Epigenome editing of the CFTR-locus for treatment of Cystic Fibrosis

Abstract

Background

Mechanisms governing the diversity of CFTR gene expression throughout the body are complex. Multiple intronic and distal regulatory elements are responsible for regulating differential CFTR expression across tissues.

Methods

Drawing on published data, 18 high-priority genomic regions were identified and interrogated for CFTR-enhancer function using CRISPR/dCas9-based epigenome editing tools. Each region was evaluated by dCas9^p300 and dCas9^KRAB for its ability to enhance or repress CFTR expression, respectively.

Results

Multiple genomic regions were tested for enhancer activity using CRISPR/dCas9 epigenome editing. dCas9^p300 mediates a significant increase in CFTR mRNA levels when targeted to the promoter and a region 44 kb upstream of the transcriptional start site in a CFTR-low expressing cell line. Multiple gRNAs targeting the promoter induced a robust increase in CFTR protein levels. In contrast, dCas9^KRAB-mediated repression is much more robust with 10 of the 18 evaluated genomic regions inducing CFTR protein knockdown. To evaluate the therapeutic efficacy of modulating CFTR gene regulation, dCas9^p300 was used to induce elevated levels of CFTR from the endogenous locus in ΔF508/ΔF508 human bronchial epithelial cells. Ussing chamber studies demonstrated a synergistic increase in ion transport in response to CRISPR-induced expression of ΔF508 CFTR mRNA along with VX809 treatment.

Conclusions

CRISPR/dCas9-based epigenome editing provides a previously unexplored tool for interrogating CFTR enhancer function. Here, we demonstrate that therapeutic interventions that increase the expression of CFTR may improve the efficacy of CFTR modulators. A better understanding CFTR regulatory mechanisms could uncover novel therapeutic interventions for the development of cystic fibrosis therapies.

Introduction

Cystic fibrosis (CF) is a life-limiting genetic disease characterized by absent or dysfunctional CF transmembrane regulator (CFTR) protein. A new era of CF precision medicine began in 2012 with the approval of the first CFTR modulator. Whereas previous treatments were limited to symptom management, CFTR modulators treat the underlying cause of CF by physically interacting with the CFTR protein to improve folding, trafficking, or stability. One of the major challenges in treating Cystic Fibrosis (CF) is the low bulk expression of CFTR in the lungs. This generally low expression may limit the potency of currently approved CFTR modulators and cause promising drug candidates to fail in clinical trials. Increasing the amount of total CFTR protein for those CFTR modulators to target could increase their therapeutic benefit. Indeed, several drug candidates have yielded promising results in CFTR-overexpressing cell lines, yet have failed to rescue the phenotype at physiological CFTR levels [1]. Thus, increasing the total amount of CFTR in the lungs of people with CF would be a significant advancement. Supporting this notion, a new class of drugs termed amplifiers are currently under development. Amplifiers aim to increase the total amount of CFTR protein. The few CFTR amplifiers that are currently under investigation were identified by broad high-throughput screens, and the mechanism has yet to be disclosed or is perhaps unknown [1].

The mechanisms governing CFTR gene regulation are largely unexplored. Millions of potential gene regulatory elements have been identified across the human genome [2, 3]. Differences in the activity of those regulatory elements cause genes, including CFTR, to be expressed in different patterns throughout the tissues of the body [4]. One class of regulatory elements is enhancers, a short region of DNA that can be bound and activated/ repressed by proteins (termed transcription factors) to control the likelihood of transcription of a particular gene. Many enhancers are specifically active in one or a small number of cell types, giving rise to cell type-specific gene expression patterns. Studies from Harris and colleagues have identified several proximal CFTR enhancers and their corresponding transcription factors [5-10], raising the possibility of therapeutic interventions that manipulate CFTR levels via gene regulation [1, 11, 12]. Building on this body of work, we demonstrate that directly manipulating gene regulation via CFTR regulatory elements may prove therapeutically beneficial for people with CF.

Due to its versatility and ease of use in screening paradigms, we use CRISPR/Cas9 epigenome-editing to characterize putative CFTR enhancer elements. Through complementary base pairing, a short guide RNA (gRNA) mediates Cas9 binding to a user-defined DNA sequence [13]. By designing the 20 bp protospacer sequence that confers DNA-binding specificity, the Cas9 enzyme can be targeted to almost any region in the genome. The natural function of Cas9 is to act as a nuclease, inducing double-stranded breaks at its genomic binding site. Mutating the endonuclease domains generates a deactivated Cas9 (dCas9) protein that has no endonuclease activity but maintains its RNA-guided DNA-binding activity [13]. dCas9, in conjunction with a gRNA, functions as a programable DNA-binding protein. As such, CRISPR/dCas9 epigenome-editing proteins can modulate gene expression without making permanent changes to the underlying DNA sequence. Covalently modified histone proteins are a prominent indicator of the activity of a regulatory element [14]. Acetylation of histone lysine residues, for example, is a classic mark of an active enhancer [14, 15]. Fusing a histone acetyltransferase domain (p300) to dCas9 is sufficient to increase the expression of target genes more than 30 kb away in the genome [16]. Similarly, fusing the Krüppel associated box (KRAB) domain to dCas9 is sufficient to recruit heterochromatin-forming proteins and repress targeted gene expression [17].

By localizing either dCas9^p300 or dCas9^KRAB to putative CFTR regulatory elements, we validated multiple genomics regions responsible for inducing and repressing CFTR expression. Finally, in patient-derived human bronchial epithelial (HBE) cells homozygous for the most common CFTR mutation (ΔF508), we demonstrate that upregulation of mutant CFTR by dCas9^p300 enhanced the effects of VX809 (Lumacaftor) treatment in Ussing chamber studies. These results demonstrate that modulating the transcriptional regulatory machinery of CFTR is a viable mechanism by which to increase CFTR levels for the treatment of CF. Increasing CFTR levels via epigenetic modulation could be achieved by a variety of therapeutic modalities including, but not limited to, small molecules that target specific transcription factors, or synthetic transcription factors that directly target the endogenous CFTR locus.

Materials and Methods

Genome accessibility data processing

Normalized DNase-seq signal tracks in counts per million (CPM) were downloaded from the ENCODE project portal for the following tissues: lung, pancreas, small intestine, and transverse colon; as well as the following cell lines: A549, Caco-2, HEK293T, HT29, and SAEC (Supplementary Table1)[2]. For tissues and cell lines in which more than one sample was assayed, mean signal was computed using wiggletools [18].

Aligned DNase-seq reads in BAM format were obtained for human epididymis epithelial (HEE) and human tracheal epithelial (HTE) cells (Supplementary Table 1). Read mappings were normalized to CPM and converted to a signal track using the deepTools utility bamCoverage with the following parameters: “--ignoreDuplicates --binsize 50 --smoothLength 100 --normalizeUsing CPM” [19]. Two samples were assayed for each of the primary cells, hence mean signal was computed using wiggletools [18].

Gene expression data processing

The most recent tissue-specific gene expression data was downloaded from GTEx Portal (GTEx Analysis v8/dbGaP Accession phs000424.v8.p2) on Mar 25, 2020 in units of transcripts per million (TPM) (Supplemental Table 2) [20]. Certain tissue subclassifications were collapsed into a more parsimonious set of tissues (e.g. {Amygdala,…,Substantia nigra} = Brain)

Cell line and primary cell gene expression was taken from a variety of sources and expression was converted to TPM (Supplementary Table 3).

gRNA selection

Putative CFTR enhancer regions were selected by identifying regions of differential chromatin accessibility across CFTR-high vs CFTR-low cells and tissues (Figure 1). Regions were prioritized based on information available in the literature characterizing these sites [4-10, 21]. From this analysis, 17 regions in addition to the promoter region were selected and evaluated for regulatory activity (Supplemental Table 4).

Figure 1: Characterization of the CFTR Locus Across Tissues and Cell Lines of Interest.

(A) CFTR expression is shown in transcripts per million (TPM) across different tissues as taken from GTEx. CFTR expression is highly variable across tissues. (B) CFTR expression is shown in TPM across different primary cell cultures and cell lines. (C) UCSC genome browser snapshot of the CFTR locus highlighting potential regulatory regions of interest. Genome accessibility is shown as assayed by DNase-seq across various tissues, primary cells, and immortalized cell lines. Bio-sample data are presented in order of low to high CFTR expression within each group. CRISPR/dCas9 targeted regions have been highlighted. CPM: Counts per million, HEE: Human Epididymis Epithelium, HTE: Human Tracheal Epithelium, SAEC: Small Airway Epithelial Cells

gRNAs were designed to span each genomic region of interest with at least 1 gRNA/ 100bp of sequence. gRNAs were optimized for lowest predicted off-target binding and highest on-target activity [22], while maintaining the desired distribution across the genomic regions of interest. This analysis resulted in design of 133 total gRNAs (Supplemental Table 5).

Plasmid Construction

Individual gRNA protospacers were cloned into pLV-hU6-gRNA (Addgene plasmid #83925). Oligonucleotides for each protospacer were synthesized (IDT-DNA), hybridized, phosphorylated, and ligated into the dual BsmBI sites using conventional cloning methods. dCas9^p300 and dCas9^KRAB were expressed from pLV-EFS-dCas9^p300-P2A-Puro (Addgene plasmid #83889) and pLV-hUbC-dCas9^KRAB -T2A-Puro (Addgene plasmid #71236) respectively. To generate an all-in-one lentivirus that co expressed a gRNA along with dCas9^p300, the hU6-gRNA cassette from pLV-hU6-gRNA was cloned between the KpnI and PacI sites of pLV-EFS-dCas9^p300-T2A-Puro.

Cell culture

HEK293T cells were obtained from the American Tissue Collection Center (ATCC, Manassas, VA, USA) and were maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% FBS, 100 Units/mL penicillin, and 100 μg/mL streptomycin. HT29 cells were obtained from Sigma-Aldrich and were maintained in McCoy’s 5A medium supplemented with 10% FBS and 1% penicillin/streptomycin. Bmi1/hTERT human bronchial epithelial cell lines, UNCN3T (wildtype CFTR) and UNCCF3T (ΔF508/ΔF508 CFTR) [23], were cultured using the CRC method as previously described [24]. Briefly, cells were co-cultured with irradiated NIH3T3 fibroblasts in the presence of the rho-kinase inhibitor, Y-27632 (Axxora).

Viral production and transduction of immortalized cell lines

All lentiviral vectors used in this study are second generation and were produced using standard viral production methods as previously described [25]. Briefly, 5.7 million HEK293T cells were plated per 10 cm dish. The following day, cells were transfected with Iipofectamine2000 (ThermoFisher) with 10 μg of transfer vector, 3 μg of pMD2G and 8 μg psPAX2. The media was changed 12–14 hours post-transfection. The viral supernatant was collected 24 and 48 hours after this media change for a total of 20 mL of virus, and passed through a 0.45 μm filter. As needed, lentivirus was concentrated with Lenti-XTM concentrator (CloneTech). Virus was snap frozen in liquid nitrogen for storage.

To transduce HEK293T and HT29s, cells were resuspended and plated into virus supplemented with 4 μg/ml polybrene. The viral supernatant was exchanged for fresh medium 12–24 hours later. To generate stable dCas9^p300 and dCas9^KRAB stable lines, cells were selected in 1ug/ml puromycin.

Flow Cytometry

Cells were fixed and permeabilized using the eBioscience Foxp3/ Transcription Factor Staining Kit using the nuclear staining protocol (Thermofisher). Cells were evaluated for Cas9 expression using the a Cas9-PE conjugated antibody at 1:50 dilution (Clone 7A9-3A3, Cell Signaling Technologies). Cells were evaluated for fluorescence using the Attune NxT flow cytometer (ThermoFisher).

Quantitative Reverse Transcription PCR

qRT-PCR samples from HEK293T and HT29s were prepped using the Cells-to-Ct 1-Step Taqman Kit (Ambion), per the manufacturer’s instructions. mRNA from UNCCF3T and UNCN3T cultures were isolated using the Qiagen RNeasy Plus kit. Equal mass of mRNA was reverse transcribed using Superscript VILO (ThermoFisher). Real-time PCR using multiplexed Taqman assays (CFTR: Hs00357011_m, TBP: Hs99999910_m1, ThermoFisher) was performed on the Quantstudio7 Detection System (ThermoFisher). The results are expressed as fold change expression of CFTR normalized to TBP using the ΔΔC_t method.

The results are expressed as fold change in CFTR mRNA expression normalized to TBP expression using the ΔΔCt method. Briefly, ΔC_T was computed as C_T,R - C_T,X where the reference gene (R) was TBP and the test gene (X) was CFTR. Log₂ fold change, or ΔΔC_T, was computed by subtracting from each well’s ΔC_T the mean ΔC_T for all control wells in which only Cas9 and no gRNA was added. Samples in which template was undetected were assigned C_T = 40. Samples with C_T values below the 95% confidence interval for mean C_T—as computed per gene and per experiment (CRISPR-activation or CRISPR-inhibition)—were discarded. One-way ANOVA was used to confirm significant effects of gRNA. Dunnett's post hoc test was used to test for significance of effects for each gRNA, comparing the distribution of log fold changes for each gRNA to that for Cas9 only.

Western Blot Analysis

Cells were lysed in RIPA buffer (Sigma-Aldrich) supplemented with protease inhibitor cocktail (Sigma-Aldrich). Protein concentration was measured using BCA protein assay (ThermoFisher) and Varioskab LUX Microplate Reader (ThermoFisher). Lysates were mixed with loading buffer; equal amounts of protein were run on Mini-PROTEAN TGX 4-15% precast polyacrylamide gels (Bio-Rad) and transferred to nitrocellulose membranes using the Trans-Blot Turbo System (Bio-Rad). Nonspecific antibody binding was blocked with Intercept TBS blocking buffer (Li-Cor) or 5% milk in TBST for 1 h at room temperature. The membranes were incubated with the following primary antibodies: anti-CFTR Clone769 (1:1000 dilution, University of North Carolina at Chapel Hill, Cystic Fibrosis Foundation) in Intercept T20 TBS (Li-Cor) or 5% milk in TBST overnight at 4 °C; anti-Actin (1:5000 dilution, Sigma-Aldrich, A2066) in Intercept T20 TBS (Li-Cor) or 5% milk in TBST overnight at 4 °C. The membranes were washed with TBST for 15 min and incubated for 30-45 min with donkey anti-mouse 680 RD (Li-Cor, 1:5000) in Intercept T20 TBS, donkey anti-rabbit 800 CW (Li-Cor, 1:5000) in Intercept T20 TBS, goat anti-mouse HRP (Abcam Ab97046, 1:5000) in 5% milk in TBST, or goat anti-rabbit HRP (Sigma A6155, 1:5000) in 5% milk. Blots were subsequently washed with TBST for 15 min. Membranes were visualized using the Odyssey CLx (Li-Cor) or the iBright FL1500 (Thermofisher).

Human Bronchial Epithelial Ussing Chamber Studies

UNCCF3T [23] cells were thawed from cryopreservation and cultured using the CRC method [24]. UNCCF3T cells were plated in irradiated NIH3T3-conditioned media (CM) supplemented with 5 μM Y-27632 for 24 hours. Conditioned media was prepared as previously described [26]. The cells were then treated with a single lentivirus co-expressing dCas9^p300 and a CFTR targeting gRNA (gRNA 40). Cells were transduced with a final concentration of 20x lentivirus for 3 hours in the presence of polybrene diluted 1:1000 (Sigma TR-1003-G). Cells were then washed with PBS and fed with fresh CM + Y. Selection with 0.5 μg/mL puromycin began 6 days after transduction when the cells were ~60% confluent. Cells were grown for one additional passage in CRC and seeded at a total seeding density of 1.5 x 10⁵ cells in 12-mm Millicell inserts (Millipore PICM01250) coated with human placental collagen (Sigma C7510) and fed UNC air-liquid interface (ALI) media supplemented with 0.5 μg/mL puromycin. Ussing chamber studies were performed on day 28 with the addition of amiloride, forskolin, VX770, CFTRinh-172, and UTP as previously described [23, 24], and a subset were treated with 5 μM VX809 for 48 hours before Ussing analysis. One-way ANOVA followed by the Tukey post hoc test was used to determine statistical significance.

Results

CFTR expression is highly variable throughout the body (Figure 1A) as well as across primary cells and immortalized cell lines (Figure 1B), and is far lower in the lung compared to the pancreas, salivary gland, colon, and small intestine. Further, CFTR expression is variable across cell subpopulations found within the lung. In fact, only 3.6% of cells in the lung contribute to total CFTR expression [27]. Ionocytes, a rare population that comprises < 1% of all lung epithelial cells [28], express CFTR at the highest levels, thus accounting for ~25% of total CFTR in the large airways [27]. By contrast, CFTR expression is markedly lower in secretory and basal cells. However, the abundance of these cells types make them the dominant CFTR expressers, contributing to ~35% and ~30% of total CFTR expression in the large airways, respectively [27]. Comparing chromatin accessibility, measured by DNase-seq, around the CFTR gene between CFTR-low and CFTR-high expressing cell types and tissues reveals a substantial diversity of chromatin structure (Figure 1C). Taken together, these data suggest CFTR expression is tightly regulated and may be regulated by tissue-specific enhancers. While regulatory elements 1 megabase from promoters have been reported [29], comprehensive studies of the genetics of gene regulation suggest that most enhancers act over shorter distances [20, 30, 31]. Therefore, we interrogating genomic regions within +/− 50kb of the CFTR gene. Candidate enhancers were identified by selecting genomic regions with differential chromatin structures across CFTR-low and CFTR-high expressing tissues and cells. By integrating DNase-seq datasets (Figure 1C), reporter assay data [7, 8], and previously interrogated genomic regions [4-6, 10, 21], we selected 18 high-priority CFTR genomic regions of interest to interrogate further (Figure 1C, Supplemental Table 4).

Enhancer activity was evaluated by localizing dCas9^KRAB (a heterochromatin-forming, repressive transcription factor) [17] or dCas9^p300 (a euchromatin-forming, activating transcription factor) [16] to each putative enhancer region of interest and evaluating the resulting change in CFTR expression. gRNAs were designed to span each putative enhancer region with at least 1 gRNA per 100 base pairs of sequence. gRNAs were optimized for lowest predicted off-target binding and highest on-target activity [22], while maintaining the desired distribution across the genomic regions of interest. This analysis resulted in the design of 133 total gRNAs (Supplemental Table 5). To evaluate the dynamics of CFTR enhancer activity, we evaluated enhancer-mediated modulation of CFTR expression in a CFTR-low (HEK293T) and a CFTR-high (HT29) cell line. Stable HEK293T-dCas9^p300 and HT29-dCas9^KRAB cell lines were generated with lentivirus and validated for Cas9 expression by flow cytometry (Supplemental Figure 1). The stable cell lines were subsequently transduced with lentivirus expressing the gRNA of interest. Seven days post transduction, cells were harvested and evaluated for CFTR expression by qRT-PCR (Figure 2) and Western Blot (Figure 3) analysis. It should be noted that variable levels of activation and repression are observed across different gRNAs targeting the same putative enhancer region, consistent with prior studies [32]. CRISPR-activation (CRISPRa) and CRISPR-inhibition (CRISPRi) gRNAs vary greatly in efficacy based on a variety of factors including sequence [33]. Thus, to compare activities across enhancer regions, we calculated an average fold change for each putative enhancer. Significant CFTR activation was observed at the mRNA level when targeting the −44 kb and promoter regions in the CFTR-low HEK293T cell line (Figure 2B). Targeting Intron 11a,b in addition to the promoter caused a significant repression of CFTR mRNA in the CFTR-high HT29 cell line (Figure 2C).

Figure 2: CRISPR/dCas9 epigenome editing of CFTR enhancer elements modulates endogenous CFTR mRNA levels.

(A) Locations and widths of targeted putative enhancer regions are displayed in linear genomic space and with respect to the CFTR gene structure. (B) HEK293T cells stably expressing dCas9^p300 and (C) HT29 cells stably expressing dCas9^KRAB were treated with lentivirus expressing a single gRNA targeted to the putative enhancer regions of interest. Seven days post transduction, cells were harvested and evaluated for CFTR expression using qRT-PCR. Mean log fold change and standard error of the mean from two independent experiments performed on different days is shown per gRNA (n = 3 or 4, * p < 0.1). With the exception of the Cas9-only control and the x-axis discontinuities between targeted regions, spacing between gRNAs is proportional to linear genomic distance and comparable across targeted regions. The mean log fold change across gRNAs for each genomic region is shown as a black bar. Guide order corresponds, from left to right, to order of guides provided in Supplemental Table 6. For log fold change and statistical significance of individual gRNAs, refer to Supplemental Table 6.

Figure 3: CRISPR/dCas9 epigenome editing of CFTR enhancer elements modulates endogenous CFTR protein levels.

HEK293T cells stably expressing dCas9^p300 were treated with lentivirus expressing a single gRNA targeted to either the −44kb genomic loci (A) or the promoter (B). HT29 cells stably expressing dCas9^KRAB were treated with lentivirus expressing a single gRNA targeted to the promoter (C) or the Intron11a,b genomic loci (D). Seven days post transduction, cells were harvested and evaluated for protein expression via western blot. Positive control: 10ug of lysate from HEK293T transduced CFTR cDNA (n = 1).

We further evaluated whether changes in CFTR mRNA expression led to changes in CFTR protein level (Figure 3). Efforts were focused on putative enhancers with at least 1 gRNA that induced a statistically significant change in CFTR mRNA. The four most potent gRNAs as determined by qRT-PCR from each region, with the exception of repressive promoter-targeting gRNAs, were selected for evaluation at the protein level. For repressive promoter-targeting gRNAs, we selected gRNAs for further evaluation that induced variable levels of mRNA repression since there were > 4 statistically significant gRNAs identified. Not surprisingly, epigenetic manipulation of the CFTR promoter caused gene activation and repression at the protein level (Figure 3B,C). We detected up to a 1.6-fold increase in CFTR protein for three of the four gRNAs targeting the −44 kb region in the CFTR-low HEK293T cells (Figure 3A). Enhancer repression had an even more robust effect than activation. Repressing Intron 11a,b with dCas9^KRAB caused a 10-fold reduction in CFTR protein across the evaluated gRNAs in CFTR-high HT29 cells (Figure 3D). Repression of eight other enhancers also reduced CFTR protein levels, consistent with observed reduction in mRNA levels (Supplemental Figure 2).

Of particular therapeutic interest is the ability to boost CFTR expression in the CFTR-high cells of the lung epithelia including ionocytes, secretory, and basal cells. To evaluate feasibility of inducing super-physiological levels of CFTR in a CFTR-high cell line, we generated a HT29-dCas9^p300 cell line. HT29-dCas9^p300 cells were transduced with a subset of promoter targeting gRNAs and a modest increase in CFTR mRNA and protein was detected (Supplemental Figure 3) suggesting it is in fact feasible to boost CFTR expression in cells with high-basal CFTR expression.

Though correlation between mRNA level and protein expression are known to vary greatly by gene [34], it was recently shown that mRNA abundance, protein expression, and CFTR function are strongly correlated in organoids [35]. Here, we saw a correlation between CFTR mRNA and protein expression for CRISPR-inhibitory perturbations (Pearson correlation coefficient = 0.51), and a weak correlation for CRISPR-activation perturbations, likely due to the narrow dynamic range of response (Pearson correlation coefficient = 0.18).

The data generated in model cell lines suggests that CFTR protein level can be modulated by targeting CRISPR/dCas9 epigenome modifiers to CFTR regulatory elements. Therefore, we hypothesized that increasing CFTR expression through modulation of the endogenous CFTR gene regulation could enhance the effects of CFTR modulator compounds. To this end, we evaluated our most potent activating CRISPR/dCas9 epigenome complex for its ability to enhance chloride transport in patient-derived HBEs. Growth enhanced ΔF508/ΔF508 HBEs (UNCCF3Ts) were transduced with an all-in-one lentiviral vector delivering dCas9^p300, a puromycin resistance gene, and either a polyT terminator gRNA (Mock) or our most potent activating gRNA 40 (CRISPRa) (Supplemental Table 6). Cells were selected with puromycin to enrich for transduced cells. Before seeding air-liquid interface (ALI) cultures, a cell pellet was harvested and evaluated for CFTR expression using qRT-PCR (Figure 4a). CFTR mRNA was significantly increased in the CRISPRa- treated HBEs, thus confirming successful transduction and selection. Puromycin selected cells were differentiated towards a mucociliary phenotype for 28 days using established methods [36].

Figure 4: Epigenome editing for therapeutic benefit in HBEs.

Δ508/Δ508 CF donor HBEs (UNCCF3Ts) were transduced with either a Mock (dCas9^p300 –gRNA) or with a CRISPRa (dCas9^p300 +gRNA40) vector. (A) Wild type (UNCN3Ts) and Δ508/Δ508 CF donor HBEs (UNCCF3Ts) cells grown in monolayer were evaluated for CFTR expression by qRT-PCR (n = 3, * p < 0.05). Samples were differentiated for 28 days in the ALI culture system and evaluated for ion transport in Ussing chambers. Cultures were pre-treated with Lumacaftor (+VX809) or DMSO-only (−VX809) for 48 hours. (B) Representative traces of n = three replicates. (C) Changes in short circuit current (ΔI_SC) measured in response to amiloride, (D) forskolin, (E) VX770, and (F) CFTRinh-172 (n = 3 or 4, * p < 0.05).

Multiple CFTR modulators are already on the market for the treatment of ΔF508-CF patients. Thus, we hypothesized that dCas9^p300 induced CFTR gene activation would increase the benefit of the CFTR potentiator, VX770 (Ivacaftor), alone or in combination with the CFTR corrector, VX809 (Lumacaftor). Mock and CRISPRa transduced cultures were pre-treated with VX809 or DMSO (−VX809) for 48 hours before Ussing Chamber studies. During Ussing analysis, cultures were sequentially treated with amiloride (Amil), forskolin (FSK), VX770, CFTR inhibitor-172 (CFTRinh-172), and Uridine-5’-Triphosphate (UTP) and the resulting changes in short circuit current (ΔIsc) were measured (Figure 4b). CRISPRa in combination with VX809 treatment resulted in a statistically significant increase in chloride transport over VX809 treatment alone (Figure 4D,F). The combined response of CRISPRa +VX809 was synergistic in nature as CRISPRa alone has little effect on chloride transport (Supplemental Table 7). VX770 further increased chloride transport in all conditions, with the greatest VX770 response in the CRISPRa +VX809 group, though the difference between Mock and CRISPRa were not significant (Figure 4E). Overall, our data shows that increased CFTR levels leads to increased CFTR-mediated ion transport in HBE cells grown in the ALI culture system. These experiments demonstrate that interventions that increase CFTR expression may ultimately increase the therapeutic efficacy of CFTR modulators.

Conclusions

The CFTR promoter is thought to act as a minimal promoter [37] that relies on distal regulatory elements to effect a complex pattern of expression across different tissues [9]. Multiple enhancer regions have been characterized to varying degrees using methods such as DNase-seq [7-9, 21], DNase footprinting [6, 8, 10, 21], chromatin conformation capture [7-9], ChIP [5-10], and reporter assays [6-8, 10]. Taken together, this body of research suggests a complex interplay between multiple genomic enhancers is responsible for CFTR gene regulation. While some transcription factors have been implicated [5, 10, 28, 38], the minimal set of factors sufficient and necessary for modulating CFTR gene expression is unknown. To our knowledge, none of these factors or their respective pathways have enabled identification of new therapeutic interventions for CF.

Here we present a new strategy for identifying and validating CFTR-specific regulatory elements. We demonstrate that enhancer-targeting CRISPR/dCas9 epigenome modulators can be utilized to induce or repress CFTR expression. Furthermore, we show that this type of intervention may result in therapeutic benefit in a disease setting. It is important to note that this study provides proof of concept that dCas9-based epigenome modulators can target and modulate CFTR gene expression. However, utilizing an array-based screening method did not enable screening of all potential gRNA targets in the putative enhancers of interest and therefore may have resulted in false negatives. In contrast, CRISPR/dCas9 high-throughput screens can be used to interrogate thousands of putative enhancers and screen all potential targeting gRNAs in a single experiment [32]. Such high-throughput functional genomic screens are a new and underexplored avenue for gaining a mechanistic understanding of CFTR gene regulation. A more thorough understanding of CFTR enhancers and their respective protein regulators may lead to identification of novel therapeutic interventions for the treatment of CF.

As a foundational study, we provide proof-of-concept that boosting CFTR expression is a promising approach to treating the most common ΔF508/ΔF508 genotype. However, we posit that increasing CFTR expression and increasing CFTR modulator efficacy would be beneficial for treating most patients with CF. For example, historically, some of the most challenging CFTR mutations to treat have been nonsense mutations (class I) that generate a premature termination codon (PTC) in the CFTR transcript. Though ribosomal readthrough of the PTC to generate full-length protein has been proposed as a treatment strategy [39, 40], nonsense mediated decay (NMD) dramatically reduces the amount of CFTR mRNA available, rendering readthrough agents clinically ineffective[41, 42]. Indeed, in a study of W1282X, the 4^th most common CFTR mutation worldwide [43], nasal cells homozygous for the mutation had at only 30–80% CFTR mRNA abundance compared with that of wildtype CFTR in non-CF subjects [44]. Studies that bypass NMD by overexpressing intron-less complementary DNA (cDNA) copies of W1282X-CFTR have demonstrated that the truncated protein can be corrected by CFTR modulators [45, 46]. However, in primary patient-derived cells, CFTR modulators are not sufficient to rescue W1282X-CFTR activity [40]. Increasing CFTR expression in people with a class I CFTR mutation might counteract the effects of NMD and increase sensitivity to read-through agents and/or other CFTR modulators. Overall, interventions that increase CFTR expression may ultimately lead to powerful adjunct therapies that may benefit most, if not all CF mutant classes. Furthermore, CRISPR/Cas9-based epigenome modulators are a valuable tool that can be used to further elucidate the endogenous CFTR signaling mechanisms and pathways.

Highlights.

• Genomic regions surrounding the CFTR locus were interrogated for enhancer function using dCas9-based epigenome editing.

• Multiple enhancers were identified that modulate CFTR expression.

• Epigenome editing of the CFTR locus in polarized CF epithelial cells increased forskolin-stimulated short circuit current.