Targeted integration (TI), utilizing sequence-specific nucleases, has been shown capable of inserting transgene sequences at specific genomic sites. In theory, TI can be achieved either via homology directed repair (HDR; utilizing flanking homology sequences) or non-homologous end joining (NHEJ)-mediated end capture.

For efficient nuclease-mediated editing utilizing a donor template for TI, there are at least two primary requirements: 1) robust sequence-specific cleavage at the target site; and 2) sufficient delivery of donor DNA template to drive the TI event, rather than the default pathway resulting in NHEJ-induced indels. Thus far, demonstrations of targeted CFTR cDNA gene insertion in the endogenous CFTR locus of primary airway basal cells have utilized CRISPR/Cas9 or ZFNs to introduce a dsDNA break together with AAV-6-mediated delivery of donor sequences. The CFTR sequences targeted for insertion were flanked by homology sequences to facilitate HDR.

Vaidyanathan et al. recently reported CRISPR/Cas9-mediated TI of the CFTR cDNA into exon 1 of the CFTR locus in airway basal cells. Due to the packaging size constraints of AAV, the CFTR cDNA was split into two separate AAV vectors with the CFTR cDNA sequences recombining in the target cells. They also incorporated the truncated CD19 (tCD19) coding sequences into one of the AAV-6 vectors. Although the initial TI efficiencies were relatively low (∼5–10%) in the basal cells, likely due to the requirement for co-delivery and recombination, the surface expression of tCD19 allowed selection of TI cells to a 60–80% purity. After differentiation in air-liquid interface (ALI) cultures, restoration of CFTR function was demonstrated (Vaidyanathan et al., 2021).

Our group has recently demonstrated proof-of-principle for this TI approach in minimally expanded CF airway basal cells. TI was first performed in intron 8 of the endogenous CFTR gene with a partial CFTR cDNA (exons 9–27) preceded by a splice acceptor and terminated with a polyadenylation signal. By generating the sequence-specific dsDNA break in intron (as opposed to exon) sequences, we sought to minimize the potential detrimental effects caused by NHEJ-induced indels at the on-target site. Electroporation-mediated delivery of the ZFN mRNA and AAV-6-mediated delivery of the donor proved to be highly efficient with TI frequencies between 50.0 and 61.8% of CFTR alleles in F508del/F508del, F508del/R553X and G542X/R785X basal cells. Importantly, upon differentiation of the targeted basal cells in ALI conditions, intron 8 TI successfully restored mature CFTR protein and channel function at therapeutically-relevant levels (Suzuki et al., 2020). We note that these high rates of TI, also achieved in targeting of intron 7, together with the restoration of CFTR protein and function, were obtained without any selection. We quantitatively assessed, in several single-cell derived clones exhibiting homozygous intron 8 TI, what fraction of CFTR mRNA transcripts from targeted alleles were spliced from the endogenous exon 8 to the transgenic exon 9 (thus including transgenic corrective sequences) vs. alternative splicing downstream to the endogenous exon 9 (including only endogenous mutant sequences). Quantitative RT-PCR demonstrated that the majority (58.0—89.9%) of CFTR transcripts in these homozygous intron 8 TI clones exhibited the desired splicing into the corrective human codon-optimized exon 9 sequences; however, the remaining transcripts reflected splicing across the corrective exon sequences directly into endogenous exon 9 sequences.

Given this efficient gene insertion ex vivo in airway basal cells, we wished to demonstrate the competence of edited cells to establish a well-differentiated airway epithelium in vivo. Thus, a bulk population of TI-8 edited F508del/R553X basal cells were seeded into denuded rat tracheas and implanted in nu/nu mice (Filali et al., 2002; Hawkins et al., 2021) (Figure 1A). We observed the development of airway epithelium in seeded tracheas. Immunostaining confirmed multipotent differentiation with the clear presence of basal, secretory, and ciliated cells (Figure 1B).

We seek to develop culture methods enabling feeder-free expansion of edited airway basal cells to obtain a sufficient cell number for transplantation while retaining CFTR function. Such methods could be applied to bulk edited cells (i.e. mixture of corrected and uncorrected cells) or to corrected single-cell derived clones. We previously documented (Suzuki et al., 2020) that basal cells expanded in SAGM medium supplemented with SMAD inhibitors and ROCK inhibitor (SAGM/SMADinh + Y) or in Pneumacult-ExPlus (P-ExPlus) were capable of significant expansion as basal cells (Figures 2A,B). However, when cultured in either medium for greater than 8 passages and then differentiated in ALI culture, the level of CFTR-dependent current decreased significantly (Figure 2C). We note that achieving the fore-mentioned 60 million cells corresponds to approximately 26 population doublings (approximately 8 passages) starting with only one basal stem cell; in our experience, airway brushing of CF airways yields a far greater number of starting cells (data not shown). We have since found that supplementing P-ExPlus with SMAD inhibitors and ROCK inhibitor (P-ExPlus/SMADinh + Y), enables even greater expansion (Figure 2A), retention of long-term basal cell markers (e.g. ITGA6, NGFR) (Figure 2B), as well as multipotential differentiation, with a more stable and thicker epithelium consisting of basal, ciliated, secretory cells and ionocytes (Figure 2D). In addition, we now have superior maintenance of CFTR channel activity (Figure 2C). This last finding is highly significant in that we are unaware of any methodology, with or without feeders, able to extensively expand human airway basal cells without significant loss of CFTR activity. Future studies will include basal cells derived from multiple donors to confirm this finding.

Whereas HDR-mediated TI requires cells to be in cell cycle, NHEJ-mediated end capture of AAV transduced sequences has also been demonstrated in non-dividing cells (Suzuki et al., 2016). Typically, NHEJ-mediated TI integrates donor sequences equally in forward or reverse directions. To maximize the desired directionality of integrated sequences via CRISPR/Cas9, gRNA and protospacer adjacent motif (PAM) recognition sequences are directionally incorporated in the donor to favor the correct orientation (known as Homology Independent TI; HITI (Suzuki et al., 2016)). A rather similar directional approach utilizing obligate heterodimer ZFNs has been developed (known as ObLiGaRe (Maresca et al., 2013; Nami et al., 2018)).

Although sequence-specific nucleases are able, in many cases, to achieve highly efficient TI (either via HDR or NHEJ end capture), such conditions frequently result in indels (in some cases significantly sized deletions) at the target site of alleles not achieving TI. In addition, off-target indels may result in unwanted and unanticipated adverse consequences. Thus, there is significant interest in developing gene insertion methodologies that do not require a dsDNA break for sequence-specific targeting, such as transposon-encoded CRISPR/Cas systems (Klompe et al., 2019), twin prime editing (Anzalone et al., 2021), and fusion of CRISPR/Cas9 nickase to both a reverse transcriptase and serine integrase (Ioannidi et al., 2021). Alternatively, there are ongoing efforts to confer sequence-specificity upon retroviral or lentiviral vectors that typically do not exhibit sequence specific integration (Yoder et al., 2021).

There are several considerations relevant to determining which CFTR intron is optimal for TI. One consideration would be to maximize the number of CF patients (or CFTR mutations) that could, in principle, benefit from site-specific TI. For example, intron 8 TI would provide correction for 89.1% of the CF-causing CFTR alleles listed in the CFTR2 database including F508del, common PTCs (e.g. G542X, R785X, and W1282X), and splicing variants (e.g. 3,849 + 10 kb C > T) (Suzuki et al., 2020), whereas exon 1 (Vaidyanathan et al., 2021) or intron 1 targeting would provide correction for nearly all CFTR mutations. A second consideration is the size of the requisite CFTR partial cDNA and whether this can be appropriately delivered with a chosen vector; thus, the use of two AAV vectors for exon 1 TI (Vaidyanathan et al., 2021). A third consideration involves the relative strength(s) of the transgene splice acceptor vs. the native splice acceptor in the immediate downstream CFTR exon. A fourth aspect comprises the need to achieve transgene integration without disrupting activity of critical cis regulatory elements (CREs) and the chromatin architecture of the CFTR. We discuss the latter two issues in the following section.

By directly integrating the partial or full-length CFTR cDNA transgene into the CFTR locus, the objective would be to obtain physiologically- and therapeutically-relevant levels of corrected CFTR. In so doing, there are several considerations:

1) How to maximize incorporation of corrective transgene sequences in the mRNA and protein? Transgenic mouse studies showed that maintaining some intron/exon structure gave rise to higher expression of the transgene. As previously mentioned, it is important to consider the relative strength(s) of the transgene splice acceptor vs. the native splice acceptor in the immediate downstream CFTR exon. If the targeted intron were immediately upstream of a strong splice acceptor sequence, it is possible that a significant amount of splicing could jump across the corrective partial CFTR cDNA. A failure to capture only corrective cDNA sequences in CFTR mRNA transcripts from successfully targeted alleles, as observed in our published study (Suzuki et al., 2020), would potentially result in reduced levels of CFTR protein and activity. If splicing across the transgene were to be an issue, this could potentially be resolved by strengthening the splice acceptor sequence, varying the poly-adenylation sequence to maximize termination of transgene transcription, or perhaps changing the insertion site within the intron.

One of the primary challenges to efficient TI of airway basal cells in vivo will be the delivery of the editing reagents. In principle, luminal delivery to the airway epithelium is possible via viral (e.g. AAVs) or non-viral (e.g. lipid nanoparticles) (Robinson et al., 2018) methodologies provided that the CF mucus barrier can be penetrated or transiently removed. Given the tight junctions that characterize the pseudostratified airway epithelium, it remains challenging to deliver editing reagents to the basolaterally located basal cells. Treatment of human airway epithelium by transiently disrupting tight junctions or inhibiting proteasomal processing facilitates AAV transduction (Coyne et al., 2000; Duan et al., 2000; Yan et al., 2004; Ellis et al., 2013). Systemic delivery of editing reagents via the bloodstream is another possibility to access the airway epithelium, including the basal cells. There have recently been significant advances in the development of various non-viral vectors, including nanoparticles, for in vivo targeting of specific organs via systemic administration. For example, it has been shown that it is possible, via systemic delivery, to successfully target delivery of Cas9 to the lung (Cheng et al., 2020). Targeting delivery of editing reagents specifically to basal cells (e.g. via recognition of surface-expressed NGFR) could perhaps maximize the efficiency of editing in these cells.

Only a low frequency of airway basal cells is generally cycling at any given time in the non-CF human lung. However, the frequency of cycling airway basal cells in CF lungs is reportedly higher, which is likely due to ongoing inflammation and repair of damage (Voynow et al., 2005). In contrast to these findings, however, Carraro et al. recently determined via scRNA-seq that the frequency of basal cells in CF airways that are cycling was less than in control non-CF airways (Carraro et al., 2021). To facilitate efficient gene insertion into the significant fraction of the CF airway basal cells that are non-dividing at any given point in time (likely to be >75%) (Voynow et al., 2005), we must also consider alternative approaches to HR-mediated TI. Since the NHEJ repair mechanism, as well as other targeting approaches, are also active in non-cycling cells, such methods may be required, e.g. HITI (Suzuki et al., 2016), prime editing (Anzalone et al., 2019), twin prime editing (Anzalone et al., 2021), transposon-encoded CRISPR/Cas system (Klompe et al., 2019), and fusion of CRISPR/Cas9 nickase to both a reverse transcriptase and serine integrase (Ioannidi et al., 2021).

DD023J (non-CF airway epithelial cells) and KK002C (CF airway epithelial cells: F508del/R553X CFTR), obtained from explanted lungs, were provided by the Tissue Procurement and Cell Culture core facility at the University of North Carolina, NC, United States. The details of culture, characterization and in vitro differentiation of airway basal cells were previously described (Suzuki et al., 2020) In addition to the two culture media previously described (Suzuki et al., 2020) Pneumacult™-Ex Plus (STEMCELL technologies, Vancouver, Canada) medium and SAGM™ medium (Lonza, Basel, Switzerland) supplemented with dual SMAD inhibitors, we tested dual SMAD inhibition medium consisting of Pneumacult™-Ex Plus supplemented with 10 µM RhoA kinase (ROCK) inhibitor Y27362 (Reagents Direct, Encinitas), 1 µM A-8301 (R&D Systems, Minneapolis, MN), and 1 µM DMH-1 (R&D Systems). In all three culture conditions, airway basal cells were cultured on pre-coated plates with laminin-enriched 804G cell-conditioned medium and split at a 1:10 ratio upon their confluency at 50—70%. DD023J cells were used to test their capabilities of long-term expansion in each medium and characterized with known basal cell surface markers CD49f and NGFR using flow cytometric analysis. Cells at several passages from each medium were differentiated in vitro under identical air liquid interface (ALI) condition.

The F508del/R553X CFTR KK002C cells, subjected to intron 8 targeted integration (Suzuki et al., 2020), were expanded in Pneumacult™-Ex Plus medium and assayed in the tracheal xenograft model. The methodology was as reported (Filali et al., 2002) and as recently utilized by our group (Hawkins et al., 2021).

Epithelial monolayers were established under ALI conditions in transwells from non-CF (DD023J) airway basal cells originally cultured in the fore-mentioned media for various number of passages. Ussing chamber experiments were performed as previously described (Suzuki et al., 2020); the acquired data, analyzed for 3-4 transwells per condition, are expressed as mean ± SD.