8 Gene Function and Bioengineering
Faizo Kasule; Walter P. Suza; and Madan K. Bhattacharyya
Introduction
A major advantage of using plant transformation to modify plants is the ability to transfer a single well-defined gene, most likely without altering the functions of other genes in the recipient plant. In contrast, plant breeding involves the simultaneous transfer of thousands of genes, and the favorable ones must then be separated from the unfavorable ones by segregation among the progenies. Transformation also allows the use of genes from other organisms that could not be introduced naturally to a plant because of crossing barriers preventing interspecific hybridization. In Plant Breeding Methods, several examples are provided that involve the use of novel genes developed through genetic engineering to confer resistance against insects and herbicides in commercial transgenic crops. It was discussed in Chapter 7 that transformation is a valuable tool for plant breeding and biotechnology. This chapter discusses the history of genome editing, as well as the various types of genome editing technologies utilized in crop biotechnology, ranging from meganucleases to zinc finger proteins, and CRISPR-Cas (clustered regularly interspaced short palindromic repeats and CRISPR-associated protein) systems, as well as prime editing.
History of Genome Editing Technology
For half a century, gene editing technologies that specifically modify genomic sequences have revolutionized various aspects of plant sciences and crop biotechnology through the elucidation of gene functions, the understanding of complex traits, and enhanced crop improvement. This has progressed through ranks from early transgenesis to the current use of precise, programmable tools (Figure 1). During the 1970s and 1980s, significant breakthroughs occurred primarily in random DNA integration, while progress in homologous recombination was limited (Mansour et al., 1988). In the 1980s, meganucleases, also known as homing endonucleases, were discovered. These create double-stranded breaks (DSBs) at rare 14-40 bp recognition sites (Jacquier & Dujon, 1985). In the same era, zinc finger proteins were discovered in 1985 in the African clawed frog (J. Miller et al., 1985). These finger-like DNA-binding motifs played a vital role in gene regulation and were applied in gene manipulation studies.
Zinc finger nucleases (ZFNs) were created in 1996 by fusing two domains: a zinc-finger-based DNA recognition domain and a non-specific Fok I endonuclease domain to cleave the targeted DNA (Kim et al., 1996a). ZFNs were widely used for gene editing in the late 1990s and a major part of the 2000s. In 2010, transcription activator-like effector nucleases (TALENs) were developed (Miller et al., 2011). Like ZFNs, TALENs utilize the Fok 1 endonuclease domain to cut the targeted DNA, and this system offers advantages for designing the cleavage of desired target sequences. This advantage over ZFNs made it a method of choice over ZFNs for a short period, until around 2010-2011.
In 1987, clustered regularly interspaced short palindromic repeats (CRISPRs) were described in the Escherichia coli genome during an analysis of genes involved in phosphate metabolism (Ishino et al., 1987). In 2012, a revolutionary gene editing tool, CRISPR/Cas9, was discovered (Jinek et al., 2012). Unlike ZFNs and TALENs, which require complex protein engineering, CRISPR/Cas9 utilizes a 20-nucleotide guide RNA to target a CRISPR-associated (Cas) nuclease corresponding site in the genome, adjacent to a protospacer adjacent motif (PAM) with the NGG sequence, for generating mutations. This makes CRISPR/Cas9 a simple and highly efficient tool for genome editing applications, making it the most widely used genome engineering system. For this reason, Jennifer Doudna and Emmanuelle Charpentier were awarded the 2020 Nobel Prize in Chemistry for their discovery of CRISPR/Cas9 as a genome editing tool.
In 2016, base editors were created that utilize a deactivated or nickase Cas9 enzyme with a deaminase activity to introduce single-nucleotide polymorphisms (SNPs) by chemically altering the targeted DNA sequence without inducing double-stranded DNA breaks (DSBs) (Komor et al., 2016). Prime editing was unveiled in 2019, which enables precise small insertions, deletions, and base swaps without introducing DSBs (Anzalone et al., 2019). Prime editors also overcome the limitations of base editors, which are restricted to only four possible base substitutions.
Types of Genome Editing Technologies in Crop Plants
Targeted gene editing is gaining popularity for crop improvement of traits for which contributing genes have been identified and characterized. There are two main bottlenecks in gene editing in crop plants: (i) lack of information about the genes that govern most traits; (ii) absence of stable transformation methods for some crop species. The main advantage of gene editing is the improvement of cultivars for one or more targeted genetic loci without changing the rest of the genome. In the backcrossing method, deleterious genes are often introduced along with the desirable trait loci. Such genes can reduce the performance of the cultivar and could result in yield reduction. This phenomenon is known as linkage drag. The precise genome editing systems enable us to replace an undesirable gene with a favorable allele, thereby avoiding linkage drag, as there is no need to introgress the favorable allele through backcrossing. In this section, we will discuss the following major genome editing technologies: (i) Zinc finger nucleases technology; (ii) TALEN technology; and (iii) CRISPR-Cas9 technology.
DNA Repair Mechanisms
Double-stranded DNA breaks (DSBs) are intentionally created during gene editing using primary genome editing tools, such as zinc finger nucleases, TALENs, and CRISPR-Cas9. DSBs are a form of DNA damage that occurs on both strands of the DNA double helix, where the DNA is cut at the same location or in its proximity. These breaks can be harmful and lead to genome instability, aberrant growth, or cell death. Therefore, the organism must employ its repair mechanisms to repair these DSBs. During the process of DNA repair, the DNA sequence at the site of the break may be altered. Many DNA repair mechanisms produce genome editing by joining the cut ends, which include Base Excision Repair, Nucleotide Excision Repair, Mismatch Repair, Homology-Directed Repair (HDR), Non-Homologous End Joining (NHEJ), and other minor repair mechanisms. By far, DSBs effect genome editing and undergo two major repair pathways in the cell: NHEJ and HDR (Figure 2).
HDR is a precise DNA repair mechanism that utilizes a homologous donor template to repair DSBs (Hussain et al., 2021). In the native HDR pathway, a homologous sequence of sister chromatids is used as a repair template to repair DSB in the S and G2 phases of the cell cycle. In contrast, an external repair template must be provided during genome editing to introduce an insertion or replacement at the targeted site. Using a DNA template to guide the repair of DSBs makes HDR highly accurate and nearly error-free. Scientists leverage HDR to introduce specific and accurate gene modifications by designing a donor template wherein the DNA sequence intended for insertion is flanked by arms homologous to the 5′ and 3′ sites of the DSB (Puchta, 2005). However, HDR efficiency is low compared to other repair pathways in plants.
NHEJ is the predominant form of plant DNA repair mechanism to join broken pieces of DNA successfully (Puchta, 2005). NHEJ might lead, in a certain fraction of cases, to genomic changes such as deletions, insertions, or various kinds of genomic rearrangements (Puchta, 2005). However, NHEJ is error-prone, often leading to insertions and deletions (indels) at the break site, which can disrupt gene function or cause mutations. This is because the NHEJ process does not rely on a homologous template for repair.
Zinc Finger Nucleases Technology
Zinc finger nuclease (ZFN) (Kim et al., 1996b) was created by fusing two proteins: (i) a zinc finger protein that binds to DNA, e.g., a transcription factor; (ii) Fok I, a type IIS restriction enzyme that recognizes asymmetric DNA sequences and cleaves next to the binding site.
The Cys 2 His 2 zinc finger proteins used for designing ZFN contain (Tyr, Phe)-Xaa-Cys-Xaa 2-4 -Cys-Xaa 3 -Phe-Xaa5 – Leu-Xaa 2 -His-Xaa 3-5 -His, amino acids, and they usually occur in tandem. Each of the Cys 2 His 2 proteins interacts with a zinc ion to form the structure termed a zinc finger that binds to the DNA double helix in sequence sequence-specific manner (Kim et al., 1996b). As shown below, we can design a Zn finger protein (Figure 3) that binds to the target site, generating a double-stranded break (Osakabe et al., 2010).
The outcomes of a double-stranded break (DBS) caused by ZFN can be repaired by two mechanisms: (1) via non-homologous end joining (NHEJ) and (2) homology-directed repair (HDR), as shown below (Urnov et al., 2010). NHEJ generates knockout mutations while HDR can lead to homologous recombination and allelic substitution (Figure 4). We can replace a bad gene with a good one.
TALEN Technology
TALEN (Transcription activator-like effector nuclease) is similar to ZFN. Concepts used in TALEN and ZFN are the same. In TALEN (Figure 5), ZF is substituted with a synthetic protein designed based on the DNA-binding properties of a class of plant pathogen effector proteins known as transcription activator-like (TAL) effector proteins (Bogdanove & Voytas, 2011). TAL proteins of a bacterial rice pathogen of the genus Xanthomonas are targeted to the nucleus of rice plants to reprogram transcription of specific genes for the pathogen’s benefit. TAL factors carry tandem, polymorphic amino acid repeats that individually specify contiguous nucleotides in DNA as targets for binding. One can generate synthetic TAL factors for sequences of the target genes based on the codes learned from the investigation of TAL factors. As shown below, the FokI nuclease is added or fused to designed TAL factors to generate the TALEN molecule for mutating a gene.
As shown above, fusion of TAL effector proteins to Fok I creates sequence-specific nucleases that enable targeted DNA cleavage for gene knockouts and genome editing. TALEN technology has been widely utilized for targeted gene mutagenesis, particularly for gene inactivation, in various organisms, including agriculturally important plants such as rice, wheat, tomato, and barley. TALENs were employed to generate stable, heritable mutations at the maize glossy2 (gl2) locus (Char et al., 2015). Transgenic lines containing mono- or di-allelic mutations were obtained from the maize genotype Hi-II at a frequency of about 10% (nine mutated events in 91 transgenic events) (Figure 6). In addition, three of the novel alleles were tested for function in progeny seedlings, where they were able to confer the glossy phenotype. Char et al. (2015) demonstrated that TALENs are an effective tool for genome mutagenesis in maize, empowering the discovery of gene function and the development of trait improvement.
CRISPR-Cas9 Technology
The Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) was first discovered in 1987 (as shown in Figure 1). The significance of CRISPR was unknown until 2007, when its immunity function was discovered. The mechanism used by the element to confer immunity became known only after another five years (2012), when a detailed understanding of this adaptive defense system was achieved (Jinek et al., 2012).
The bacteria and archaea use CRISPR to defend against the invading viruses, termed as bacteriophages (Figure 7). Following viral infection, bacteria employ a special CRISPR-associated nuclease 9 (Cas9) to generate a double-strand break (DSB) in the target bacteriophage DNA molecules. Thus, viruses become ineffective (inactive). Cas9 is guided to the target sequence by a short RNA fragment known as a guide RNA (gRNA), which is complementary to a viral genome segment, thereby generating a DSB (Le Rhun et al., 2019).
Parallel to viral DNA cleavage, a short viral DNA fragment is captured and integrated into the host genome at one end of the CRISPR locus, at the end of the leader end, to form what is known as a “spacer”. The spacers flanked by short palindromic repeats (20–50 bp) serve as a genetic memory for past infections and form a growing library of immunity. Therefore, the sequence is used to generate the gRNA, which rapidly activates this defense mechanism against any subsequent infection by the same bacteriophage (Figure 7).
CRISPRs are a diverse family of DNA repeats with a common architecture. Each CRISPR locus consists of a series of short repeat sequences (20–50 bp) separated by unique spacer sequences of a similar length. The number of repeat sequences varies; some bacteria may have up to 100 repeat segments. Spacers are highly variable and derived from foreign DNA.
Invading DNA is fragmented and is selected for integration based on the PAM (protospacer adjacent motif) sequence. The CRISPR locus acquires new DNA at the Leader end of the locus and provides an immunity “library” of foreign DNAs. The CRISPR-Cas immune response, a bacterial defense mechanism, operates in three stages: Acquisition, expression, and Interference.
In the acquisition phase, when a virus infects a bacterium, bacterial Cas1 and Cas2 proteins recognize foreign phage DNA containing protospacers adjacent to PAM sequences and insert them into the CRISPR array as spacers. In the expression phase, the CRISPR array is transcribed into a pre-crRNA, which is then processed into individual CRISPR RNAs (crRNAs) by RNase III, assisted by tracrRNA in type II systems. Each crRNA carries a unique spacer and guides the Cas nuclease. crRNAs pair with tracrRNA or exist as synthetic single guide RNAs (sgRNAs) (Figure 7). During the interference phase, the crRNA-Cas complex (crRNP) scans for matching sequences in the incoming foreign DNA and directs Cas9 to cleave and degrade the target phage DNA, thereby establishing immunity against those viruses.
This system is similar to antibody production in humans. Once an antibody is produced against a pathogen following infection or exposure to a vaccine, the information is retained in the cells’ memory, allowing the rapid production of the antibody against the pathogen in subsequent infections.
CRISPR-Cas9, a tool for genome editing, was discovered by Dr. Jennifer Doudna and Dr. Emmanuelle Charpentier (Figure 8).
Two steps to consider in designing a CRISPR-Cas9 system for editing a gene are (Asmamaw & Zawdie, 2021): (i) the design of gRNA, and (ii) the nuclease to be used should be specific to the application (Figure 9).
The CRISPR-Cas9 Genetic Scissors
The gRNA that targets specifically the gene of interest should be designed. The ideal scenario would be zero non-specific cuts, with the gRNA being highly specific to the target site in an exon. You can edit members of a gene family simultaneously. Following DSB, cells repair the DNA using the non-homologous end-joining (NHEJ) mechanism (Figure 10). During this process, indels are created, leading to frameshift mutations and gene knockouts (KO) (Addgene, 2017).
We can also conduct a knock-in (KI) mutation by repairing the DSB through the homology-directed repair (HDR) mechanism. To facilitate or induce HDR-mediated repair, copies of homologous DNA molecules with a desirable mutation are provided as a template. The mutation can be a single-point mutation to change an amino acid. It can correct disease-causing mutations in humans and generate herbicide resistance in plants. For example, we can replace an amino acid in a plant gene encoding acetolactate synthase (ALS), which is involved in the biosynthesis of branched-chain amino acids, to make the enzyme resistant to several herbicides, including imidazolinones and sulfonylureas. In KI experiments, we must carefully check the templates for PAM sequences to ensure that they are not treated as targets.
Point or insertion-deletion mutations through CRISPR/Cas9 are predictable. The only weakness is the mutations that may be caused by the system in non-targeted genomic regions. Many sequences are similar to the 16 bp target sequence in a complex genome. To minimize this, careful analysis of the genome is essential to confirm that the target sequence is unique and not located within coding regions. One can always breed the desirable CRISPR/Cas9-induced mutations in crop plants through backcrossing.
Another way to eliminate non-specific mutation is to use gRNA and Cas9 protein in test tubes and apply the two as ribonucleoprotein (RNP) complexes to the cells, thereby generating a mutation. This approach has been reported to be an effective strategy (Liang et al., 2017). The complex is unstable and does not continue to function, thereby preventing additional off-target mutations from occurring. At the same time, the regenerated gene-edited lines are free from integration of any foreign DNA and selectable antibiotic resistance genes (Liang et al., 2017).
Strategies for delivering single guide RNA (sgRNA) and Cas9 ribonucleoprotein complex CRISPR/Cas9 genome editing have recently been reviewed by Zhang et al. (2021). Svitashev et al. (2016) demonstrated that Cas9 and gRNA in RNP complexes can be delivered into maize embryo cells via particle bombardment. The resulting regenerated plants had specific targeted gene mutations and gene edits at high frequencies (Figure 11). They showed that this completely DNA- and selectable marker-free method generated mutant plants at high frequencies. Therefore, RNP delivery provides a precise and efficient tool for plant breeders to create improved crop varieties more quickly and reliably than traditional breeding methods.
CRISPR/Cas9 has been utilized to generate targeted gene knockouts in teosinte (Zea mays ssp. parviglumis) using a biolistic delivery method (Zobrist et al., 2023). Leaf‐derived callus explants contained short indel mutations generated by a single guide RNA (sgRNA) and precise sequence deletion using two sgRNAs (Figure 12). Selectable markers, including mutant ALS and bialaphos resistance, enabled the recovery of edited callus with an average transformation frequency of 4.17%. Visual markers (tdTomato, GUS) confirmed expression in T1 and T2 progeny. Genes such as Glossy2 (Gl2) and Kernel Row Number2 (KRN2) were successfully edited, demonstrating that CRISPR/Cas9 can introduce heritable, beneficial traits. This approach allows plant breeders to harness teosinte’s genetic diversity for yield improvement, trait discovery, and de novo domestication of maize.
Marker-free HDR-mediated insertion of the golden rice trait
In the original golden rice reported in 2000 by Ye et al. (2000) the phytoene synthase (psy) was from the daffodil flower. In a subsequent version, Paine et al. (2005) demonstrated that replacing the daffodil gene with the maize psy gene increased carotenoid content up to 23-fold (maximum 37 μg/g) compared to the original Golden Rice. The genes used by Paine et al. (2005) were used in the CRISPR-mediated generation of the marker-free golden rice by Dong et al. (2020), shown below. They delivered the gRNA-Cas9 construct and a selectable marker in a second plasmid, along with the plasmid carrying the carotenoid cassette that contained the two beta-carotenoid pathway enzyme genes. They targeted the gene cassette to a genomic safe harbor (where genes can be incorporated without causing any damage to the genome) (Figures 14 and 15).
In the Golden rice study, embryonic calli was bombarded, and 55 T 0 plants were regenerated (Dong et al., 2020). They performed genetic segregation analysis on the T1 generation to obtain rice plants homozygous for the carotenoid cassette at Target B, which lack the Cas9-gRNA module. identified T1 individual 48A-7 as being homozygous for the inserted carotenoid cassette. Interestingly, this integration did not follow perfect homology-directed repair; the cassette was inserted via non-homologous end joining and was flipped in orientation. This was an unexpected outcome, but it still produced the desired phenotype. In plant breeding, such work demonstrates the practical challenges and potential of precision gene insertion, showing that even rare, imperfect recombination events can produce stable traits for crop improvement.
Cas12 nucleases
These form several variants of the type-V CRISPR systems and are often characterized by the presence of a single RuvC-like nuclease domain that mediates the cleavage of both target DNA strands, unlike Cas9, which requires an HNH domain (cleaving the DNA strand complementary to the crRNA) and a RuvC domain (cleaving the complementary strand). Cas 12 nucleases like Cas12a(Cpf1), Cas12b, Cas12e, and Cas12f recognize T-rich PAM sequences, and they produce staggered DNA breaks with 5 ́ overhangs, which improves insertion (Xin et al., 2022). This allows for the targeting of genomic sequences that are not targetable by G-rich PAM-dependent nucleases, such as Cas9 (NGG). Furthermore, Cas12 nucleases can process multiple crRNAs simultaneously from an array, thereby simplifying multiplex gene editing compared to Cas9. However, overall, Cas9 achieves a higher editing frequency in plants for biallelic or homozygous mutations, while Cas12a is efficient at specific target site editing (Lee et al., 2019).
Read: Scientists have harnessed TnpB, an RNA-guided DNA endonuclease from the radiation-resistant bacterium Deinococcus radiodurans, which can cleave targeted DNA sequences with 34% efficiency in both dicots and monocots. The system has been utilized to develop resilient rice varieties and holds promise for future applications of the TnpB system in crop improvement for trait enhancement, including climate resilience, disease and pest resistance, yield improvement, and nutritional enrichment (Karmakar et al., 2024).
For more information, read: A miniature alternative to Cas9 and Cas12: Transposon-associated TnpB mediates targeted genome editing in plants.
Base editing
This technology, developed by Davis Liu and Akihiko Kondo in 2016, enables the direct, irreversible conversion of one base of DNA into another at a specific target site without introducing double-stranded breaks (DSBs) (Komor et al., 2016). Base editing combines two components, including Cas9 nickase (nCas9), a catalytically inactive form of the Cas9 enzyme, or a transcription activator-like effector (TALE) repeat array fused to a nucleoside deaminase enzyme that removes and converts one base to a different base. DNA base editors are categorized as cytosine base editors (CBEs), which substitute a cytosine with a Thymine (C→T), and adenosine base editors (ABEs), which introduce a substitution of adenine with guanine (A→G) (Figure 16). New base editors, which can introduce cytosine-to-guanine (CGBEs) and adenine-to-cytosine (ACBEs) substitutions, have been developed, although they are still of limited use.
A guide RNA (gRNA) directs the base editor to the matching DNA sequence for CRISPR base editors. The Cas domain then unwinds the DNA sequence, thereby exposing a short stretch of single-stranded DNA where the tethered deaminase chemically changes one base to another. The base editing technology can revert a single base change or SNP with high accuracy, without disrupting the gene, thereby minimizing insertions and deletions (Indels). The approach has been applied to rice, wheat, maize, and tomatoes.Because base editors produce fewer indels and prevent harmful outcomes associated with DSB repair, they are more efficient than standard Cas nucleases. However, their precision is limited by: (1) base editors deaminate within a small window of 4–5 nucleotides (nt), which causes C or A nucleotides very close to the target C or A can also undergo conversion, resulting in ‘bystander editing’, (2) also base editing is restricted by Cas domain which requires a PAM located about 15 ± 2 nucleotides from the intended base change, (3) Some base editors can induce off-target mutations in DNA and RNA, (4) Current base editor designs only perform six of the 12 possible types of point mutation, hence cannot make large insertions, deletions, and most base transversions. Gaps still exist in base editing capabilities, despite engineering efforts that have reduced off-target activity and expanded the compatibility of PAMs.
Prime editing
Prime editing enables precise small insertions, deletions, and base swaps in a unique and precise manner. Problems associated with the use of base editing motivated researchers to develop prime editing. The latter differs from traditional Cas9 in that this system performs targeted DNA modification for all 12 possible base substitutions without inducing double-stranded breaks (Anzalone et al., 2019).This Technology was developed by Andrew Anzalone, a postdoctoral fellow in David Liu’s lab in 2019, and it utilizes the fusion of a Cas9 nickase (nCas9) to an engineered reverse transcriptase (RT) and guided by a prime editing guide RNA (pegRNA), which specifies the target site for the edit and contains a programmable RNA template for the desired DNA sequence change (Figure 17).
Prime editing (PE) surpasses the base editing limitation to only four substitution types, offering greater precision and flexibility. To mediate editing, the PE system nicks the target site on the genome and extends a new DNA sequence from this nick site using the pegRNA as a template (Figure 18). This edited DNA strand is then incorporated into the genome through endogenous cellular processes that can be promoted by also nicking the non-edited strand (Figure 18).
The prime editing mechanism involves the fusion of a Cas9 nickase and a reverse transcriptase (RT) with a prime editing guide RNA (pegRNA) to form a complex. PegRNA directs the complex to the complementary DNA target site. The Cas9 nickase creates a single-strand break (nick), three bp upstream of the PAM site in one of the DNA strands, exposing a 3′-hydroxyl group without making a double-stranded break. The primer binding site (PBS) of pegRNA anneals to the exposed 3′ end, and the reverse transcriptase enzyme, also part of the prime editor complex, extends the nicked DNA strand using the RNA template sequence (edit site) located within the pegRNA.
This synthesizes a new DNA flap containing the desired edit. Flap resolution and repair occur when the newly synthesized 3′ flap, containing the edited sequence, competes with and displaces the original 5′ flap. Cellular nucleases remove the unwanted 5′ flap, and DNA ligases seal the nick, incorporating the edited DNA into the genome. Complementary strand correction occurs when a second gRNA is used to nick the opposite strand, allowing the edit to be copied to both strands and completing the duplex. Cellular nucleases then remove unedited DNA, fully integrating the new sequence.
Using an improved prime editing system, Gupta et al. (2023) developed two strategies to confer resistance to bacterial blight (BB) caused by Xanthomonas oryzae pathovar oryzae (Xoo). The first approach involved the Knock-in of TAL effector binding elements (EBE) derived from the BB-susceptible gene SWEET14 into the promoter of a dysfunctional executor R gene, xa23. This enables TALE-dependent activation of xa23 and BB resistance, achieving a 47.2% editing rate in T 0 plants with 18% biallelic editing. The second approach involved editing the transcription factor TFIIA gene TFIIAγ5, which is required for TAL effector-dependent BB susceptibility, converting it to a resistant xa5 allele. This resulted in an editing efficiency of 88.5% and 30% biallelic edits among T 0 plants. Both strategies produced T 1 plants with broad resistance against multiple Xoo strains. These plants showed significantly shorter lesion lengths compared to the wild type. These approaches, which utilize prime editing, hold promise for reducing BB disease occurrence in rice and can be applied to other crops.
Chapter Summary
Genetic engineering techniques are crucial for agricultural biotechnology, enabling the application of functional genomics to study complex traits and accelerate crop improvement. This technology is not new and has progressed through various stages, from early transgenesis using methods like meganucleases and zinc finger nucleases to the current use of precise, programmable tools like CRISPR/Cas9 systems. Among these technologies, the CRISPR/Cas system is the most adapted technology due to its simplicity and effectiveness. CRISPR/Cas reagents can be introduced into plant cells using either Agrobacterium– or biolistic-mediated delivery methods. Edited plants that carry either the CRISPR reagents or selectable marker genes can be backcrossed to non-transgenic plants to segregate the transgene from the edited plant. CRISPR/Cas9 reagents can also be delivered as an in vitro-assembled RNP complex for DNA-free editing. Improving methods to recover edited events from this method is a goal for academics and industry alike. Other toolkits, such as base and prime editing, enable single-nucleotide changes and small indels, respectively, without introducing double-stranded breaks, thereby reducing off-target effects and unintended mutations. Collectively, these tools facilitate the precise manipulation of plant genomes to improve crop traits, including disease and pest resistance, as well as other attributes, for which encoding genes are identified. Base and prime editing can facilitate allele swapping experiments without requiring backcross breeding of desirable alleles that may co-inherit many linked undesirable genes, resulting in a yield penalty. Thus, genome editing holds considerable promise for the precision breeding of crop plants in sustainable agriculture.
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