Patent Application
20230070731
Incorporated herein by reference in its entirety is the Sequence Listing submitted via EFS-Web as a text file named SEQLIST_UPNK102.txt., created Jan. 20, 2021 and having a size of 235,835 bytes.
This invention relates to the fields of gene therapy and base editing. More specifically, the invention provides split DNA deaminase encoding constructs which exhibit controllable and efficient base editing while reducing undesirable off target effects. Methods employing such constructs and kits comprising the same, are also disclosed.
Several publications and patent documents are cited throughout the specification in order to describe the state of the art to which this invention pertains. Each of these citations is incorporated herein by reference as though set forth in full.
Base editing of the immunoglobulin locus by AID, the ancestral member of the AID/APOBEC family of cytosine deaminase enzymes, normally initiates maturation of antibody responses in B-cells, while APOBEC3 enzymes provide protection against retroviruses. Out of their physiological context, when DNA deaminases are directed towards a specific genomic locus by catalytically-impaired Cas9, their base editing activity can be used to introduce targeted mutations at a desired locus. While this system offers a potentially powerful means to edit the genome for biological or therapeutic purposes, base editors have at least two natural constraints that could limit their broader application. First, the enzymes have naturally evolved to be constrained deaminases with low overall catalytic activity, as hyperactivation is associated with increased oncogenic mutations. Second, AID/APOBECs are known to act outside of their targets, promoting cancer mutagenesis, chromosomal translocations, and resistance to chemotherapy. When the natural regulatory constraints are lost, overexpression of a functionally intact deaminase in a gene editing complex poses similar risks to the genome. In existing base editors, the DNA deaminases are targeted, but they are not regulated which increases undesirable off-target activity which is not mitigated by linking it to a targeting module like dCas9. As the deaminase is active, overexpressed and present in the nucleus, the active enzyme will be able to access ssDNA intermediates normally exposed in the process of DNA replication, transcription, and repair, much as it does in cancers. Indeed, an increase in genome-wide mutation at activation induced deaminase (AID) preferred hotspots has been shown with expression of AID-containing ZFN and TALE base editors, and recent work has shown widespread genome-wide action by the most commonly employed BE3 base editors.
Added concerns arise from evidence of off-target deaminase activity on RNA, highlighting the need to regulate where and when the deaminases are active. Although many biological goals can be achieved with current base editors, the therapeutic utility of base editing approaches in human patients will be limited if off target activity is not addressed.
It is clear that a need exists in the art for improved base editors whose activity can be regulated to permit action with greater precision at the targeted site with minimal off target effects.
The present invention provides precise base editor complexes and methods of use thereof for efficient and controllable site-specific editing at sites of interest in targeted DNA and RNA sequences. The base editor complexes described herein comprise different protein modules which act in concert to effect inducible and specific gene editing. The modules are fused using appropriate linker sequences and comprise at least a targeting module (TM) which localizes the complex to a particular genomic site of interest. The tethered modifying module (MM) edits the local DNA. In certain aspects skewing downstream repair pathways via inclusion of accessory modules (MMX) can improve efficiency. Via inclusion of a specific binding pair into the complex, the present invention provides for regulatory, small molecule control over based editors by exploiting knowledge of DNA deaminase structure and function to split DNA deaminases into inactive components that can only be reconstituted at the desired site of action. In other embodiments, both the targeting module and the modifying modules are split and reassembled upon dimerization of the specific binding pair. In yet another aspect, the complex comprises two distinct targeting molecules, e.g., two distinct dCas9/sgRNAs, for enhanced specificity, each of which is linked to one part of the split deaminase.
In one embodiment, a first fusion protein for precise small molecule control of targeted base editing comprising an optional accessory module, a targeting module, a first portion of a split deaminase operably linked to a first member of a specific binding pair, and a second fusion protein comprising a second portion of a split deaminase which is operably linked to a second member of a specific binding pair is provided, wherein said specific binding pair members dimerize upon contact with a dimerization agent causing two portions of the split deaminase enzyme to reform thereby resulting in formation of small molecule inducible base editor complex which edits a site of interest on a nucleic acid bound by the targeting module.
In another aspect, a first fusion protein comprising a first portion of a split deaminase, operably linked to a first portion of a split targeting module, said targeting module being operably linked to a first member of a specific binding pair, and a second fusion protein comprising a second portion of a split deaminase operably linked to a second portion of a split targeting module operably linked to a second specific binding pair member is provided, wherein said specific binding pair members dimerize upon contact with a dimerization agent, causing two portions of a split deaminase enzyme and the two portions of the targeting module to reform thereby resulting in formation of small molecule inducible base editor complex which edits a site of interest on a nucleic acid bound by the targeting module.
In another embodiment, a first fusion protein comprising a targeting module operably linked to a first member of a specific binding pair which is operably linked to a first portion of a split deaminase and second fusion protein comprising a second member of a specific binding pair, operably linked to a second portion of a split deaminase which is operably linked to a separate second targeting module. The two targeting modules are approximated close to one another at the nucleic acid target, with the specific binding pair members dimerizing upon contact with a dimerization agent, wherein dimerization causes two portions of a split deaminase enzyme to reform thereby resulting in formation of small molecule inducible base editor complex which edits a site of interest on a nucleic acid bound by the two co-localizing targeting modules with reduced off target effects.
In certain aspects, the targeting molecule is selected from nCas9, dCas9, dCas12, nCas12, xCas9, Cas13, transcription activator effector-like effectors (TALENs), and zinc finger nucleases (ZFNs), and comprises at least one sequence which directs said base editing complex to the site to be edited.
Deaminase proteins useful in the base editing complexes described herein can be selected from rat or human APOBEC1, human APOBEC3A, APOBEC3B, APOBEC3C, APOBEC3DE, APOBEC3F, APOBEC3G, Activation-induced cytidine deaminase (AID), CDA from lamprey, mutant version of Adenosine Deaminases (TadA) engineered to act on DNA, and Adenosine Deaminase acting on dsRNA (ADAR) or proteins having at least 90% identity with these proteins.
The fusion proteins may also comprise accessory molecules for reducing efficiency. Such molecules include, without limitation, UGI, 2x UGI, and μ-GAM.
In preferred embodiments the fusion proteins are present in a cell, and the cell is contacted with an effective amount of a dimerization agent, thereby causing the specific binding pair to dimerize. Specific binding pairs included in the base editing complex include, without limitation, FKBP and FRB wherein binding is induced by contact with dimerization agent rapamycin or a rapamycin analog, FKBP-F36V and FKBP-F36V wherein binding is induced by dimerization agent AP1903, BCLxl and scAZI, where binding is induced with dimerization agent ABT737, and CRY2 and CIB1 where binding is induced by light. In other embodiments, the first and second binding pairs are GFP 1-10 and GFP11 wherein binding occurs spontaneously.
Another embodiment of the invention includes a method of deaminating one or more selected bases in a target nucleic acid comprising contacting the target nucleic acid with the fusion proteins and dimerization agent described above. Also provided are host cells comprising the fusion proteins encoding the base editing complexes of the invention.
In another aspect a composition comprising the fusion proteins described above in a suitable biological carrier.
The invention also provides one or more isolated nucleic acids encoding the fusion proteins described above. Exemplary nucleic acids encoding the base editing complexes of the invention are shown in
The compositions of the invention can further comprise one or more of a liposome, a nanoparticle, a pharmaceutically acceptable carrier, and a buffer.
In yet another aspect, a method of deaminating one or more selected bases in a target nucleic acid is disclosed. An exemplary method comprises contacting a cell harboring the target nucleic acid with the base editing complex encoding nucleic acids described above under conditions where said complex is expressed, and a dimerization agent, thereby causing reformation of the deaminase and deaminating the base of interest in said target nucleic acid.
Also disclosed is a method for producing a small molecule inducible base editor complex in a cell for editing a target nucleic acid bound by an sgRNA, comprising introducing the expression vectors described above and a dimerization agent into said cell under conditions where said split deaminase reforms upon binding between said operably linked specific binding pair members, thereby catalyzing base editing at the site bound by said sgRNA.
Finally, kits for practicing the methods described above are also provided.
The recent repurposing of natural base editors for targeted genome editing has transformative potential (3). The typical formula for a base-editing (BE) complex (
Targeted base editing has applications across biology and medicine. While CRISPR/Cas9 based approaches are effective in generating knockout by causing dsDNA breaks, these result in heterogenous knockouts given unpredictable dsDNA break repair pathways and can also promote unwanted translocations. Base editors, by contrast, have the possibility of precisely introducing stop codons (CRISPR-Stop) to knockout genes without heterogeneity (42-44). Furthermore, base editors can make precise point mutations to correct disease alleles or make neomorphic protein variants, which is not possible with Cas9 alone in the absence of homology directed repair. Base editing can therefore be used to make knockouts more precisely, to reverse targeted mutations, and to edit primary cells or hosts with less risk.
By exploiting what we know about the mechanism, structure and function of DNA deaminases, existing base editors have been transformed into more effective and therapeutically useful reagents.
The terms “polynucleotide”, “nucleotide”, “nucleotide sequence”, “nucleic acid”, and “oligonucleotide” are used interchangeably in this disclosure. They refer to a polymeric form of nucleotides of any length, either deoxyribonucleotides or ribonucleotides, or analogs thereof. Polynucleotides may have any three-dimensional structure, and may perform any function, known or unknown. The following are non-limiting examples of polynucleotides: single-, double-, or multi-stranded DNA or RNA, genomic DNA, cDNA, DNA-RNA hybrids, or a polymer comprising purine and pyrimidine bases or other natural, chemically or biochemically modified, non-natural, or derivatized nucleotide bases. The terms “polynucleotide” and “nucleic acid” should be understood to include, as applicable to the embodiment being described, single-stranded (such as sense or antisense) and double-stranded polynucleotides. A polynucleotide may comprise one or more modified nucleotides, such as methylated nucleotides and nucleotide analogs. If present, modifications to the nucleotide structure may be imparted before or after assembly of the polymer. The sequence of nucleotides may be interrupted by non-nucleotide components. A polynucleotide may be further modified after polymerization, such as by conjugation with a labeling component.
The term “exogenous” nucleic acid can refer to a nucleic acid that is not normally or naturally found in or produced by a given bacterium, organism, or cell in nature. The term “endogenous” nucleic acid can refer to a nucleic acid that is normally found in or produced by a given bacterium, organism, or cell in nature.
The term “recombinant” is understood to mean that a particular nucleic acid (DNA or RNA) or protein is the product of various combinations of cloning, restriction, or ligation steps resulting in a construct having a structural coding or non-coding sequence distinguishable from endogenous nucleic acids found in natural systems.
The terms “construct”, “cassette”, “expression cassette”, “plasmid”, “vector”, or “expression vector” is understood to mean a recombinant nucleic acid, generally recombinant DNA, which has been generated for the purpose of the expression or propagation of a nucleotide sequence(s) of interest, or is to be used in the construction of other recombinant nucleotide sequences.
As used herein, a “modulating module” (MM) refers to the deaminase module of the base editors described herein. Exemplary MMs include for example, AID, APOBEC3 enzymes and TadA.
A “targeting module” localizes the base editing complex to the genomic region to be edited. Targeting modules can include for example, dCas9, nCas9, dCas12, ZFNs and TALENs.
An “accessory module” can optionally be included which are useful for controlling down stream repair pathways, thereby influencing efficiency of editing. Suitable accessory modules can encode a uracil glycosylase inhibitor (UGI) in one or multiple copies or μGAM for example.
The term “promoter” or “promoter polynucleotide” is understood to mean a regulatory sequence/element or control sequence/element that is capable of binding/recruiting an RNA polymerase and initiating transcription of sequence downstream or in a 3′ direction from the promoter. A promoter can be, for example, constitutively active, or always on, or inducible in which the promoter is active or inactive in the presence of an external stimulus. Example of promoters include T7 promoters or U6 promoters.
“Deamination” is the removal of an amino group from a molecule. Enzymes that catalyze this reaction are called deaminases. Deaminases include, without limitation, APOBEC1, APOBEC3A, APOBEC3B, APOBEC3C, APOBEC3DE, APOBEC3F, APOBEC3G, Activation-induced cytidine deaminase (AID), CDA from lamprey, Adenosine Deaminases acting on tRNA (TadA), and Adenosine Deaminase acting on dsRNA (ADAR). More broadly this deaminase family includes homologs from various species all of which are thought to catalyze similar reactions on nucleic acids as described in Krishnan et al. (Proc Natl Acad Sci USA. 2018; 115(14):E3201-E3210 and Iyer et al. (Nucleic Acids Res. 2011 December; 39(22):9473-97).
An “adapter or adaptor”, or a “linker” for use in the compositions and methods described herein is a short, chemically synthesized, single-stranded or double-stranded oligonucleotide that can be ligated to the ends of other DNA or RNA molecules. Double stranded adapters can be synthesized to have blunt ends to both terminals or to have sticky end at one end and blunt end at the other, or sticky ends at both ends. For instance, a double stranded DNA adapter can be used to link the ends of two other DNA molecules (i.e., ends that do not have “sticky ends”, that is complementary protruding single strands by themselves). It may be used to add sticky ends to cDNA allowing it to be ligated into the plasmid much more efficiently. Two adapters could base pair to each other to form dimers. A conversion adapter is used to join a DNA insert cut with one restriction enzyme, say EcoRl, with a vector opened with another enzyme, Bam Hl. This adapter can be used to convert the cohesive end produced by Bam Hl to one produced by Eco Rl or vice versa. One of its applications is ligating cDNA into a plasmid or other vectors instead of using Terminal Deoxynucleotide Transferase enzyme to add poly A to the cDNA fragment.
Alternatively, the linker may be a peptide linker such as those that occur between protein domains. Short peptide linkers are often composed of flexible residues like glycine and serine so that the adjacent protein domains are free to move relative to one another. Exemplary linkers include without limitation, 2 amino acid GS linkers, 6 amino acid (GS)x linker, 10 amino acid (GS)x linker, short linkers (Gly-Gly-Ser-Gly; SEQ ID NO: 1), Middle linkers (Gly-Gly-Ser-Gly; SEQ ID NO: 1) x2 and long linkers (Gly-Gly-Ser-Gly; SEQ ID NO: 1) x3, flexible linkers 2x(GGGS; SEQ ID NO: 2), 2x (GGGGS(SEQ ID NO: 3) and 13 amino acid linkers (GGGS GGGGS GGGS; SEQ ID NO:4).
The term “operably linked” can mean the positioning of components in a relationship which permits them to function in their intended manner. For example, a promoter can be linked to a polynucleotide sequence to induce transcription of the polynucleotide sequence.
The terms “sequence identity” or “identity” refers to a specified percentage of residues in two nucleic acid or amino acid sequences that are identical when aligned for maximum correspondence over a specified comparison window, as measured by sequence comparison algorithms or by visual inspection. When sequences differ in conservative substitutions, the percent sequence identity may be adjusted upwards to correct for the conservative nature of the substitution. Sequences that differ by such conservative substitutions are said to have “sequence similarity” or “similarity.” Means for making this adjustment are well known to those of skill in the art. Typically this involves scoring a conservative substitution as a partial rather than a full mismatch, thereby increasing the percentage sequence identity.
The term “comparison window” refers to a segment of at least about 20 contiguous positions in which a sequence may be compared to a reference sequence of the same number of contiguous positions after the two sequences are aligned optimally. In a refinement, the comparison window is from 15 to 30 contiguous positions in which a sequence may be compared to a reference sequence of the same number of contiguous positions after the two sequences are aligned optimally. In another refinement, the comparison window is usually from about 50 to about 200 contiguous positions in which a sequence may be compared to a reference sequence of the same number of contiguous positions after the two sequences are aligned optimally.
The terms “complementarity” or “complement” refer to the ability of a nucleic acid to form hydrogen bond(s) with another nucleic acid sequence by either traditional Watson-Crick or other non-traditional types. A percent complementarity indicates the percentage of residues in a nucleic acid molecule which can form hydrogen bonds (e.g., Watson-Crick base pairing) with a second nucleic acid sequence (e.g., 4, 5, and 6 out of 6 being 66.67%, 83.33%, and 100% complementary). “Perfectly complementary” means that all the contiguous residues of a nucleic acid sequence will hydrogen bond with the same number of contiguous residues in a second nucleic acid sequence. “Substantially complementary” as used herein refers to a degree of complementarity that is at least 40%, 50%, 60%, 62.5%, 70%, 75%, 80%, 85%, 90%, 95%, 97%, 98%, 99%, or 100%, or percentages in between over a region of 4, 5, 6, 7, and 8 nucleotides, or refers to two nucleic acids that hybridize under stringent conditions.
In general, a CRISPR system is characterized by elements that promote the formation of a CRISPR complex at the site of a target sequence (also referred to as a protospacer in the context of an endogenous CRISPR system). In the context of formation of a CRISPR complex, “target sequence” refers to a sequence to which a guide sequence is designed to have complementarity, where hybridization between a target sequence and a guide sequence promotes the formation of a CRISPR complex. Full complementarity is not necessarily required, provided there is sufficient complementarity to cause hybridization and promote formation of a CRISPR complex. A target sequence may comprise any polynucleotide, such as DNA or RNA polynucleotides. In some embodiments, a target sequence is located in the nucleus or cytoplasm of a cell. In some embodiments, the target sequence may be within an organelle of a eukaryotic cell, for example, mitochondrion or chloroplast. A sequence or template that may be used for recombination into the targeted locus comprising the target sequences is referred to as an “editing template” or “editing polynucleotide” or “editing sequence”. In aspects of the invention, an exogenous template polynucleotide may be referred to as an editing template. In an aspect of the invention the recombination is homologous recombination.
In general, a guide sequence is any polynucleotide sequence having sufficient complementarity with a target polynucleotide sequence to hybridize with the target sequence and direct sequence-specific binding of a CRISPR complex to the target sequence. In some embodiments, the degree of complementarity between a guide sequence and its corresponding target sequence, when optimally aligned using a suitable alignment algorithm, is about or more than about 85%, 90%, 95%, 97.5%, 99%, or more. Optimal alignment may be determined with the use of any suitable algorithm for aligning sequences, non-limiting example of which include the Smith-Waterman algorithm, the Needleman-Wunsch algorithm, algorithms based on the Burrows-Wheeler Transform (e.g. the Burrows Wheeler Aligner), ClustalW, Clustal X, BLAT, Novoalign (Novocraft Technologies, ELAND (Illumina, San Diego, Calif.), SOAP (available at soap.genomics.org.cn), and Maq (available at maq.sourceforge.net).
A “zinc finger nuclease” as used herein refers to artificial restriction enzymes generated by fusing a zinc finger DNA-binding domain to a DNA-cleavage domain. Zinc finger domains can be engineered to target specific desired DNA sequences and this enables zinc-finger nucleases to target unique sequences within complex genomes. By taking advantage of endogenous DNA repair machinery, these reagents can be used to precisely alter the genomes of higher organisms.
“Transcription activator-like effector nucleases” (TALEN) are restriction enzymes that can be engineered to cut specific sequences of DNA. They are made by fusing a TAL effector DNA-binding domain to a DNA cleavage domain (a nuclease which cuts DNA strands). Transcription activator-like effectors (TALEs) can be engineered to bind to practically any desired DNA sequence, so when combined with a nuclease, DNA can be cut at specific locations. The restriction enzymes can be introduced into cells, for use in gene editing or for genome editing in situ, a technique known as genome editing with engineered nucleases. Alongside zinc finger nucleases and CRISPR/Cas9, TALENs are also suitable for use in the base editing complexes of the invention.
Several aspects of the invention relate to vector systems comprising one or more vectors, or vectors as such. Vectors can be designed for expression of editing complexes of the invention (e.g. nucleic acid transcripts, proteins, or enzymes) in prokaryotic or eukaryotic cells. For example, base editing transcripts can be expressed in bacterial cells such as Escherichia coli, insect cells (using baculovirus expression vectors), yeast cells, or mammalian cells. Suitable host cells are discussed further in Goeddel, Gene Expression Technology: Methods in Enzymology 185, Academic Press. San Diego, Calif. (1990). Alternatively, the recombinant expression vector can be transcribed and translated in vitro, for example using T7 promoter regulatory sequences and T7 polymerase.
In some embodiments, a vector is capable of driving expression of one or more sequences in mammalian cells using a mammalian expression vector. Examples of mammalian expression vectors include pCDM8 (Seed, 1987. Nature 329: 840) and pMT2PC (Kaufman, et al., 1987. EMBO J. 6: 187-195). When used in mammalian cells, the expression vector's control functions are typically provided by one or more regulatory elements. For example, commonly used promoters are derived from polyoma, adenovirus 2, cytomegalovirus, simian virus 40, and others disclosed herein and known in the art. For other suitable expression systems for both prokaryotic and eukaryotic cells see, e.g., Chapters 16 and 17 of Sambrook, et al., Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y., 1989.
In some embodiments, the recombinant mammalian expression vector is capable of directing expression of the nucleic acid encoding the base editing complex preferentially in a particular cell type (e.g., tissue-specific regulatory elements are used to express the nucleic acid). Tissue-specific regulatory elements are known in the art. Non-limiting examples of suitable tissue-specific promoters include the albumin promoter (liver-specific; Pinkert, et al., 1987. Genes Dev. 1: 268-277), lymphoid-specific promoters (Calame and Eaton, 1988. Adv. Immunol. 43: 235-275), in particular promoters of T cell receptors (Winoto and Baltimore, 1989. EMBO J. 8: 729-733) and immunoglobulins (Baneiji, et al., 1983. Cell 33: 729-740; Queen and Baltimore, 1983. Cell 33: 741-748), neuron-specific promoters (e.g., the neurofilament promoter; Byrne and Ruddle, 1989. Proc. Natl. Acad. Sci. USA 86: 5473-5477), pancreas-specific promoters (Edlund, et al., 1985. Science 230: 912-916), and mammary gland-specific promoters (e.g., milk whey promoter, U.S. Pat. No. 4,873,316 and European Application Publication No. 264,166). Developmentally-regulated promoters are also encompassed, e.g., the murine hox promoters (Kessel and Gruss, 1990. Science 249: 374-379) and the α-fetoprotein promoter (Campes and Tilghman, 1989. Genes Dev. 3: 537-546).
In some aspects, the invention provides methods comprising delivering one or more polynucleotides, such as or one or more vectors as described herein (e.g., encoding all or portions of the base editing complexes discussed below), one or more transcripts thereof, and/or one or proteins transcribed therefrom, to a host cell. In some aspects, the invention further provides cells produced by such methods, and organisms (such as animals, plants, or fungi) comprising or produced from such cells. In some embodiments, a CRISPR enzyme in combination with (and optionally complexed with) a guide sequence, a zinc finger nuclease or a TALEn is delivered to a cell. Conventional viral and non-viral based gene transfer methods can be used to introduce nucleic acids in mammalian cells or target tissues. Such methods can be used to administer nucleic acids encoding components of a base editing system to cells in culture, or in a host organism. Non-viral vector delivery systems include DNA plasmids, RNA (e.g. a transcript of a vector described herein), naked nucleic acid, and nucleic acid complexed with a delivery vehicle, such as a liposome. Viral vector delivery systems include DNA and RNA viruses, which have either episomal or integrated genomes after delivery to the cell. For a review of gene therapy procedures, see Anderson, Science 256:808-813 (1992); Nabel & Felgner, TIBTECH 11:211-217 (1993); Mitani & Caskey, TIBTECH 11:162-166 (1993); Dillon, TIBTECH 11:167-175 (1993); Miller, Nature 357:455-460 (1992); Van Brunt, Biotechnology 6(10):1149-1154 (1988); Vigne, Restorative Neurology and Neuroscience 8:35-36 (1995); Kremer & Perricaudet, British Medical Bulletin 51(1):31-44 (1995); Haddada et al., in Current Topics in Microbiology and Immunology Doerfler and Bihm (eds) (1995); and Yu et al., Gene Therapy 1:13-26 (1994).
Methods of non-viral delivery of nucleic acids include lipofection, nucleofection, microinjection, biolistics, virosomes, liposomes, immunoliposomes, polycation or lipid:nucleic acid conjugates, naked DNA, artificial virions, and agent-enhanced uptake of DNA. Lipofection is described in e.g., U.S. Pat. Nos. 5,049,386, 4,946,787; and 4,897,355) and lipofection reagents are sold commercially (e.g., Transfectam™ and Lipofectin™). Cationic and neutral lipids that are suitable for efficient receptor-recognition lipofection of polynucleotides include those of Feigner, WO 91/17424; WO 91/16024. Delivery can be to cells (e.g. in vitro or ex vivo administration) or target tissues (e.g. in vivo administration).
The preparation of lipid:nucleic acid complexes, including targeted liposomes such as immunolipid complexes, is well known to one of skill in the art (see, e.g., Crystal, Science 270:404-410 (1995); Blaese et al., Cancer Gene Ther. 2:291-297 (1995); Behr et al., Bioconjugate Chem. 5:382-389 (1994); Remy et al., Bioconjugate Chem. 5:647-654 (1994); Gao et al., Gene Therapy 2:710-722 (1995); Ahmad et al., Cancer Res. 52:4817-4820 (1992); U.S. Pat. Nos. 4,186,183, 4,217,344, 4,235,871, 4,261,975, 4,485,054, 4,501,728, 4,774,085, 4,837,028, and 4,946,787).
The use of RNA or DNA viral based systems for the delivery of nucleic acids take advantage of highly evolved processes for targeting a virus to specific cells in the body and trafficking the viral payload to the nucleus. Viral vectors can be administered directly to patients (in vivo) or they can be used to treat cells in vitro, and the modified cells may optionally be administered to patients (ex vivo). Conventional viral based systems could include retroviral, lentivirus, adenoviral, adeno-associated and herpes simplex virus vectors for gene transfer. Integration in the host genome is possible with the retrovirus, lentivirus, and adeno-associated virus gene transfer methods, often resulting in long term expression of the inserted transgene. Additionally, high transduction efficiencies have been observed in many different cell types and target tissues.
The tropism of a retrovirus can be altered by incorporating foreign envelope proteins, expanding the potential target population of target cells. Lentiviral vectors are retroviral vectors that are able to transduce or infect non-dividing cells and typically produce high viral titers. Selection of a retroviral gene transfer system would therefore depend on the target tissue. Retroviral vectors are comprised of cis-acting long terminal repeats with packaging capacity for up to 6-10 kb of foreign sequence. The minimum cis-acting LTRs are sufficient for replication and packaging of the vectors, which are then used to integrate the therapeutic gene into the target cell to provide permanent transgene expression. Widely used retroviral vectors include those based upon murine leukemia virus (MuLV), gibbon ape leukemia virus (GaLV), Simian Immuno deficiency virus (SIV), human immuno deficiency virus (HIV), and combinations thereof (see, e.g., Buchscher et al., J. Virol. 66:2731-2739 (1992); Johann et al., J. Virol. 66:1635-1640 (1992); Sommnerfelt et al., Virol. 176:58-59 (1990); Wilson et al., J. Virol. 63:2374-2378 (1989); Miller et al., J. Virol. 65:2220-2224 (1991); PCT/US94/05700). In applications where transient expression is preferred, adenoviral based systems may be used. Adenoviral based vectors are capable of very high transduction efficiency in many cell types and do not require cell division. With such vectors, high titer and levels of expression have been obtained. This vector can be produced in large quantities in a relatively simple system. Adeno-associated virus (“AAV”) vectors may also be used to transduce cells with target nucleic acids, e.g., in the in vitro production of nucleic acids and peptides, and for in vivo and ex vivo gene therapy procedures (see, e.g., West et al., Virology 160:38-47 (1987); U.S. Pat. No. 4,797,368; WO 93/24641; Kotin, Human Gene Therapy 5:793-801 (1994); Muzyczka, J. Clin. Invest. 94:1351 (1994). Construction of recombinant AAV vectors are described in a number of publications, including U.S. Pat. No. 5,173,414; Tratschin et al., Mol. Cell. Biol. 5:3251-3260 (1985); Tratschin, et al., Mol. Cell. Biol. 4:2072-2081 (1984); Hermonat & Muzyczka, PNAS 81:6466-6470 (1984); and Samulski et al., J. Virol. 63:03822-3828 (1989).
Packaging cells are typically used to form virus particles that are capable of infecting a host cell. Such cells include 293 cells, which package adenovirus, and ψ2 cells or PA317 cells, which package retrovirus. Viral vectors used in gene therapy are usually generated by producing a cell line that packages a nucleic acid vector into a viral particle. The vectors typically contain the minimal viral sequences required for packaging and subsequent integration into a host, other viral sequences being replaced by an expression cassette for the polynucleotide(s) to be expressed. The missing viral functions are typically supplied in trans by the packaging cell line. For example, AAV vectors used in gene therapy typically only possess ITR sequences from the AAV genome which are required for packaging and integration into the host genome. Viral DNA is packaged in a cell line, which contains a helper plasmid encoding the other AAV genes, namely rep and cap, but lacking ITR sequences. The cell line may also be infected with adenovirus as a helper. The helper virus promotes replication of the AAV vector and expression of AAV genes from the helper plasmid. The helper plasmid is not packaged in significant amounts due to a lack of ITR sequences. Contamination with adenovirus can be reduced by, e.g., heat treatment to which adenovirus is more sensitive than AAV.
In some embodiments, a host cell is transiently or non-transiently transfected with one or more vectors described herein. In some embodiments, a cell is transfected as it naturally occurs in a subject. In some embodiments, a cell that is transfected is taken from a subject. In some embodiments, the cell is derived from cells taken from a subject, such as a cell line.
In one aspect, the invention provides for methods of modifying a target polynucleotide in a eukaryotic cell, which may be in vivo, ex vivo or in vitro. In some embodiments, the method comprises sampling a cell or population of cells from a human or non-human animal, and modifying the cell or cells. Culturing may occur at any stage ex vivo. The cell or cells may be re-introduced into the human or non-human animal.
In other embodiment, proteins comprising the base editing complex can be delivery directly into cells via use of nanoparticles, RNPs and other methods known to the skilled artisan.
DNA deaminases serve important roles in immune defense and other processes. Exemplary AID/APOBEC enzymes are immune enzymes. AID plays a role in somatic hypermutation, the mechanism by which antibody encoding genes are mutated and affinity matured. The related APOBEC3 enzymes are also known to target retroviruses for deamination.
As mentioned above, a family of deaminases exists and includes adenosine deaminase enzymes like TadA, which catalyzes A to I mutation in tRNAs, and whose mutant variants can act on DNA rather than RNA. Notably, each of these DNA deaminases possess comparable secondary structures facilitating identification of suitable splitting sites which can be effectively reassembled when tagged with proteins or agents having specific binding affinity for one another which spontaneously reassemble when in proximity. Strategies for splitting DNA deaminase based on secondary structure within “families of deaminases” are described herein.
The ability to precisely edit specific bases has broad biotechnological potential in many practical and therapeutic approaches. While base editing of the human genome is the most exciting and promising of these approaches, many other applications exist, for example in modification of epigenetic sequences, agriculture and the biofuel industry. Other intriguing applications include directed somatic hypermutation for generation of improved antibodies and other therapeutic proteins.
In order to more precisely control and target base editing, the split DNA deaminases described herein are constructed such that reassembly is effected by the binding of a small molecule to an added domain that induces split deaminases to spontaneously reassemble, thereby reforming the split enzyme into an active and efficient deaminase. This inventive approach enables simultaneous spatiotemporal and small molecule control over activation of the mutator enzyme conferring a number of advantages including introduction of mutations at a precise time and location which has the benefit of decreasing off target, undesired activities or delaying the introduction of mutations until a time when it is desirable.
Our strategy entailed first identifying a control point for insertion of the spontaneously reassembling binding partners, then splitting the enzyme into demonstrably inactive parts which can effectively and spontaneously reassemble when tagged with proteins that spontaneously come together, finally providing an inducer element which alters the protein partners from ones which spontaneously reassemble to those that come together only in the presence of an inducing agent, and demonstrating that small-molecule inducible, precise base editing has been achieved.
The secondary structure of the DNA deaminase fold was examined to identify “control points” or insertion sites for small regulatory elements which would allow for small-molecule control over the deaminase reassembly and activity.
In initial studies, a foreign protein/domain was inserted into an enhanced hyperactive version of AID described in U.S. patent application Ser. No. 16/025,261 (See for example, SEQ ID NO: 20 of the '261 patent application) which is incorporated herein by reference as though set forth in full. This mutant version of the human DNA deaminase AID, involved in antibody maturation was assessed and the loop regions which appeared to be tolerant to insertion of control elements (e.g., Green Fluorescent Protein GFP fragments) were identified as described hereinbelow. Several candidate locations for insertion were identified. We used an E. coli-based rifampin mutagenesis assay to evaluate activity (See Kohli et al., J Biol Chem. (2009) 284:pages 22898-904). In this assay we measure the activity level of a mutator by measuring how many E. coli can be turned resistant to the antibiotic rifampin when the enzyme is turned on.
This study led us to focus on inserting into the loop between alpha2 and beta3, but other candidate sites are also suitable as shown in
Having identified candidate sites for insertion, we then assessed whether the protein could be split into two inactive components. We split the GFP between beta stand 10 and 11, which is known as split GFP, and resulted in splitting of AID into N- and C-terminal halves. We showed that the split enzymes are inactive by themselves in the rifampin based resistance assay (see
The analogous approach with other AID/APOBEC family members has also been assessed as described herein, including APOBEC3A. See
Using this split A3A, we also tested the system employing the spontaneously reassembling split deaminase in mammalian cell lines (HEK293 and HeLa cells). When we express two inactive APOBEC3A splits together, GFP spontaneously reassembles and we see DNA damage to the mammalian genome, as measured by a DNA damage marker. See
Having established the split sites and the feasibility of spontaneous reassembly with split GF, the last steps of tool development for split base editor development was switching from split, spontaneously reassembling GFP to two proteins which can reassemble under small molecule control, and moving from the DNA deaminase domain by itself to a more complex scaffold of a base editor complex. Here, the dimerization domain is exemplified by FKBP-FRB, which can be brought together with rapamycin, and use of the the Cas9-based base editor platform. Other small molecules for this purpose, include, without limitation those shown in
Using the seBEa scaffold with split AID, A3A, or evolved APOBEC1 (evoA1) we have achieved the goal of small molecule control over base editing. Using an assay measuring inactivation of a single copy of GFP in cell lines (See
Now that identified suitable sites for splitting deaminases, these can be substituted in the editing constructs described herein. Notably, other DNA deaminases can be split at analogous sites between alpha2 and beta3. Existing base editors constructs can be altered in split engineered base editors by the insertion of a DNA cassette into at the split site, as schematized in
We envision various combinations of the DNA deaminase with different Targeting Modules beyond nCas9, in different orders (e.g. Cas9-deaminase, instead of Deaminase-Cas9, etc), and with various accessory modules. Each of these could be joined with linkers of various lengths or make-up. See
The following methods are provided to facilitate the practice of the present invention.
HEK293T d2GFP contains a single integrated copy of destabilized GFP in its genome. The cell line was maintained in Dulbecco's Modified Eagle's Medium with L-Glutamine, 4.5 g/L Glucose and Sodium Pyruvate (Corning) supplemented with 10% (v/v) bovine calf serum (CS) and 1% (v/v) Penicillin-Streptomycin mix, at 37° C. with 5% CO2.
For mammalian base editing constructs, the intact or split-engineered constructs were cloned into the scaffold of pCMV_BE4max (Addgene Plasmid #112093), which contains rat APOBEC1. The parent plasmid contains a NotI restriction site. An additional XmaI restriction site was added into pCMV_BE4max using the Q5 Site-Directed Mutagenesis Kit (NEB) to facilitate cloning. The deaminase sequences were amplified from their respective pET41 plasmids, introducing a region of overlap. AID′ differs from AID* in that it contains a smaller subset of mutations, including K10E, T82I, D118A, R119G, K120R, A121R, and E156G. To facilitate cloning of seBE constructs, gene fragments were synthesized (IDT) containing DeaminaseN-FRB, the T2A self-cleaving peptide between the two fragments, and FKBP12-DeaminaseC. The associated strategy for linkers between domains was derived from that recently employed to split human TET247. Using the gene fragments, all BE4max and seBE plasmids were then constructed using Gibson Assembly Master Mix (NEB), merging the relevant gene fragments with the NotI/XmaI digested vector. Notably the intact AID′-BE4max and A3A-BE4max lack the N-terminal NLS present in BE4max vectors. A3A-seBE contains a missense mutation (M13I) as a result of a PCR error, which does not appear to impact activity.
The evoA1-seBE4max-IRES construct, where the two split protein fragments are independently translated, was cloned into the scaffold of evoA1-seBE4max. The IRES sequence fragment was amplified from Addgene Plasmid #10559448 with Phusion High-Fidelity DNA Polymerase (NEB). The vector backbone of evoA1-seBE4max was amplified, excluding the T2A sequence. The vector and IRES sequence fragment were then joined using the In-Fusion HD Cloning system (TBUSA).
The sgRNA expression plasmids were constructed using oligonucleotide cassettes for cloning. Briefly, the primers listed in the Supplementary Information were annealed and phosphorylated using T4 Polynucleotide Kinase (NEB) according to the manufacturer's instructions and further purified using the oligo clean and concentrator kit (Zymo Research). Next, LRcherry2.1 plasmid49 or LRG plasmid (Addgene #65656) were incubated with restriction enzyme Esp3I (Thermo Fisher Scientific) at 37° C. for 2 hours to remove a short filler sequence, and further agarose gel purified. The sgRNA cassettes were then ligated in place of the filler using T4 DNA ligase (NEB).
The mutation frequency of various DNA deaminases, including insertion constructs, were determined using a modified version of previously reported rifampin mutagenesis assay (Kohli, JBC 2009). Plasmids encoding the deaminase variant were transformed into BL21(DE3) E. coli, that already harbor a plasmid encoding uracil DNA glycosylase inhibitor (UGI) on a pETcoco2 plasmid. Overnight cultures grown in LB with kanamycin (30 ng/mL) and chloramphenicol (25 ng/mL) from single colonies were diluted to an A600 of 0.2 and grown for 1 hr at 37° C. before inducing deaminase expression with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG). After 4 hrs of additional growth, aliquots of cultures were separately plated on Luria Bertani (LB) agar plates containing rifampicin (100 μg/mL) and plasmid-selective antibiotics. The mutation frequencies were then calculated by the ratio of rifampicin resistant colonies to total population. For bacterial work with AID*, the parent pET41 plasmid with AID* combines three different sets of previously described29-31 mutations that increase activity or solubility (K10E, F42E, T82I, D118A, R119G, K120R, A121R, H130A, R131E, F141Y, F145E, and E156G) in a construct with an N-terminal maltose binding protein tag (MBP). The plasmids named AID*-INS contain an insertion of optGFP flanked by linkers at each position within a specified loop of AID*. The N-terminal fragment of AID (AID*N) and C-terminal fragment of AID (AID*C) were generated by PCR amplification from the AID* parent plasmid with primers listed in Supplementary Table 2. A sequence containing linker-optGFP-linker was obtained as a gene fragment (Integrated DNA Technologies, IDT) and amplified with primers provided below, which add flanking regions that permit overlap extension PCR. Overlap extension PCR was performed to fuse the three fragments encoding AID*N, linker-optGFP-linker, and the AID*C, using 10 cycles of amplification without primers to permit fusion of fragments, followed by amplification of the entire AID*N-optGFP-AID*C sequence with the outer primers. PCR products from the overlap extension PCR were TA cloned (Invitrogen). Sequence-confirmed inserts were then digested with SalI and AvrII and ligated into the digested parent plasmid with T4 DNA ligase (NEB). The control plasmids containing unmutated AID (AID-WT) or its catalytically inactive analog, AID(E58A), were previously reported30.
For bacterial work with split AID*, AID*-SPL2N and AID*-SPL2C were created using AID*-INS2 as a scaffold in the pET41 backbone. To create AID*-SPL2N, the parent plasmid (AID*-INS2) was digested with KpnI and AvrII to remove the C-terminal region of AID*. Then, an oligonucleotide cassette containing a stop codon (TAG) was ligated into the digested vector. To create AID*-SPL2C, the parent plasmid (AID*-INS2) was digested with XbaI and KpnI to remove AID*-SPL2N. Then, a cassette containing a start codon (ATG) was ligated into the digested vector. The AID*-SPL2 plasmid, co-expressing the N-terminal and C-terminal fragments, from separate promoters was created using AID*-INS2 as a scaffold. A gene fragment was synthesized containing the C-terminal region of AID*-SPL2N, the transcriptional terminator, T7 RNA polymerase promoter and N-terminal region of AID*-SPL2C. This fragment was ligated into a KpnI/AvrII digested AID*-INS2 parent vector.
For bacterial expression of A3A constructs with insertion of optGFP, cloning was performed in_the scaffold of MBP-A3A-His-pET41 backbone45, 46 (Addgene #109231) using restriction enzymes EagI and AvrII. The appropriate optGFP-containing insert was synthesized as a gene fragment (IDT), digested with EagI/AvrII (NEB), and ligated into the similarly digested parent plasmid.
For mammalian expression of A3A constructs, plasmids were cloned into a pLEXm backbone._A3A-INS2,_A3A-SPL2N, and A3A-SPL2C were amplified from the pET41 construct, adding flanking regions of overlap with the pLEXm plasmid backbone. The final plasmids were then constructed using Gibson Assembly Master Mix (NEB), merging the amplified gene fragments with the EcoRI/XhoI (NEB) digested parent vector. The catalytically inactive variant A3A(E72A)-INS2 was created using Q5 Site-Directed Mutagenesis Kit (NEB).
For in vitro assays, purified intact, optGFP-inserted, or split DNA deaminases were expressed in BL21(DE3) cells that co-express the Trigger Factor (TF) chaperone, as previously described33. Briefly, 600 mL cultures were grown to an OD600 of 0.6 at 37° C. Cultures were shifted to 16° C. for 16 hours after induction with 1 mM IPTG. For AID variants, the pelleted cells were resuspended in 50 mM Tris-Cl (pH 7.5) 150 mM NaCl, 10% glycerol (wash buffer) and lysed through sonication. The soluble fraction was filtered after high-speed centrifugation and incubated with 3 mL of Amylose Resin (NEB) for 1 hr at 4° C. The resin was washed extensively prior to elution with wash buffer plus 10 mM maltose. Total protein was quantified by comparison to a BSA standard curve. For A3A variants, the pelleted cells were resuspended in 50 mM Tris-Cl (pH 7.5) 150 mM NaCl, 10% glycerol, 25 mM imidazole (wash buffer) and lysed through sonication. The soluble fraction was filtered after high-speed centrifugation and incubated with 3 mL of HisPur cobalt resin (Thermo) for 1 hr at 4° C. The resin was washed extensively prior to elution with wash buffer with 150 mM imidazole.
For the in vitro assay, a fluorescein (FAM)-labeled oligonucleotide substrate was used containing a single cytosine, along with a product control oligonucleotide containing uracil at the same location. For AID variants, the oligonucleotide substrate was co-incubated with 3-fold dilutions of the purified AID variant (520 nM to 0.6 nM) and 25 U of uracil DNA glycosylase (NEB). The reaction was performed in 20 mM Tris-HCl (pH 8.0), 1 mM DTT and 1 mM EDTA at 37° C. for 1 hr. For A3A, the oligonucleotide substrate was co-incubated with 3-fold dilutions of the purified A3A variant (18 nM to 10 pM) and 25 U of uracil DNA glycosylase. The reaction was performed in 350 mM succinic acid, sodium dihydrogen phosphate, and glycine (SPG) buffer (pH 5.5) and 0.1% Tween-20 at 37° C. for 30 min. Deamination reactions were terminated by incubation at 95° C. for 10 min. The samples were heat denatured by using 2× bromophenol blue loading dye containing 0.6 M NaOH to cleave abasic sites and 0.03 M EDTA. Samples were run on a preheated 20% acrylamide/Tris-Borate-EDTA(TBE)/urea gel at 50° C., and imaged using FAM filters on a Typhoon imager (GE Healthcare). Product formation was quantified using ImageJ by taking the ratio of substrate to product under each condition. Product formation as a function of enzyme concentration was fit to a sigmoidal dose-response curve and used to determine the EC50, defined as the amount of enzyme that converts 50% of the substrate to product under the fixed reaction conditions.
HEK293T cells were transiently transfected with A3A-INS2, A3A(E72A)-INS2 or co-transfected with A3A-SPL2N and A3A-SPL2C constructs for 24 hours prior to incubation with γH2AX antibody (BD Pharmigen, 647) and flow cytometry analysis. Cells were gated on FITC and APC using the Fortessa Flow Cytometer (BD Biosciences), and results were analyzed using FlowJo. Statistical analysis was performed using GraphPad Prism. U2OS cells plated on coverslips were transiently transfected with A3A-INS, A3A(E72A)-INS2 or co-transfected with A3A-SPL2N A3A-SPL2N constructs for 24 hours prior to incubation with γH2AX antibody (Millipore Sigma) and immunofluorescent staining with Alexa Fluor 568 (Invitrogen) and DAPI. Stained cells were imaged with a Nikon MR confocal microscope and analyzed using Image J. HEK293T and U2OS cells were cultured in Dulbecco's Modified Eagle Medium (Gibco) media supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin.
Base Editing Assay Using d2GFP Inactivation by Flow Cytometry
HEK293T cells were lentivirally-transduced with a constitutively expressed destabilized GFP (d2GFP) reporter (derived from Addgene #14760) and selected for individual clones that contained a single copy of integrated d2gfp. The cell line was maintained in Dulbecco's Modified Eagle Medium with L-glutamine, 4.5 g/L glucose and sodium pyruvate (Corning) supplemented with 10% (v/v) bovine calf serum (CS) and 1% (v/v) penicillin-streptomycin mix, at 37° C. with 5% CO2. The HEK293T d2GFP cells were seeded on 24-well plates and transfected at approximately 60% confluency. 660 ng of intact BE4max or seBE4max constructs and 330 ng of LRcherry2.1 sgRNA expression plasmids were transfected using 1.5 μL of Lipofectamine 2000 CD (Invitrogen) per well according to manufacturer's protocol. Negative control samples include LRcherry2.1 plasmid lacking a protospacer (labeled as no sgRNA samples). The d2gfp-targeting sgRNA exposes a window where base editing can result in the introduction of a Q158X nonsense mutation in d2gfp. For seBE experiments, 24 hrs after transfection, rapamycin (Research Products International) was added to select wells at a final concentration of 200 nM. Transfected cells were harvested at day 3 after transfection, ensuring single-cell suspension. The percentage of d2GFP-negative and mCherry-positive (sgRNA+) cells was determined by flow cytometry with Guava Easycyte 10HT instrument (Millipore). Flow cytometry analysis was performed using FlowJo Software Version 10.7.1 (FloJo, LCC).
Genomic DNA was also collected from cells using the DNeasy Blood & Tissue Kit (Qiagen) according to manufacturer's instructions for amplification across the d2gfp locus and deep sequencing as described below. Total RNA was isolated using Direct-zol™ RNA Miniprep Plus kit (Zymo Research #R2072) following the manufacturer's protocol for sequencing as described below. For RNA-seq analysis, negative control transfections included d2gfp-targeting LRcherry2.1 plasmid without any base editor construct.
For editing of diverse genomic loci, HEK293T cells (lacking the single copy d2gfp) were used and maintained as above. The transfection protocol was performed as described above, with the exception that different sgRNAs were used to targeting of other loci. In each case, the sgRNAs expose a window where base editing can result in the introduction of point mutations in DNA modifying enzymes that lead to either missense or nonsense mutations. As with the d2GFP editing assay, 24 hrs after transfection, rapamycin (Research Products International) was added to select wells at a final concentration of 200 nM. Transfected cells were harvested at day 3 after transfection, ensuring single-cell suspension. Genomic DNA was collected using the DNeasy Blood & Tissue Kit (Qiagen) according to manufacturer's instructions for sequencing analysis as described below.
Target loci of interest were PCR-amplified from 100 ng genomic DNA (primer pairs in Supplementary Sequences) using KAPA HiFi HotStart Uracil+ Ready Mix (Kapa Biosystems) or Phusion High-Fidelity DNA Polymerase (New England Biolabs, NEB). PCR products were then purified (Qiagen).
Some samples were deep-sequenced by Amplicon-EZ Next Generation Sequencing (Genewiz). Alternatively, indexed DNA libraries were prepared using the NEBNext Ultra II DNA Library Prep Kit for Illumina with the following specifications. After adapter ligation and 4 cycles of PCR enrichment, indexed amplicon concentration was quantified by Qubit dsDNA HS Assay Kit (ThermoFisher), and size distribution was determined on a Bioanalyzer 2100 (Agilent) with the DNA 1000 Kit (Agilent). Indexed PCR amplicons with different barcodes were pooled together in an equimolar ratio for paired-end sequencing by MiSeq (Illumina) with the 300-cycle MiSeq Reagent Nano Kit v2 (Illumina). Raw reads were automatically demultiplexed by MiSeq Reporter. Demultiplexed read qualities were evaluated by FastQC v0.11.9 as described on the world wide web at bioinformatices.babraham.ac.uk/projects/fastqc. Low-quality sequence (Phred quality score <28) and adapters were trimmed via Trim Galore v0.6.5 as described on the world wide web bioinformatics.babraham.ac.uk/projects/trim_galore/ prior to analysis with CRISPResso2. Sequencing yielded ˜13,000 median aligned reads per sample (5th percentile ˜4,000, 95th percentile ˜63,000). The reported data (
Total RNA, isolated as described above, was analyzed for quality using the RNA 6000 Nano Bioanalyzer kit (Agilent). Only RNA with an RNA integrity number (RIN) ≥8 was used for subsequent library construction. RNA-seq was performed on 500 ng-1 μg of total RNA according to the Genewiz Illumina Hi-seq protocol for poly(A)-selected samples (2×150 bp pair-end sequencing, 350M raw reads per lane). The resulting reads were analyzed using the RADAR pipeline (RNA-editing Analysis-pipeline to Decode All twelve-types of RNA-editing events51. RNA edits that were present in the sgRNA-only samples were removed with analysis performed only on unique editing events present in the samples.
SEQUENCES Suitable for Use in the Base Editing Complexes Described Herein.
All oligonucleotides were purchased from Integrated DNA Technologies (IDT).
Primers used for generating sgRNA transfection plasmids. LRche2.1T vector was used as a template as noted in the methods section.
Primers used to add XmaI restriction site to pCMV_ABEmax and pCMV_BE4max.
Primers used for generating split BE3, split BE4max and split monomer ABEmax transfection plasmids. The same forward primer (splitCD FRB/FKBP Forward) was used to generate all 5 constructs.
Split Deaminases Gene Block Fragments
Myc-NLS-A3An-FRB-T2A-FKBP12-A3Ac-FlagTag
Myc-NLS-AIDn-FRB-T2A-FKBP12-AID-FlagTag
Myc-NLS-evorA1n-FRB-T2A-FKBP12-evorA1c
Myc-NLS-TadAn-FRB-T2A-FKBP12-TadAc
Primers Used for d2GFP Loci Sequencing:
d2GFP Sequence
Linker-GFP-Linker Sequence
Primers used to clone AIDn fragments. AIDn Forward primer was used to generate all AIDn fragments. Select sequence for insert 2 are shown as these were the sites carried forward.
Primers used to clone AIDc fragments. AIDc Reverse primer was used to generate all AIDc fragments.
Primers used to clone linker-GFP-linker fragments.
Primers Used for Overlap Extension PCR
The following examples are provided to illustrate certain embodiments of the invention. They are not intended to limit the invention in any way.
DNA deaminase enzymes have been converted into efficient and controllable genome editors, thereby overcoming constraints that will otherwise limit their scientific and therapeutic potential.
Members of the zinc-dependent nucleic acid deaminase family have evolved distinctively to act on a variety of substrates serving different biological roles, while retaining the same core structure. Activation induced deaminase (AID) mutates cytosine bases to uracil in the immunoglobulin locus of B-cells, initiating somatic hypermutation and antibody maturation. Related APOBEC3 DNA deaminases mutate and restrict foreign retroviruses, and more distantly related deaminases can even act on adenosine in tRNA. Nature's enzymatic toolbox for introducing base transition mutations, while powerful, has been subjected to several evolutionary requirements, given the threat that purposeful mutators pose to genomic stability. These requirements include constrained sub-optimal deaminase activity and several layers of regulatory control. Despite these constraints, DNA deaminases can act aberrantly on the genome when mis-regulated, and their activity is known to contribute to genomic instability and to promote cancer mutagenesis.
The ability to target DNA deaminases to specific loci has opened up new frontiers with the potential to transform biology and medicine by allowing for precise gene editing without introducing double-stranded DNA (dsDNA) breaks. In the base editing complex, catalytically-inactive Cas9 (dCas9) is partnered with a DNA deaminase. Unable to generate dsDNA breaks, dCas9 functions as a ‘genomic GPS’ bringing the deaminase to a specific locus dictated by a single-guide RNA (sgRNA), where dCas9 binding also exposes a window of single-stranded DNA (ssDNA) that can then be edited by the DNA deaminase. The tethered DNA deaminase can then act on the exposed single-stranded DNA to induce C:G to T:A mutations in the case of AID/APOBEC cytosine base editors (CBEs) or A:T to G:C mutations with evolved TadA adenosine base editors (ABEs)4, 5. In the case of CBEs, the fusion of one or more protein inhibitors of uracil repair (UGIs) further promotes C:G to T:A transitions over other outcomes6. Alternatively, more processive DNA deaminases can facilitate targeted diversification in place of precise transition mutations7, 8. In their physiological roles in immune defense, AID/APOBEC enzymes are highly regulated at multiple levels, including via transcriptional control, alternative splicing, post-translational modification, and interaction partners9, 10. Efficient regulation is imperative, as DNA deaminases also pose risks to the genome11, 12. Mistargeting of AID and its APOBEC3 (A3) relatives results in mutations and translocations in a variety of cancers13-17. These known pathological activities help explain why BEs, which contain unregulated deaminases, have more recently been shown to have significant sgRNA-independent off-target activities. Indeed, genome-wide transition mutations occur more frequently after CBE or ABE exposure, and transcriptome-wide mutations increase due to off-target deaminase activity on RNA18-23.
Although different AID/APOBEC family members have been explored, initial efforts largely focused on rat APOBEC1 as the base editor, in concert with accessory modules that skew downstream repair pathways to favor the desired transition mutations. Notably, while mutations can be localized within the ssDNA exposed by dCas9, editing efficiency remains a major challenge.
Current strategies have increased efficiency by using a nickase-Cas9 (nCas9), but at the cost of imprecision, tolerating more insertions/deletions (indels). Furthermore, recent work has uncovered substantial off-target effects from deaminases, which can mutate DNA or RNA independent of Cas9 binding. Both the power and challenges of base editing are captured by recent advances in the correction of pathologic point mutations, the generation of knockouts via targeted stop codon introduction, and broad applications in discovery platforms in the lab. In such settings, base editing can be used to great effect, but can also lead to off-target action, given the absence of regulatory control over the editing enzymes.
Inducible editing activity of split engineered base editors is described in the present example. Our strategy for moving to controllable mammalian base editing complexes involves use of molecules which are capable of dimerization in response to dimerization inducing molecules, for example the rapamycin-regulated dimerization of FKBP and FRB. In this system, proteins linked to FKBP and FRB (e.g., portions of a split deaminase) can be approximated to one another by the addition of rapamycin or related analogs (rapalogs). The seBEs described herein link the split deaminase elements with the targeting dCas9 module, although many possible permutations are described and are shown in the Figures.
To advance towards a split DNA deaminase, we looked to precedents from the larger deaminase family that share a characteristic α/β deaminase fold27, including pyrimidine salvage enzymes that have been split via rational manipulation of loop regions28. Our strategy involved two steps: identifying sites that tolerate insertion of GFP, and then splitting GFP to test if the DNA deaminase can be split and spontaneously reconstituted. Building on the known structure of AID29, we focused first on a variant containing several hyperactivating mutations30, 31 (AID*) that could potentiate efficient genome editing. We targeted five loops in AID* for insertion (
To test for insertional tolerance, we expressed constructs in E. coli and measured deaminase activity with a rifampin-based mutagenesis assay. In this assay, DNA deaminase expression promotes untargeted mutagenesis, and the frequency of acquired rifampin resistance (RifR) is a well-established means to assess overall deaminase activity30, 34. Using this approach, AID(WT) expression increases RifR 12-fold relative to a catalytically inactive mutant AID(E58A), while hyperactive AID* shows a 265-fold RifR increase (
We hypothesized that strategies may differ based on the location of the tolerated split in the DNA deaminase, which will in turn influence choice of linkers and the order of linkage between the different elements in the editing complex. Having demonstrated insertion tolerance, we next evaluated if the insertion tolerant site could be used to split the DNA deaminase. We had initially inserted optGFP because this variant can be used to split GFP in the loop between the last two β-strands (β10-β11), with co-expression of two fragments leading to spontaneous GFP reconstitution32. With therefore next split AID*-INS2 between β10 and β11 of optGFP, resulting in a construct pair of AID*N-optGFP1-10 (AID*-SPL2N) and GFP11-AID*C (AID*-SPL2C). As predicted, either AID* fragment alone showed no increase in RifR (
Given the shared structural architecture of AID/APOBEC family enzymes, we hypothesized that the α2-β3 loop might prove to be a generalizable split site. To this end, we examined if human A3 enzyme APOBEC3A (A3A)25, 35, 36 could also be split into two inactive fragments that can be reconstituted. We first validated that A3A tolerated optGFP insertion at its α2-β3 loop in vitro (
Our controllable split-engineered base editor (seBE) design requires a transition from spontaneous split GFP reassembly to switchable chemical-induced protein dimerization (CID) of deaminase fragments. To achieve CID, we employed the common rapamycin-regulated heterodimerization of FK506 binding protein 12 (FKBP12) and FKBP rapamycin binding domain (FRB)38. To explore generalizability of the seBE strategy, we generated three distinct seBE variants in the scaffold of BE4max39, containing an alternative hyperactive variant of AID (AID′), evolved APOBEC1 (evoA1), or A3A followed by Cas9 nickase (nCas9) and tandem UGIs. The distinctive features of these deaminase variants can permit exploration of different applications: AID is processive and primed for diversity generation7, evoA1 has been shown to be highly precise40, and A3A demonstrates high C to T conversion efficiency25, 35, 36. Starting from intact BE4max scaffolds, we created seBEs by inserting an artificial gene encoding FRB and FKBP12 at the loop between α2 and β3 with fragments separated by a T2A self-cleaving polypeptide (
To measure editing efficiency, we derived a HEK293T reporter cell line with a single copy of destabilized GFP (d2GFP) stably integrated (
To more rigorously assess editing footprints, we deep-sequenced the d2gfp locus for each condition (
We next aimed to explore whether seBEs permit controllable editing for alternative targets across the genome. We focused our analysis on APOBEC1 constructs given their observed precision and frequent application in the field. We first targeted seven loci involving epigenetic regulators and analyzed on-target base editing efficiency with seBE4max and BE4max constructs. Across sites, the intact evoA1-BE4max average editing efficiency was 44% (
A strength of the seBE strategy is that the system is well poised for modifications to alter either the nature or the degree of regulatory control. For example, we noted that while editing was readily induced by rapamycin with the seBEs, low-level activity was still observable in the absence of rapamycin. We hypothesized that this editing could have resulted from incomplete ribosome skipping with the T2A self-cleaving peptide, which would yield an intact editor. To further increase the dynamic range of small-molecule inducible editing, we generated an evoA1-seBE4max-IRES construct, where the two polypeptides were expressed from two independent promoters, one from a CMV promoter and the other from an internal ribosome entry sequence (IRES) (
Notably, split deaminases can address multiple off target problems: (1) the existence of an unregulated, constitutively active deaminase that can mutate sites beyond the one targeted by dCas9 and (2) binding of dCas9 to sites outside of the intended sgRNA target. Our seBE-a strategy allows for temporal deaminase control. In cases where off target activity is to be minimized in seBE-a constructs, nuclear localization signals (NLS) can be introduced into either or both constructs perturbing localization and thereby reducing off-target RNA deamination activity. Next, in our seBEb design, we will exploit split Cas9, whereby a Cas9N-FKBP and FRB-Cas9C can be successfully approximated with rapamycin (71). The seBE-b constructs (
While we have already shown success in the generation of split enzyme base editors (see
The generalizability of this strategy is captured in
In sum, we have demonstrated a generalizable strategy for small-molecule regulation over DNA deaminase activity. Although we focus on BE applications, these split sites could be used to study conditional control over isolated DNA deaminases, as in antibody somatic hypermutation or cancer mutagenesis. Given that the α2-β3 loop tolerates insertion of either split GFP or FKBP/FRB, we anticipate extensions to other CID strategies, such as those using rapalogs, abscisic acid, or photo-inducible protein dimerization systems24. seBEs are also anticipated to function with editor scaffolds beyond BE4max, including those using Cas proteins other than nCas9, or with two different targeting modules to minimize sgRNA-dependent off-target activities, akin to recently developed split dsDNA deaminase editors43 or the dimeric Cas9-FokI heterodimerization systems44. Finally, we note that small-molecule inducible seBEs could allow for the potentially powerful ability to controllably induce base edits in more complex settings, including in vivo, analogous to conditional systems that allow for tissue or time-specific gene knockouts.
While certain of the preferred embodiments of the present invention have been described and specifically exemplified above, it is not intended that the invention be limited to such embodiments. Various modifications may be made thereto without departing from the scope and spirit of the present invention, as set forth in the following claims.
This application claims priority of U.S. Provisional application Nos. 62/965,886 and 62/966,303 filed Jan. 25, 2020 and Jan. 27, 2020 respectively, the entire contents being incorporated herein by reference as though set forth in full.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/US21/14252 | 1/20/2021 | WO |
| Number | Date | Country | |
|---|---|---|---|
| 62965886 | Jan 2020 | US | |
| 62966303 | Jan 2020 | US |
