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. 2021 Dec 10;188(4):1757–1768. doi: 10.1093/plphys/kiab574

Advances in gene editing without residual transgenes in plants

Yubing He 1,2,3, Michael Mudgett 4, Yunde Zhao 5,✉,†
PMCID: PMC8968301  PMID: 34893903

Abstract

Transgene residuals in edited plants affect genetic analysis, pose off-target risks, and cause regulatory concerns. Several strategies have been developed to efficiently edit target genes without leaving any transgenes in plants. Some approaches directly address this issue by editing plant genomes with DNA-free reagents. On the other hand, DNA-based techniques require another step for ensuring plants are transgene-free. Fluorescent markers, pigments, and chemical treatments have all been employed as tools to distinguish transgenic plants from transgene-free plants quickly and easily. Moreover, suicide genes have been used to trigger self-elimination of transgenic plants, greatly improving the efficiency of isolating the desired transgene-free plants. Transgenes can also be excised from plant genomes using site-specific recombination, transposition or gene editing nucleases, providing a strategy for editing asexually produced plants. Finally, haploid induction coupled with gene editing may make it feasible to edit plants that are recalcitrant to transformation. Here, we evaluate the strengths and weaknesses of recently developed approaches for obtaining edited plants without transgene residuals.

New technologies enable efficient isolation of target gene-edited plants without any transgene residuals.

Introduction

CRISPR (Clustered Regularly Inter-spaced Short Palindromic Repeats)-mediated gene editing has become a widely used tool for genetic studies and crop improvement. The main goal in plant gene editing is to generate stable, heritable, and nonmosaic modifications in a genome so that the resulting trait/phenotype can be reliably maintained and propagated into subsequent generations (Chen et al., 2019; Hua et al., 2019; Mao et al., 2019; Zhang et al., 2019; Zhu et al., 2020; Gao, 2021; Huang et al., 2021). Therefore, it is essential to remove any gene editing components once the target gene editing has been accomplished (Metje-Sprink et al., 2018; Miroshnichenko et al., 2019; He and Zhao, 2020). The continuous presence of gene editing machinery after editing is complete poses several risks. The transgenes make it difficult to determine whether the observed gene editing events are heritable or not because the genetic changes in offspring can be either inherited from the previous generation or newly generated by the gene editing machinery at the current generation (He et al., 2018). The transgenes can also make genetic analysis less predictable. For example, when a line that has been edited and that still contains the gene editing machinery is crossed to a different genetic background, the transgenes will be able to generate changes at the WT (Wild Type) allele of the target gene in F1 plants, complicating the analysis of the offspring in subsequent generations. Likewise, the transgenes can make it challenging to analyze the progeny from multiplexed gene editing experiments because it is rare that all target genes have been edited to homozygosity at the T0 generation (Ma et al., 2015; Miao et al., 2018; Zhao et al., 2018). CRISPR can generate various new mutant combinations in the subsequent generations, requiring sequencing and genotyping at each generation. The continued presence of gene editing elements will also increase the risk of off-target effects (Zhang et al., 2014, 2018). Furthermore, the removal of transgenes is likely a prerequisite for gaining government approval for commercial applications of gene-edited plants (Huang et al., 2016; El-Mounadi et al., 2020; Schmidt et al., 2020; Turnbull et al., 2021).

Three general strategies have been employed to achieve gene editing without leaving any transgene residuals in edited plants: (1) avoid using foreign DNA components or foreign DNA integration in gene editing experiments; (2) eliminate transgenes after target genes have been edited; and (3) cross a gene editing donor plant to edit a transgene-free acceptor plant and induce haploid formation to remove the donor genome in the F1 generation. All three strategies have been successful with a wide range of effectiveness. In this article, we summarize the recently reported technologies that enable efficient isolation of gene-edited yet transgene-free plants. We also evaluate the advantages and disadvantages of the new technologies for achieving transgene-free genome editing.

Gene editing without foreign DNA components or foreign DNA integration

Gene editing without delivering any foreign DNA components into plant cells

DNA-free gene editing strategies typically rely on generating CRISPR gene editing components including nucleases such as Cas9 (CRISPR associated protein 9) and guide RNAs (gRNAs) in heterologous systems or in vitro. Ribonucleoprotein (RNP) complexes consisting of the nuclease and gRNAs are then assembled in vitro before they are delivered into plant cells. Alternatively, a mixture of Cas9 mRNA and gRNAs, which are produced in vitro, can be delivered into plant cells so that Cas9 mRNA can be translated into the Cas9 nuclease. The DNA-free CRISPR reagents are delivered into plant cells using particle bombardment (Svitashev et al., 2016; Zhang et al., 2016; Liang et al., 2017, 2018), PEG (Polyethylene glycol)-mediated transfection (Woo et al., 2015; Toda et al., 2019), or liposome-based transformation (Liu et al., 2020c). Because no foreign DNA is introduced into the edited plants, they are inherently transgene-free. An additional benefit that the DNA-free methods offer is that the variabilities of nuclease expression levels caused by promoter strength and tissue specificity can be avoided. The main disadvantage of the DNA-free methods is a lack of selection pressure such as antibiotic resistance, which can be used to eliminate cells that did not receive RNP reagents. Consequently, the majority of the regenerated plants from such experiments are not transformed, requiring a huge amount of effort to identify the edited plants if the gene editing outcome does not result in obvious phenotypes. These methods are also not suitable for certain types of gene editing experiments that require a DNA component such as homology-directed repair (HDR), which requires a repair template (Sun et al., 2016). This difficulty can potentially be overcome by using RNA transcripts as HDR templates (Li et al., 2019). The RNP or mRNA methods usually use protoplasts, immature embryos, and calli for transformation. However, protocols for regenerating plants from protoplasts and immature embryos have not been reliably established for most plant species. The advantages and challenges of RNP-mediated genome editing have recently been summarized (Zhang et al., 2021).

Transient production of gene editing components using Agrobacterium tumefaciens-mediated transformation without DNA integration

Agrobacterium-mediated infiltration has been widely used to transiently express proteins in Nicotiana benthamiana leaves to analyze promoter activity, protein–protein interaction, and protein subcellular localization (Krenek et al., 2015). Agrobacterium has also been used to transiently produce Cas9 and gRNAs without integrating the transgenes into plant genomes (Chen et al., 2018; Danilo et al., 2019; Veillet et al., 2019; Bánfalvi et al., 2020). The advantage of this approach is that it is relatively easy to operate without the requirement of any special equipment. Almost all plant molecular biologists have experience in handling Agrobacterium and Agrobacterium-mediated transformation. The main disadvantage of this approach is that most edited plants are mosaic and additional generations are needed in order to identify transgene-free and heritable modifications. Similar to the RNP methods, transient expression of gene editing components is usually conducted without selection pressure, making it difficult to differentiate edited plants from unedited plants. Moreover, some of the gene-edited plants produced by this method will have stable T-DNA insertions, which need to be identified and eliminated through genotyping, further increasing the workload. Modifying the components in Agrobacterium responsible for T-DNA integration may improve the efficiency of this approach.

Plant virus-mediated gene editing

Viruses have been widely used to deliver cargo into plant cells. Virus-induced gene silencing (VIGS) is a powerful genetic tool for studying gene functions in plants (Liu et al., 2002). Viruses can transmit and propagate in plant cells, but the viral genomes are rarely integrated into plant genomes, rendering them a useful tool for delivering gene editing components into plant cells (Ali et al., 2015, 2018; Hu et al., 2019; Luo et al., 2021). For example, tobacco rattle virus (TRV), an RNA virus, is an effective VIGS vector and is widely used in functional genomics research in many plant species (Liu et al., 2002). A gRNA production unit can be placed into the TRV genome and the resulting virus can infect plants to deliver gRNAs into plant cells (Ali et al., 2015). Because TRV can only carry a small cargo such as a gRNA-producing unit, it is not feasible to place both the nuclease gene and the gRNA unit in TRV. In practice, the gRNA-producing virus is used to infect plants that have already expressed Cas9. In this sense, the end product is not considered transgene-free.

In addition to RNA virus vectors, DNA virus vectors can also be used for targeted gene editing in plants. Geminiviruses, a family of DNA viruses, were used to deliver gRNAs into transgenic plants that express Cas9, thereby inducing systemic gene mutations and achieving systemic gene knockout in plants (Yin et al., 2015; Butler et al., 2016). Geminiviruses have a wide range of hosts, including both monocots and dicots. The tomato golden mosaic virus (Kjemtrup et al., 1998), maize streak virus (Shen and Hohn, 1995), and bean yellow dwarf virus (Halley-Stott et al., 2007) are all geminiviruses and they may be modified for gene editing.

As discussed above, the main challenge in using TRV RNA virus and Geminiviruses is that only small cargo such as a gRNA unit can be inserted into the virus genome, making it necessary to use plants that have already expressed a nuclease. This challenge can potentially be overcome by either reducing the size of gene editing nucleases or testing other types of viruses. Notably, newly discovered nucleases such as CasΦ (786 amino acids; Pausch et al., 2020), Cas12f1 and its variants (400–600 amino acids; Karvelis et al., 2020; Kim et al., 2021; Wu et al., 2021; Xu et al., 2021), TnpB (400 amino acids; Karvelis et al., 2021), and IscBs (about 400 amino acids; Altae-Tran et al., 2021) can functionally perform gene editing with much less coding sequence than the widely used SpCas9 (1,368 amino acids). The availability of such small nucleases may eventually make it feasible to pack both nuclease and gRNA units into commonly used viruses. On the other hand, a negative-strand RNA virus-based vector modified from the Sonchus yellow net rhabdovirus (SYNV) was very recently demonstrated to have the capacity to carry a large cargo (Ma et al., 2020). Both Cas9 and gRNA units were inserted into the SYNV genome, and the modified vectors were delivered into plant cells by Agrobacterium-mediated infiltration. It was clearly shown that the virus vectors were effective in generating heritable mutations in N. benthamiana. A drawback of SYNV-based vectors is that the host plants are restricted to several species in the Asteraceae, Solanaceae, and Chenopodiaceae families

Viruses have been successfully used to accomplish gene editing in N. benthamiana, Arabidopsis (Arabidopsis thaliana), wheat (Triticum), maize (Zea mays), soybean (Glycine max), and other plants (Zaidi and Mansoor, 2017). The major advantage of virus-induced gene editing is that it does not require tissue culture and transformation, enabling large-scale gene editing projects. It is well known that many nonmodel plants and some model crop plants including maize and cotton (Gossypium) are difficult to transform. Viruses offer a promising gene editing approach without transformation. Recently, elements of endogenous mobile RNAs have been used to improve the efficiency of virus-mediated gene editing. For example, N. benthamiana FT RNA is known to migrate to the shoot apical meristem to induce flowering after being transcribed in the leaf vascular bundle tissue. Fusions of FT mRNA and gRNA delivered into N. benthamiana using TRV greatly increased the frequency of heritable mutations in gene editing targets (Ellison et al., 2020; Lei et al., 2021; Li et al., 2021b).

Eliminate transgenes after the target gene has been edited

The commonly used gene editing approach by plant biologists is to generate transgenic plants that are capable of producing all of the necessary gene editing components. The Cas9-producing unit, gRNA units, and selection markers are placed in a T-DNA plasmid, which is then transformed into plants through Agrobacterium-mediated transformation (Hiei et al., 1994). T-DNA fragments are then randomly inserted into the plant genome where Cas9 and gRNA can be produced (Alonso et al., 2003; Sallaud et al., 2004). Once the target gene has been successfully edited, the transgenes need to be removed so that stable and heritable modifications are obtained. T-DNA insertions in the first-generation transgenic plants (T0) are usually heterozygous, allowing for the elimination of the transgenes by genetic segregation if the plants reproduce sexually (Figure 1A(1)). This approach is labor-intensive and time-consuming. For crops, there are often constraints on plant growth space because growing transgenic plants is tightly regulated, and they can only be grown in restricted spaces. However, several new technologies have been reported recently, which can greatly reduce the time and labor required to completely eliminate the transgenes after the gene editing task has been completed.

Figure 1.

Figure 1

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Strategies for eliminating transgenes after gene editing. Plants are transformed through tissue culture by Agrobacterium-mediated transformation under antibiotic/herbicide selection or using visual markers to generate T0 plants in which T-DNA fragments have been randomly integrated into the plant genome. A, Eliminate T-DNA by genetic segregation. The T-DNA insertion locus segregates according to Mendelian genetics when T0 transgenic plants undergo sexual reproduction by self-pollination. At least three-fourths of the seeds from T0 plants contain the T-DNA insertion. 1, No selection-pressure is used to identify transgene-free plants. T1 plants are transplanted into soil and genotyped using PCR-based methods for the presence of the transgenes. This method is labor-intensive and time-consuming. 2, Transgenic seeds can be differentiated from transgene-free seeds based on fluorescent proteins or visible markers such as RUBY. This method eliminates the need to grow and genotype the transgenic plants, reducing the workload by at least 75%. 3, During the positive selection of transgene-free T1 plants, transgenic plants are killed by chemical treatments whereas transgene-free plants can survive. B, Removal of transgenes by excision. Transgenes in T0 plants can be deleted by inducible excision, resulting in mosaic T0 plants. Tissue from the mosaic T0 plants can be used to regenerate plants through tissue culture. This method is especially useful for plants that are propagated asexually. Once the transgene-free plants are identified, it can be determined whether target genes have been edited. C, Self-elimination of transgenic plants using suicide genes. The TKC technology enables T0 plants to produce only transgene-free seeds.

Fluorescent marker-assisted transgene removal

Growing and genotyping the second-generation (T1) plants using PCR (Polymerase Chain Reaction)-based methods are the most time-consuming and labor-intensive parts of the process of identifying target gene-edited and transgene-free plants through genetic segregation. At least 75% of the T1 plants contain transgenes, meaning that three-fourths of the effort put into identifying transgene-free and target gene-edited T1 plants is wasted. If all transgenic seeds produced by T0 plants can be easily identified, one can focus on growing and analyzing only the nontransgenic T1 plants, greatly reducing the space needed for growing the plants and the labor needed for genotyping. Fluorescent proteins such as GFP and mCherry can serve as both positive and negative selection markers for transgenes. The expression of mCherry under the control of the Arabidopsis seed-specific promoter At2S3 makes transgenic seeds bright red, providing an effective approach to distinguish transgenic seeds from nontransgenic seeds (Figure 1A(2)). The mCherry-based method has led to the reliable identification of target gene-edited and transgene-free Arabidopsis plants with a much-reduced labor requirement (Gao et al., 2016; Yu and Zhao, 2019; Ouyang et al., 2020). A further improvement of using fluorescent proteins in gene editing was reported recently. Wang and Chen (2020) used GFP as a marker for both the presence of transgenes and an indication of Cas9 expression levels, which directly correlate with gene editing efficiency. They placed a P2A linker between Cas9 and GFP, allowing the same promoter to drive the expression of both genes. GFP intensity then served as a proxy for Cas9 expression levels. The Cas9–P2A–GFP design allows for the identification of the most efficient gene editing lines without conducting PCR reactions, overcoming the laborious and time-consuming process of isolating gene-edited Cas9-free plants. Fluorescent proteins have been successfully used as markers in gene editing experiments in rice (Oryza sativa; Aliaga-Franco et al., 2019), tomato (Solanum lycopersicum; Aliaga-Franco et al., 2019), maize (Xu et al., 2021; Yan et al., 2021), and other plants. Although fluorescent marker-assisted transgene removal reduces the workload of screening and identifying nontransgenic mutant plants by at least 75%, this approach still requires equipment for visualizing fluorescent proteins and manual labor for differentiating transgenic seeds from transgene-free seeds (He et al., 2017). Recently, a machine for high-throughput fluorescent sorting of rice grains has been developed to assist the selection of plants with/without transgenic fragments (Chang et al., 2016). Using such machines in fluorescent marker-assisted transgene removal will further improve the efficiency.

Pigment-assisted transgene removal

Plants produce many colorful pigments that can be adopted as markers for the presence of transgenes. Anthocyanins, which have a bright and vivid purple color, can be used as markers for the presence of transgenes (Li et al., 2014). Pigment markers have several advantages over fluorescent markers and antibiotic resistance markers. First, visualization of pigments does not require any special equipment or chemical treatments. Additionally, pigments can serve as both positive and negative selection markers. The presence of transgenes can also be easily and continuously monitored throughout plant developmental stages. Combining a CRISPR/Cas9 gene editing cassette with a pigment-production unit can facilitate the identification of gene-edited and transgene-free plants (Figure 1A(2)).

Anthocyanin biosynthesis pathways are very complex and require multiple enzymes (Zhu et al., 2017; Zheng et al., 2019). It is not practical to place an entire anthocyanin biosynthesis pathway in the gene editing vector. Fortunately, anthocyanin biosynthesis is under the control of a MYB transcription factor (Zhang et al., 2014). The overexpression of the MYB gene in rice, N. benthamiana, and Arabidopsis leads to the upregulation of anthocyanin biosynthesis genes and accumulation of anthocyanins (Borevitz et al., 2000; Shin et al., 2006). Combining CRISPR units and a MYB-overexpression unit in the same plasmid offers the convenience to directly monitor the presence of transgenes and to also infer the expression levels of Cas9 (Liu et al., 2019; He et al., 2020). Anthocyanin-assisted CRISPR technology allows the identification of transgenic events in very early stages of plant transformation during callus differentiation. The co-production of anthocyanin and Cas9 enables easy selection of highly efficient CRISPR/Cas9 gene editing lines. In subsequent generations, gene-edited plants without transgenic DNA components can be visually selected. A main caveat of the anthocyanin-based approach is that the plants must carry the complete anthocyanin synthesis pathway, which is not the case for many species (Zheng et al., 2019). Therefore, the application of anthocyanin biosynthesis as a visible marker by overexpressing the MYB transcription factor is not easily adopted for many plants.

To overcome the disadvantages of anthocyanin biosynthesis as a marker, we developed the RUBY reporter based on the betalain biosynthesis pathway (He et al., 2020). Betalains have a vivid red color. Betalain synthesis uses the amino acid tyrosine as the starting material and only requires three enzymes: the cytochrome P450 enzyme CYP76AD1, l,4-dihydroxyphenylalanine (l-DOPA) dioxygenase (DODA), and betanidin glucosyl-transferase. The three genes are placed in tandem and are linked by 2A peptides to generate the artificial gene RUBY. RUBY has served as a very convenient marker for transformation and gene expression in N. benthamiana, rice, and Arabidopsis (He et al., 2020). Using RUBY for marker-assisted gene editing is advantageous over anthocyanin because all necessary enzymes are contained in the RUBY gene itself. The system is suitable for any plant species since the transformed plant only needs to provide the substrate tyrosine, which is present in every cell.

Chemical-induced positive selection of transgene-free plants

Rice cytochrome P450 CYP81A6 encodes a key enzyme for degrading the herbicide bentazon (Pan et al., 2006). Plants overexpressing CYP81A6 are resistant to bentazon treatments, whereas the disruption of CYP81A6 leads to bentazon hypersensitivity and plant death. Lu et al. took advantage of the unique property of CYP81A6 and developed an effective approach to isolate target-gene-edited and transgene-free rice plants (Lu et al., 2017). They coupled the gene editing unit with an RNA interference unit that targets the CYP81A6 gene in the same plasmid. Plants with the CYP81A6 RNAi unit are hypersensitive to bentazon, providing an efficient way to kill all transgenic plants at the T1 generation. The positive selection of transgene-free plants makes large scale screening feasible (Figure 1A(3)). A major disadvantage of this approach is that this strategy may not be suitable for all plant species.

Similar methods using hygromycin and mannose for selecting transgene-free, gene-edited plants have been developed (Wu et al., 2019; Li et al., 2020). When transgene-free plants are treated with hygromycin, oxidative stress occurs in the plants, which leads to an increased concentration of hydrogen peroxide. Transgenic plants containing the hygromycin phosphotransferase gene are unaffected. Hydrogen peroxide and other reactive oxygen species can be visualized using 3,3′-diaminobenzidine, providing a visual readout of transgene-carrier status. A disadvantage of this method is that the hygromycin-treated transgene-free plants are stressed and may not be able recover to a normal healthy state (Wu et al., 2019). Similarly, transgene-free plants grow slower in the presence of mannose due to inhibited ATP production (Hu et al., 2016). Mannose treatment allows for selection against transformed plants containing a Mannose-6-phosphate isomerase gene, with an easier recovery compared to hygromycin treatment (Li et al., 2020).

Transgene elimination by enzymatic reactions

Another approach to removing transgenes from the genome after a target gene has been edited is through transgene deletion via site-specific recombinases such as the flippase/flippase recognition target (FLP-FRT) and Cre-Lox recombination systems (Figure 1B; Wang et al., 2014). FLP is a recombinase that specifically recognizes the 34-bp FRT sequence, and performs recombination reactions such as deletion, insertion, inversion, and translocation between target sites. When two FRT sites are arranged in the same direction on a strand of DNA, the DNA fragment between the two sites can be excised under the action of the FLP recombinase (Amin et al., 1991). If gene editing machinery is flanked by FRT sites, the CRISPR unit can be deleted from the plant genome when FLP recombinase is present (Figure 2A; PompiLi et al., 2020). If the expression of the FLP recombinase is under the control of an inducible promoter such as a heat-inducible promoter, the CRISPR elements can be deleted upon heat induction after gene editing has been achieved. The nuclease activity of Cas9 can also be used to delete the transgenes. Adding two Cas9 target sites (cleavage target sites, CTS) inside the left and right border sequences in a T-DNA plasmid can enable the deletion of the entire T-DNA insertion by Cas9. Both FLP/FRT and Cas9/CTS systems were tested in transgenic apple (Malus domestica) and grapevine (Vitis vinifera) plants (Figure 2A; Dalla Costa et al., 2020). It was shown that the number of T-DNA insertions was decreased in transgenic apple plants, but not in grapevine, upon the induction of FLP expression or Cas9 expression, suggesting that T-DNA excision took place in the apple genome. Several obstacles in using the enzyme-based excision approach were encountered. It was found that the T-DNA boundaries varied among the transgenic lines and the excision target sequences were often mutated by Cas9-mediated NHEJ (Non-Homologous End Joining) due to leakage of Cas9 expression, rendering them resistant to excision. The system could be further improved by inducing the CTS guide RNA via an inducible promoter rather than Cas9, since Cas9 expression is required for both the gene editing and the T-DNA excision. Inducible gRNA production can be achieved by using our previously reported ribozyme-based strategy (Gao and Zhao, 2014). A similar excision-based approach using a piggyBac transposon-mediated transgenesis system was recently employed to delete the CRISPR/Cas9 gene editing elements from the host genome by means of transposition (Figure 2A; Nishizawa-Yokoi and Toki, 2021). The piggyBac transposon/transposase system was able to generate transgene-free, gene-edited rice plants. The above gene excision systems are valuable for editing plants that are propagated through vegetative tissue. One concern with these techniques is that the excision by enzymes may leave a scar at the excision site, which may interfere with endogenous gene functions or cause regulatory concerns.

Figure 2.

Figure 2

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Mechanisms of transgene deletion and self-elimination of transgenic plants. A, Enzymatic site-specific excision of transgenes. A CRISPR cassette and recombinase/nuclease/transposase unit are in the same plasmid, and they are integrated into the plant genome by Agrobacterium-mediated transformation. After target genes are edited by Cas/gRNA, all transgenes between the T-DNA borders are excised from the host genome by enzymes. Such excisions may leave a scar at the excision site. B, TKC technology. The TKC plasmid consists of suicide units, a selection marker, a CRISPR gene editing cassette, T-DNA borders (data not shown), and other T-DNA components (data not shown). The REG2::BARNASE and 35S::CMS2 suicide units cause death of embryos and pollen, respectively, preventing the production of any transgenic seeds from the T0 plants.

Self-elimination of transgenic seeds/plants using suicide genes

One method for saving time and labor in establishing transgene-free plants is to prevent transformed plants from producing transgenic progeny in the first place. The transgene killer CRISPR (TKC) system (He et al., 2018) relies on two temporally separated events (Figure 1C). First, Cas9 and gRNAs are produced for editing target genes during tissue culture and plant regeneration. Then, when T0 plants transition to reproductive growth, suicide genes that are linked to the Cas9 and gRNA unit are expressed, triggering the death of all transgene-containing sperms, eggs, and embryos. All seeds/grains harvested from the T0 plants are thus transgene-free. Two toxic genes are used in the TKC system: the rice cytoplasmic sterility gene Cytoplasmic Male Sterility 2 (CMS2) gene and the Bacillus subtilis ribonuclease BARNASE. It was previously reported that expression of the CMS2 gene in pollen leads to the failure of pollen development, causing male sterility, while BARNASE expression effectively kills plant cells. BARNASE was expressed under the control of the Rice Embryo Globulin-2 (REG2) promoter, which is active during early embryonic development in rice, and CMS2 was driven by the cauliflower mosaic virus 35S (CaMV 35S) promoter. Linking the two CMS2/BARNASE-based suicide cassettes with the CRISPR/Cas9 gene editing element cassette in the same plasmid enables targeted gene editing followed by self-elimination of transgenic cells (Figure 2B). The TKC (Transgene Killer CRISPR) system was extremely effective in rice, achieving almost 100% elimination of the transgenes without reducing gene editing efficiency (He et al., 2018).

The TKC system is especially suitable for multiplexed gene editing, which often fails to edit all targets with similar efficiency (Yang et al., 2021). Consequently, T0 plants usually contain a mixture of zygosities at the target gene loci. This makes genetic analysis of subsequent generations difficult since residual gene editing machinery may be reactivated in the T1 generation. With TKC technology, genotypes of T1 plants are defined and the edited events transmit to the T2 generation according to the predicted Mendelian genetics.

Several advancements to the TKC system have been reported since the first generation of the technology was published. The improvements mainly involve using different promoters or alternative suicide genes (He et al., 2019). Because the CaMV35S promoter has been widely used in rice and in other plants, many genetic materials may already contain the CaMV35S promoter, which can interfere with suicide gene expression. Thus, replacing the CaMV35S promoter with native plant promoters such as those from UBIQUITIN genes has been beneficial. Yu et al. used the pollen-specific Zea mays Alpha-Amylase 1 (ZmAA1) to eliminate transgenic pollen from T0 plants, providing an alternative to CMS2 (Yu et al., 2021). It increased the transgene-free seed proportion from 25% to 50% in the case of a single T-DNA insertion event.

Coupled gene editing with haploid induction to generate gene-edited, transgene-free plants

Haploid induction (HI) technology can fix the genotype of crops in a short period of time and has long been used in crop breeding (Chang and Coe, 2009). Studies have found that mutations in specific genes in plants can induce haploid production, or the generation of offspring with only one set of chromosomes. The disruption of MATRILINEAL/PHOSPHOLIPASE A1/NOT LIKE DAD (MATL/PLA1/NLD), which encodes a sperm cell-specific phospholipase, resulted in the production of defective male gametophytes in maize (Kelliher et al., 2017; Liu et al., 2017; Dong et al., 2018), rice (Yao et al., 2018; Wang et al., 2019), wheat (Liu et al., 2020a, 2020b), and millet (Panicum sumatrense; Cheng et al., 2021). Similar results were obtained in maize and Arabidopsis by CRISPR-mediated disruption of DUF679 domain membrane protein (DMP; Zhong et al., 2019, 2020). In addition, CRISPR–Cas9-mediated deletion of the N-terminal α-helix of centromere-specific histone 3 (CENH3) resulted in the generation of Arabidopsis HI lines (Kuppu et al., 2020). Genome editing of wheat TaCENH3α resulted in a HI rate of 7%. Editing the restored frameshift alleles of heterozygous genotypes triggered a higher paternal HI rate (Lv et al., 2020) than homozygous combinations. Similarly, it was also found that maize plants heterozygous for the cenh3 null allele could efficiently induce haploid production as either the male or the female parent of a cross (the effective HI rate reached 20%; Wang et al., 2021).

It is speculated that the chromosomes from one parent may be lost during the process of HI in F1 plants. Therefore, if the to-be-eliminated parent genome contains a gene editing cassette, haploid-inducing gametes could carry expressed Cas9 and gRNAs, which perform gene editing on the remaining parent genome post-fertilization (Figure 3A). According to this principle, the haploid-induced editing (HI-Edit; Kelliher et al., 2019) and the haploid inducer-mediated genome editing (Wang et al., 2019) systems were successfully developed. In both cases, haploid-inducing pollen carrying gene editing machinery was crossed to an elite line to generate maternally edited, transgene-free haploid progeny.

Figure 3.

Figure 3

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Coupled HI and gene editing to generate edited plants without transgene residuals. The CRISPR constructs including gene editing and HI cassettes are delivered into easily transformable plant varieties using Agrobacterium-mediated transformation or gene gun to generate a haploid-inducing and editing donor line. The donor line enables the production of Cas9 protein and gRNAs to edit the target genes of recipient lines after fertilization by intraspecific cross (A) or distance hybridization (B). Post-fertilization, the male genome is eliminated to form transgene-free and gene-edited haploids. The haploid lines can be treated with chemicals to induce chromosome doubling, resulting in diploid progeny in which target genes have been edited and which are transgene-free.

Recently, double HI lines have also successfully induced transgene-free double haploid gene-edited plants in rapeseed (Brassica napus; Li et al., 2021a). In this study, the gene editing vector was transformed into the octoploid Brassica double HI line Y3380, and a selected transgenic line Y3380-15 was used to pollinate B. oleracea and B. napus. The editing efficiency of the mutant materials in B. oleracea and B. napus was 1.89%–12.11% and 12.92%–12.97%, respectively. The mutations produced by the octoploid double HI line can be inherited stably without transgenes, but no homozygous mutation type has been detected, which indicates that the editing mechanism mediated by the octoploid double HI line may not be the same as MTL in grass crops.

Obtaining transgene-free gene-edited recipient plants through distance hybridization

Earlier studies have shown that haploid wheat can be created by crossing maize with wheat via distant hybridization (Laurie and Bennett, 1988). By combining the HI-Edit system and intergeneric hybridization, scientists successfully edited the target genes in wheat using maize pollen produced by a transgenic maize plant carrying the Cas9 and gRNA cassettes (Kelliher et al., 2019). In the process of embryonic cell division of the hybrid offspring of wheat and maize, the maize chromosomes are eliminated due to the asynchronous process of DNA replication, aggregation, and centromere formation, resulting in transgene-free haploid wheat (Figure 3B; Budhagatapal Li et al., 2020). Wheat is especially suitable for editing by HI because wheat is difficult to transform and has a strong ability to accept pollen from many plants in the same family.

Future directions

Isolation of plants that have been edited without any residual transgenes can be accomplished through multiple approaches. In model plants such as rice and Arabidopsis, recent technologies have made both gene editing and the removal of transgenes very efficient with much reduced time and labor requirement (He and Zhao, 2020). We would like to point out that any DNA-based methods can potentially leave small fragments of foreign DNA in the genome, which cannot be eliminated by the strategies discussed above. It is imperative to analyze multiple alleles resulting from independent transformation events in order to rule out the contributions from such random insertions. Likewise, it would be prudent to sequence the whole genome before any edited plants are commercially grown.

However, gene editing without leaving transgene residuals in plants is still very challenging in crop plants that are propagated asexually. Current methods for editing asexually reproduced plants are either very inefficient or transgene removal is difficult. RNPs and transient production of Cas9/gRNAs for editing plants can achieve transgene-free plants, but editing efficiency is very low due to a lack of selection pressure for transformed cells. On the other hand, transgenic approaches can lead to high editing efficiency, but the transgenes cannot be genetically segregated out. Site-directed recombination can be used to remove the transgenes, but recombination often leaves a scar. Several strategies may be employed to overcome the challenges in editing clonally propagated plants. Using a visible marker such as RUBY will make it more efficient to identify transgene-free sects after the recombinase or Cas9 is induced to delete the transgenes (He et al., 2020). Combining morphogenic agents such as BABY BOOM (Lowe et al., 2016), CRISPR components, and RUBY to directly edit meristematic cells may also achieve the goal of transgene-free gene editing.

HI and distance hybridization offers great potential for generating edited transgene-free plants. So far, the approach has only been demonstrated in monocots and Arabidopsis. The same principle should also be applicable in dicotyledonous crop plants. One advantage of some dicots including some Brassica species is that they often show self-incompatibility (Takayama and Isogai, 2005). The donor species can be modified according to the mechanism of reproductive isolation, so that its pollen can deceive the recipient species. As long as the DNA of the donor plant can be sent into the embryo sac of the recipient plant, it can express gene editing elements. Because the chromosomes cannot be effectively paired, the genome of the donor plant is degraded to achieve the goal of transgene-free plants.

ADVANCES.

  • Target genes can be edited using DNA-free reagents or without foreign DNA integration, resulting in transgene-free edited plants.

  • Marker-assisted transgene removal isolates transgene-free, target gene-modified plants. Fluorescent proteins and plant-produced pigments facilitate the identification of transgene-free plants and indicate the expression levels of nucleases required for gene editing.

  • Transgenes can be excised from plant genomes using recombinases, CRISPR nucleases, and transposases after gene editing.

  • Transgene-Killer CRISPR temporally separates expression of gene editing components and suicide genes, enabling self-elimination of all transgenic progeny and reducing the time and labor needed for isolating transgene-free plants.

  • Gene editing coupled with HI generates edits in a single set of transgene-free chromosomes through standard crossing techniques.

Acknowledgment

We thank Ms Yichao Li for help in drawing the figures.

Funding

Y.H. is partially supported by grants from the Jiangsu Science and Technology Development Program (BE2018381), the Jiangsu Agricultural Science and Technology Independent Innovation Fund [CX(21)3105] and [CX(21)1002], and The Fundamental Research Funds for the Central Universities (JCQY201903). M.M. is partially supported by an National Institutes of Health training grant.

Confli ct of interest statement. None declared.

Contributor Information

Yubing He, State Key Laboratory of Crop Genetics and Germplasm Enhancement, Nanjing Agricultural University, Nanjing 210095, China; Collaborative Innovation Center for Modern Crop Production Co-sponsored by Province and Ministry, Nanjing Agricultural University, Nanjing 210095, China; Excellence and Innovation Center, Jiangsu Academy of Agricultural Science, Nanjing 210014, China.

Michael Mudgett, Section of Cell and Developmental Biology, University of California San Diego, La Jolla, California 92093-0116, USA.

Yunde Zhao, Section of Cell and Developmental Biology, University of California San Diego, La Jolla, California 92093-0116, USA.

 

All authors contributed to writing the paper.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (https://academic.oup.com/plphys/pages/general-instructions) is: Yunde Zhao (yundezhao@ucsd.edu).

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