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Selective silencing of euchromatic L1s revealed by genome-wide screens for L1 regulators

Nature volume 553, pages 228–232 (2018)Cite this article

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Abstract

Transposable elements, also known as transposons, are now recognized not only as parasitic DNA, the spread of which in the genome must be controlled by the host, but also as major players in genome evolution and regulation1,2,3,4,5,6. Long interspersed element-1 (LINE-1, also known as L1), the only currently autonomous mobile transposon in humans, occupies 17% of the genome and generates inter- and intra-individual genetic variation, in some cases resulting in disease1,2,3,4,5,6,7. However, how L1 activity is controlled and the function of L1s in host gene regulation are not completely understood. Here we use CRISPR–Cas9 screening strategies in two distinct human cell lines to provide a genome-wide survey of genes involved in the control of L1 retrotransposition. We identify functionally diverse genes that either promote or restrict L1 retrotransposition. These genes, which are often associated with human diseases, control the L1 life cycle at the transcriptional or the post-transcriptional level in a manner that can depend on the endogenous L1 nucleotide sequence, underscoring the complexity of L1 regulation. We further investigate the restriction of L1 by the protein MORC2 and by the human silencing hub (HUSH) complex subunits MPP8 and TASOR8. HUSH and MORC2 can selectively bind evolutionarily young, full-length L1s located within transcriptionally permissive euchromatic environments, and promote deposition of histone H3 Lys9 trimethylation (H3K9me3) for transcriptional silencing. Notably, these silencing events often occur within introns of transcriptionally active genes, and lead to the downregulation of host gene expression in a HUSH-, MORC2-, and L1-dependent manner. Together, these results provide a rich resource for studies of L1 retrotransposition, elucidate a novel L1 restriction pathway and illustrate how epigenetic silencing of transposable elements rewires host gene expression programs.

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Figure 1: Genome-wide screen for L1 activators and suppressors in K562 cells.
Figure 2: HUSH and MORC2 silence L1 transcription to inhibit retrotransposition.
Figure 3: HUSH and MORC2 target young full-length L1s in euchromatic environments.
Figure 4: HUSH or MORC2 binding at L1s decreases active host gene expression.

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References

  1. Lander, E. S. et al. Initial sequencing and analysis of the human genome. Nature 409, 860–921 (2001)

    Article  ADS  CAS  PubMed  Google Scholar 

  2. Levin, H. L. & Moran, J. V. Dynamic interactions between transposable elements and their hosts. Nat. Rev. Genet. 12, 615–627 (2011)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Beck, C. R., Garcia-Perez, J. L., Badge, R. M. & Moran, J. V. LINE-1 elements in structural variation and disease. Annu. Rev. Genomics Hum. Genet. 12, 187–215 (2011)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. Mita, P. & Boeke, J. D. How retrotransposons shape genome regulation. Curr. Opin. Genet. Dev. 37, 90–100 (2016)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  5. Goodier, J. L. Restricting retrotransposons: a review. Mob. DNA 7, 16 (2016)

    Article  PubMed  PubMed Central  Google Scholar 

  6. Chuong, E. B., Elde, N. C. & Feschotte, C. Regulatory activities of transposable elements: from conflicts to benefits. Nat. Rev. Genet. 18, 71–86 (2017)

    Article  CAS  PubMed  Google Scholar 

  7. Philippe, C. et al. Activation of individual L1 retrotransposon instances is restricted to cell-type dependent permissive loci. eLife 5, e13926 (2016)

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  8. Tchasovnikarova, I. A. et al. Epigenetic silencing by the HUSH complex mediates position–effect variegation in human cells. Science 348, 1481–1485 (2015)

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  9. Moran, J. V. et al. High frequency retrotransposition in cultured mammalian cells. Cell 87, 917–927 (1996)

    Article  CAS  PubMed  Google Scholar 

  10. Morgens, D. W. et al. Genome-scale measurement of off-target activity using Cas9 toxicity in high-throughput screens. Nat. Commun. 8, 15178 (2017)

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  11. Morgens, D. W., Deans, R. M., Li, A. & Bassik, M. C. Systematic comparison of CRISPR/Cas9 and RNAi screens for essential genes. Nat. Biotechnol. 34, 634–636 (2016)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Suzuki, J. et al. Genetic evidence that the non-homologous end-joining repair pathway is involved in LINE retrotransposition. PLoS Genet. 5, e1000461 (2009)

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  13. Chance, P. F. et al. Linkage of the gene for an autosomal dominant form of juvenile amyotrophic lateral sclerosis to chromosome 9q34. Am. J. Hum. Genet. 62, 633–640 (1998)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Németh, A. H. et al. Autosomal recessive cerebellar ataxia with oculomotor apraxia (ataxia-telangiectasia-like syndrome) is linked to chromosome 9q34. Am. J. Hum. Genet. 67, 1320–1326 (2000)

    PubMed  PubMed Central  Google Scholar 

  15. Albulym, O. M. et al. MORC2 mutations cause axonal Charcot–Marie–Tooth disease with pyramidal signs. Ann. Neurol. 79, 419–427 (2016)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. Schottmann, G., Wagner, C., Seifert, F., Stenzel, W. & Schuelke, M. MORC2 mutation causes severe spinal muscular atrophy-phenotype, cerebellar atrophy, and diaphragmatic paralysis. Brain 139, 1–4 (2016)

    Article  Google Scholar 

  17. Brégnard, C. et al. Upregulated LINE-1 activity in the Fanconi anemia cancer susceptibility syndrome leads to spontaneous pro-inflammatory cytokine production. EBioMedicine 8, 184–194 (2016)

    Article  PubMed  PubMed Central  Google Scholar 

  18. Ostertag, E. M., Prak, E. T., DeBerardinis, R. J., Moran, J. V. & Kazazian, H. H. Jr. Determination of L1 retrotransposition kinetics in cultured cells. Nucleic Acids Res. 28, 1418–1423 (2000)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Han, J. S. & Boeke, J. D. A highly active synthetic mammalian retrotransposon. Nature 429, 314–318 (2004)

    Article  ADS  CAS  PubMed  Google Scholar 

  20. Wagstaff, B. J., Barnerssoi, M. & Roy-Engel, A. M. Evolutionary conservation of the functional modularity of primate and murine LINE-1 elements. PLoS ONE 6, e19672 (2011)

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  21. Tchasovnikarova, I. A. et al. Hyperactivation of HUSH complex function by Charcot–Marie–Tooth disease mutation in MORC2. Nat. Genet. 49, 1035–1044 (2017)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  22. Moissiard, G. et al. MORC family ATPases required for heterochromatin condensation and gene silencing. Science 336, 1448–1451 (2012)

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  23. Pastor, W. A. et al. MORC1 represses transposable elements in the mouse male germline. Nat. Commun. 5, 5795 (2014)

    Article  ADS  CAS  PubMed  Google Scholar 

  24. Garcia-Perez, J. L. et al. LINE-1 retrotransposition in human embryonic stem cells. Hum. Mol. Genet. 16, 1569–1577 (2007)

    Article  CAS  PubMed  Google Scholar 

  25. Han, J. S., Szak, S. T. & Boeke, J. D. Transcriptional disruption by the L1 retrotransposon and implications for mammalian transcriptomes. Nature 429, 268–274 (2004)

    Article  ADS  CAS  PubMed  Google Scholar 

  26. Saint-André, V., Batsché, E., Rachez, C. & Muchardt, C. Histone H3 lysine 9 trimethylation and HP1γ favor inclusion of alternative exons. Nat. Struct. Mol. Biol. 18, 337–344 (2011)

    Article  PubMed  CAS  Google Scholar 

  27. Khan, H., Smit, A. & Boissinot, S. Molecular evolution and tempo of amplification of human LINE-1 retrotransposons since the origin of primates. Genome Res. 16, 78–87 (2006)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  28. Buecker, C. et al. Reorganization of enhancer patterns in transition from naive to primed pluripotency. Cell Stem Cell 14, 838–853 (2014)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  29. Taylor, M. S. et al. Affinity proteomics reveals human host factors implicated in discrete stages of LINE-1 retrotransposition. Cell 155, 1034–1048 (2013)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Brouha, B. et al. Evidence consistent with human L1 retrotransposition in maternal meiosis I. Am. J. Hum. Genet. 71, 327–336 (2002)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. Gasior, S. L., Roy-Engel, A. M. & Deininger, P. L. ERCC1/XPF limits L1 retrotransposition. DNA Repair (Amst.) 7, 983–989 (2008)

    Article  CAS  Google Scholar 

  32. Deans, R. M. et al. Parallel shRNA and CRISPR–Cas9 screens enable antiviral drug target identification. Nat. Chem. Biol. 12, 361–366 (2016)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  33. Bassik, M. C. et al. A systematic mammalian genetic interaction map reveals pathways underlying ricin susceptibility. Cell 152, 909–922 (2013)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Cong, L. et al. Multiplex genome engineering using CRISPR/Cas systems. Science 339, 819–823 (2013)

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  35. Coufal, N. G. et al. L1 retrotransposition in human neural progenitor cells. Nature 460, 1127–1131 (2009)

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  36. Shukla, R. et al. Endogenous retrotransposition activates oncogenic pathways in hepatocellular carcinoma. Cell 153, 101–111 (2013)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Carreira, P. E. et al. Evidence for L1-associated DNA rearrangements and negligible L1 retrotransposition in glioblastoma multiforme. Mob. DNA 7, 21–34 (2016)

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  38. Doucet, A. J., Wilusz, J. E., Miyoshi, T., Liu, Y. & Moran, J. V. A. A 3′ poly(A) tract is required for LINE-1 retrotransposition. Mol. Cell 60, 728–741 (2015)

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Otsu, N. A threshold selection method from gray-level histograms. IEEE Trans. Syst. Man Cybern. 9, 62–66 (1979)

    Article  Google Scholar 

  40. Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550 (2014)

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  41. Bajpai, R. et al. CHD7 cooperates with PBAF to control multipotent neural crest formation. Nature 463, 958–962 (2010)

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  42. Rada-Iglesias, A. et al. A unique chromatin signature uncovers early developmental enhancers in humans. Nature 470, 279–283 (2011)

    Article  ADS  CAS  PubMed  Google Scholar 

Download references

Acknowledgements

We thank J. Moran for the LRE-GFP plasmid and A. Engel for the codon-optimized L1 construct; D. Fuentes, A. Spencley, R. Srinivasan, J. Mohammed, V. Bajpai, K. Tsui, G. Hess, D. Morgens and G. Cornelis for assistance and discussions; K. Cimprich, A. Fire and A. Urban for comments on the manuscript; and L. Bruhn, S. Altschuler, B. Borgo, P. Sheffield and C. Carstens (Agilent) for discussions and oligonucleotide synthesis. This work was funded by grants from the Jane Coffin Childs Memorial Fund for Medical Research (N.L.), National Science Foundation DGE-114747 (C.H.L.), National Institutes of Health (NIH) R01HG008150, 1UM1HG009436-01 and NIH 1DP2HD084069-01 (M.C.B.), NIH R01 GM112720, Stinehart Reed Award and Howard Hughes Medical Institute (J.W.).

Author information

Author notes

  1. Edward Grow

    Present address: Huntsman Cancer Institute, University of Utah, Salt Lake City, Utah, 84112-5550, USA

  2. Nian Liu and Cameron H. Lee: These authors contributed equally to this work.

Authors and Affiliations

  1. Department of Chemical and Systems Biology, Stanford School of Medicine, Stanford University, Stanford, 94305, California, USA

    Nian Liu, Tomek Swigut, Bo Gu & Joanna Wysocka

  2. Department of Genetics, Stanford School of Medicine, Stanford University, Stanford, 94305, California, USA

    Cameron H. Lee, Edward Grow & Michael C. Bassik

  3. Stanford University Chemistry, Engineering, and Medicine for Human Health (ChEM-H), Stanford School of Medicine, Stanford University, Stanford, 94305, California, USA

    Michael C. Bassik

  4. Institute of Stem Cell Biology and Regenerative Medicine, Stanford School of Medicine, Stanford University, Stanford, 94305, California, USA

    Joanna Wysocka

  5. Department of Developmental Biology, Stanford School of Medicine, Stanford University, Stanford, 94305, California, USA

    Joanna Wysocka

  6. Howard Hughes Medical Institute, Stanford School of Medicine, Stanford University, Stanford, 94305, California, USA

    Joanna Wysocka

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Contributions

N.L., C. H.L., T.S., J.W. and M.C.B. designed and performed experiments, analysed data and wrote the manuscript. E.G., C.H.L., J.W. and M.C.B. initiated the K562 genome-wide screen. B.G. analysed smFISH data. J.W. and M.C.B. supervised the study.

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Correspondence to Michael C. Bassik or Joanna Wysocka.

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Reviewer Information Nature thanks D. Bourc’his and the other anonymous reviewer(s) for their contribution to the peer review of this work.

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Extended data figures and tables

Extended Data Figure 1 Genome-wide CRISPR–Cas9 screen for L1 regulators in K562 cells.

a, Schematic of L1-G418R and L1–GFP reporters used in this work. b, PCR assay on genomic DNA using primers that flank the engineered intron within the G418R cassette. Two experiments repeated independently with similar results. The spliced PCR bands were not observed before dox induction in either K562 or HeLa cells, suggesting that the L1-G418R reporter was not activated before the screening. However, there may exist an extremely low level of reporter leakiness that is below the detection limits of the PCR assay. c. Fluorescence-activated cell sorting experiments show that the L1–GFP cells have no GFP signals without dox induction (0 out of approximately 300,000 cells), and begin to produce GFP after dox induction. Therefore, the level of reporter leakiness without dox induction is insignificant. Two experiments repeated independently with similar results. d. CasTLE analysis of genome-wide screens in K562 cells, with 20,488 genes represented as individual points. Genes falling under 10% FDR coloured in blue, analysed by CasTLE likelihood ratio test11. n = 2 biologically independent screens. e. HeLa cells with L1-G418R are resistant to G418 after dox induction (seven days of dox induction followed by ten days of G418 selection). Live cells in equal volumes were counted in a single (n = 1) fluorescence-activated cell sorting experiment. The centre value indicates the total number of live cells. Error bar, square root of total events assuming Poisson distribution of counts.

Extended Data Figure 2 A secondary screen identifies functionally diverse L1 regulators in K562 cells.

a, The reproducibility between two independent secondary screens (n = 2) in K562 cells. R-squared value is from a linear regression model. b. The K562 secondary screen recovers more sgRNAs than the K562 genome-wide screen, suggesting a higher detection sensitivity in the secondary screen. c, Comparison of the secondary screen data (252 genes from n = 2 independent screens) with the genome-wide screen data (n = 2 independent screens) in K562 cells. The R-squared value is from a linear regression model. d, Volcano plot showing K562 secondary screen results (252 genes from two independent screens), with genes previously implicated in L1 biology coloured in red. e, Classification-diverse L1 activators and suppressors identified in K562 cells by their known biological processes. f, The maximum effect size (centre value) of indicated DNA repair genes, estimated by CasTLE from two independent K562 secondary screens with ten different sgRNAs per gene. Error bars, 95% credible intervals of the estimated effect size.

Extended Data Figure 3 Screen for L1 regulators in HeLa cells and and L1- sequence-dependent L1 regulators.

a, CasTLE analysis of two independent genome-wide screens in HeLa cells, with 20,514 genes represented as individual points. Genes at 10% FDR cutoff coloured in red, analysed by CasTLE likelihood ratio test11. b, The maximum effect size (centre value) estimated by CasTLE from two independent HeLa secondary screens with ten different sgRNAs per gene. Bars, 95% credible interval. L1 activators are shown in red; L1 suppressors are shown in blue. Genes for which the credible interval includes zero are coloured in grey and are considered non-effective against L1. c, Scatter plots showing the secondary screen hits identified in K562 cells and HeLa cells (252 genes from two independent screens in each cell line), with a Venn diagram comparing hits in the two cell lines shown on the right. d, The maximum effect size (centre value) of indicated heterochromatin regulators, estimated by CasTLE from two independent HeLa secondary screens with ten different sgRNAs per gene. Error bars, 95% credible intervals of the estimated effect size. e, The maximum effect size (centre value) of indicated DNA repair genes, estimated by CasTLE from two independent HeLa secondary screens with ten different sgRNAs per gene. Error bars, 95% credible intervals of the estimated effect size. f, The (opt)-L1–GFP reporter retrotransposed more frequently than did L1–GFP in K562. The GFP+ fraction of cells with the indicated L1 reporter after 15 days of dox induction was normalized to the L1–GFP sample. Box plots show median and IQR, whiskers are 1.5 × IQR. n = 6 biologically independent replicates. g, The GFP+ fraction of dox-induced control and mutant cell pools with the L1–GFP reporter or (opt)-L1–GFP reporter. Experiments were performed as in Fig. 1e. Chromatin regulators (for example, TASOR, MORC2, MPP8 and SAFB) did not suppress the (opt)-L1–GFP reporter, in which 24% of the L1 ORF nucleotide sequence is altered, without changes in the encoded amino acid sequence19,20, indicating that their L1 regulation depends on the native nucleotide sequence of L1Hs. h, K562 secondary screen with the (opt)-L1-G418R reporter (252 genes from n = 2 independent screens) revealed genes that regulate retrotransposition dependent or non-dependent on the native L1 nucleotide sequence. The K562 secondary screen candidates identified with L1-G418R (252 genes from n = 2 independent screens) are labelled in blue. A Venn diagram comparing hits identified from the two L1 reporters is shown on the top right.

Extended Data Figure 4 MORC2, MPP8 and TASOR silence L1 transcription.

a, Relative genomic copy number of newly integrated L1–GFP reporters in the indicated mutant K562 pools after dox induction. The PspGI-assisted qPCR assay used here was designed to selectively detect spliced GFP rather than the unspliced version (see Methods). The L1–GFP copies were normalized to β-actin DNAs; data were then normalized to the control. As a putative L1 activator, SLTM shows an opposite effect on the DNA copy number compared with L1 suppressors. Centre value is the median; n = 3 technical replicates per gene. b, RNA-seq data in control K562 cells show that most heterochromatin regulators in Fig. 2a are expressed, supporting the selective effect of HUSH and MORC2 in L1 regulation. c, Western blots validating the effects of knockout in independent knockout K562 cell clones. Control samples were loaded at four different amounts (200%, 100%, 50% and 25% of knockout clones). Three experiments, repeated independently with similar results. To obtain knockout clones, we sorted pools of mutant K562 cells (those used in Fig. 1e, f) into 96-well plates, expanded the cells and screened for knockout clones through western blotting. Of note is that all K562 knockout clones were derived from the same starting L1–GFP reporter line, and therefore do not differ in reporter transgene integrations among the clones. d, Representative images of smFISH assays targeting ACTB mRNAs and RNA transcripts from L1–GFP reporters in control and knockout K562 clones after five days of dox induction. No signal was observed from L1–GFP reporters without dox induction (data not shown). Two experiments repeated independently with similar results. See also e and Fig. 2b (showing L1–GFP mRNA only). e, Quantification of the L1–GFP transcription level from the indicated number of K562 cells, determined by smFISH assays (d and Fig. 2b). The number of L1–GFP mRNA transcripts is normalized to the number of β-actin mRNAs within each K562 cell. Box plots show median and IQR, whiskers are 1.5 × IQR. P value, two-sided Wilcoxon test. 95% credible interval for median from 1,000× bootstrap: control: 0.059–0.082; MORC2: 0.106–0.123; MPP8: 0.264–0.410; TASOR: 0.514–0.671. f, MORC2, MPP8 and TASOR knockouts increase the genomic copy number of newly integrated L1–GFP reporters. PspGI-assisted qPCR assays were performed as in a, but using clonal knockout K562 clones instead of mutant cell pools. Data are normalized to the control. n = 3 technical replicates; centre value is median. g, MORC2 knockout, MPP8 knockout, and TASOR knockout increase the expression of endogenous L1s. RT–qPCR experiments were performed as in Fig. 1f, but using clonal knockout K562 clones instead of mutant cell pools. n = 2 biological replicates × 3 technical replicates; centre value as median. The primers do not target the L1–GFP reporter and the cell lines were not dox-induced, so these RT–qPCR assays will not detect L1–GFP transcripts. h, Western blots showing the effects of depletion of MORC2, MPP8 and TASOR in the mutant pools of K562 cells (left) and in the mutant pools of H9 hES cells without transgenic L1 reporters (right), independently repeated twice with similar results. i, Northern blots showing increased transcription from the L1–GFP reporter in knockout K562 clones (same cell lines as in c) after five days of dox induction, independently repeated twice with similar results. As observed in Fig. 2b, whereas HUSH knockout significantly increases L1–GFP transcription, MORC2 knockout leads to only a modest increase. This is probably because the L1–GFP reporter does not contain the native L1 5′ UTR sequence, to which MORC2 binds strongly (see Extended Data Fig. 6f, g). The 5-kb and 1.9-kb marks on the membrane refer to the 28S rRNA and 18S rRNA bands, respectively. j, Northern blots showing that disruption of MORC2, MPP8 and TASOR increases the expression level of endogenous L1Hs in hES cells, using the same cell lines as in h. Size marker indicated as in i. Independently repeated twice with similar results. k, Western blots showing protein abundance of L1_ORF1p and HSP90 in the mutant pools of K562 cells and hES cells (same cell line as shown in h), independently repeated twice with similar results. Experiments were performed without dox induction of the transgenic L1 reporter. Owing to the strong signal of bands from the knockout samples, the blots were exposed for a very short time and the band signals in the control samples were relatively very weak compared to the knockout samples; this is was also the case in i, j.

Extended Data Figure 5 The binding profiles of MORC2, MPP8 and TASOR revealed by ChIP–seq in K562 cells.

a, Using a paired-end sequencing strategy for ChIP–seq, together with the sequence divergence within native L1 elements, we could map ChIP–seq reads to individual L1 instances in the genome. Genome browser snapshots of MORC2 ChIP–seq reads alignment over L1PA7 (left) and L1Hs (right). The experiment was repeated once with similar results. The colour scale indicates the mapping quality score MAPQ for each read pair. MAPQ = 10log10p, where p is the probability that true alignment belongs elsewhere. With the exception of L1Hs, which is the youngest and least sequence-divergent family, the bodies of L1 repeats are uniquely mappable. In case of L1Hs, the 5′ UTR is still mappable to determine the level of L1Hs in control and knockout clones. b, Genome browser snapshots for MPP8 (blue), TASOR (orange) and MORC2 (purple) ChIP–seq read densities from control and corresponding knockout K562 clones at two representative genomic loci. The experiment was repeated once with similar results. LINE element occurrences are indicated by blue rectangles at the bottom of the plot. Four instances of long L1 elements are named, indicating the L1 families to which they belong. Note the complete absence of ChIP–seq signals from knockout lines, and selectivity towards some but not other L1 instances. Of note is that, whereas MPP8 and MORC2 ChIP signals were robust, TASOR ChIPs showed relatively weak enrichments (either owing to poor antibody quality or genuine biological properties); for this reason, a subset of our downstream analyses is focused on MORC2 and MPP8. c, In addition to full-length L1, the HUSH complex and MORC2 bind 3′ UTR of KRAB zinc-finger (KRAB-ZNF) genes. Genome browser snapshots of ChIP–seq read densities over representative examples, from both control and corresponding knockout K562 clones. The experiment was repeated once with similar results. d, The HUSH complex and MORC2 preferentially bind expressed KRAB-ZNF genes over other ZNF genes. Heat maps of MPP8 (left) and MORC2 (centre) signals over 2,600 ZNF genes, centred at the 3′ end of the genes and sorted first by the presence of the KRAB domain and then by the MPP8 ChIP signal. The upper 1,600 genes are KRAB-ZNF genes, the lower 1,000 genes are non-KRAB ZNF genes. The heat maps on the right code the absolute expression level of each gene in RPKM scale from the K562 RNA-seq data (far right).

Extended Data Figure 6 HUSH and MORC2 preferentially bind full-length L1 instances in human ES cells, mouse ES cells and K562 cells.

a, Widespread genomic co-binding of MPP8 and MORC2 in hES cells. Heat map representation of ChIP–seq results at 57,000 genomic loci, centred on MPP8 and MORC2 summits and sorted by MORC2 ChIP–seq signal. Plotted is the normalized ChIP read density from hES cells. b, Heat maps of MORC2/MPP8 ChIP–seq density over the indicated repeat classes, centred and sorted as in a. The HUSH complex and MORC2 bind predominantly to L1 elements in hES cells, in particular to the primate-specific L1P families, suggesting that HUSH- and MORC2-dependent silencing is relevant in many embryonic and somatic cell types. c, L1 families that encompass active L1 copies, such as L1Md-T and L1Md-A, are significantly enriched among MPP8 binding sites in mouse ES cells. L1Md_Gf is also enriched, but not shown owing to the low number of instances. Thus, HUSH-mediated L1 regulation appears to be conserved among species. Of note is that MPP8 is also strongly enriched at IAP elements, a class of murine endogenous retroviruses that remain currently mobile in the mouse genome. d, MPP8 ChIP–seq heat maps in mouse embryonic stem cells featuring retrotransposition-competent L1Md-T, L1Md-A and L1Md-Gf. e, MPP8 preferentially binds full-length L1Md-A and L1Md-T in mouse ES cells. Plotted is the size distribution of the indicated L1 instances that overlap with MPP8 ChIP–seq peaks, or remaining L1s that do not overlap with such ChIP–seq signals. Box plots show median and IQR, whiskers are 1.5 × IQR. f, Aggregate plots of MORC2 (red) and MPP8 (black) ChIP–seq signals over 500 full-length, MPP8-bound L1PAs, centred on the L1 5′ end. g, Aggregate plots of MORC2 (red) and MPP8 (black) ChIP–seq signals on L1Hs (L1PA1). Similar to the binding profile on L1PA (f), MPP8 and MORC2 occupy the whole body of L1Hs, with MORC2 additionally binding L1Hs 5′ UTR. It is to be noted that ChIP–seq fragments are much less likely to be uniquely mapped—and thus removed by the alignment criteria—within the L1Hs non-5′ UTR region, owing to their minimal sequence divergence (Extended Data Fig. 5a).

Extended Data Figure 7 HUSH and MORC2 collaborate in binding target L1s.

a, A representative genome browser view of normalized ChIP–seq read densities over L1 elements. The experiment was repeated once with similar results. The loss of MPP8 and TASOR results in no detectable binding by MORC2, MPP8 and TASOR, whereas the loss of MORC2 results in partially diminished recruitment of HUSH complex subunits. b, Heat maps of MPP8 (left), TASOR (centre) and MORC2 (right) ChIP–seq signals subtracted for the ChIP signal from corresponding knockout lines. Heat maps are centred on MPP8 and MORC2 peaks, separated by the presence or absence of underlying L1 and then sorted by MPP8 ChIP signal strength. The loss of MORC2 has only a partial effect on the recruitment of MPP8 and TASOR to the L1 elements, whereas the loss of either MPP8 or TASOR abrogates MORC2 recruitment.

Extended Data Figure 8 HUSH and MORC2 preferentially bind intronic L1s within actively transcribed genes.

a. Genes that contain MPP8- or MORC2-bound intronic L1s are expressed at significantly higher levels in control K562 cells, compared to genes that contain intronic full-length L1s unbound by MPP8 or MORC2. P value, two-sided Mann–Whitney-Wilcoxon test. Box plots show median and IQR, whiskers are 1.5 × IQR. b. The promoters of genes that contain MPP8- or MORC2-bound intronic full-length L1s are marked by transcriptionally permissive H3K27ac in wild-type K562 cells. H3K27ac ChIP–seq data are taken from a K562 epigenome pilot study, accession number PRJEB8620. TSS, transcription start site. c, Genes selectively occupied by MORC2 or MPP8 either in K562 or in hES cells exhibit higher gene expression in the corresponding cell line (P = 4.3 × 10−107 for MPP8 binding; 5.0 × 10−92 for MORC2 binding, Kruskal–Wallis test). Boxplots defined as in a. RNA-seq datasets for hES cells are from SRA entries SRR2043329 and SRR2043330. d, ChIP–qPCR assays quantifying HUSH and MORC2 binding to an inducible L1 transgene in K562 cells before or after its transcriptional induction by dox. Transcriptional induction increases binding of MORC2 and MPP8 to the L1 transgene. n = 2 biological replicates × 3 technical replicates; centre value is median.

Extended Data Figure 9 HUSH and MORC2 facilitate H3K9me3 at their L1 targets for transcription repression.

a, A concordant subset (approximately 1%) of 111,499 H3K9me3 sites in the genome lose the H3K9me3 signal in MORC2 knockout, MPP8 knockout and TASOR knockout K562 clones. Two independent lines each for wild-type, MORC2 knockout, TASOR knockout or MPP8 knockout clone. Plotted is the log2 fold change in the H3K9me3 ChIP signal in the TASOR knockout relative to the control (x axis) and the log2 fold change in the H3K9me3 ChIP signal in the MORC2 knockout relative to the control (y axis). Points are colour-coded with blue sites having significant H3K9me3 loss in MPP8 knockout, red sites significantly gaining a signal in MPP8 knockout, and grey sites having no detectable change. Sites that significantly lose the H3K9me3 signal in a knockout line are more likely to show a corresponding loss in other knockout lines. Odds ratios: 26.23 with 95% confidence intervals (23,66, 29.10) for MORC2 versus MPP8; 21.70 with 95% confidence intervals (19.75, 23.83) for TASOR versus MPP8; 122.53 with 95% confidence intervals (109.21, 137.43) for TASOR versus MORC2. P = 0 each case, two-sided Fisher's exact test. b, Genomic sites that exhibit the strongest loss of H3K9me3 in MORC2, MPP8 or TASOR knockouts are preferentially L1-occupied by MORC2, MPP8 or TASOR respectively. Boxplots of log2 fold change in H3K9me3 relative to control for MPP8 knockout (left), MORC2 knockout (centre) and TASOR knockout (right). Box plots show median and IQR, whiskers are 1.5 × IQR. MPP8- and MORC2-bound L1s show a significant loss of H3K9me3 (P values, two-sided Mann–Whitney–Wilcoxon test). c, Averaged distribution of H3K9me3 ChIP–seq signals in control and knockout K562 clones over the host genes that contain the MORC2-targeted intronic full-length L1s, centred on the TSS of the host genes. d, Genome browser track showing MORC2 binding at the intronic full-length L1Hs within CDH8 in both K562 and hES cells. The experiment was repeated once with similar results. e, Genome browser track showing MORC2 binding at the intronic full-length L1PA2 within DNAH3 in both K562 and hES cells. The experiment was repeated once with similar results. f, Depletion of MORC2 or HUSH increases the expression of CDH8 in both K562 (n = 2 biological replicates × 3 technical replicates) and hES cells (n = 3 technical replicates), as measured by RT–qPCR assay. The CDH8 expression level was normalized to β-actin mRNA. All samples were then normalized to the control sample. Centre value is median. g, Depletion of MORC2 or HUSH increases the expression of DNAH3 in both K562 (n = 2 biological replicates × 3 technical replicates) and hES cells (n = 3 technical replicates), as measured by RT–qPCR assay. The DNAH3 expression level was normalized to β-actin mRNA. All samples were then normalized to the control sample. Centre value is median.

Extended Data Figure 10 HUSH and MORC2 binding at intronic L1s results in the decreased expression of active host genes.

a, Genome browser tracks illustrating that the loss of HUSH and MORC2 causes decreased H3K9me3 over the intronic L1PA5 element and concomitant increase in the expression of host gene RABL3. The experiment was repeated once with similar results. b, The loss of HUSH or MORC2 leads to increased Pol II signals at the 5′ UTR and decreased Pol II signals within L1 bodies at HUSH-bound L1PA elements (orange bars). Heat maps show Pol II density change in knockout K562 clones compared to control, centred on the L1 5′ end and sorted by MPP8 ChIP signal. c, Deletion of the intronic L1 within RABL3 causes increased RABL3 expression. Top, an agarose gel analysis of the PCR assay with primers flanking the HUSH- or MORC2-bound intronic L1; two experiments repeated independently with similar results. Bottom, RT–qPCR analysis of RABL3 expression. The RABL3 expression level was normalized to β-actin mRNA. All samples were then normalized to the wild-type sample. n = 2 biological replicates × 3 technical replicates (centre value is median). d, Depletion of MORC2, MPP8 or TASOR increases RABL3 expression. RT–qPCR data normalized as in c. n = 2 biological replicates × 3 technical replicates (centre value is median).

Supplementary information

Supplementary Information

This file contains the uncropped scans with size marker indications. (PDF 1044 kb)

Life Sciences Reporting Summary (PDF 92 kb)

Supplementary Table 1

This table contains genome-wide screen results in K562 cells and HeLa cells. (XLSX 5858 kb)

Supplementary Table 2

This table contains the secondary screen results in K562 cells and HeLa cells. (XLSX 150 kb)

Supplementary Table 3

The sequence of sgRNAs in this study. (XLSX 28 kb)

Supplementary Table 4

This table contains the sequences of oligonucleotides used in this work. (XLSX 24 kb)

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Liu, N., Lee, C., Swigut, T. et al. Selective silencing of euchromatic L1s revealed by genome-wide screens for L1 regulators. Nature 553, 228–232 (2018). https://doi.org/10.1038/nature25179

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