RNA along the chromosome as shown by RNA FISH (FIG. 5K, top panels)(Jeon and Lee, 2011), it abolished ATRX localization to the Xist locus (FIG. 5K, bottom graph). Repeat A is therefore essential for ATRX-Xist chromatin interactions in vivo.

We conclude that the Xi allele of Xist is a hotspot of ATRX binding. These data lead to a model in which ATRX binds the Repeat A motif and reconfigures the RepA/Xist transcript to promote PRC2 binding (FIG. 5L), with poising of ATRX at the Repeat A DNA potentially aiding ATRX-RNA interactions. Following nucleation at the Xist hotspot, the ATRX-RNA-PRC2 ternary complex spreads outwardly along the Xi.

We next investigated whether ATRX's effect on PRC2 localization extended beyond the Xi. We classified 21,677 annotated unique RefSeq genes into equal quartiles (Q1-Q4) based on ATRX coverage (FIG. 6A; Tables 4, 5), with Q1 having high positive ATRX coverage and Q2 moderate coverage. Interestingly, Q1 and Q2 harbored a large number of PRC2 target genes, together accounting for 63% of all autosomal PRC2 target genes. By contrast, Q3 and Q4 had near-zero or negative ATRX coverage and harbored fewer PRC2 target genes. To determine whether loss of ATRX affected PRC2 recruitment to genes marked by ATRX binding, we compared EZH2 and H3K27me3 coverages in ATRX+ versus ATRX-deficient cells (FIG. 6B). Indeed, ATRX KD affected PRC2 localization and function, with the 2- to 4-fold drop in EZH2 and H3K27me3 coverages being highly significant (FIG. 6B). Scatterplots showed an obvious shift in EZH2 and H3K27me3 coverage for Q1 PRC2 targets (FIG. 6C, purple dots; Table SI). We also examined probability density functions for EZH2 and H3K27me3 to determine the likelihood that a given gene in ATRX-deficient cells will have decreased EZH2 and H3K27me3 coverage based on the population behavior of ATRX-PRC2 target genes (FIG. 6D). The distribution for ATRX KD is left- and upwardly shifted, indicating that the probability of a target gene in ATRX KD having a low EZH2 (or H3K27me3) coverage is greater than that of the same gene in ATRX+. To examine how EZH2 and H3K27me3 patterns change within gene bodies, we performed metagene analysis (FIG. 6E). In ATRX+, EZH2 and H3K27me3 patterns showed characteristic gene-body enrichment. Upon ATRX KD, EZH2 and H3K27me3 densities dramatically decreased in Q1 and Q2 genes, with decrease occurring at TSS's and within gene bodies. Minimal changes were observed for Q3 and Q4 genes (FIG. 6E and data not shown). Analysis of biological replicates showed good reproducibility (FIG. 12A-B, data not shown). Collectively, these data demonstrate that ATRX is also required to target PRC2 to autosomal genes.

Given the loss of PRC2 from genic regions upon ATRX KD, we asked whether PRC2 became ectopically localized elsewhere. Examination of global EZH2 and H3K27me3 ChIP-seq patterns revealed an intriguing shift to intergenic space (FIG. 6F). In ATRX+, EZH2 was bound to 3178 sites, roughly distributed equally among gene bodies, promoters, and intergenic space. On the other hand, in ATRX KD, EZH2 shifted away from promoters and expanded into intergenic space. Thus, ATRX is required to specify PRC2 localization to genes in general.

Curiously, the distribution pattern for H3K27me3 peaks did not change in ATRX-depleted cells, as the percentages occurring in TSS, gene bodies, and intergenic space remained the same (FIG. 6F, purple pie charts). At the same time, bulk H3K27me3 levels also did not change on ATRX KD in MEFs and ES cells (FIG. 2G and data not shown). In spite of unchanged overall levels of H3K27me3 and genomic ChIP-seq density, there was an apparent loss of H3K27me3 peaks in many regions (FIG. 6F). Combined, these observations suggest a significant change in H3K27me3 patterns concurrent with mistargeting of PRC2. At the Polycomb target, Gpr26, loss of ATRX resulted in loss of EZH2 binding to the promoter and a relative reduction in H3K27me3 across the gene body (FIG. 6G). At Tmem200a, loss of EZH2 binding at the promoter was accompanied by a new EZH2-binding site ˜70 kb upstream in intergenic space. However, in spite of the newly acquired EZH2 site, H3K27me3 densities were not increased in the intergenic region. Altogether, these data demonstrate that loss of ATRX function causes improper PRC2 targeting, shifting bulk PRC2 to ectopic intergenic space where chaotic or unstable PRC2 accumulation results in decreased selectivity of H3K27 methylation. We conclude that ATRX is a specificity determinant for PRC2 localization and function.

To determine whether failure of PRC2 targeting results in gene expression changes, we tested specific Polycomb targets (FIG. 7A,B; 13A,B). Within the Hoxc and Hoxd clusters, EZH2 and H3K27me3 were highly enriched in ATRX+ MEFs. Upon ATRX KD, EZH2 and H3K27me3 levels were significantly reduced, as evidenced by both decreased peak numbers as well as overall coverages across the locus. Quantitative RT-PCR analysis revealed concomitant gene upregulation (FIG. 7C, 13C). At Parp12, Rnf19b, and Spock2, EZH2 bound the TSS region in ATRX+ cells (FIG. 7B). On ATRX KD, EZH2 and H3K27me3 were consistently lost at the promoters (FIG. 7B,D; biological replicate shown in FIG. 13D) and gene expression increased 2- to 5-fold at all loci (FIG. 7C). In contrast, expression of Zc3 h11a—a control that is not a Polycomb target—did not change significantly after ATRX KD. We conclude that ATRX plays a global role in gene regulation by controlling PRC2 localization.

The ATRX peaks in the mm9 mouse assembly coordinates were converted to human coordinates using Galaxy Server's LiftOver utility (The liftOver utility effectively maps one genome to another, allowing rapid identification of regions of interest between successive assemblies of the same species or between two distinct species; available online at usegalaxy.org). This tool is based in the Liftover utility from the UC Santa Cruz Genome Browser. Mouse-to-human LiftOver of the mouse chromosome coordinates and strand of these mouse Peaks was performed in the Galaxy server to generate orthologous human chromosome coordinates. The parameters used were “0.1 minimum ratio of bases that must remap” from mm9 to Hg19 as is the default for the program when performing a Liftover between different species. This process and LiftOver chains are generally described in Kent et al., Proc. Nat'l Acad. Sci., 100(20) 11484-11489 (2003).