Patent Application
20210395730
Described herein are engineered base editor variants that have reduced or negligible RNA editing activity, and methods of using the same.
Engineered base editors have recently emerged as a powerful technology for efficiently introducing single base changes in DNA1. Cytosine base editors (CBEs) are fusion proteins that induce targeted cytosine (C) to uracil (U) alterations in single-stranded DNA by using catalytically inactive or nickase versions of CRISPR-Cas nucleases to direct Apolipoprotein B mRNA editing enzyme, catalytic polypeptide-like (APOBEC) cytosine deaminases to cytosines that lie within an “editing window” in the R-loop induced by the CRISPR-Cas RNA-protein complex2. The most commonly used CBEs are the BE32 and BE43 fusions, which comprise the rat APOBEC1 (rAPOBEC1) cytosine deaminase fused to a nickase version of Cas9 (and also harbor one or two uracil glycosylase inhibitor (UGI) domains that minimize base excision repair of deaminated cytosines). rAPOBEC1-based CBEs have been used successfully in a wide variety of organisms and cell types to induce C to T changes in DNA2-10. Other cytosine deaminases such as human APOBEC3A11, 12 an engineered form of human APOBEC3A11, APOBEC3G3, CDA13, and AID3, 13-15 have also been used to create additional CBEs that function efficiently in human cells, hamster cells, yeast, rice, and tomato cells.
Described herein are cytosine base editors that have reduced RNA editing activity. The base editors comprise a cytoside deaminase, e.g., an APOBEC1, bearing one or more mutations that decrease RNA editing activity while preserving DNA editing activity, wherein the mutations are at amino acid positions that correspond to residues P29, R33, K34, E181, and/or L182 of rat apolipoprotein B mRNA editing enzyme, catalytic polypeptide-like 1 (rAPOBEC1, SEQ ID NO:67), and a programmable DNA binding domain, and optionally further comprise a uracil glycosylase inhibitor (UGI).
In some embodiments, the cytosine deaminase comprises one or more mutations corresponding to APOBEC1 mutations at positions: P29F, P29T, R33A, K34A, R33A+K34A (double mutant), E181Q and/or L182A of SEQ ID NO:67 (rAPOBEC1, Rattus norvegicus APOBEC1) or an orthologue thereof. In some embodiments, the cytosine deaminase comprises one or more mutations corresponding to a mutation listed in table D.
In some embodiments, the base editors further comprise one or more mutations at APOBEC1 residues corresponding to E24, V25; R118, Y120, H121, R126; W224-K229; P168-1186; L173+L180; R15, R16, R17, to K15-17 & A15-17; Deletion E181-L210; P190+P191; Deletion L210-K229 (C-terminal); and/or Deletion S2-L14 (N-terminal) of SEQ ID NO:67 or an orthologue thereof.
In some embodiments, the cytosine base editor comprise a linker between the cytosine deaminase and the programmable DNA binding domain.
In some embodiments, the programmable DNA binding domain is selected from the group consisting of engineered C2H2 zinc-fingers, transcription activator effector-like effectors (TALEs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) Cas RNA-guided nucleases (RGNs) and variants thereof. In some embodiments, the programmable DNA binding domain is an engineered C2H2 zinc-finger or TALEs that directs the base editor to edit a target sequence in Table E.
In some embodiments, the CRISPR RGN is an ssDNA nickase or is catalytically inactive, e.g., a Cas9 or Cas12a that is catalytically inactive or has ssDNA nickases activity.
Also provided herein are base editing systems comprising (i) the cytosine base editors described herein, wherein the programmable DNA binding domain is a CRISPR Cas RGN or a variant thereof; and (ii) at least one guide RNA compatible with the base editor that directs the base editor to a target sequence. In some embodiments, the guide RNA targets a sequence shown in Table E.
Also provided are isolated nucleic acids encoding the cytosine base editors; vectors comprising the isolated nucleic acids; and isolated host cells, preferably mammalian host cells, comprising the nucleic acids. In some embodiments, the isolated host cell expresses a cytosine base editor.
Further, provided herein are methods for deaminating a selected cytidine in a nucleic acid, the method comprising contacting the nucleic acid with a cytosine base editor or base editing system as described herein. In some embodiments, the method includes the use of a guide RNA that targets a sequence shown in Table E. In some embodiments, the nucleic acid is in a living cell. In some embodiments, the nucleic acid is genomic DNA, e.g., in a living cell.
In some embodiments, the cell is in a mammal, e.g., a human or a veterinary subject (e.g., dog, cat, cow, horse, pig, sheep, or goat).
Also provided are compositions comprising a purified cytosine base editor or base editing system as described herein. In some embodiments, the composition comprises one or more ribonucleoprotein (RNP) complexes.
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.
troglodytes]
mulatta]
Although CBEs can efficiently induce C to T edits in DNA, the rAPOBEC1 protein (present in the most commonly used CBEs) was originally actually discovered based on its ability to induce C to U edits in RNA (
Thus, described herein are variants of APOBEC1 bearing mutations that exhibit reduced RNA editing (RRE) activities (also referred to herein as SElective Curbing of Unwanted RNA Editing (SECURE) variants) while maintaining DNA deamination activities, optionally fused to an engineered DNA binding domain such as a CRISPR-Cas nuclease modified to either be a nickase or catalytically inactive, to enable DNA base editing with reduced RNA mutation profiles.
In some embodiments, the APOBEC is APOBEC1 from rat, or from a different species, e.g., a different mammalian species such as human. The APOBEC family members have high sequence homology.
Thus described herein are base editors comprising cytosine deaminases with mutations that reduce undesirable RNA editing activity. In general, these base editors have mutations as described herein. In some embodiments, they have mutations that correspond to residues P29, R33, K34, E181, and/or L182 of rAPOBEC1. Alternatively, or in addition, they may have mutations at E24, V25; R118, Y120, H121, R126; W224-K229; P168-1186; L173+L180; R15, R16, R17, to K15-17&A15-17; Deletion E181-L210; P190+P191; Deletion L210-K229 (C-terminal); and/or Deletion S2-L14 (N-terminal). In preferred embodiments, the mutations correspond to P29F, P29T, R33A, K34A, R33+K34A (double mutant), E181Q and/or L182A of SEQ ID NO:67 (rat APOBEC1).
The wild type sequence of rAPOBEC1, also known as C->U-editing enzyme APOBEC-1 [Rattus norvegicus], and available in GenBank at NP_037039.1, is as follows:
Other exemplary cytosine deaminase sequences are shown in
In some embodiments, the cytosine deaminase is evoFERNY (Thuronyi et al., Nature Biotechnology volume 37, pages 1070-1079 (2019)) and the R33 equivalent mutation can be made at R12.
In some embodiments, the cytosine deaminase is evoAPOBEC1 (Thuronyi et al., Nature Biotechnology volume 37, pages 1070-1079 (2019)) and the R33 and/or R34 equivalent mutations can be made at R33/R34.
In some embodiments, the base editors do not include catalytically dead cytosine deaminase variants, e.g. E63A, W90S, and C93A. (Harris et al, 2002, PMID: 12453430).
In some embodiments, the base editors include programmable DNA binding domains such as engineered C2H2 zinc-fingers, transcription activator effector-like effectors (TALEs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) Cas RNA-guided nucleases (RGNs) and their variants, including ssDNA nickases (nCas9) or their analogs and catalytically inactive dead Cas9 (dCas9) and its analogs, and any engineered protospacer-adjacent motif (PAM) or high-fidelity variants (e.g., as shown inTable D). A programmable DNA binding domain is one that can be engineered to bind to a selected target sequence.
CRISPR-Cas Nucleases
Although herein we refer to Cas9, in general any Cas9-like nickase could be used (including the related Cpf1/Cas12a enzyme classes), unless specifically indicated.
S. pyogenes Cas9
S. aureus Cas9
S. thermophilus
S. pasteurianus
C. jejuni Cas9
F. novicida Cas9
P. lavamentivorans
C. lari Cas9
Pasteurella
multocida Cas9
F. novicida Cpf1
M. bovoculi Cpf1
L. bacterium N2006
These orthologs, and mutants and variants thereof as known in the art, can be used in any of the fusion proteins described herein. See, e.g., WO 2017/040348 (which describes variants of SaCas9 and SpCas 9 with increased specificity) and WO 2016/141224 (which describes variants of SaCas9 and SpCas 9 with altered PAM specificity).
The Cas9 nuclease from S. pyogenes (hereafter simply Cas9) can be guided via simple base pair complementarity between 17-20 nucleotides of an engineered guide RNA (gRNA), e.g., a single guide RNA or crRNA/tracrRNA pair, and the complementary strand of a target genomic DNA sequence of interest that lies next to a protospacer adjacent motif (PAM), e.g., a PAM matching the sequence NGG or NAG (Shen et al., Cell Res (2013); Dicarlo et al., Nucleic Acids Res (2013); Jiang et al., Nat Biotechnol 31, 233-239 (2013); Jinek et al., Elife 2, e00471 (2013); Hwang et al., Nat Biotechnol 31, 227-229 (2013); Cong et al., Science 339, 819-823 (2013); Mali et al., Science 339, 823-826 (2013c); Cho et al., Nat Biotechnol 31, 230-232 (2013); Jinek et al., Science 337, 816-821 (2012)). The engineered CRISPR from Prevotella and Francisella 1 (Cpf1, also known as Cas12a) nuclease can also be used, e.g., as described in Zetsche et al., Cell 163, 759-771 (2015); Schunder et al., Int J Med Microbiol 303, 51-60 (2013); Makarova et al., Nat Rev Microbiol 13, 722-736 (2015); Fagerlund et al., Genome Biol 16, 251 (2015). Unlike SpCas9, Cpf1/Cas12a requires only a single 42-nt crRNA, which has 23 nt at its 3′ end that are complementary to the protospacer of the target DNA sequence (Zetsche et al., 2015). Furthermore, whereas SpCas9 recognizes an NGG PAM sequence that is 3′ of the protospacer, AsCpf1 and LbCp1 recognize TTTN PAMs that are found 5′ of the protospacer (Id.).
In some embodiments, the present system utilizes a wild type or variant Cas9 protein from S. pyogenes or Staphylococcus aureus, or a wild type or variant Cpf1 protein from Acidaminococcus sp. BV3L6 or Lachnospiraceae bacterium ND2006 either as encoded in bacteria or codon-optimized for expression in mammalian cells and/or modified in its PAM recognition specificity and/or its genome-wide specificity. A number of variants have been described; see, e.g., WO 2016/141224, PCT/US2016/049147, Kleinstiver et al., Nat Biotechnol. 2016 August; 34(8):869-74; Tsai and Joung, Nat Rev Genet. 2016 May; 17(5):300-12; Kleinstiver et al., Nature. 2016 Jan. 28; 529(7587):490-5; Shmakov et al., Mol Cell. 2015 Nov. 5; 60(3):385-97; Kleinstiver et al., Nat Biotechnol. 2015 December; 33(12):1293-1298; Dahlman et al., Nat Biotechnol. 2015 November; 33(11):1159-61; Kleinstiver et al., Nature. 2015 Jul. 23; 523(7561):481-5; Wyvekens et al., Hum Gene Ther. 2015 July; 26(7):425-31; Hwang et al., Methods Mol Biol. 2015; 1311:317-34; Osborn et al., Hum Gene Ther. 2015 February; 26(2):114-26; Konermann et al., Nature. 2015 Jan. 29; 517(7536):583-8; Fu et al., Methods Enzymol. 2014; 546:21-45; and Tsai et al., Nat Biotechnol. 2014 June; 32(6):569-76, inter alia. Concerning rAPOBEC1 itself, a number of variants have been described, e.g. Chen et al, RNA. 2010 May; 16(5):1040-52; Chester et al, EMBO J. 2003 Aug. 1; 22(15):3971-82; Teng et al, J Lipid Res. 1999 April; 40(4):623-35.; Navaratnam et al, Cell. 1995 Apr. 21; 81(2):187-95; MacGinnitie et al, J Biol Chem. 1995 Jun. 16; 270(24):14768-75; Yamanaka et al, J Biol Chem. 1994 Aug. 26; 269(34):21725-34. The guide RNA is expressed or present in the cell together with the Cas9 or Cpf1. Either the guide RNA or the nuclease, or both, can be expressed transiently or stably in the cell or introduced as a purified protein or nucleic acid.
In some embodiments, the Cas9 also includes one of the following mutations, which reduce nuclease activity of the Cas9; e.g., for SpCas9, mutations at D10A or H840A (which creates a single-strand nickase).
In some embodiments, the SpCas9 variants also include mutations at one of each of the two sets of the following amino acid positions, which together destroy the nuclease activity of the Cas9: D10, E762, D839, H983, or D986 and H840 or N863, e.g., D10A/D10N and H840A/H840N/H840Y, to render the nuclease portion of the protein catalytically inactive; substitutions at these positions could be alanine (as they are in Nishimasu al., Cell 156, 935-949 (2014)), or other residues, e.g., glutamine, asparagine, tyrosine, serine, or aspartate, e.g., E762Q, H983N, H983Y, D986N, N863D, N863S, or N863H (see WO 2014/152432).
In some embodiments, the Cas9 is fused to one or more Uracil glycosylase inhibitor (UGI) protein sequences; an exemplary UGI sequence is as follows: TNLSDIIEKETGKQLVIQESILMLPEEVEEVIGNKPESDILVHTAYDESTDENVMLLTSD APEYKPWALVIQDSNGENKIKML (SEQ ID NO:134; Uniprot: P14739). Typically, the UGIs are at the C-terminus of a BE fusion protein, but can also be positioned at the N-terminus, or between the DNA binding domain and the deaminase domain. Linkers as known in the art can be used to separate domains.
S. pyogenes
S. pyogenes
S. pyogenes
S. pyogenes
S. pyogenes
S. pyogenes
S. pyogenes
S. pyogenes
S. pyogenes
S. aureus Cas9
TAL Effector Repeat Arrays
Transcription activator like effectors (TALEs) of plant pathogenic bacteria in the genus Xanthomonas play important roles in disease, or trigger defense, by binding host DNA and activating effector-specific host genes. Specificity depends on an effector-variable number of imperfect, typically ˜33-35 amino acid repeats. Polymorphisms are present primarily at repeat positions 12 and 13, which are referred to herein as the repeat variable-diresidue (RVD). The RVDs of TAL effectors correspond to the nucleotides in their target sites in a direct, linear fashion, one RVD to one nucleotide, with some degeneracy and no apparent context dependence. In some embodiments, the polymorphic region that grants nucleotide specificity may be expressed as a tri residue or triplet.
Each DNA binding repeat can include a RVD that determines recognition of a base pair in the target DNA sequence, wherein each DNA binding repeat is responsible for recognizing one base pair in the target DNA sequence. In some embodiments, the RVD can comprise one or more of: HA for recognizing C; ND for recognizing C; HI for recognizing C; HN for recognizing G; NA for recognizing G; SN for recognizing G or A; YG for recognizing T; and NK for recognizing G, and one or more of: HD for recognizing C; NG for recognizing T; NI for recognizing A; NN for recognizing G or A; NS for recognizing A or C or G or T; N* for recognizing C or T, wherein * represents a gap in the second position of the RVD; HG for recognizing T; H* for recognizing T, wherein * represents a gap in the second position of the RVD; and IG for recognizing T.
TALE proteins may be useful in research and biotechnology as targeted chimeric nucleases that can facilitate homologous recombination in genome engineering (e.g., to add or enhance traits useful for biofuels or biorenewables in plants). These proteins also may be useful as, for example, transcription factors, and especially for therapeutic applications requiring a very high level of specificity such as therapeutics against pathogens (e.g., viruses) as non-limiting examples.
Methods for generating engineered TALE arrays are known in the art, see, e.g., the fast ligation-based automatable solid-phase high-throughput (FLASH) system described in U.S. Ser. No. 61/610,212, and Reyon et al., Nature Biotechnology 30,460-465 (2012); as well as the methods described in Bogdanove & Voytas, Science 333, 1843-1846 (2011); Bogdanove et al., Curr Opin Plant Biol 13, 394-401 (2010); Scholze & Boch, J. Curr Opin Microbiol (2011); Boch et al., Science 326, 1509-1512 (2009); Moscou & Bogdanove, Science 326, 1501 (2009); Miller et al., Nat Biotechnol 29, 143-148 (2011); Morbitzer et al., T. Proc Natl Acad Sci USA 107, 21617-21622 (2010); Morbitzer et al., Nucleic Acids Res 39, 5790-5799 (2011); Zhang et al., Nat Biotechnol 29, 149-153 (2011); Geissler et al., PLoS ONE 6, e19509 (2011); Weber et al., PLoS ONE 6, e19722 (2011); Christian et al., Genetics 186, 757-761 (2010); Li et al., Nucleic Acids Res 39, 359-372 (2011); Mahfouz et al., Proc Natl Acad Sci USA 108, 2623-2628 (2011); Mussolino et al., Nucleic Acids Res (2011); Li et al., Nucleic Acids Res 39, 6315-6325 (2011); Cermak et al., Nucleic Acids Res 39, e82 (2011); Wood et al., Science 333, 307 (2011); Hockemeye et al. Nat Biotechnol 29, 731-734 (2011); Tesson et al., Nat Biotechnol 29, 695-696 (2011); Sander et al., Nat Biotechnol 29, 697-698 (2011); Huang et al., Nat Biotechnol 29, 699-700 (2011); and Zhang et al., Nat Biotechnol 29, 149-153 (2011); all of which are incorporated herein by reference in their entirety.
Zinc Fingers
Zinc finger (ZF) proteins are DNA-binding proteins that contain one or more zinc fingers, independently folded zinc-containing mini-domains, the structure of which is well known in the art and defined in, for example, Miller et al., 1985, EMBO J., 4:1609; Berg, 1988, Proc. Natl. Acad. Sci. USA, 85:99; Lee et al., 1989, Science. 245:635; and Klug, 1993, Gene, 135:83. Crystal structures of the zinc finger protein Zif268 and its variants bound to DNA show a semi-conserved pattern of interactions, in which typically three amino acids from the alpha-helix of the zinc finger contact three adjacent base pairs or a “subsite” in the DNA (Pavletich et al., 1991, Science, 252:809; Elrod-Erickson et al., 1998, Structure, 6:451). Thus, the crystal structure of Zif268 suggested that zinc finger DNA-binding domains might function in a modular manner with a one-to-one interaction between a zinc finger and a three-base-pair “subsite” in the DNA sequence. In naturally occurring zinc finger transcription factors, multiple zinc fingers are typically linked together in a tandem array to achieve sequence-specific recognition of a contiguous DNA sequence (Klug, 1993, Gene 135:83).
Multiple studies have shown that it is possible to artificially engineer the DNA binding characteristics of individual zinc fingers by randomizing the amino acids at the alpha-helical positions involved in DNA binding and using selection methodologies such as phage display to identify desired variants capable of binding to DNA target sites of interest (Rebar et al., 1994, Science, 263:671; Choo et al., 1994 Proc. Natl. Acad. Sci. USA, 91:11163; Jamieson et al., 1994, Biochemistry 33:5689; Wu et al., 1995 Proc. Natl. Acad. Sci. USA, 92: 344). Such recombinant zinc finger proteins can be fused to functional domains, such as transcriptional activators, transcriptional repressors, methylation domains, and nucleases to regulate gene expression, alter DNA methylation, and introduce targeted alterations into genomes of model organisms, plants, and human cells (Carroll, 2008, Gene Ther., 15:1463-68; Cathomen, 2008, Mol. Ther., 16:1200-07; Wu et al., 2007, Cell. Mol. Life Sci., 64:2933-44).
One existing method for engineering zinc finger arrays, known as “modular assembly,” advocates the simple joining together of pre-selected zinc finger modules into arrays (Segal et al., 2003, Biochemistry, 42:2137-48; Beerli et al., 2002, Nat. Biotechnol., 20:135-141; Mandell et al., 2006, Nucleic Acids Res., 34:W516-523; Carroll et al., 2006, Nat. Protoc. 1:1329-41; Liu et al., 2002, J. Biol. Chem., 277:3850-56; Bae et al., 2003, Nat. Biotechnol., 21:275-280; Wright et al., 2006, Nat. Protoc., 1:1637-52). Although straightforward enough to be practiced by any researcher, recent reports have demonstrated a high failure rate for this method, particularly in the context of zinc finger nucleases (Ramirez et al., 2008, Nat. Methods, 5:374-375; Kim et al., 2009, Genome Res. 19:1279-88), a limitation that typically necessitates the construction and cell-based testing of very large numbers of zinc finger proteins for any given target gene (Kim et al., 2009, Genome Res. 19:1279-88).
Combinatorial selection-based methods that identify zinc finger arrays from randomized libraries have been shown to have higher success rates than modular assembly (Maeder et al., 2008, Mol. Cell, 31:294-301; Joung et al., 2010, Nat. Methods, 7:91-92; Isalan et al., 2001, Nat. Biotechnol., 19:656-660). In preferred embodiments, the zinc finger arrays are described in, or are generated as described in, WO 2011/017293 and WO 2004/099366. Additional suitable zinc finger DBDs are described in U.S. Pat. Nos. 6,511,808, 6,013,453, 6,007,988, and 6,503,717 and U.S. patent application 2002/0160940.
In some embodiments, the components of the fusion proteins are at least 80%, e.g., at least 85%, 90%, 95%, 97%, or 99% identical to the amino acid sequence of a exemplary sequence (e.g., as provided herein), e.g., have differences at up to 1%, 2%, 5%, 10%, 15%, or 20% of the residues of the exemplary sequence replaced, e.g., with conservative mutations, e.g., including or in addition to the mutations described herein. In preferred embodiments, the variant retains a desired activity of the parent, e.g., deaminase activity, and/or the ability to interact with a guide RNA and/or target DNA, optionally with improved specificity or altered substrate specificity.
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-homologous 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 “homology”). 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. Percent identity between two polypeptides or nucleic acid sequences is determined in various ways that are within the skill in the art, for instance, using publicly available computer software such as Smith Waterman Alignment (Smith, T. F. and M. S. Waterman (1981) J Mol Biol 147:195-7); “BestFit” (Smith and Waterman, Advances in Applied Mathematics, 482-489 (1981)) as incorporated into GeneMatcher PIus™ Schwarz and Dayhof (1979) Atlas of Protein Sequence and Structure, Dayhof, M. O., Ed, pp 353-358; BLAST program (Basic Local Alignment Search Tool; (Altschul, S. F., W. Gish, et al. (1990) J Mol Biol 215: 403-10), BLAST-2, BLAST-P, BLAST-N, BLAST-X, WU-BLAST-2, ALIGN, ALIGN-2, CLUSTAL, or Megalign (DNASTAR) software. In addition, those skilled in the art can determine appropriate parameters for measuring alignment, including any algorithms needed to achieve maximal alignment over the length of the sequences being compared. In general, for proteins or nucleic acids, the length of comparison can be any length, up to and including full length (e.g., 5%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 95%, or 100%). For purposes of the present compositions and methods, at least 80% of the full length of the sequence is aligned.
For purposes of the present disclosure, 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.
Conservative substitutions typically include substitutions within the following groups: glycine, alanine; valine, isoleucine, leucine; aspartic acid, glutamic acid, asparagine, glutamine; serine, threonine; lysine, arginine; and phenylalanine, tyrosine.
Also provided herein are isolated nucleic acids encoding the base editor fusion proteins, vectors comprising the isolated nucleic acids, optionally operably linked to one or more regulatory domains for expressing the variant proteins, and host cells, e.g., mammalian host cells, comprising the nucleic acids, and optionally expressing the variant proteins. In some embodiments, the host cells are stem cells, e.g., hematopoietic stem cells.
In some embodiments, the fusion proteins include a linker between the DNA binding domain (e.g., ZFN, TALE, or nCas9) and the BE domains. Linkers that can be used in these fusion proteins (or between fusion proteins in a concatenated structure) can include any sequence that does not interfere with the function of the fusion proteins. In preferred embodiments, the linkers are short, e.g., 2-20 amino acids, and are typically flexible (i.e., comprising amino acids with a high degree of freedom such as glycine, alanine, and serine). In some embodiments, the linker comprises one or more units consisting of GGGS (SEQ ID NO:135) or GGGGS (SEQ ID NO:136), e.g., two, three, four, or more repeats of the GGGS (SEQ ID NO:135) or GGGGS (SEQ ID NO:136) unit. Other linker sequences can also be used.
In some embodiments, the deaminase fusion protein includes a cell-penetrating peptide sequence that facilitates delivery to the intracellular space, e.g., HIV-derived TAT peptide, penetratins, transportans, or hCT derived cell-penetrating peptides, see, e.g., Caron et al., (2001) Mol Ther. 3(3):310-8; Langel, Cell-Penetrating Peptides: Processes and Applications (CRC Press, Boca Raton Fla. 2002); El-Andaloussi et al., (2005) Curr Pharm Des. 11(28):3597-611; and Deshayes et al., (2005) Cell Mol Life Sci. 62(16):1839-49.
Cell penetrating peptides (CPPs) are short peptides that facilitate the movement of a wide range of biomolecules across the cell membrane into the cytoplasm or other organelles, e.g. the mitochondria and the nucleus. Examples of molecules that can be delivered by CPPs include therapeutic drugs, plasmid DNA, oligonucleotides, siRNA, peptide-nucleic acid (PNA), proteins, peptides, nanoparticles, and liposomes. CPPs are generally 30 amino acids or less, are derived from naturally or non-naturally occurring protein or chimeric sequences, and contain either a high relative abundance of positively charged amino acids, e.g. lysine or arginine, or an alternating pattern of polar and non-polar amino acids. CPPs that are commonly used in the art include Tat (Frankel et al., (1988) Cell. 55:1189-1193, Vives et al., (1997) J. Biol. Chem. 272:16010-16017), penetratin (Derossi et al., (1994) J. Biol. Chem. 269:10444-10450), polyarginine peptide sequences (Wender et al., (2000) Proc. Natl. Acad. Sci. USA 97:13003-13008, Futaki et al., (2001) J. Biol. Chem. 276:5836-5840), and transportan (Pooga et al., (1998) Nat. Biotechnol. 16:857-861).
CPPs can be linked with their cargo through covalent or non-covalent strategies. Methods for covalently joining a CPP and its cargo are known in the art, e.g. chemical cross-linking (Stetsenko et al., (2000) J. Org. Chem. 65:4900-4909, Gait et al. (2003) Cell. Mol. Life. Sci. 60:844-853) or cloning a fusion protein (Nagahara et al., (1998) Nat. Med. 4:1449-1453). Non-covalent coupling between the cargo and short amphipathic CPPs comprising polar and non-polar domains is established through electrostatic and hydrophobic interactions.
CPPs have been utilized in the art to deliver potentially therapeutic biomolecules into cells. Examples include cyclosporine linked to polyarginine for immunosuppression (Rothbard et al., (2000) Nature Medicine 6(11):1253-1257), siRNA against cyclin B1 linked to a CPP called MPG for inhibiting tumorigenesis (Crombez et al., (2007) Biochem Soc. Trans. 35:44-46), tumor suppressor p53 peptides linked to CPPs to reduce cancer cell growth (Takenobu et al., (2002) Mol. Cancer Ther. 1(12):1043-1049, Snyder et al., (2004) PLoS Biol. 2:E36), and dominant negative forms of Ras or phosphoinositol 3 kinase (P13K) fused to Tat to treat asthma (Myou et al., (2003) J. Immunol. 171:4399-4405).
CPPs have been utilized in the art to transport contrast agents into cells for imaging and biosensing applications. For example, green fluorescent protein (GFP) attached to Tat has been used to label cancer cells (Shokolenko et al., (2005) DNA Repair 4(4):511-518). Tat conjugated to quantum dots have been used to successfully cross the blood-brain barrier for visualization of the rat brain (Santra et al., (2005) Chem. Commun. 3144-3146). CPPs have also been combined with magnetic resonance imaging techniques for cell imaging (Liu et al., (2006) Biochem. and Biophys. Res. Comm. 347(1):133-140). See also Ramsey and Flynn, Pharmacol Ther. 2015 Jul. 22. pii: S0163-7258(15)00141-2.
Alternatively or in addition, the deaminase fusion proteins can include a nuclear localization sequence, e.g., SV40 large T antigen NLS (PKKKRRV (SEQ ID NO:137)) and nucleoplasmin NLS (KRPAATKKAGQAKKKK (SEQ ID NO:138)). Other NLSs are known in the art; see, e.g., Cokol et al., EMBO Rep. 2000 Nov. 15; 1(5): 411-415; Freitas and Cunha, Curr Genomics. 2009 December; 10(8): 550-557.
In some embodiments, the deaminase fusion proteins include a moiety that has a high affinity for a ligand, for example GST, FLAG or hexahistidine sequences. Such affinity tags can facilitate the purification of recombinant deaminase fusion proteins.
The deaminase fusion proteins described herein can be used for altering the genome of a cell. The methods generally include expressing or contacting the deaminase fusion proteins in the cells; in versions using one or two Cas9s, the methods include using a guide RNA having a region complementary to a selected portion of the genome of the cell. Methods for selectively altering the genome of a cell are known in the art, see, e.g., U.S. Pat. No. 8,993,233; US 20140186958; U.S. Pat. No. 9,023,649; WO/2014/099744; WO 2014/089290; WO2014/144592; WO144288; WO2014/204578; WO2014/152432; WO2115/099850; U.S. Pat. No. 8,697,359; US20160024529; US20160024524; US20160024523; US20160024510; US20160017366; US20160017301; US20150376652; US20150356239; US20150315576; US20150291965; US20150252358; US20150247150; US20150232883; US20150232882; US20150203872; US20150191744; US20150184139; US20150176064; US20150167000; US20150166969; US20150159175; US20150159174; US20150093473; US20150079681; US20150067922; US20150056629; US20150044772; US20150024500; US20150024499; US20150020223; US20140356867; US20140295557; US20140273235; US20140273226; US20140273037; US20140189896; US20140113376; US20140093941; US20130330778; US20130288251; US20120088676; US20110300538; US20110236530; US20110217739; US20110002889; US20100076057; US20110189776; US20110223638; US20130130248; US20150050699; US20150071899; US20150050699; US20150045546; US20150031134; US20150024500; US20140377868; US20140357530; US20140349400; US20140335620; US20140335063; US20140315985; US20140310830; US20140310828; US20140309487; US20140304853; US20140298547; US20140295556; US20140294773; US20140287938; US20140273234; US20140273232; US20140273231; US20140273230; US20140271987; US20140256046; US20140248702; US20140242702; US20140242700; US20140242699; US20140242664; US20140234972; US20140227787; US20140212869; US20140201857; US20140199767; US20140189896; US20140186958; US20140186919; US20140186843; US20140179770; US20140179006; US20140170753; WO/2008/108989; WO/2010/054108; WO/2012/164565; WO/2013/098244; WO/2013/176772; US 20150071899; Makarova et al., “Evolution and classification of the CRISPR-Cas systems” 9(6) Nature Reviews Microbiology 467-477 (1-23) (June 2011); Wiedenheft et al., “RNA-guided genetic silencing systems in bacteria and archaea” 482 Nature 331-338 (Feb. 16, 2012); Gasiunas et al., “Cas9-crRNA ribonucleoprotein complex mediates specific DNA cleavage for adaptive immunity in bacteria” 109(39) Proceedings of the National Academy of Sciences USA E2579-E2586 (Sep. 4, 2012); Jinek et al., “A Programmable Dual-RNA-Guided DNA Endonuclease in Adaptive Bacterial Immunity” 337 Science 816-821 (Aug. 17, 2012); Carroll, “A CRISPR Approach to Gene Targeting” 20(9) Molecular Therapy 1658-1660 (September 2012); U.S. Appl. No. 61/652,086, filed May 25, 2012; Al-Attar et al., Clustered Regularly Interspaced Short Palindromic Repeats (CRISPRs): The Hallmark of an Ingenious Antiviral Defense Mechanism in Prokaryotes, Biol Chem. (2011) vol. 392, Issue 4, pp. 277-289; Hale et al., Essential Features and Rational Design of CRISPR RNAs That Function With the Cas RAMP Module Complex to Cleave RNAs, Molecular Cell, (2012) vol. 45, Issue 3, 292-302.
For methods in which the deaminase fusion proteins are delivered to cells, the proteins can be produced using any method known in the art, e.g., by in vitro translation, or expression in a suitable host cell from nucleic acid encoding the deaminase fusion protein; a number of methods are known in the art for producing proteins. For example, the proteins can be produced in and purified from yeast, E. CO/i, insect cell lines, plants, transgenic animals, or cultured mammalian cells; see, e.g., Palomares et al., “Production of Recombinant Proteins: Challenges and Solutions,” Methods Mol Biol. 2004; 267:15-52. In addition, the deaminase fusion proteins can be linked to a moiety that facilitates transfer into a cell, e.g., a lipid nanoparticle, optionally with a linker that is cleaved once the protein is inside the cell. See, e.g., LaFountaine et al., Int J Pharm. 2015 Aug. 13; 494(1):180-194.
To use the deaminase fusion proteins described herein, it may be desirable to express them from a nucleic acid that encodes them. This can be performed in a variety of ways. For example, the nucleic acid encoding the deaminase fusion can be cloned into an intermediate vector for transformation into prokaryotic or eukaryotic cells for replication and/or expression. Intermediate vectors are typically prokaryote vectors, e.g., plasmids, or shuttle vectors, or insect vectors, for storage or manipulation of the nucleic acid encoding the deaminase fusion for production of the deaminase fusion protein. The nucleic acid encoding the deaminase fusion protein can also be cloned into an expression vector, for administration to a plant cell, animal cell, preferably a mammalian cell or a human cell, fungal cell, bacterial cell, or protozoan cell.
To obtain expression, a sequence encoding a deaminase fusion protein is typically subcloned into an expression vector that contains a promoter to direct transcription. Suitable bacterial and eukaryotic promoters are well known in the art and described, e.g., in Sambrook et al., Molecular Cloning, A Laboratory Manual (3d ed. 2001); Kriegler, Gene Transfer and Expression: A Laboratory Manual (1990); and Current Protocols in Molecular Biology (Ausubel et al., eds., 2010). Bacterial expression systems for expressing the engineered protein are available in, e.g., E. coli, Bacillus sp., and Salmonella (Palva et al., 1983, Gene 22:229-235). Kits for such expression systems are commercially available. Eukaryotic expression systems for mammalian cells, yeast, and insect cells are well known in the art and are also commercially available.
The promoter used to direct expression of a nucleic acid depends on the particular application. For example, a strong constitutive promoter is typically used for expression and purification of fusion proteins. In contrast, when the deaminase fusion protein is to be administered in vivo for gene regulation, either a constitutive or an inducible promoter can be used, depending on the particular use of the deaminase fusion protein. In addition, a preferred promoter for administration of the deaminase fusion protein can be a weak promoter, such as HSV TK or a promoter having similar activity. The promoter can also include elements that are responsive to transactivation, e.g., hypoxia response elements, Gal4 response elements, lac repressor response element, and small molecule control systems such as tetracycline-regulated systems and the RU-486 system (see, e.g., Gossen & Bujard, 1992, Proc. Natl. Acad. Sci. USA, 89:5547; Oligino et al., 1998, Gene Ther., 5:491-496; Wang et al., 1997, Gene Ther., 4:432-441; Neering et al., 1996, Blood, 88:1147-55; and Rendahl et al., 1998, Nat. Biotechnol., 16:757-761).
In addition to the promoter, the expression vector typically contains a transcription unit or expression cassette that contains all the additional elements required for the expression of the nucleic acid in host cells, either prokaryotic or eukaryotic. A typical expression cassette thus contains a promoter operably linked, e.g., to the nucleic acid sequence encoding the deaminase fusion protein, and any signals required, e.g., for efficient polyadenylation of the transcript, transcriptional termination, ribosome binding sites, or translation termination. Additional elements of the cassette may include, e.g., enhancers, and heterologous spliced intronic signals.
The particular expression vector used to transport the genetic information into the cell is selected with regard to the intended use of the deaminase fusion protein, e.g., expression in plants, animals, bacteria, fungus, protozoa, etc. Standard bacterial expression vectors include plasmids such as pBR322 based plasmids, pSKF, pET23D, and commercially available tag-fusion expression systems such as GST and LacZ.
Expression vectors containing regulatory elements from eukaryotic viruses are often used in eukaryotic expression vectors, e.g., SV40 vectors, papilloma virus vectors, and vectors derived from Epstein-Barr virus. Other exemplary eukaryotic vectors include pMSG, pAV009/A+, pMTO10/A+, pMAMneo-5, baculovirus pDSVE, and any other vector allowing expression of proteins under the direction of the SV40 early promoter, SV40 late promoter, metallothionein promoter, murine mammary tumor virus promoter, Rous sarcoma virus promoter, polyhedrin promoter, or other promoters shown effective for expression in eukaryotic cells.
The vectors for expressing the deaminase fusion protein can include RNA Pol III promoters to drive expression of the guide RNAs, e.g., the H1, U6 or 7SK promoters. These human promoters allow for expression of deaminase fusion protein in mammalian cells following plasmid transfection.
Some expression systems have markers for selection of stably transfected cell lines such as thymidine kinase, hygromycin B phosphotransferase, and dihydrofolate reductase. High yield expression systems are also suitable, such as using a baculovirus vector in insect cells, with the gRNA encoding sequence under the direction of the polyhedrin promoter or other strong baculovirus promoters.
The elements that are typically included in expression vectors also include a replicon that functions in E. coli, a gene encoding antibiotic resistance to permit selection of bacteria that harbor recombinant plasmids, and unique restriction sites in nonessential regions of the plasmid to allow insertion of recombinant sequences.
Standard transfection methods are used to produce bacterial, mammalian, yeast or insect cell lines that express large quantities of protein, which are then purified using standard techniques (see, e.g., Colley et al., 1989, J. Biol. Chem., 264:17619-22; Guide to Protein Purification, in Methods in Enzymology, vol. 182 (Deutscher, ed., 1990)). Transformation of eukaryotic and prokaryotic cells are performed according to standard techniques (see, e.g., Morrison, 1977, J. Bacteriol. 132:349-351; Clark-Curtiss & Curtiss, Methods in Enzymology 101:347-362 (Wu et al., eds, 1983).
Any of the known procedures for introducing foreign nucleotide sequences into host cells may be used. These include the use of calcium phosphate transfection, polybrene, protoplast fusion, electroporation, nucleofection, liposomes, microinjection, naked DNA, plasmid vectors, viral vectors, both episomal and integrative, and any of the other well-known methods for introducing cloned genomic DNA, cDNA, synthetic DNA or other foreign genetic material into a host cell (see, e.g., Sambrook et al., supra). It is only necessary that the particular genetic engineering procedure used be capable of successfully introducing at least one gene into the host cell capable of expressing the deaminase fusion protein.
In methods wherein the fusion proteins include a Cas9 domain, the methods also include delivering at least one gRNA that interacts with the Cas9, or a nucleic acid that encodes a gRNA.
Alternatively, the methods can include delivering the deaminase fusion protein and guide RNA together, e.g., as a complex. For example, the deaminase fusion protein and gRNA can be can be overexpressed in a host cell and purified, then complexed with the guide RNA (e.g., in a test tube) to form a ribonucleoprotein (RNP), and delivered to cells. In some embodiments, the deaminase fusion protein can be expressed in and purified from bacteria through the use of bacterial expression plasmids. For example, His-tagged deaminase fusion protein can be expressed in bacterial cells and then purified using nickel affinity chromatography. The use of RNPs circumvents the necessity of delivering plasmid DNAs encoding the nuclease or the guide, or encoding the nuclease as an mRNA. RNP delivery may also improve specificity, presumably because the half-life of the RNP is shorter and there's no persistent expression of the nuclease and guide (as you′d get from a plasmid). The RNPs can be delivered to the cells in vivo or in vitro, e.g., using lipid-mediated transfection or electroporation. See, e.g., Liang et al. “Rapid and highly efficient mammalian cell engineering via Cas9 protein transfection.” Journal of biotechnology 208 (2015): 44-53; Zuris, John A., et al. “Cationic lipid-mediated delivery of proteins enables efficient protein-based genome editing in vitro and in vivo.” Nature biotechnology 33.1 (2015): 73-80; Kim et al. “Highly efficient RNA-guided genome editing in human cells via delivery of purified Cas9 ribonucleoproteins.” Genome research 24.6 (2014): 1012-1019.
The present invention also includes the vectors and cells comprising the vectors, as well as kits comprising the proteins and nucleic acids described herein, e.g., for use in a method described herein.
Methods of Use
The base editors described herein can be used to deaminate a selected cytosine in a nucleic acid sequence, e.g., in a cell, e.g., a cell in an animal (e.g., a mammal such as a human or veterinary subject), or a synthetic nucleic acid substrate. The methods include contacting the nucleic acid with a base editor as described herein. Where the base editor includes a CRISPR Cas9 or Cas12a protein, the methods further include the use of one or more guide RNAs that direct binding of the base editor to a sequence to be deaminated.
For example, the base editors described herein can be used for in vitro, in vivo or in situ directed evolution, e.g., to engineer polypeptides or proteins based on a synthetic selection framework, e.g. antibiotic resistance in E. coli or resistance to anti-cancer therapeutics being assayed in mammalian cells (e.g. CRISPR-X Hess et al, PMID: 27798611 or BE-plus systems Jiang et al, PMID: 29875396).
In addition, the base editors can be used to base-edit a therapeutically relevant sequence, to treat a subject. Table E provides a list of disease-associated gene variants that could be base-edited therapeutically with an NGG PAM positioned appropriately. See, e.g., Komor et al, Nature 2016).
The invention is further described in the following examples, which do not limit the scope of the invention described in the claims.
Methods
The following materials and methods were used in the Examples set forth below.
Molecular Cloning
Expression plasmids were constructed by selectively amplifying desired DNA sequences using the PCR method such that they had significant overlapping ends and using isothermal assembly (or “Gibson Assembly”, NEB) to assemble them in the desired order in a CAG or CMV expression vectors. PCR was conducted using Phusion HF polymerase (NEB). Cas9 gRNAs were cloned into the pUC19-based entry vector BPK1520 (via BsmBl) under control of a U6 promoter.
Cell Culture and Transfections
HEK293T cells (CRL-3216, obtained from ATCC) were grown in culture using media consisting of Advanced Dulbeccos Modified Medium (Gibco) supplemented with 10% FBS (Gibco) and 1% penicillin-streptomycin solution (Gibco). Cells were passaged at ˜80% confluency every 2-3 days to maintain an actively growing population and avoid anoxic conditions. HepG2 cells (HB80-65, obtained from ATCC) were grown in Eagle's Minimum Essential Medium (ATCC) supplemented with 10% FBS and 0.5% penicillin-streptomycin solution (Gibco). Cells were passaged at ˜80% confluency every 4 days. Both cell lines were used for experiments until passage 20 for HEK293T and passage 12 for HepG2. Cells were tested for mycoplasma bi-weekly.
For sorting experiments, transfections with 50 ug of transfection quality DNA (Qiagen Maxiprep) encoding desired BE3-P2A-EGFP fusion proteins or controls and gRNAs (75:25%) were conducted by seeding 6×106 HEK293T or 15×106 HepG2 into TC-treated 150 mm plates 18-24 h prior to transfection to yield ˜80% confluency on the day of transfection. Cells were transfected at 60-80% confluency using TransIT-293 (HEK293T, Mirus) or tranfeX (HepG2, ATCC) reagents according to the manufacturers' protocols. To ensure maximal correlation of negative controls to BE overexpression, cells of the same passage were transfected with nCas9-UGI-NLS (negative control) and base editors in parallel. RNA and gDNA was harvested after cell sorting. For experiments validating DNA on-target activity of SECURE-BE variants, 1.5×104 HEK293T cells were seeded into the wells of a 96-well plate and transfected 18-24 h after seeding with 220 ng DNA (BE3/nCas9-UGI:gRNA ration of 75:25%). In this context, gDNA was harvested 72 h post-transfection.
FACS & RNA/DNA Harvest
Sorting of negative control and BE expressing cells as well as RNA/DNA harvest were carried out on the same day. Cells were sorted on an BD FACSARIAII 36-40 h after transfection. We gated on the cell population on forward/sideward scatter after exclusion of doublets. We then sorted all GFP-positive cells and/or top 5% of cells with the highest FITC signal into pre-chilled 100% FBS and 5% of mean fluorescence intensity (MFI)-matched cells for nCas9-UGI negative controls, matching the MFI/GeoMean of top 5% of BE3-transfected cells. We used MFI-matching for these controls, as the nCas9-UGI-P2A-EGFP plasmid is smaller than BE3-P2A-EGFP—due to the lack of rAPOBEC1—and thus yields higher transfection efficiency and overall higher FITC signal. After sorting, cells were spun down, lysed using DNA lysis buffer (Laird et al, 1991) with DTT and Proteinase K or RNA lysis buffer (Macherey-Nagel). gDNA was extracted using magnetic beads (made from FisherSci Sera-Mag SpeedBeads Carboxyl Magnetic Beads, hydrophobic according to Rohland & Reich, 2012), after over-night lysis. RNA was extracted with Macherey-Nagel's NucleoSpin RNA Plus kit.
High-throughput Amplicon Sequencing, RT-PCR & Base Editing Data Analysis
Target site genomic DNA was amplified using gene-specific DNA primers flanking desired target sequence. These primers included illumina-compatible adapter-flaps. The amplicons were molecularly indexed with NEBNext Dual Index Primers (NEB) or index primers with the same or similar sequence ordered from IDT. Samples were combined into libraries and sequenced on the Illumina MiSeq machine using the MiSeq Reagent Kit v2 or Micro Kit v2 (Illumina). Sequencing results were analyzed using a batch version of the software CRISPResso 2.0 beta (crispresso.rocks). Reverse transcription was performed using the High Capacity RNA-to-cDNA kit (Thermo Fisher) following the manufacturer's instructions. Amplicon PCR and library preparation for Next-Generation Sequencing (NGS) off of cDNA was done as described above for gDNA (e.g. for the apoB amplicon around C6666). If possible, we used exon-exon junction spanning primers to exclude amplification of gDNA traces.
RNA-Seq and Single Nucleotide Variant Calling
RNA library preparation was performed using Illumina's TruSeq Stranded Total RNA Gold Kit with initial input of 500 ng of extracted RNA per sample, using SuperScript III for first-strand synthesis (Thermo Fisher). rRNA depletion was confirmed during library preparation on a High Resolution QIAxcel (Qiagen) automated electrophoresis device and/or by fluorometric quantitation using the Qubit HS RNA kit before and after depletion (Thermo Fisher). For indexing, we used IDT-Illumina Unique Dual Indeces (Illumina). Libraries were pooled based on qPCR quantification (NEBNext Library Quant Kit for Illumina) and loaded onto a NextSeq (at MGH Cancer Center, PE 2×150, 500/550 MidOutput Cartridge) or HiSeq2500 in High Output mode (Broad Institute, PE 2×76). Illumina fastq sequencing reads were aligned to the human hg38 reference genome with STAR (Dobin et al., 2013, PMID: 23104886) and processed with GATK best practices (McKenna et al., 2010, PMID: 20644199: DePristo et al., 2011, PMID: 21478889). RNA variants were called using HaplotypeCaller, and empirical editing efficiencies were established on PCR-de-duplicated alignment data.
Variant loci in BE overexpression experiments were further required to have comparable read coverage in the corresponding control experiment (read coverage for SNV in control >90th percentile of read coverage across all SNVs in overexpression). Additionally, the above loci were required to have a consensus of at least 99% of reads calling the reference allele in control.
The rAPOBEC1 amino acid sequence was obtained from uniprot and entered into the Phyre2 interface (Kelley LA et al. Nature Protocols 10, 845-858 2015) to obtain a protein model prediction. Three-dimensional distribution of residues in this predicted model were analyzed using the software PyMOL (Schrödinger). DNA and RNA binding was predicted using the DRNApred web interface (Yan&Kurgan, NAR 2017).
rAPOBEC1 was aligned to other APOBEC1 homologues or other members of the human APOBEC family using Geneious 7 software.
HEK293T (2.5×106 cells) cells were seeded into 100 mm TC-treated culture dishes (Fisher) 24 h prior to transfection. Cells were transfected in triplicate with 16.5 μg of BE3, BE3(E63Q), SECURE-BE3 or negative control plasmids as well as 5.5 μg of guide RNA expression plasmid (RNF2 site1), and 66 μL TransIT-293T. Cells were incubated for 36 h post-transfection, followed by sorting for GFP-positive cells (as described in FACS Methods). After sorting, cells were counted using a LUNA-FL Cell Counter (Logos Biosystems) with Acridine Orange/Propidium Iodide Stain. 5×103 viable cells were seeded into 96-well solid white TC treated microplates (Corning) in 100 μL DMEM; each condition was seeded into 3 wells for technical triplicates per biological replicate (n=3 biologically independent samples), and 4 plates of cells were prepared from this experiment for 4 different endpoints (d1-d4). At 24 h, 48 h, 72 h, and 96 h post-sorting, cell viability was determined using the CellTiter-Glo Luminescent Cell Viability Assay reagent (Promega). After the plate was equilibrated at room temperature for 30 minutes, 100 μL of 1:5 diluted CellTiter-Glo reagent were directly added to each well (adapted from ref. 45). After 2 minutes of plate shaking on the Synergy HT microplate reader (BioTek), plates were incubated at room temperature for 10 minutes, and read with the Synergy HT for luminescence. The luminescence background (average of 8 empty wells per plate) was subtracted from all luminescence values generated in the respective plate. Cells were not seeded at the edge of the plate (columns 1 and 12 as well as rows A and H).
We fit a linear mixed effects model using the R nlme package with log 2 (RLU) as the outcome to assess the effect on cell viability of each base editor variant compared to nCas9-UGI-NLS. A random effect for biological replicate was used to account for the correlation between technical replicates. P-values represent the significance of the fixed effect coefficient encoding the base editor in the mixed-effects models.
To test whether BE3 might be capable of editing cytosines in RNA, we first assessed whether this base editor fusion could edit the C6666 nucleotide in APOB mRNA previously shown to be edited by isolated rAPOBEC1. To do this, we transfected human HepG2 cells with a plasmid that expressed a BE3-P2A-EGFP fusion protein (the P2A sequence mediates a post-translational cleavage that releases EGFP from the BE3 part of the fusion) (Methods). At 36 hours after transfection, we then used flow cytometry to sort out the highest expressing (top 5%) of GFP-positive cells and isolated total RNA from these cells. As a negative control, we transfected HepG2 cells in parallel with a plasmid that expressed a nickase Cas9 (nCas9)-UGI-P2A-EGFP fusion protein (i.e., a plasmid identical to the BE3-P2A-EGFP expression plasmid but lacking the rAPOBEC1 and XTEN-linker within the BE3 part of the fusion protein) and also sorted these for the top 5% GFP-positive cells and isolated total RNA. We assessed the RNA sequence of the human APOB transcript that encompasses the C6666 previously shown to be deaminated by rAPOBEC1 in these samples using reverse transcription followed by targeted amplicon sequencing of this region (Methods). Consistent with previous studies of isolated rAPOBEC1 overexpression, we found that BE3 not only edited C6666 to a U with high efficiency (˜55%) in the APOB mRNA transcript but that it also edited other proximal Cs that were preceded by an A as well (
To test whether BE3 might edit Cs in other mRNA transcripts, transcriptome-wide experiments using ultra-deep RNA-seq were performed in two human cell lines (HEK293T and HepG2 cells). In these experiments (as illustrated in
These edited Cs were distributed throughout the human genome (
Given the extensive transcriptome-wide RNA editing induced by BE3, we sought to create variants of this base editor that would diminish this unwanted activity while retaining the desired capability to perform targeted DNA base editing. We reasoned that the introduction of mutations into the APOBEC1 part of a base editor might accomplish this. A previously published study described a series of 16 different amino acid substitutions in APOBEC1 that had been suggested to confer reduced RNA binding capability, reduced binding to auxiliary co-factors or reduced dimerization potential25-29 in isolated APOBEC1; however, these mutants had not been characterized for their RNA editing activities in the context of a base editor fusion nor had they been characterized for the desired retention of DNA editing capabilities in the context of a base editor fusion. As a result, it was unknown and unclear which mutations in the context of a base editor would have the desired combined properties of reduced RNA editing but preserved targeted DNA editing.
To begin to assess phenotypic behavior of the 16 previously described APOBEC1 mutations on base editor activities, we constructed a series of 16 BE3 fusions harboring the following amino acid substitutions in the APOBEC1 part of the protein (numbering of amino acid residues refers to the rAPOBEC1 sequence): R17A, P29F, P29T, R33A, K34A, R33A+K34A (double mutant), H61A, H61C, V62A, E63Q, E181Q, L182A, I185A, L187A, L189A, and P190A+P191A (double mutant). These variants were initially screened for their abilities to induce targeted DNA edits using three gRNAs targeted to different endogenous human genes (
We next assessed these ten BE3 variants for their RNA editing activities. We initially examined their abilities to edit the C6666 base and other adjacent Cs within the APOB mRNA transcript in human cells. To do this, HepG2 cells were transfected with plasmid expressing wild-type BE3 or a BE3 variant. RNA was harvested after 24 h (no sorting), followed by reverse transcription and targeted amplicon sequencing of a 200 bp region encompassing C6666 on the APOB transcript (Methods). This experiment revealed that seven of these BE3 variants (P29F, P29T, R33A, K34A, R33+K34A (double mutant), E181Q and L182A) showed relative reductions in RNA editing activities at these cytosines compared with wild-type BE3 (
We next performed transcriptome-wide analysis of RNA editing with overexpression of six of these variants in human cells, excluding E181Q due to its low DNA editing capabilities. This was done by transfecting HEK293T cells with plasmids expressing wild-type BE3 or a BE3 variant as P2A fusions to EGFP and a RNF2-targeted gRNA, sorting for the top 5% of GFP expressing cells 36 hours after transfection, isolating total RNA, and carrying out RNA-seq with 20 million reads/sample (using NextSeq) (Methods). This experiment demonstrated that all six variants showed substantially reduced transcriptome-wide RNA editing activities relative to wild-type BE3 and that the P29F and R33A+K34A variants in particular had activities similar to a BE3 harboring a E63Q active site mutation previously shown to completely abolish cytosine deaminase activity of APOBEC126, 28 (Table 2).
To more rigorously characterize RNA editing by these two variants, we performed RNA-seq experiments with the RNF2 gRNA using transfected HEK293T cells sorted for high-level expression of wild-type BE3, BE3-R33A, BE3-R33A/K34A, or a catalytically impaired BE3-E63Q mutant (Navaratnam et al, Cell. 1995 Apr. 21; 81(2):187-95). For these studies, we used high expression conditions (top 5% sorting) to enable the most sensitive detection of any residual RNA editing by these variants. We observed dramatic reductions in the number of transcriptome-wide C-to-U edits with BE3-R33A inducing only hundreds and BE3-R33A/K34A inducing 26 or fewer of such edits (
Importantly, examination of the on-target RNF2 DNA site in these same cells showed that all six variants retained DNA base editing activities and also perhaps possessed a more narrowed editing window (
We next sought to characterize targeted DNA editing activities of the six BE3 variants as well as E181Q with a larger series of gRNAs and under conditions in which we did not select cells for overexpression via sorting of GFP positive cells. To do this, we transfected HEK293T cells with plasmids expressing one of 12 different gRNAs and wild-type BE3 or a BE3 variant, harvested genomic DNA 72 hours following transfection without flow sorting, and examined the target DNA site for evidence of base editing using targeted amplicon sequencing with MiSeq (Methods). These experiments show that the BE3 variants harboring the R33A, K34A, R33A+K34A, or L182A mutations consistently show high targeted DNA editing activities comparable to wild-type BE3 across a range of sides, in some cases again showing a more narrowed window of editing at these sites as well as reduced insertion/deletion (indel) profiles as seen on VEGFA site 2. We conclude that the base editor variants described here possess reduced RNA editing activities while still retaining targetable sequence-specific DNA editing activities and we therefore refer to these as SElective Curbing of Unwanted RNA Editing (SECURE) base editor variants.
In addition to the SECURE base editor variants described and characterized above, we hypothesize that a number of additional APOBEC1 mutations may on their own confer the desired differential RNA vs. DNA editing activities to base editors and/or may help to improve the activity profiles of the variants we have already tested. No structural information is currently available for APOBEC1. However, as described in Methods, we built a structural model of APOBEC1 using Phyre2 (Kelley LA et al. Nature Protocols 10, 845-858 2015; PMID 25950237) and then predicted DNA- and RNA-binding residues using the DRNApred web interface (Yan&Kurgan, NAR 2017; PMID 28132027) (
Amino acid residues whose mutation may be expected to yield base editor SECURE variants. These positions were chosen based on an APOBEC1 structural model and RNA/DNA binding predictions or based on previous description in the literature as residues whose mutation reduced the RNA editing or binding activities of isolated APOBEC1.
The observation of extensive RNA edits by both cytosine and adenine base editors has important implications for research and therapeutic applications of these technologies. Confounding effects of unwanted RNA editing will need to be accounted for in research studies, especially if stable base editor expression (even in the absence of a gRNA) is used. For human therapeutic applications, the duration and level of BE expression should be kept to the minimums needed. Our data suggest that safety assessments for human therapeutics may need to include an analysis of the potential functional consequences of transcriptome-wide RNA edits. The short timeframe of our transient transfection experiments did not permit us to assess the longer-term functional consequences of widespread RNA editing but initial in silico and experimental analyses we have performed suggest that some edits may have phenotypic impacts on cells (
We transfected HEK293T cells in triplicate with plasmids expressing the RNF2 gRNA and either nCas9 UGI-NLS, wild-type BE3, BE3-R33A, BE3-R33A/K34A, and BE3-E63Q (each as 2A fusions to GFP). GFP-positive cells were sorted 36 hours post-transfection (all GFP-sorting, see Methods) and then equal numbers of viable sorted cells (as determined by acridine orange/propidium iodide staining) were plated into three technical replicate wells per biological replicate for four timepoints (Methods). At various timepoints post-plating (days 1, 2, 3, and 4), we performed a cell viability assay (CellTiter-Glo) for each biological replicate (n=3) in technical triplicates (Methods). In this assay, mean luminescence RLU values are an indirect measure of ATP content, which is directly proportional to the number of viable cells. The results of these experiments (
We note that there are several reasons why this experimental setup might detect only a modest effect of wild-type BE3 on cell viability: First, the negative impacts of transfection and FACS procedures on cell health are likely more substantial than that of the base editor. The effects of these experimental procedures are controlled for with the nCas9-UGI and the BE3-E63Q negative controls but it is likely that a large proportion of the dynamic range of the cell viability assay is lost due to the early toxicity induced by those two procedures. Second, because we are performing transient transfection, there will be a great deal of heterogeneity in the numbers, frequencies, and combinations of RNA edits induced in any given cell in the population. Hence, it may be challenging to observe any toxic effects due to this heterogeneity and an inducible, stable expression system will likely be better suited to detect cell viability effects. Finally, it is also possible that both pro- and anti-proliferative edits may exist in the same or different cells and this might therefore offset any anti-proliferative effects as well.
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It is to be understood that while the invention has been described in conjunction with the detailed description thereof, the foregoing description is intended to illustrate and not limit the scope of the invention, which is defined by the scope of the appended claims. Other aspects, advantages, and modifications are within the scope of the following claims.
This application claims the benefit of U.S. Provisional Patent Application Ser. No. 62/744,026, filed on Oct. 10, 2018. The entire contents of the foregoing are hereby incorporated by reference.
This invention was made with Government support under Grant No. HG009490 awarded by the National Institutes of Health. The Government has certain rights in the invention.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/US2019/055705 | 10/10/2019 | WO | 00 |
| Number | Date | Country | |
|---|---|---|---|
| 62744026 | Oct 2018 | US |
