Precision gene editing technology and applications in nephrology

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The expanding field of precision gene editing is empowering researchers to directly modify DNA. Gene editing is made possible using synonymous technologies: a DNA-binding platform to molecularly locate user-selected genomic sequences and an associated biochemical activity that serves as a functional editor. The advent of accessible DNA-targeting molecular systems, such as zinc-finger nucleases, transcription activator-like effectors (TALEs) and CRISPR–Cas9 gene editing systems, has unlocked the ability to target nearly any DNA sequence with nucleotide-level precision. Progress has also been made in harnessing endogenous DNA repair machineries, such as non-homologous end joining, homology-directed repair and microhomology-mediated end joining, to functionally manipulate genetic sequences. As understanding of how DNA damage results in deletions, insertions and modifications increases, the genome becomes more predictably mutable. DNA-binding platforms such as TALEs and CRISPR can also be used to make locus-specific epigenetic changes and to transcriptionally enhance or suppress genes. Although many challenges remain, the application of precision gene editing technology in the field of nephrology has enabled the generation of new animal models of disease as well as advances in the development of novel therapeutic approaches such as gene therapy and xenotransplantation.

Key points

  • Zinc-finger nucleases, transcription activator-like effector nucleases and CRISPR systems are powerful tools that are enabling new applications of genome engineering in diverse systems.

  • Targeted double-stranded breaks in DNA activate diverse repair processes, such as non-homologous end joining, homology-directed repair and microhomology-mediated end joining, which can be utilized to modify the nucleotide sequence of DNA.

  • Use of non-nuclease genomic tools enables the editing of single bases and locus-specific epigenetic targeting to modify gene expression.

  • Applications of precision gene editing in nephrology include the generation of animal models to investigate kidney development and disease mechanisms as well as the development of targeted gene therapies.

  • Genome editing in the kidney is challenging owing to anatomical barriers to gene delivery, limitations of vector size and immune responses against viral vectors, modified cells and editing proteins.

  • Despite these challenges, precision gene editing has great potential to accelerate basic science in nephrology and to advance clinical practice through the development of novel therapies for renal diseases.

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Fig. 1: Programmable DNA platforms that recognize double-stranded DNA.
Fig. 2: Cost and utilization of precision gene editing in scientific research.
Fig. 3: Utilization of DNA repair pathways for precision gene editing.
Fig. 4: Precision epigenetic modulation.


  1. 1.

    Kim, Y. G., Cha, J. & Chandrasegaran, S. Hybrid restriction enzymes: zinc finger fusions to Fok I cleavage domain. Proc. Natl Acad. Sci. USA 93, 1156–1160 (1996).

  2. 2.

    Carroll, D. Genome engineering with zinc-finger nucleases. Genetics 188, 773–782 (2011).

  3. 3.

    Christian, M. et al. Targeting DNA double-strand breaks with TAL effector nucleases. Genetics 186, 757–761 (2010).

  4. 4.

    Joung, J. K. & Sander, J. D. TALENs: a widely applicable technology for targeted genome editing. Nat. Rev. Mol. Cell Biol. 14, 49–55 (2013).

  5. 5.

    Doudna, J. A. & Charpentier, E. The new frontier of genome engineering with CRISPR-Cas9. Science 346, 1258096 (2014).

  6. 6.

    Sternberg, S. H. & Doudna, J. A. Expanding the biologists toolkit with CRISPR-Cas9. Mol. Cell 58, 568–574 (2015).

  7. 7.

    Peng, Y. et al. Making designer mutants in model organisms. Development 141, 4042–4054 (2014).

  8. 8.

    Porteus, M. H. & Baltimore, D. Chimeric nucleases stimulate gene targeting in hu-man cells. Science 300, 763–763 (2003).

  9. 9.

    Bibikova, M., Beumer, K., Trautman, J. K. & Carroll, D. Enhancing gene targeting with designed zinc finger nucleases. Science 300, 764–764 (2003).

  10. 10.

    Wolfe, S. A., Nekludova, L. & Pabo, C. O. DNA recognition by Cys(2)His(2) zinc finger proteins. Annu. Rev. Biophys. Biomol. Struct. 29, 183–212 (2000).

  11. 11.

    Pavletich, N. P. & Pabo, C. O. Zinc finger dna recognition - crystal-structure of a Zif268-DNA complex at 2.1-A. Science 252, 809–817 (1991).

  12. 12.

    Desjarlais, J. R. & Berg, J. M. Redesigning the DNA-binding specificity of a zinc finger protein - a data base-guided approach. Proteins 12, 101–104 (1992).

  13. 13.

    Segal, D. J., Dreier, B., Beerli, R. R. & Barbas, C. F. Toward controlling gene expression at will: selection and design of zinc finger domains recognizing each of the 5′-GNN-3′ DNA target sequences. Proc. Natl Acad. Sci. USA 96, 2758–2763 (1999).

  14. 14.

    Beerli, R. R. & Barbas, C. F. Engineering polydactyl zinc-finger transcription factors. Nat. Biotechnol. 20, 135–141 (2002).

  15. 15.

    Kim, H. J., Lee, H. J., Kim, H., Cho, S. W. & Kim, J.-S. Targeted genome editing in human cells with zinc finger nucleases constructed via modular assembly. Genome Res. 19, 1279–1288 (2009).

  16. 16.

    Bhakta, M. S. et al. Highly active zinc-finger nucleases by extended modular assembly. Genome Res. 23, 530–538 (2013).

  17. 17.

    Gupta, A. et al. An optimized two-finger archive for ZFN-mediated gene targeting. Nat. Methods 9, 588–590 (2012).

  18. 18.

    Laoharawee, K. et al. Dose-dependent prevention of metabolic and neurologic disease in murine MPS II by ZFN-mediated in vivo genome editing. Mol. Ther. 26, 1127–1136 (2018).

  19. 19.

    Bibikova, M. et al. Stimulation of homologous recombination through targeted cleavage by chimeric nucleases. Mol. Cell. Biol. 21, 289–297 (2001).

  20. 20.

    Bibikova, M., Golic, M., Golic, K. G. & Carroll, D. Targeted chromosomal cleavage and mutagenesis in Drosophila using zinc-finger nucleases. Genetics 161, 1169–1175 (2002).

  21. 21.

    Urnov, F. D. et al. Highly efficient endogenous human gene correction using designed zinc-finger nucleases. Nature 435, 646–651 (2005).

  22. 22.

    Doyon, Y. et al. Heritable targeted gene disruption in zebrafish using designed zinc-finger nucleases. Nat. Biotechnol. 26, 702–708 (2008).

  23. 23.

    Miller, J. C. et al. An improved zinc-finger nuclease architecture for highly specific genome editing. Nat. Biotechnol. 25, 778–785 (2007).

  24. 24.

    Szczepek, M. et al. Structure-based redesign of the dimerization interface reduces the toxicity of zinc-finger nucleases. Nat. Biotechnol. 25, 786–793 (2007).

  25. 25.

    Doyon, Y. et al. Enhancing zinc-finger-nuclease activity with improved obligate heterodimeric architectures. Nat. Methods 8, 74–79 (2011).

  26. 26.

    Cornu, T. I. et al. DNA-binding specificity is a major determinant of the activity and toxicity of zinc-finger nucleases. Mol. Ther. 16, 352–358 (2008).

  27. 27.

    Haendel, E.-M., Alwin, S. & Cathomen, T. Expanding or restricting the target site repertoire of zinc-finger nucleases: the inter-domain linker as a major determinant of target site selectivity. Mol. Ther. 17, 104–111 (2009).

  28. 28.

    Gabriel, R. et al. An unbiased genome-wide analysis of zinc-finger nuclease specificity. Nat. Biotechnol. 29, 816–823 (2011).

  29. 29.

    Boch, J. et al. Breaking the code of DNA binding specificity of TAL-type III effectors. Science 326, 1509–1512 (2009).

  30. 30.

    Cermak, T. et al. Efficient design and assembly of custom TALEN and other TAL effector-based constructs for DNA targeting. Nucleic Acids Res. 39, e82 (2011).

  31. 31.

    Deng, D. et al. Structural basis for sequence-specific recognition of DNA by TAL effectors. Science 335, 720–723 (2012).

  32. 32.

    Moscou, M. J. & Bogdanove, A. J. A simple cipher governs DNA recognition by TAL effectors. Science 326, 1501 (2009).

  33. 33.

    Bogdanove, A. J. & Voytas, D. F. TAL effectors: customizable proteins for DNA targeting. Science 333, 1843–1846 (2011).

  34. 34.

    Mussolino, C. et al. A novel TALE nuclease scaffold enables high genome editing activity in combination with low toxicity. Nucleic Acids Res. 39, 9283–9293 (2011).

  35. 35.

    Mali, P. et al. CAS9 transcriptional activators for target specificity screening and paired nickases for cooperative genome engineering. Nat. Biotechnol. 31, 833–838 (2013).

  36. 36.

    Smith, C. et al. Whole-genome sequencing analysis reveals high specificity of CRISPR/Cas9 and TALEN-based genome editing in human iPSCs. Cell Stem Cell 15, 13–14 (2014).

  37. 37.

    Hockemeyer, D. et al. Genetic engineering of human pluripotent cells using TALE nucleases. Nat. Biotechnol. 29, 731–734 (2011).

  38. 38.

    Lamb, B. M., Mercer, A. C. & Barbas, C. F. Directed evolution of the TALE N-terminal domain for recognition of all 5′ bases. Nucleic Acids Res. 41, 9779–9785 (2013).

  39. 39.

    Deng, D. et al. Recognition of methylated DNA by TAL effectors. Cell Res. 22, 1502–1504 (2012).

  40. 40.

    Kim, Y. et al. A library of TAL effector nucleases spanning the human genome. Nat. Biotechnol. 31, 251–258 (2013).

  41. 41.

    Zhang, F. et al. Efficient construction of sequence-specific TAL effectors for modulating mammalian transcription. Nat. Biotechnol. 29, 149–153 (2011).

  42. 42.

    Reyon, D. et al. FLASH assembly of TALENs for high-throughput genome editing. Nat. Biotechnol. 30, 460–465 (2012).

  43. 43.

    Heigwer, F. et al. E-TALEN: a web tool to design TALENs for genome engineering. Nucleic Acids Res. 41, e190 (2013).

  44. 44.

    Ma, A. C. et al. FusX: a rapid one-step transcription activator-like effector assembly system for genome science. Hum. Gene Ther. 27, 451–463 (2016).

  45. 45.

    Jinek, M. et al. A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337, 816–821 (2012).

  46. 46.

    Makarova, K. S., Grishin, N. V., Shabalina, S. A., Wolf, Y. I. & Koonin, E. V. A putative RNA-interference-based immune system in prokaryotes: computational analysis of the predicted enzymatic machinery, functional analogies with eukaryotic RNAi, and hypothetical mechanisms of action. Biol. Direct 1, 7 (2006).

  47. 47.

    Barrangou, R. et al. CRISPR provides acquired resistance against viruses in prokaryotes. Science 315, 1709–1712 (2007).

  48. 48.

    Makarova, K. S. et al. Evolution and classification of the CRISPR-Cas systems. Nat. Rev. Microbiol. 9, 467–477 (2011).

  49. 49.

    Sapranauskas, R. et al. The Streptococcus thermophilus CRISPR/Cas system provides immunity in Escherichia coli. Nucleic Acids Res. 39, 9275–9282 (2011).

  50. 50.

    Barrangou, R. & Marraffini, L. A. CRISPR-Cas systems: prokaryotes upgrade to adaptive immunity. Mol. Cell 54, 234–244 (2014).

  51. 51.

    Bolotin, A., Ouinquis, B., Sorokin, A. & Ehrlich, S. D. Clustered regularly interspaced short palindrome repeats (CRISPRs) have spacers of extrachromosomal origin. Microbiology 151, 2551–2561 (2005).

  52. 52.

    Mojica, F. J. M., Diez-Villasenor, C., Garcia-Martinez, J. & Soria, E. Intervening sequences of regularly spaced prokaryotic repeats derive from foreign genetic elements. J. Mol. Evol. 60, 174–182 (2005).

  53. 53.

    Brouns, S. J. J. et al. Small CRISPR RNAs guide antiviral defense in prokaryotes. Science 321, 960–964 (2008).

  54. 54.

    Mojica, F. J. M., Diez-Villasenor, C., Garcia-Martinez, J. & Almendros, C. Short motif sequences determine the targets of the prokaryotic CRISPR defence system. Microbiology 155, 733–740 (2009).

  55. 55.

    Gasiunas, G., Barrangou, R., Horvath, P. & Siksnys, V. Cas9-crRNA ribonucleoprotein complex mediates specific DNA cleavage for adaptive immunity in bacteria. Proc. Natl Acad. Sci. USA 109, E2579–E2586 (2012).

  56. 56.

    Sternberg, S. H., Redding, S., Jinek, M., Greene, E. C. & Doudna, J. A. DNA interrogation by the CRISPR RNA-guided endonuclease Cas9. Nature 507, 62–67 (2014).

  57. 57.

    Wiedenheft, B., Sternberg, S. H. & Doudna, J. A. RNA-guided genetic silencing systems in bacteria and archaea. Nature 482, 331–338 (2012).

  58. 58.

    Jinek, M. et al. Structures of Cas9 endonucleases reveal RNA-mediated conformational activation. Science 343, 1247997 (2014).

  59. 59.

    Mali, P., Esvelt, K. M. & Church, G. M. Cas9 as a versatile tool for engineering biology. Nat. Methods 10, 957–963 (2013).

  60. 60.

    Cencic, R. et al. Protospacer adjacent motif (PAM)-distal sequences engage CRISPR Cas9 DNA target cleavage. PLoS ONE 9, e109213 (2014).

  61. 61.

    Anders, C., Niewoehner, O., Duerst, A. & Jinek, M. Structural basis of PAM-dependent target DNA recognition by the Cas9 endonuclease. Nature 513, 569–573 (2014).

  62. 62.

    Belhaj, K., Chaparro-Garcia, A., Kamoun, S. & Nekrasov, V. Plant genome editing made easy: targeted mutagenesis in model and crop plants using the CRISPR/Cas system. Plant Methods 9, 39 (2013).

  63. 63.

    Cho, S. W., Kim, S., Kim, J. M. & Kim, J.-S. Targeted genome engineering in human cells with the Cas9 RNA-guided endonuclease. Nat. Biotechnol. 31, 230–232 (2013).

  64. 64.

    Dickinson, D. J., Ward, J. D., Reiner, D. J. & Goldstein, B. Engineering the Caenorhabditis elegans genome using Cas9-triggered homologous recombination. Nat. Methods 10, 1028–1034 (2013).

  65. 65.

    Cradick, T. J., Fine, E. J., Antico, C. J. & Bao, G. CRISPR/Cas9 systems targeting beta-globin and CCR5 genes have substantial off-target activity. Nucleic Acids Res. 41, 9584–9592 (2013).

  66. 66.

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

  67. 67.

    Cho, S. W., Lee, J., Carroll, D., Kim, J.-S. & Lee, J. Heritable gene knockout in Caenorhabditis elegans by direct injection of Cas9-sgRNA ribonucleoproteins. Genetics 195, 1177–1180 (2013).

  68. 68.

    Gratz, S. J. et al. Genome engineering of Drosophila with the CRISPR RNA-guided Cas9 nuclease. Genetics 194, 1029–1035 (2013).

  69. 69.

    Fu, Y. et al. High-frequency off-target mutagenesis induced by CRISPR-Cas nucleases in human cells. Nat. Biotechnol. 31, 822–826 (2013).

  70. 70.

    Hsu, P. D. et al. DNA targeting specificity of RNA-guided Cas9 nucleases. Nat. Biotechnol. 31, 827–832 (2013).

  71. 71.

    Cho, S. W. et al. Analysis of off-target effects of CRISPR/Cas-derived RNA-guided endonucleases and nickases. Genome Res. 24, 132–141 (2014).

  72. 72.

    Tsai, S. Q. & Joung, J. K. Defining and improving the genome-wide specificities of CRISPR-Cas9 nucleases. Nat. Rev. Genet. 17, 300–312 (2016).

  73. 73.

    Peng, R. X., Lin, G. G. & Li, J. M. Potential pitfalls of CRISPR/Cas9-mediated genome editing. FEBS J. 283, 1218–1231 (2016).

  74. 74.

    Duan, J. Z. et al. Genome-wide identification of CRISPR/Cas9 off-targets in human genome. Cell Res. 24, 1009–1012 (2014).

  75. 75.

    Kuscu, C., Arslan, S., Singh, R., Thorpe, J. & Adli, M. Genome-wide analysis reveals characteristics of off-target sites bound by the Cas9 endonuclease. Nat. Biotechnol. 32, 677–683 (2014).

  76. 76.

    Frock, R. L. et al. Genome-wide detection of DNA double-stranded breaks induced by engineered nucleases. Nat. Biotechnol. 33, 179–186 (2015).

  77. 77.

    Fu, Y., Sander, J. D., Reyon, D., Cascio, V. M. & Joung, J. K. Improving CRISPR-Cas nuclease specificity using truncated guide RNAs. Nat. Biotechnol. 32, 279–284 (2014).

  78. 78.

    Guilinger, J. P., Thompson, D. B. & Liu, D. R. Fusion of catalytically inactive Cas9 to Fokl nuclease improves the specificity of genome modification. Nat. Biotechnol. 32, 577–582 (2014).

  79. 79.

    Qi, L. S. et al. Repurposing CRISPR as an RNA-guided platform for sequence-specific control of gene expression. Cell 152, 1173–1183 (2013).

  80. 80.

    Tsai, S. Q. et al. Dimeric CRISPR RNA-guided Fokl nucleases for highly specific genome editing. Nat. Biotechnol. 32, 569–576 (2014).

  81. 81.

    Ran, F. A. et al. Double nicking by RNA-guided CRISPR Cas9 for enhanced genome editing specificity. Cell 154, 1380–1389 (2013).

  82. 82.

    Slaymaker, I. M. et al. Rationally engineered Cas9 nucleases with improved specificity. Science 351, 84–88 (2016).

  83. 83.

    Kleinstiver, B. P. et al. High-fidelity CRISPR-Cas9 nucleases with no detectable genome-wide off-target effects. Nature 529, 490–495 (2016).

  84. 84.

    Zetsche, B. et al. Cpf1 is a single RNA-guided endonuclease of a class 2 CRISPR-Cas system. Cell 163, 759–771 (2015).

  85. 85.

    Kim, D. et al. Genome-wide analysis reveals specificities of Cpf1 endonucleases in human cells. Nat. Biotechnol. 34, 863–868 (2016).

  86. 86.

    East-Seletsky, A., O’Connell, M. R., Burstein, D., Knott, G. J. & Doudna, J. A. RNA targeting by functionally orthogonal type VI-A CRISPR-Cas enzymes. Mol. Cell 66, 373–383 (2017).

  87. 87.

    East-Seletsky, A. et al. Two distinct RNase activities of CRISPR-C2c2 enable guide-RNA processing and RNA detection. Nature 538, 270–273 (2016).

  88. 88.

    Yin, P. et al. Structural basis for the modular recognition of single-stranded RNA by PPR proteins. Nature 504, 168–171 (2013).

  89. 89.

    Kim, Y., Kweon, J. & Kim, J.-S. TALENs and ZFNs are associated with different mutation signatures. Nat. Methods 10, 185–185 (2013).

  90. 90.

    Xiao, A. et al. Chromosomal deletions and inversions mediated by TALENs and CRISPR/Cas in zebrafish. Nucleic Acids Res. 41, e141 (2013).

  91. 91.

    Nakade, S. et al. Microhomology-mediated end-joining-dependent integration of donor DNA in cells and animals using TALENs and CRISPR/Cas9. Nat. Commun. 5, 5560 (2014).

  92. 92.

    Chu, V. T. et al. Increasing the efficiency of homology-directed repair for CRISPR-Cas9-induced precise gene editing in mammalian cells. Nat. Biotechnol. 33, 543–548 (2015).

  93. 93.

    Maruyama, T. et al. Increasing the efficiency of precise genome editing with CRISPR-Cas9 by inhibition of nonhomologous end joining. Nat. Biotechnol. 33, 538–542 (2015).

  94. 94.

    Sakuma, T., Nakade, S., Sakane, Y., Suzuki, K. T. & Yamamoto, T. MMEJ-assisted gene knock-in using TALENs and CRISPR-Cas9 with the PITCh systems. Nat. Protoc. 11, 118–133 (2016).

  95. 95.

    Hoeijmakers, J. H. J. Genome maintenance mechanisms for preventing cancer. Nature 411, 366–374 (2001).

  96. 96.

    Komor, A. C., Kim, Y. B., Packer, M. S., Zuris, J. A. & Liu, D. R. Programmable editing of a target base in genomic DNA without double-stranded DNA cleavage. Nature 533, 420–424 (2016).

  97. 97.

    Auer, T. O., Duroure, K., De Cian, A., Concordet, J.-P. & Del Bene, F. Highly efficient CRISPR/Cas9-mediated knock-in in zebrafish by homology-independent DNA repair. Genome Res. 24, 142–153 (2014).

  98. 98.

    Greene, E. C. DNA sequence alignment during homologous recombination. J. Biol. Chem. 291, 11572–11580 (2016).

  99. 99.

    Qi, Z. et al. DNA sequence alignment by microhomology sampling during homologous recombination. Cell 160, 856–869 (2015).

  100. 100.

    Zu, Y. et al. TALEN-mediated precise genome modification by homologous recombination in zebrafish. Nat. Methods 10, 329–331 (2013).

  101. 101.

    Radecke, S., Radecke, F., Cathomen, T. & Schwarz, K. Zinc-finger nuclease-induced gene repair with oligodeoxynucleotides: wanted and unwanted target locus modifications. Mol. Ther. 18, 743–753 (2010).

  102. 102.

    Sakuma, T. & Yamamoto, T. Magic wands of CRISPR-lots of choices for gene knock-in. Cell Biol. Toxicol. 33, 501–505 (2017).

  103. 103.

    Richardson, C. D. et al. CRISPR-Cas9 genome editing in human cells works via the Fanconi anemia pathway. Preprint at bioRxiv (2017).

  104. 104.

    Danner, E. et al. Control of gene editing by manipulation of DNA repair mechanisms. Mamm. Genome 28, 262–274 (2017).

  105. 105.

    Bothmer, A. et al. Characterization of the interplay between DNA repair and CRISPR/Cas9-induced DNA lesions at an endogenous locus. Nat. Commun. 8, 13905 (2017).

  106. 106.

    Chen, F. et al. High-frequency genome editing using ssDNA oligonucleotides with zinc-finger nucleases. Nat. Methods 8, 753–755 (2011).

  107. 107.

    Kostyrko, K. & Mermod, N. Assays for DNA double-strand break repair by microhomology-based end-joining repair mechanisms. Nucleic Acids Res. 44, e56 (2016).

  108. 108.

    Ahrabi, S. et al. A role for human homologous recombination factors in suppressing microhomology-mediated end joining. Nucleic Acids Res. 44, 5743–5757 (2016).

  109. 109.

    McVey, M. & Lee, S. E. MMEJ repair of double-strand breaks (director’s cut): deleted sequences and alternative endings. Trends Genet. 24, 529–538 (2008).

  110. 110.

    Lu, G. Q. et al. Ligase I and ligase III mediate the DNA double-strand break ligation in alternative end-joining. Proc. Natl Acad. Sci. USA 113, 1256–1260 (2016).

  111. 111.

    Nakamae, K. et al. Establishment of expanded and streamlined pipeline of PITCh knock-in - a web-based design tool for MMEJ-mediated gene knock-in, PITCh designer, and the variations of PITCh, PITCh-TG and PITCh-KIKO. Bioengineered 8, 302–308 (2017).

  112. 112.

    Aida, T. et al. Gene cassette knock-in in mammalian cells and zygotes by enhanced MMEJ. BMC Genomics 17, 979 (2016).

  113. 113.

    Yao, X. et al. Homology-mediated end joining-based targeted integration using CRISPR/Cas9. Cell Res. 27, 801–814 (2017).

  114. 114.

    Bennardo, N., Cheng, A., Huang, N. & Stark, J. M. Alternative-NHEJ Is a mechanistically distinct pathway of mammalian chromosome break repair. PLoS Genet. 4, e1000110 (2008).

  115. 115.

    Yang, L. H. et al. Corrigendum: engineering and optimising deaminase fusions for genome editing. Nat. Commun. 8, 16169 (2017).

  116. 116.

    Gaudelli, N. M. et al. Programmable base editing of A·T to G·C in genomic DNA without DNA cleavage. Nature 551, 464–471 (2017).

  117. 117.

    Zong, Y. et al. Precise base editing in rice, wheat and maize with a Cas9-cytidine deaminase fusion. Nat. Biotechnol. 35, 438–440 (2017).

  118. 118.

    Zhang, Y. H. et al. Programmable base editing of zebrafish genome using a modified CRISPR-Cas9 system. Nat. Commun. 8, 118 (2017).

  119. 119.

    Li, Z. et al. APOBEC signature mutation generates an oncogenic enhancer that drives LMO1 expression in T-ALL. Leukemia 31, 2057–2064 (2017).

  120. 120.

    Kouno, T. et al. Crystal structure of APOBEC3A bound to single-stranded DNA reveals structural basis for cytidine deamination and specificity. Nat. Commun. 8, 15024 (2017).

  121. 121.

    Kim, Y. B. et al. Increasing the genome-targeting scope and precision of base editing with engineered Cas9-cytidine deaminase fusions. Nat. Biotechnol. 35, 371–376 (2017).

  122. 122.

    Brachova, P., Alvarez, N. S., Van Voorhis, B. J. & Christenson, L. K. Cytidine deaminase Apobec3a induction in fallopian epithelium after exposure to follicular fluid. Gynecol. Oncol. 145, 577–583 (2017).

  123. 123.

    Billon, P. et al. CRISPR-mediated base editing enables efficient disruption of eukaryotic genes through induction of STOP codons. Mol. Cell 67, 1068–1079 (2017).

  124. 124.

    Yang, L. H. et al. Engineering and optimising deaminase fusions for genome editing. Nat. Commun. 7, 13330 (2016).

  125. 125.

    Kungulovski, G. & Jeltsch, A. Epigenome editing: state of the art, concepts, and perspectives. Trends Genet. 32, 101–113 (2016).

  126. 126.

    Smith, A. E. & Ford, K. G. Specific targeting of cytosine methylation to DNA sequences in vivo. Nucleic Acids Res. 35, 740–754 (2007).

  127. 127.

    Thakore, P. I., Black, J. B., Hilton, I. B. & Gersbach, C. A. Editing the epigenome: technologies for programmable transcription and epigenetic modulation. Nat. Methods 13, 127–137 (2016).

  128. 128.

    Xu, X. X. et al. A CRISPR-based approach for targeted DNA demethylation. Cell Discov. 2, 16009 (2016).

  129. 129.

    Carvin, C. D., Parr, R. D. & Kladde, M. P. Site-selective in vivo targeting of cytosine-5 DNA methylation by zinc-finger proteins. Nucleic Acids Res. 31, 6493–6501 (2003).

  130. 130.

    Maeder, M. L. et al. Targeted DNA demethylation and activation of endogenous genes using programmable TALE-TET1 fusion proteins. Nat. Biotechnol. 31, 1137–1142 (2013).

  131. 131.

    Liu, Y. D. et al. Zinc finger protein 618 regulates the function of UHRF2 (ubiquitin-like with PHD and ring finger domains 2) as a specific 5-hydroxymethylcytosine reader. J. Biol. Chem. 291, 13679–13688 (2016).

  132. 132.

    Vojta, A. et al. Repurposing the CRISPR-Cas9 system for targeted DNA methylation. Nucleic Acids Res. 44, 5615–5628 (2016).

  133. 133.

    Sterner, D. E. & Berger, S. L. Acetylation of histones and transcription-related factors. Microbiol. Mol. Biol. Rev. 64, 435–459 (2000).

  134. 134.

    Seto, E. & Yoshida, M. Erasers of histone acetylation: the histone deacetylase enzymes. Cold Spring Harb. Perspect. Biol. 6, a018713 (2014).

  135. 135.

    Hilton, I. B. et al. Epigenome editing by a CRISPR-Cas9-based acetyltransferase activates genes from promoters and enhancers. Nat. Biotechnol. 33, 510–517 (2015).

  136. 136.

    Kwon, D. Y., Zhao, Y. T., Lamonica, J. M. & Zhou, Z. Locus-specific histone deacetylation using a synthetic CRISPR-Cas9-based HDAC. Nat. Commun. 8, 15315 (2017).

  137. 137.

    Looman, C., Abrink, M., Mark, C. & Hellman, L. KRAB zinc finger proteins: an analysis of the molecular mechanisms governing their increase in numbers and complexity during evolution. Mol. Biol. Evol. 19, 2118–2130 (2002).

  138. 138.

    Krishna, S. S., Majumdar, I. & Grishin, N. V. Structural classification of zinc fingers. Nucleic Acids Res. 31, 532–550 (2003).

  139. 139.

    Guilliere, F. et al. Solution structure of an archaeal DNA binding protein with an eukaryotic zinc finger fold. PLoS ONE 8, e52908 (2013).

  140. 140.

    Erkes, A., Reschke, M., Boch, J. & Grau, J. Evolution of transcription activator-like effectors in Xanthomonas oryzae. Genome Biol. Evol. 9, 1599–1615 (2017).

  141. 141.

    Chavez, A. et al. Highly efficient Cas9-mediated transcriptional programming. Nat. Methods 12, 326–328 (2015).

  142. 142.

    Thakore, P. I. et al. Highly specific epigenome editing by CRISPR-Cas9 repressors for silencing of distal regulatory elements. Nat. Methods 12, 1143–1149 (2015).

  143. 143.

    Shen, F., Triezenberg, S. J., Hensley, P., Porter, D. & Knutson, J. R. Transcriptional activation domain of the herpesvirus protein VP16 becomes conformationally constrained upon interaction with basal transcription factors. J. Biol. Chem. 271, 4827–4837 (1996).

  144. 144.

    Graslund, T., Li, X. L., Magnenat, L., Popkov, M. & Barbas, C. F. Exploring strategies for the design of artificial transcription factors. J. Biol. Chem. 280, 3707–3714 (2005).

  145. 145.

    Hofherr, A. et al. Efficient genome editing of differentiated renal epithelial cells. Pflugers Arch. 469, 303–311 (2017).

  146. 146.

    Freedman, B. S. et al. Modelling kidney disease with CRISPR-mutant kidney organoids derived from human pluripotent epiblast spheroids. Nat. Commun. 6, 8715 (2015).

  147. 147.

    Ben, J., Elworthy, S., Ng, A. S., van Eeden, F. & Ingham, P. W. Targeted mutation of the talpid3 gene in zebrafish reveals its conserved requirement for ciliogenesis and Hedgehog signalling across the vertebrates. Development 138, 4969–4978 (2011).

  148. 148.

    Jiang, D. et al. CRISPR/Cas9-induced disruption of wt1a and wt1b reveals their different roles in kidney and gonad development in Nile tilapia. Dev. Biol. 428, 63–73 (2017).

  149. 149.

    Jaffe, K. M. et al. c21orf59/kurly controls both cilia motility and polarization. Cell Rep. 14, 1841–1849 (2016).

  150. 150.

    Marusugi, K. et al. Functional validation of tensin2 SH2-PTB domain by CRISPR/Cas9-mediated genome editing. J. Vet. Med. Sci. 78, 1413–1420 (2016).

  151. 151.

    Brophy, P. D. et al. A gene implicated in activation of retinoic acid receptor targets is a novel renal agenesis gene in humans. Genetics 207, 215–228 (2017).

  152. 152.

    Wang, X. et al. Generation and phenotypic characterization of Pde1a mutant mice. PLoS ONE 12, e0181087 (2017).

  153. 153.

    Ye, H. et al. Modulation of polycystic kidney disease severity by phosphodiesterase 1 and 3 subfamilies. J. Am. Soc. Nephrol. 27, 1312–1320 (2016).

  154. 154.

    Chen, C. C., Geurts, A. M., Jacob, H. J., Fan, F. & Roman, R. J. Heterozygous knockout of transforming growth factor-beta1 protects Dahl S rats against high salt-induced renal injury. Physiol. Genomics 45, 110–118 (2013).

  155. 155.

    Mattson, D. L. et al. Genetic mutation of recombination activating gene 1 in Dahl salt-sensitive rats attenuates hypertension and renal damage. Am. J. Physiol. Regul. Integr. Comp. Physiol. 304, R407–R414 (2013).

  156. 156.

    Zhou, X. et al. Heterozygous disruption of renal outer medullary potassium channel in rats is associated with reduced blood pressure. Hypertension 62, 288–294 (2013).

  157. 157.

    He, J. et al. PKD1 mono-allelic knockout is sufficient to trigger renal cystogenesis in a mini-pig model. Int. J. Biol. Sci. 11, 361–369 (2015).

  158. 158.

    De Tomasi, L. et al. Mutations in GREB1L cause bilateral kidney agenesis in humans and mice. Am. J. Hum. Genet. 101, 803–814 (2017).

  159. 159.

    Sanna-Cherchi, S. et al. Exome-wide association study identifies GREB1L mutations in congenital kidney malformations. Am. J. Hum. Genet. 101, 789–802 (2017).

  160. 160.

    Johnson, B. G. et al. Uromodulin p. Cys147Trp mutation drives kidney disease by activating ER stress and apoptosis. J. Clin. Invest. 127, 3954–3969 (2017).

  161. 161.

    Yang, L. H. et al. Genome-wide inactivation of porcine endogenous retroviruses (PERVs). Science 350, 1101–1104 (2015).

  162. 162.

    Niu, D. et al. Inactivation of porcine endogenous retrovirus in pigs using CRISPR-Cas9. Science 357, 1303–1307 (2017).

  163. 163.

    Petersen, B. & Niemann, H. Molecular scissors and their application in genetically modified farm animals. Transgenic Res. 24, 381–396 (2015).

  164. 164.

    Long, C. et al. Postnatal genome editing partially restores dystrophin expression in a mouse model of muscular dystrophy. Science 351, 400–403 (2016).

  165. 165.

    Nelson, C. E. et al. In vivo genome editing improves muscle function in a mouse model of Duchenne muscular dystrophy. Science 351, 403–407 (2016).

  166. 166.

    Tabebordbar, M. et al. In vivo gene editing in dystrophic mouse muscle and muscle stem cells. Science 351, 407–411 (2016).

  167. 167.

    Cirak, S. et al. Exon skipping and dystrophin restoration in patients with Duchenne muscular dystrophy after systemic phosphorodiamidate morpholino oligomer treatment: an open-label, phase 2, dose-escalation study. Lancet 378, 595–605 (2011).

  168. 168.

    Hildebrandt, F. Decade in review–genetics of kidney diseases: genetic dissection of kidney disorders. Nat. Rev. Nephrol. 11, 635–636 (2015).

  169. 169.

    Samulski, R. J. & Muzyczka, N. AAV-mediated gene therapy for research and therapeutic purposes. Annu. Rev. Virol. 1, 427–451 (2014).

  170. 170.

    Hwang, M. et al. TGF-beta1 siRNA suppresses the tubulointerstitial fibrosis in the kidney of ureteral obstruction. Exp. Mol. Pathol. 81, 48–54 (2006).

  171. 171.

    Zincarelli, C., Soltys, S., Rengo, G. & Rabinowitz, J. E. Analysis of AAV serotypes 1–9 mediated gene expression and tropism in mice after systemic injection. Mol. Ther. 16, 1073–1080 (2008).

  172. 172.

    Hillestad, M. L., Guenzel, A. J., Nath, K. A. & Barry, M. A. A. Vector-host system to fingerprint virus tropism. Hum. Gene Ther. 23, 1116–1126 (2012).

  173. 173.

    Chung, D. C. et al. Adeno-associated virus-mediated gene transfer to renal tubule cells via a retrograde ureteral approach. Nephron Extra 1, 217–223 (2011).

  174. 174.

    Ellis, B. L., Hirsch, M. L., Porter, S. N., Samulski, R. J. & Porteus, M. H. Zinc-finger nuclease-mediated gene correction using single AAV vector transduction and enhancement by Food and Drug Administration-approved drugs. Gene Ther. 20, 35–42 (2013).

  175. 175.

    Yang, J. et al. Targeting of macrophage activity by adenovirus-mediated intragraft overexpression of TNFRp55-Ig, IL-12p40, and vIL-10 ameliorates adenovirus-mediated chronic graft injury, whereas stimulation of macrophages by overexpression of IFN-gamma accelerates chronic graft injury in a rat renal allograft model. J. Am. Soc. Nephrol. 14, 214–225 (2003).

  176. 176.

    Brunetti-Pierri, N. & Ng, P. Gene therapy with helper-dependent adenoviral vectors: lessons from studies in large animal models. Virus Genes 53, 684–691 (2017).

  177. 177.

    Yang, Y. P., Su, Q. & Wilson, J. M. Role of viral antigens in destructive cellular immune responses to adenovirus vector-transduced cells in mouse lungs. J. Virol. 70, 7209–7212 (1996).

  178. 178.

    Raper, S. E. et al. Fatal systemic inflammatory response syndrome in a ornithine transcarbamylase deficient patient following adenoviral gene transfer. Mol. Genet. Metab. 80, 148–158 (2003).

  179. 179.

    Jooss, K., Yang, Y., Fisher, K. J. & Wilson, J. M. Transduction of dendritic cells by DNA viral vectors directs the immune response to transgene products in muscle fibers. J. Virol. 72, 4212–4223 (1998).

  180. 180.

    Chirmule, N. et al. Immune responses to adenovirus and adeno-associated virus in humans. Gene Ther. 6, 1574–1583 (1999).

  181. 181.

    Ertl, H. C. J. & High, K. A. Impact of AAV capsid-specific T-cell responses on design and outcome of clinical gene transfer trials with recombinant adeno-associated viral vectors: an evolving controversy. Hum. Gene Ther. 28, 328–337 (2017).

  182. 182.

    Mitani, K., Graham, F. L., Caskey, C. T. & Kochanek, S. Rescue, propagation, and partial-purification of a helper virus-dependent adenovirus vector. Proc. Natl Acad. Sci. USA 92, 3854–3858 (1995).

  183. 183.

    Fisher, K. J., Choi, H., Burda, J., Chen, S. J. & Wilson, J. M. Recombinant adenovirus deleted of all viral genes for gene therapy of cystic fibrosis. Virology 217, 11–22 (1996).

  184. 184.

    Morral, N. et al. High doses of a helper-dependent adenoviral vector yield supraphysiological levels of alpha(1)-antitrypsin with negligible toxicity. Hum. Gene Ther. 9, 2709–2716 (1998).

  185. 185.

    Morral, N. et al. Administration of helper-dependent adenoviral vectors and sequential delivery of different vector serotype for long-term liver-directed gene transfer in baboons. Proc. Natl Acad. Sci. USA 96, 12816–12821 (1999).

  186. 186.

    Charlesworth, C. T. et al. Identification of pre-existing adaptive immunity to Cas9 proteins in humans. Preprint at bioRxiv (2018).

  187. 187.

    US National Library of Medicine. (2018).

  188. 188.

    US National Library of Medicine. (2018).

  189. 189.

    US National Library of Medicine. (2018).

  190. 190.

    Hotta, A. & Yamanaka, S. in Annual Review of Genetics Vol. 49 (ed. Bassler, B. L.) 47–70 (Annual Reviews, 2015).

  191. 191.

    US National Library of Medicine. (2018).

  192. 192.

    Ahn, J. D. et al. Transcription factor decoy for AP-1 reduces mesangial cell proliferation and extracellular matrix production in vitro and in vivo. Gene Ther. 11, 916–923 (2004).

  193. 193.

    Isaka, Y. et al. Gene therapy by skeletal muscle expression of decorin prevents fibrotic disease in rat kidney. Nat. Med. 2, 418–423 (1996).

  194. 194.

    Higuchi, N. et al. Hydrodynamics-based delivery of the viral interleukin-10 gene suppresses experimental crescentic glomerulonephritis in Wistar-Kyoto rats. Gene Ther. 10, 1297–1310 (2003).

  195. 195.

    Ka, S. M. et al. Decoy receptor 3 inhibits renal mononuclear leukocyte infiltration and apoptosis and prevents progression of IgA nephropathy in mice. Am. J. Physiol. Renal Physiol. 301, F1218–F1230 (2011).

  196. 196.

    Choi, Y. K. et al. Suppression of glomerulosclerosis by adenovirus-mediated IL-10 expression in the kidney. Gene Ther. 10, 559–568 (2003).

  197. 197.

    Chao, J. & Chao, L. Experimental kallikrein gene therapy in hypertension, cardiovascular and renal diseases. Pharmacol. Res. 35, 517–522 (1997).

  198. 198.

    Yang, C. C., Hsu, S. P., Chen, K. H. & Chien, C. T. Effect of adenoviral catalase gene transfer on renal ischemia/reperfusion injury in rats. Chin. J. Physiol. 58, 420–430 (2015).

  199. 199.

    Ravichandran, K., Ozkok, A., Wang, Q., Mullick, A. E. & Edelstein, C. L. Antisense-mediated angiotensinogen inhibition slows polycystic kidney disease in mice with a targeted mutation in Pkd2. Am. J. Physiol. Renal Physiol. 308, F349–357 (2015).

  200. 200.

    Zheng, X. et al. Attenuating ischemia-reperfusion injury in kidney transplantation by perfusing donor organs with siRNA cocktail solution. Transplantation 100, 743–752 (2016).

  201. 201.

    Ding, Z. et al. Adenovirus-mediated anti-sense ERK2 gene therapy inhibits tubular epithelial-mesenchymal transition and ameliorates renal allograft fibrosis. Transpl. Immunol. 25, 34–41 (2011).

  202. 202.

    Nakamura, H. et al. Introduction of DNA enzyme for Egr-1 into tubulointerstitial fibroblasts by electroporation reduced interstitial alpha-smooth muscle actin expression and fibrosis in unilateral ureteral obstruction (UUO) rats. Gene Ther. 9, 495–502 (2002).

  203. 203.

    Terada, Y. et al. Gene transfer of Smad7 using electroporation of adenovirus prevents renal fibrosis in post-obstructed kidney. Kidney Int. 61, S94–S98 (2002).

  204. 204.

    Lan, H. Y. et al. Inhibition of renal fibrosis by gene transfer of inducible Smad7 using ultrasound-microbubble system in rat UUO model. J. Am. Soc. Nephrol. 14, 1535–1548 (2003).

  205. 205.

    Liu, X., Shen, W., Yang, Y. & Liu, G. Therapeutic implications of mesenchymal stem cells transfected with hepatocyte growth factor transplanted in rat kidney with unilateral ureteral obstruction. J. Pediatr. Surg. 46, 537–545 (2011).

  206. 206.

    Qiao, X. et al. Intermedin is upregulated and attenuates renal fibrosis by inhibition of oxidative stress in rats with unilateral ureteral obstruction. Nephrology (Carlton) 20, 820–831 (2015).

  207. 207.

    Ozbek, E. et al. Role of mesenchymal stem cells transfected with vascular endothelial growth factor in maintaining renal structure and function in rats with unilateral ureteral obstruction. Exp. Clin. Transplant 13, 262–272 (2015).

  208. 208.

    Ren, Y. et al. CTGF siRNA ameliorates tubular cell apoptosis and tubulointerstitial fibrosis in obstructed mouse kidneys in a Sirt1-independent manner. Drug Des. Devel. Ther. 9, 4155–4171 (2015).

  209. 209.

    Bolar, N. A. et al. Heterozygous loss-of-function SEC61A1 mutations cause autosomal-dominant tubulo-interstitial and glomerulocystic kidney disease with anemia. Am. J. Hum. Genet. 99, 174–187 (2016).

  210. 210.

    Huang, Y. H. et al. DNA epigenome editing using CRISPR-Cas SunTag-directed DNMT3A. Genome Biol. 18, 176 (2017).

  211. 211.

    Jin, C. et al. HV1 acts as a sodium sensor and promotes superoxide production in medullary thick ascending limb of Dahl salt-sensitive rats. Hypertension 64, 541–550 (2014).

  212. 212.

    Endres, B. T. et al. Mutation of Plekha7 attenuates salt-sensitive hypertension in the rat. Proc. Natl Acad. Sci. USA 111, 12817–12822 (2014).

  213. 213.

    Mullins, L. J. et al. Mineralocorticoid excess or glucocorticoid insufficiency: renal and metabolic phenotypes in a rat Hsd11b2 knockout model. Hypertension 66, 667–673 (2015).

  214. 214.

    Cowley, A. W. Jr et al. Evidence of the importance of Nox4 in production of hypertension in Dahl salt-sensitive rats. Hypertension 67, 440–450 (2016).

  215. 215.

    Anderson, B. R. et al. In vivo modeling implicates APOL1 in nephropathy: evidence for dominant negative effects and epistasis under anemic stress. PLoS Genet. 11, e1005349 (2015).

  216. 216.

    Yoshino, H. et al. microRNA-210-3p depletion by CRISPR/Cas9 promoted tumorigenesis through revival of TWIST1 in renal cell carcinoma. Oncotarget 8, 20881–20894 (2017).

  217. 217.

    Yang, L. et al. Genome-wide inactivation of porcine endogenous retroviruses (PERVs). Science 350, 1101–1104 (2015).

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The authors have received research funding from the Mayo Foundation and the US National Institutes of Health (grants GM63904 and P30DK084567 (S.C.E.) and P30DK090728 (C.R.S., P.C.H. and S.C.E.)).

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Z.W.J., W.A.C.G., M.A.B., P.C.H. and C.R.S. researched the data for the article. Z.W.J., J.M.C., M.A.B., P.C.H., C.R.S. and S.C.E. made substantial contributions to discussions of the content. Z.W.J., G.M.G., W.A.C.G., M.A.B., P.C.H. and C.R.S. wrote the article and Z.W.J., J.M.C., G.M.G., M.A.B., P.C.H., C.R.S. and S.C.E. reviewed and edited the manuscript before submission.

Correspondence to Stephen C. Ekker.

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Programmable DNA nucleases

A DNA-binding platform that can be customized to bind to a specific DNA sequence and introduce a DSB in this targeted manner.

Transposon elements

DNA sequences that can be translocated within the genome by transposase proteins.

CpG site

A cytosine residue directly followed by a guanine residue in a DNA strand. Cytosine residues in CpG sites can be directly methylated by DNA methyltransferase.

Morpholino oligomers

Synthetic modified oligomers that are capable of sterically inhibiting translation of specific RNAs in a targetable manner.

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