- Open Access
A non-viral genome editing platform for site-specific insertion of large transgenes
Stem Cell Research & Therapy volume 11, Article number: 380 (2020)
The precise, functional and safe insertion of large DNA payloads into host genomes offers versatility in downstream genetic engineering-associated applications, spanning cell and gene therapies, therapeutic protein production, high-throughput cell-based drug screening and reporter cell lines amongst others. Employing viral- and non-viral-based genome engineering tools to achieve specific insertion of large DNA—despite being successful in E. coli and animal models—still pose challenges in the human system. In this study, we demonstrate the applicability of our lambda integrase-based genome insertion tool for human cell and gene therapy applications that require insertions of large functional genes, as exemplified by the integration of a functional copy of the F8 gene and a Double Homeobox Protein 4 (DUX4)-based reporter cassette for potential hemophilia A gene therapy and facioscapulohumeral muscular dystrophy (FSHD)-based high-throughput drug screening purposes, respectively. Thus, we present a non-viral genome insertion tool for safe and functional delivery of large seamless DNA cargo into the human genome that can enable novel designer cell-based therapies.
Previously, we have demonstrated the utility of our phage λ-integrase platform to generate seamless vectors and subsequently achieve functional integration of large-sized DNA payloads at defined loci in the human genome. To further explore this tool for therapeutic applications, we used pluripotent human embryonic stem cells (hESCs) to integrate large seamless vectors comprising a ‘gene of interest’. Clonal cell populations were screened for the correct integration events and further characterized by southern blotting, gene expression and protein activity assays. In the case of our hemophilia A-related study, clones were differentiated to confirm that the targeted locus is active after differentiation and actively express and secrete Factor VIII.
The two independent approaches demonstrated specific and functional insertions of a full-length blood clotting F8 expression cassette of ~ 10 kb and of a DUX4 reporter cassette of ~ 7 kb in hESCs.
We present a versatile tool for site-specific human genome engineering with large transgenes for cell/gene therapies and other synthetic biology and biomedical applications.
Genetic insertions of large transgenes find utility in the design of gene therapies for monogenic diseases, innovative cell therapies, and in imparting multifunctionality to cells for biosynthetic applications . A simple approach for the integration of large multi-transgene cassettes larger than 10 kb into the human genome remains a niche application domain where most of the tools (both viral- and non-viral-based) struggle to make an impact. This is due to problems of lack of specificity, undesirable genotoxicity, low efficiency and safety concerns. For example, adeno-associated viruses (AAVs) have a packaging limit of 4.7 kb, and within its capacity, it has shown promising clinical outcomes with long-term expression of truncated variants of F8 (4371 bp) and F9 (1257 bp) in hemophilia A and B patients, respectively. Although AAVs usually express transgenes as an episome, chromosomal integration still occurs either via homologous or non-homologous recombination pathways and can produce long-term effects [2, 3]. On the other hand, lentiviral-based vectors have superior payload capacity and carry inserts up to 18 kb; however, it is known that functional output and packaging efficiency significantly reduces as the load size increases > 8 kb [4,5,6,7,8,9,10,11]. Furthermore, viral-based transgenesis is cost and labour extensive and can lead to potential accentuating effects such as genotoxicity, oncogenicity and adverse humoral immune responses [12,13,14,15]. In contrast, non-viral CRISPR/Cas9 tools and other endonuclease-based genome editing (ZFNs and TALENs) systems are specific towards their target sequences, but their capability to routinely integrate payloads is somewhat limited to ~ 5 kb in size . This is due to their inherent mechanistic principle of entirely relying on host-encoded recombination pathways such as homologous recombination that can be impaired in certain human cell types, especially in hES and somatic cells [17,18,19,20,21,22].
The most commonly used tool for large DNA transgenesis employs transposons that have been shown to integrate 8–10 kb DNA payloads . However, their utility has been hindered by random transgene integration. To overcome these challenges, conventional genome engineering tools must be refined to successfully achieve functional insertion of large transgenes into the human genome. Several studies have employed combinatorial strategies of different editing tools to achieve specific insertion of large DNA . Transposons are being explored in combination with CRISPR/Cas, called CRISPR-associated transposase system (CAST), to enable large DNA (~ 10 kb) integration at specific genomic locations and has, so far, only been validated in E. coli [24, 25]. However, another approach where piggyBac transposase was fused to catalytically inactive dCas9 demonstrated a successful ‘proof-of-concept’ in achieving the integration of the transgene at the CCR5 safe harbour site in HEK293 cells, thus enabling targeted delivery of large DNA cargos in the future [24, 26]. In addition, the CRISPR Cas systems have been paired with different homologous and non-homologous end joining (NHEJ) repair strategies to achieve large DNA knock-ins, an effective strategy in some eukaryotes but not in human systems [27,28,29,30]. Therefore, there is a void in the current genome editing toolbox to meet the need of functional large transgene insertions into the human genome safely at specific locations. Such an approach could substantially improve and enable downstream applications, spanning from engineered cell-based high-throughput drug screening, stem cells for regenerative medicine and cancer immunotherapies amongst others.
Important aspects of genome engineering include both integration of the desired DNA payload and disposing of undesired non-functional sequences, such as bacterial plasmid backbones that can elicit humoral responses due to immunogenic CpG motifs [31,32,33,34,35,36,37]. To achieve this, an alternative class of tools, site-specific recombinases (SSRs), are being employed to generate seamless vectors via intramolecular recombination using their respective recombination sites within the plasmid [38,39,40]. Thus, seamless vectors are circular supercoiled molecules obtained by eliminating the prokaryotic sequences that reduce the size of the vector by about 3 kb. This strategy can enable higher DNA uptake and reduced cell toxicity [41, 42]. In the context of human genome engineering, none of the SSRs tools has dual capability to produce and subsequently target specific endogenous sequences in the human genome. We previously reported a derivative of the phage lambda integrase (λ-Int) system which is proficient in targeting at endogenous Long INterspersed Elements (LINE-1) in the human genome with seamless vectors [43,44,45]. The derivative λ-Int system deploys self-sufficient intramolecular recombination to generate seamless vectors and executes specific human genome insertion by subsequent intermolecular recombination [44, 45]. Using this enhanced strategy, we also demonstrated specific targeting and sustained expression of CD19 chimeric antigen receptors (CARs) in hESCs for potential cancer immunotherapy-related applications .
The wild-type λ-Int system normally integrates the ~ 48 kb circular phage genome into the host genome. Here, we used the ability of our engineered λ-Int to perform large DNA insertions at specific genomic sites in human cells through our seamless vector approach, and exemplify the utility of our transgenesis tool for potential gene therapy approaches in hemophilia A and drug screening for FSHD disease. We demonstrate functional seamless transgenesis of both the ~ 10 kb full-length F8 gene and a ~ 7 kb multi-reporter cassette into specific LINE-1 sequences in hESCs. The demonstrated simplicity of our genome engineering tool provides the basis for broadly based economical applications in the future.
Materials and methods
The hESC line ‘Genea 019’ (Genea Biocells) was used in this study. The cells were cultured in BioCoat Collagen I-coated Plates (Corning) and maintained at 37 °C in 5% humidified CO2 and O2 atmosphere in M2 media (Genea Biocells). Media was supplied with serum and additionally supplemented with penicillin and streptomycin at 25 U/ml each (Gibco). Passaging solution and neutralization solution (Genea Biocells) were used for routine passaging of cells.
To generate F8 expressing pattP4X-pEF1a-FLF8-IRES-Neo-attH4X, full-Length F8 was amplified from F8 expressing piggyBac vector (kindly provided by Prof. Akitsu Hotta, Kyoto University) using high-fidelity DNA polymerase and cloning primers 5.1F and 5.1R. The amplified F8 PCR product was cloned in the AflII linearized pEF1a-IRES-Neo vector (Plasmid #28019, Addgene) to generate pEF1a-F8-IRES-Neo. The EF1a-F8-IRES-Neo cassette was amplified using high fidelity DNA polymerase and cloning primers 7.1F and 7.1R and finally cloned into the master plasmid pattP4X-attH4X using PstI.
To generate pattP4X-16BS-mNeon-PGKss-Puro-bpa-attH4X, a linear fragment comprising of 16BS-mNeon flanked by PstI sites was synthesized (GenScript, USA) and cloned into the master plasmid using In-Fusion HD Cloning kit (Takara), eventually adding 16BS-mNeon cassette in between attP4X and attH4X sequence. PGKss-Puro was then added to this plasmid by PCR amplification of the PGKss-Puro-bpa cassette from pattP4X-PGKss-Puro-bpa-attH4X (in-house), using the primers PGK_fwd_HR and Puro_bpa rev_HR. The PCR product was cloned into pattP4X-16BS-mNeon-attH4X using NheI as per the protocol of In-Fusion HD Cloning kit (Takara Bio USA), adding PGKss-Puro-bpa cassette downstream of 16BS-mNeon cassette.
Cloning was performed using Q5 High Fidelity DNA Polymerase (New England Biolabs) and In-Fusion HD cloning kit (Takara). E. coli DH5α cells were used for transformation. Plasmids were extracted using QIAprep Spin miniprep kit (Qiagen) and EndoFree plasmid maxi kit (Qiagen).
Generation of seamless vector via in vitro recombination using Int-h/218
The integrase-mediated in vitro recombination reaction for seamless vector generation was modified from the method described in . Briefly, recombination was carried out in a reaction mixture (20 μl) containing 500 ng substrate vector, 10 mM TE buffer, pH 8.0, 150 mM KCl, 57 ng/μl of purified single chain Integration Host Factor (scIHF)  and partially purified Int-h/218 (33.25 ng/μl) [43, 47]. Sixty (30 μg DNA in total) reactions were incubated at 37 °C for 60 min and terminated by adding 0.5% SDS. Reactions were pooled and DNA was phenol/chloroform/isoamyl alcohol extracted and precipitated overnight using sodium acetate-ethanol. The reaction mixture containing unrecombined substrate plasmid and catenated circular DNA were digested with a suitable restriction enzyme (single cutter on the bacterial sequence of plasmid) and T5 exonuclease (NEB M0363) at 37 °C. The seamless vector was purified from the digestion mixture using phenol-chloroform extraction and ethanol precipitation of DNA.
Transfection and antibiotic selection
Parental hESCs (250,000 cells/well) were seeded in 6-well plates overnight at 50% confluency. The following day, the cells were reverse co-transfected with the substrate or seamless vector along with Int-C3/Inactive Int expression plasmid using FuGENE HD Transfection Reagent (Promega) at a ratio of 1:3 (DNA: Reagent) using previously published protocol . Forty-eight hours post-transfection, transfected cells were collected and replated onto 10 cm dishes. After 13–14 days of 300 ng/ml of puromycin or 100 μg/ml of neomycin (stock solution of 50 mg/ml in water, Gibco, Life Technologies) selection, surviving colonies were manually lifted, dissociated into single cells and reseeded for expansion initially in 96-well plates and later in 24-well plates.
PCR screening to identify recombination events
Genomic DNA was isolated from parental hESCs and clones using the DNeasy Blood & Tissue Kit (Qiagen). Approximately 50 ng of genomic DNA from parental hESCs and clones was used as a template to amplify left and right recombination junctions. PCR was performed using GoTaq Flexi DNA polymerase (Promega) according to the manufacturer’s instructions. Primer sets were specific to vector and genomic DNA sequences adjacent to the site of integration. Primer positions and amplicon sizes are shown in figures (primer sequences are listed in Supplementary Table S1). PCR amplicons were gel extracted using QIAquick gel extraction kit (Qiagen) and examined by sequencing.
Southern blot hybridization
Genomic DNA was isolated from parental hESCs and clones using the DNeasy Blood & Tissue Kit (Qiagen). Approximately 20 μg of each DNA was digested with a suitable restriction enzyme (New England Biolabs) overnight at 37 °C. Genomic DNA fragments were separated by electrophoresis on a 0.8% agarose gel in 1x TAE (Tris-Acetate-Boric acid) buffer, with 1 kb DNA marker ladder (New England Biolabs) and transferred onto a positively charged nylon membrane (GE Healthcare) via capillary transfer method. The DNA on the membrane was UV crosslinked and the membrane was probed at 48 °C with PCR-amplified DIG-labelled NeoR probe using the DIG-High Prime DNA Labelling and Detection Starter Kit II (Roche) as per the manufacturers’ protocol. The probe-target hybrids on the blot were detected by an AP-conjugated DIG-Antibody (Roche) using CSPD (Roche) as a substrate for chemiluminescence. The blots were exposed to X-Ray film (Kodak) and developed on a Kodak X-OMAT 2000 Processor.
Total RNA from parental hESCs and clones was isolated using TRIzol reagent (Invitrogen). The RNA quality and quantity were assessed by Nanodrop UV-VIS spectrophotometer (Thermo Fisher Scientific). One microgram of total RNA from each sample was reverse transcribed to cDNA using the QuantiTect Reverse Transcription Kit (Qiagen). Using the QuantiNova SYBR Green PCR Kit (Qiagen), RT-qPCR was performed on the CFX96 Touch Real-Time PCR Detection System (Bio-Rad). The actin gene was amplified as an endogenous reference gene. Expression of the target gene was normalized to actin gene expression and represented as fold change using comparative CT method (2-ΔΔCT method) .
FVIII activity assays
Parental hESCs and clones were seeded in 96-well plates at ~ 70% confluence and culture supernatants were collected after 24 h. activity was determined by a fluorometric assay using the Factor VIIIa Activity Assay as per the manufacturer’s instructions. The assay was performed in a Corning 96-well microplate with a black flat bottom and the readings were recorded at kinetic mode (Ex/Em = 360/450 nm) using BioTek Cytation 5 cell imaging multimode reader for 8 h at 37 °C. The Factor VIII activity was normalized to cell viability and represented as fold change compared to parental hESCs.
Cell viability was measured by MTT assay that quantifies the reduction of tetrazolium dye - MTT (3-[4,5-dimethyl thiazole-2-yl]-2,5-diphenyl tetrazolium bromide) in viable cells by mitochondrial NADPH-dependent cellular oxidoreductase enzymes . MTT reagent (Sigma-Aldrich) was prepared at a concentration of 5 mg/ml in PBS. After collecting supernatants for Factor VIII activity, MTT reagent (10 μl) was added in wells (clones and parental hESCs) and incubated for 3 h at 37 °C. The medium in each well was replaced with DMSO to solubilize the purple-coloured formazan dye. The plate was mixed thoroughly and read for absorbance at 570 nm using BioTek Cytation 5 cell imaging multimode reader.
Differentiation of hESCs
Parental hESCs and clones were differentiated with retinoic acid (RA; Sigma-Aldrich) over a period of 14 days as described previously . Briefly, cells were initially cultured in DMEM containing 1 μM RA for 48 h and subsequently maintained in DMEM without RA for 12 days. Culture supernatants were used to measure Factor VIII activity and cells were collected for gene expression analysis.
Statistical tests were performed using Graph Pad Prism6 software. Student’s unpaired t test was applied to compare between two groups. Data is represented as mean ± SEM and p value < 0.05 was considered statistically significant.
Production of seamless F8 targeting vector for site-specific transgenesis
We recently presented a phage λ integrase (Int)-mediated site-specific transgenesis platform capable of inserting large functional multi-transgene cassettes into a specific endogenous sequence, termed attH4x, within a subset of human LINE-1 . The attH4x sequence is present at about 900 locations throughout the human genome. An important improvement of our platform was the inclusion of supercoiled seamless target vectors devoid of prokaryotic DNA elements. This was achieved by using Int for in vitro/in vivo site-specific intramolecular recombination between two directly repeated recombination sequences (so-called attachment (att) sites) flanking the desired transgene expression cassette in a supercoiled parental substrate vector [44, 45]. Thus, besides eliminating unwanted bacterial sequences from the target vector, this approach also reduces the vector size and can enhance transfection efficiency, reduce innate immune responses and contribute to sustained gene expression in human cells [33, 50,51,52].
As a first step towards future autologous cell replacement therapies for hemophilia A, we employed this seamless vector transgenesis platform for site-specific integration of a functional, full-length F8 expression cassette (10.1 kb) into the attH4X sequence in hESCs. The seamless target vector carries the attL4X recombination site and the EF1α promoter-driven F8 gene expression cassette followed by an internal ribosome entry site (IRES)-driven neomycin resistance marker (NeoR). Targeted recombination into the genomic attH4X will generate attL4X and attH4X sequences flanking the inserted F8 gene expression cassette (Fig. 1a). We used a modification of the previously published in vitro vector production protocol using purified Int  that now includes linearization of both the supercoiled bacterial backbone and remaining un-recombined substrate vector by restriction digest in conjunction with the degradation of linear and nicked DNA by phage T5 exonuclease. Simultaneous digestion of the in vitro recombination reaction products by restriction enzyme and T5 exonuclease greatly facilitated the production of sufficient amounts of highly purified supercoiled seamless F8 vector (Fig. 1b).
Targeted integration of F8 seamless expression vectors
The in vitro manufactured seamless vector containing the F8 expression cassette plus selection marker was co-introduced into hESCs together with Int expression vector to establish F8 knock-in clones. Importantly, since the intramolecular recombination reaction on the substrate vector can also occur inside cells before intermolecular recombination with the genome (Fig. 1a), we also tested this alternate route of integration and introduced the unrecombined substrate vector to determine whether in vitro seamless vector production can be bypassed by intramolecular recombination inside the cell. In parallel, this would also explore the possibility of insertion the entire substrate vector into genomic attH4X via recombination with attP4X (Fig. 1c).
Substrate and seamless vectors were co-transfected in hESCs with either an expression vector for variant Int-C3 or a catalytically inactive integrase Int INA . Two days after co-transfection, G418 selection was applied resulting in stable cell clones after 15 days. Importantly, transfection with Int INA resulted in 50% fewer clones compared to Int-C3. A total of fifteen and nine hESC clones were obtained by co-transfection of catalytically active Int-C3 with the substrate and seamless vector, respectively (Fig. 2a). Viable clones were expanded, and genomic DNA was subjected to junction PCR analysis using consensus genomic primers (cs_attH4X_F1/F2 and cs_attH4X_R1) designed to bind adjacent to attH4X sites within the corresponding LINE-1 (Fig. 2b) [44, 45]. Accordingly, successful integration of the F8 expression cassette in any of the LINE-1 loci will result in PCR amplicons specific for left and right recombinant junctions using combinations of the genomic (LINE-1) and cassette-specific primers in F8 or NeoR (Fig. 2b).
Co-transfection with substrate vector and Int-C3 can convert the episomal substrate vector into a seamless vector via intramolecular recombination. Hence, either the entire substrate (via attP4X) or the smaller seamless vector (via attL4X) can recombine with the genomic attH4X sequence (Fig. 1a, c). Analyzing the respective outcome after co-transfection of the entire substrate vector with Int-C3 expression vector by PCR would result in the same product for the left recombination junction but yields two distinct products for the right junction PCR, thus allowing us to distinguish between the two scenarios. PCR screening for substrate vector transfections revealed only integration events of the seamless vector via attL4X and genomic recombination. We found that three out of eight clones (B6, B7 and B8) were positive for PCR analysis of both junctions (Fig. 2b) indicating that Int-C3 had first intramolecularly recombined the transfected substrate vector and subsequently integrated the seamless vector into the genomic attH4X of LINE-1.
Transfection with in vitro generated seamless vector resulted in four out of nine viable clones that were positive for right junction PCR; two clones (F1 and F9) were tested positive for both junctions. As shown in Fig. 2b, semi-nested PCRs were performed in order to obtain sufficient products from all left junctions for sequencing, whereas right junction PCR amplicons were identified in primary PCRs (Fig. 2b). PCR products obtained using LINE-1-specific primers were subjected to sequence analysis to identify the genomic locus of F8 cassette integration. The corresponding targeted LINE-1 loci were subsequently verified by PCR/sequencing using chromosome-specific primers (Fig. 2b, right panel). Our combined results demonstrate that at least five clones (B6, B7, B8, F1, F9) harboured the complete F8 expression cassette and that three different LINE-1 loci were targeted by Int-mediated recombination (Supplementary Table 2).
Single copy F8 seamless vector insertion at endogenous attH4X sites
We employed Southern blot hybridization to confirm seamless vector insertions at the identified loci and, furthermore, to determine if only a single copy of the F8 expression cassette has been site-specifically integrated into the LINE-1. Two restriction endonucleases with recognition sites within the cassette and in the vicinity of the three predicted targeted LINE-1 loci were independently used for digestion of genomic DNA. Using a vector-internal probe hybridizing to NeoR, it was possible to identify single-copy insertions at the three loci based on restriction fragment patterns (Fig. 3a).
The Southern blots obtained with NsiI and KpnI-digested genomic DNA from four out of the five above-mentioned clones, and untargeted hESCs DNA as control, clearly revealed single-copy integration of the seamless cassette for each clone/locus (Fig. 3a) and confirmed the stable integration of the seamless vector in intron 2 of CDCA7L (Cell Division Cycle Associated b 7 Like; Chr7) in clones F1 and F9, intron 4 of CCDC141 (Coiled-Coil Domain Containing 141, Chr2) in clone B6 and intron 7 of DMD (Duchenne Muscular Dystrophy, ChrX) in clone B8 (Fig. 3a, b, Supplementary Table 2). With respect to clone B7, the Southern blot data suggested the existence of restriction site polymorphism near the targeted LINE-1 locus (data not shown) and hence was not analysed further. Altogether these findings demonstrate the ability of our transgenesis tool to target endogenous attH4X sites within LINE-1 elements with a 10.1-kb-sized therapeutic gene expression cassette. As exemplified by the independent targeting of the CDCA7L locus on chromosome 7 (for clones F1 and F9), the data also revealed the possible existence of hot-spot recombination loci for targeted transgene insertion mediated by mutant phage lambda Int-C3 .
F8 expression and catalytic FVIII activity in LINE-1 targeted clones
We next investigated if the targeted loci permitted sustained transgene expression. Quantitative RT-PCR analysis was performed to analyse the F8 mRNA expression levels of the four F8 transgenic clones (F1, F9, B6 and B8) normalized to the endogenous F8 levels in untargeted hESCs. We observed a significant increase in the amount of F8 mRNA in all transgenic clones (Fig. 4a). We included untargeted hESCs transiently transfected with the substrate F8 expression vector (1 μg) as a positive control, which, expectedly, showed the highest expression levels (Fig. 4a). These data demonstrated that the EF1α-F8-IRES-NeoR expression cassette is sustainably expressed in hESCs from these three targeted LINE-1 loci.
We also determined if the produced F8 mRNA was translated into protein and secreted from hESCs into the media in a biologically active form. We examined FVIII activity by a fluorometric assay in hESC culture supernatants, using again transiently transfected (100 ng) hESCs as positive and parental hESCs as negative controls. The fluorometric assay measures the ability of activated FVIIIa to generate Factor Xa in the presence of calcium and phospholipids, which further proteolytically cleaves a specific substrate to release a fluorophore that can be quantified. The FVIII activity was normalized to untargeted hESCs and to cell viability as measured by MTT assays to account for possible differences in cell density and growth rates of clones. Coinciding with the observed increase in F8 mRNA expression, we found a significant increase in FVIII activity with all targeted hESCs clones and transiently transfected cells (Fig. 4b). Interestingly, we also noted that untargeted hESCs did express a substantial level of biologically active FVIII protein when compared with unexposed cell culture media as negative control, which may open interesting possibilities for non-recombinant FVIII production at a larger scale using hESC fermenters. Taken together, these results clearly indicated that the LINE-1-targeted cell clones, regardless of the transgene locus, produced biologically active FVIII and that clone B8 exhibited both the highest F8 mRNA expression and protein activity.
Since many future applications of hESCs and induced pluripotent stem cells (iPSCs) will likely involve differentiation of stem cells into specific desired cell types, e.g. platelets, we next tested how F8 transgene expression might be affected by the differentiation status of our targeted hESC clones. Hence, we employed an established retinoic acid (RA)-induced differentiation protocol which typically results in a mixture of various cell lineages and differentiation states when hESCs are cultured in DMEM containing 1 μM RA for 48 h and subsequently maintained in DMEM w/o RA for 12 days . The results showed that the expression of the F8 transgene cassette in the four differentiated cell clones was substantially reduced when compared to undifferentiated hESCs, but remained significantly higher in the two clones that carry the transgene in the same genomic locus (clones F1 and F9) compared to the endogenous F8 transcript levels in parental differentiated cells (Fig. 4c). Control qRT-PCRs measuring expression of the key pluripotency factor genes Oct4, Nanog and Sox2 confirmed that the most cells in the transgenic hESC clones and parental hESCs had lost their pluripotent stem cell state (Fig. 4d–f). Furthermore, FVIII activity tests revealed that differentiated cells from clone F1 are still secreting biologically active clotting factor when compared to differentiated untargeted cells (Fig. 4g).
λ-Int-mediated reporter insertion for drug screening applications in FSHD disease
The human DUX4 gene is located within a D4Z4 sequence repeat array in the subtelomeric region of chromosome 4q35. It is known that contraction of these D4Z4 macro-satellite sequences is associated with decreased cytosine methylation and an open chromatin structure, leading to infrequent sporadic expression of the DUX4 gene in the skeletal muscle that results in facioscapulohumeral muscular dystrophy (FSHD) [54,55,56] (Fig. 5a). Given that DUX4 expression is difficult to detect in FSHD muscle cells, we employed our transgenesis system to generate a seamless vector comprising of a cassette harbouring a DUX4-responsive artificial promoter with 16 DUX4 binding sites upstream of a reporter gene (mNeon/fluorescent protein) and a downstream antibiotic selection cassette (PuroR driven by the PGK promoter: Fig. 5b). The mNeon expression as a readout was first validated with the episomal reporter by co-transfecting a DUX4 protein-expressing construct (pCMV-DUX4) into hESCs (Fig. 5b and data not shown). In order to generate the stable DUX4 reporter cell lines, our transgenesis platform was used to integrate the seamless reporter vector into LINE-1 of hESCs (Fig. 6a). PCR analysis confirmed both left and right junctions indicating specific and complete integration of the reporter cassette in three transgenic cell clones (M27, T13, T25) (Fig. 6b, Supplementary Table 2). The functionality of the inserted reporter in these clones was confirmed by ectopic expression of the DUX4 protein using pCMV-DUX4 expression vector.
As shown by fluorescence microcopy, activation of reporter expression in the three clonal cell lines can be achieved via transient expression of DUX4 (Fig. 6c). Importantly, the transfection efficiency in these reporter cell lines is sufficient to transiently express DUX4 and activate the reporter in a sufficient number of cells for downstream applications. For example, potential high-throughput small compound screening can be performed on DUX4-activated cells (within a 24–48 h time window) to identify molecules that antagonize DUX4-mediated activation of the mNeon reporter (Fig. 6d) and thereby identify potential lead compounds.
Genetic engineering attributes that offer flexibility for large transgene insertions equivalent to 10 kb or more can have profound implications for cell/gene therapy and synthetic biology applications. However, as the genomic transgene insert size increases, multiple genotoxic effects due to random integrations, epigenetic silencing and chromosomal aberrations, amongst others, represent potential complications. Therefore, both versatility and safety features of genome editing tools are critical, especially for gene therapy applications of monogenic diseases that necessitate large transgene insertions for curative outcomes. Hemophilia A (F8 coding sequence − 7 kb), DMD (Dystrophin coding sequence − 14 kb) and skin disease Recessive Dystrophic Epidermolysis Bullosa (COL7A1 coding sequence − 9 kb) are examples of diseases for which replacement corrections of dysfunctional large genes could yield clinical benefits. In order to validate the utility of our previously reported λ-Int-based seamless transgenesis tool [44, 45] in achieving large DNA transgenesis, we have demonstrated here its use in the insertion of the full-length F8 gene for hemophilia A as an example of a disease model.
Gene therapies for hemophilia A provides a tractable alternative to the present standard of care confined to prophylaxis, management of bleeding incidences and replacement therapy that includes repeated infusion of clotting factors to replace the missing/low endogenous FVIII protein [57,58,59,60,61,62]. Ideally, replacing the dysfunctional F8 gene with a functional copy would be the most desirable way to benefit more than 400,000 affected hemophilia A patients worldwide [63, 64], but such genome engineering pursuits are extremely challenging owing to the large size of the gene [64, 65]. Hence, truncated F8 variants as a substitute have been pursued to mimic FVIII-mediated physiological coagulation effects. AAV and other vectors have been widely used as a carrier for the truncated version of the F8 gene; however, certain safety issues persist [64, 66,67,68]. An example of remaining adverse virus-mediated oncogenic effects has been concretely pointed in a canine model of hemophilia administered with AAV gene therapy in a decade long follow-up study, wherein DNA payload insertion was evidenced near genes that regulate cell growth [69, 70]. Many precedented ex vivo pioneering studies [71,72,73,74,75,76] have also been attempted to either genetically correct or introduce a separate functional copy of truncated F8 into different types of cells by lentiviral, transposons and CRISPR Cas systems with a fair degree of success, yet still requiring significant improvements. In addition, lentivirus-based transduction of truncated F8 variants into patient-derived iPSCs and directed differentiation to megakaryocyte  and endothelial cell-lineage  for functional FVIII production have achieved some success, albeit some adverse effects of random integrations linger. CRISPR Cas tools were also used to correct F8 chromosomal inversions in patient-derived iPSCs and subsequent liver endothelial differentiation, an approach that could only benefit a subset of hemophilia patients who harbour such inversions . Contrastingly, a CRISPR-Cas-mediated universal gene-correction knock-in strategy of introducing BDD-F8 gene at the endogenous F8 locus of hemophilia A patient-derived iPSCs differentiated into endothelial cells also did not yield optimal levels of FVIII . This could be because the human F8 locus is located on the X-chromosome and only one copy has been inserted at this locus which did not allow sufficient expression and yield of the FVIII protein. In addition, deletion of the protein’s B-domain results in a reduced rate of FVIII secretion, which could be attributed to misfolding and degradation of the BDD-FVIII protein compared to the full-length FVIII protein. Furthermore, this approach is marred with common issues of CRISPR, including indels, chromosomal aberrations and translocations . A plausible direction of genome-editing strategies may involve introducing the F8 coding sequence into putative safe harbour and high expression loci, such as AAVS1 or CCR5, but such approaches need to be rigorously evaluated. To this end, non-viral tools like transcription activator-like effector nickases (TALENickases) identified the multicopy ribosomal DNA (rDNA) locus as a safe and effective target for F8 gene integrations and expression in hemophilia A-affected iPSCs. Unfortunately, they achieved a significant increase in the FVIII protein in the lysates of the targeted iPSCs but failed to achieve desirable FVIII protein in cell supernatants, indicating potential problems with folding and secretion of the FVIII protein .
To address the complex issues with hemophilia A gene therapy designs, we conceived a non-viral-based transgenesis of F8 at potentially safe harbour sites in human ESC genome. We took advantage of our previously reported λ-Int system to generate seamless vectors harbouring the full-length F8 gene using in vitro site-specific intramolecular recombination between two DNA recombination sequences (attH4X and attP4X) [44, 45] flanking the F8 expression cassette in a 14-kb supercoiled parental substrate plasmid. Our seamless vector approach should minimize potential adverse host immune responses to bacterial sequences [31,32,33,34,35,36,37]. The attL4x harbouring ~ 10.1 kb F8 seamless expression vector is then targeted to attH4x in the hESC genome. This approach also reduces the vector size, which, in turn, enhances DNA transfer. Our transgenesis strategy is potentially superior to Piggy Bac transposon-mediated full-length F8 insertion with respect to controlled and specific transgene insertion at predetermined LINE-1 sites . The Piggy Bac system offers no control over integration sites, which bears a potential risk for insertional mutagenesis and unwanted genotoxicities [77,78,79]. A paralleled approach in our study of introducing the substrate plasmid for Int-C3 to catalyze intracellular intramolecular recombination to convert the episomal substrate vector into a seamless vector before integration into LINE-1 elements is an important advance since it greatly simplifies the entire platform by eliminating seamless vector production at a larger scale in vitro. However, further experiments need to verify that the circular bacterial backbone DNA that is generated by intramolecular recombination inside cells is not randomly inserted in the host cells’ genome.
Our proof-of-concept study with transgenesis of F8 resulted in five hESC clones (B6, B7, B8, F1, F9) that harboured the complete F8 expression cassette in three different LINE-1 loci. Southern blot and sequencing analysis confirmed stable single copy integrations at so-called LINE-1 hot spots in four clones, a feature that will further simplify our platform technology and can be exploited in the future with other transgene constructs. Interestingly, the targeting site in clone F1 is identical to hotspots documented in our previous report . This locus lies on chromosome 7 and is part of an intron 2 of CDCA7L responsible for regulation of cell division and apoptosis signalling pathway. We confirmed the expression and activity of the F8 transgene from this targeted locus. We also showed that F8 transgene expression can be retained in differentiated hESCs, an important validation for our technology’s use in future stem cell and cell therapy approaches. The fact that we can target several endogenous attH4X sequences in parallel and test for functional transgene expression in differentiated cells represents an additional bonus of our transgenesis method to eventually generate the desired transgenic cell product.
In a second approach, we expanded the applicability of our platform for the further development of reporter cell lines for drug screening applications. We had previously generated a hESC-derived pluripotency reporter cell line that has already been successfully used in safety assessments of lead compounds for the treatment of tuberculosis [44, 80]. Here, we employed a seamless transgenesis approach for hESC-derived reporter cells related to FSHD disease. FSHD is a genetic muscle disorder caused by the loss of transcriptional repression of DUX4 gene, resulting in its aberrant expression and subsequent progressive muscle wasting predominantly in the face, shoulder blades and upper arms [81, 82]. The DUX4 protein is a transcription factor that targets a large set of genes and initiates a cascade of downstream signalling pathways that inhibit myogenesis and induces oxidative stress and cell death in FSHD skeletal muscle [83,84,85]. Various efforts are underway to model the disease in cultured cells for further studies of FSHD and to identify molecules that would interfere with pathogenic DUX4 expression or activity [84,85,86,87]. Given the high transfection efficiency that we can achieve with hES cells and their ability to differentiate into muscle lineage, herein, we reported the development of an alternative hESC-based reporter system comprised of large gene(s) cassette that can be adapted for high-throughput screening of drugs for FSHD disease. We constructed a DUX4 target gene reporter comprising of binding sites of DUX4 driving the mNeon gene that responds to DUX4 stimulation. We demonstrated that ectopic expression of DUX4 protein triggered the expression of the fluorescent reporter. We think it is feasible that these cell lines can be employed for high-throughput drug screening to identify small lead compounds that suppress DUX4’s activity as a transcriptional activator.
We presented a simple λ-Int transgenesis platform as a non-viral alternative to achieve large transgenic insertions into the human genome for cell/gene therapy and synthetic biology applications, including drug screening.
Availability of data and materials
All data generated during this study are included in this published article and its supplementary information file. Research findings are available from the corresponding author upon reasonable request.
Human Embryonic Stem Cells
Double Homeobox Protein 4
Facioscapulohumeral muscular dystrophy
CRISPR-associated transposase system
Non-homologous end joining
- LINE-1 :
Long INterspersed Elements
Chimeric Antigen Receptors
single chain Integration Host Factor
Internal Ribosome Entry Site
- CDCA7L :
Cell Division Cycle Associated- 7 Like
- CCDC141 :
Coiled-Coil Domain Containing 141
- DMD :
Duchenne Muscular Dystrophy
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We would like to thank Prof. Akitsu Hotta, Kyoto University, Japan, for providing F8 expressing piggyBac vector and our collaborator GENEA, Sydney, Australia, for providing human ESCs. The authors also acknowledge the funding agencies for their financial support.
This work was supported through grants from the Singapore-MIT Alliance for Research and Technology, Grant Award Numbers M4062347.080 and M4062198.080 to H.M., and the National Research Foundation (NRF) Singapore, NRF Competitive Research Programme (CRP), Grant Award Number NRF-CRP21-2018-0002 to P.D. Funding for open access charge: National Research Foundation Competitive Research Programme, Singapore (NRF-CRP21-2018-0002).
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H.M. and P.D. filed USA Patent Application No. 15/629,334 entitled “Site-Specific DNA Recombination” related to the technology and are co-founders and shareholders of LambdaGen Pte. Ltd. N.C., A.M.R. and S.R. declare no conflict of interest.
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Chaudhari, N., Rickard, A.M., Roy, S. et al. A non-viral genome editing platform for site-specific insertion of large transgenes. Stem Cell Res Ther 11, 380 (2020). https://doi.org/10.1186/s13287-020-01890-6
- Genome engineering
- Gene therapy
- Embryonic stem cells
- Large transgene
- FVIII clotting protein
- Lambda integrase
- Site-specific tyrosine recombinase