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Cystic fibrosis is characterized by deficiency of the cystic fibrosis transmembrane conductance regulator (CFTR), a Cl− transporter. The packaging constraints of adeno-associated viral (AAV) vectors preclude delivery of both an active promoter and CFTR cDNA to target cells. We hypothesized that segmental trans-splicing, in which two AAV vectors deliver the 5′ and 3′ halves of the CFTR cDNA, could mediate splicing of two pre-mRNAs into a full-length, functional CFTR mRNA. Using a segmental trans-splicing 5′ donor–3′ acceptor pair that split the CFTR cDNA between exons 14a and 14b, cotransfection of donor and acceptor plasmids into CFTR− cells resulted in full-length CFTR message and protein. Microinjection of plasmids into CFTR− cells produced cAMP-activated Cl− conductance. Vectors created with an engineered human serotype, AAV6.2, were used to deliver CFTR donor and acceptor constructs, resulting in full-length CFTR mRNA and protein as well as cAMP-activated Cl− conductance in CFTR− cells, including human CF airway epithelial IB3-1 cells. Thus, segmental trans-splicing can be used with AAV vectors to mediate expression of CFTR, a strategy potentially applicable to individuals with CF.
Cystic fibrosis, a common autosomal recessive disorder in the white population, is caused by mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) gene (Riordan, 1989; Rommens et al., 1989; Welsh and Smith, 1993; Riordan et al., 2008). In the lung, these CFTR mutations result in the absence of or reduction in airway epithelial apical chloride channel activity, leading to salt and water imbalances across the epithelium, with resulting accumulation of mucus and colonization by pathogens (Riordan, 2008). Gene transfer studies have attempted to restore CFTR function to human airway epithelium via transfer of the normal CFTR cDNA (Zabner et al., 1993; Crystal et al., 1994; Sorscher et al., 1994; Wilson et al., 1994; Caplen et al., 1995; Knowles et al., 1995; Flotte et al., 1996, 2007; Harvey et al., 1999; Wagner et al., 1999; Moss et al., 2004, 2007; Griesenbach et al., 2006). Although the data from in vitro models are compelling, an efficient method for persistent, efficient expression of CFTR in the airway epithelium remains elusive owing to multiple barriers to gene transfer to the airway in vivo, including innate and acquired immunity to vectors, physical barriers, inefficient interaction of vectors with airway cells, limits on the genetic payload of some vectors, and limits on the persistence of others (Griesenbach et al., 2006; Flotte et al., 2007).
Adeno-associated viral (AAV) gene transfer vectors are capable of mediating persistent gene expression in the airway epithelium in immunocompetent animals, and thus have been the focus of interest for gene therapy for the respiratory manifestations of CF (Flotte et al., 1993b; Halbert et al., 1997, 2001; Zabner et al., 2000; Sirninger et al., 2004; Limberis and Wilson, 2006; Fischer et al., 2007). However, AAV vectors are small, unable to accommodate an expression cassette containing the full 4.5-kb CFTR cDNA with an active promoter and relevant 3′ noncoding sequences. Although a number of strategies for overcoming this packaging constraint have been proposed, the challenge of using AAV vectors to deliver a large cDNA remains (Flotte et al., 1993a,b; Zhang et al., 1998; Duan et al., 2000a, 2001; Yan et al., 2000; Liu et al., 2002, 2005; Sirninger et al., 2004; Ostedgaard et al., 2005). Toward this end, the present study uses a strategy known as pre-mRNA “segmental trans-splicing” (STS), which relies on two pre-mRNAs sharing an intronic hybridization domain that can lead the spliceosome apparatus to mediate a splicing event that will combine the 5′ upstream (“donor”) sequence of one mRNA with the 3′ downstream (“acceptor”) sequence of the second mRNA (Pergolizzi et al., 2003; Pergolizzi and Crystal, 2004; Liu et al., 2005; Nakayama et al., 2005). To create an intact, full-length CFTR cDNA, we used two nonfunctional, vector-encoded partial human CFTR cDNAs, one of which encodes the 5′ untranslated and upstream coding sequence through exon 14a and one of which encodes the downstream coding sequence starting with exon 14b and ending with 3′ untranslated region signals for RNA processing. After demonstration that plasmids encoding the 5′ and 3′ CFTR cDNA fragments could together direct a functional CFTR mRNA, fragments were used to create separate AAV 5′ donor and 3′ acceptor vectors. Using AAV serotype 6.2, an AAV6 derivative with a single amino acid change in the capsid (F129L) that results in highly efficient transduction of respiratory epithelium (Rutledge et al., 1998; Limberis et al., 2009), these vectors were used to successfully alter cells lacking CFTR (CFTR− cells) to become functional CFTR-expressing cells (CFTR+ cells), including epithelial cells derived from an individual with CF, supporting the concept that segmental trans-splicing may be a strategy that can be used to treat the pulmonary manifestations of CF.
Fischer rat thyroid (FRT) cells stably expressing a Cl−-sensitive yellow fluorescent protein (YFP-H148Q) and FRT cells stably transfected with CFTR cDNA were kindly provided by A.S. Verkman (University of California, San Francisco, San Francisco, CA) (Galietta et al., 2001a,b). The cells were cultured in Coon's modified F-12 medium (Sigma-Aldrich, St. Louis, MO) supplemented with 5% (v/v) fetal bovine serum (FBS; Hyclone, Logan, UT), penicillin (100U/ml), and streptomycin (100μg/ml; Invitrogen, Carlsbad, CA). HEK-293 cells and 293orf6 cells (Graham et al., 1977; Brough et al., 1996) were cultured in Dulbecco's modified Eagle's medium (DMEM; Invitrogen) supplemented with 10% (v/v) FBS, sodium pyruvate (1%), penicillin (50U/ml), and streptomycin (50μg/ml). IB3-1 cells were provided by P. Zeitlin (Johns Hopkins University, Baltimore, MD) and A. Prince (Columbia University College of Physicians and Surgeons, New York, NY) (Zeitlin et al., 1991). IB3-1 cells were maintained in LHC-8 medium (Invitrogen) supplemented with 10% (v/v) FBS, penicillin (50U/ml), and streptomycin (50μg/ml). All cells were kept in a humidified incubator at 37°C and 5% CO2.
The CFTR donor and acceptor plasmids were based on the segmental trans-splicing plasmid derived from pcDNA3.1(+) (Invitrogen) (Pergolizzi et al., 2003) (Fig. 1). Briefly, polymerase chain reaction (PCR)-amplified CFTR coding sequences were cloned into the pcDNA plasmid. The engineered segmental trans-splicing junction of CFTR was located between exon 14a and exon 14b. The donor CFTR segmental trans-splicing construct (pCFTR Donor) included the cytomegalovirus immediate-early promoter/enhancer, CFTR exons 1–14a, the 5′ end of intron 6 cloned from the human vascular endothelial growth factor gene, a unique 160-bp hybridization domain that is not found in the human genome, and a polyadenylation signal (Pergolizzi et al., 2003). The acceptor CFTR segmental trans-splicing construct (pCFTR Acceptor) included the cytomegalovirus immediate-early promoter/enhancer, the complementary sequence of the unique 160-bp hybridization domain in the donor construct, an optimized branch point, a polypyrimidine tract, a splice acceptor, CFTR exons 14b–24, and a polyadenylation signal (Pergolizzi et al., 2003). A positive control plasmid was synthesized to contain the full-length CFTR cDNA (pCFTR). All the plasmids were prepared with kits (Qiagen, Valencia, CA) according to the manufacturer's instructions.
The constructs (8μg/plate) were transfected into HEK-293 cells (2.5×106cells) plated in 60-mm plates, using Lipofectamine 2000 (Invitrogen) according to the manufacturer's protocol. Forty-eight hours after transfection, the cells were lysed. Total RNA was isolated with RNeasy mini kits (Qiagen) and reverse transcribed (SuperScript III reverse transcriptase; Invitrogen) using an oligo(dT) primer (Invitrogen). To demonstrate that the exon 14a–exon 14b junction was made by the segmental trans-splicing process, PCR was performed with SuperMix High Fidelity (Invitrogen) and the following primers: forward probe (TCA CCG AAA GAC AAC AGC ATC C); reverse probe (TGG TAA GAG GCA GAA GGT CAT CC). PCR was carried out in a 35-cycle reaction, using a 5-min initial 95°C denaturing step. Each repeating cycle consisted of three steps: 30sec at 94°C, 30sec at 60°C, and 45sec at 72°C. PCR products were analyzed on a 1.5% agarose Tris–acetate gel stained with ethidium bromide.
HEK-293 cells (6.5×106cells) plated in 100-mm plates were transfected with pcDNA3.1, pCFTR Donor, pCFTR Acceptor, pCFTR Donor plus pCFTR Acceptor, or pCFTR (24μg/dish). Forty-eight hours later, the transfected cells were lysed with radioimmunoprecipitation (RIPA) buffer (Sigma-Aldrich) according to the manufacturer's instructions. The protein lysate was immunoprecipitated with 1.6μg of anti-CFTR monoclonal antibody (clone MAB1660; R&D Systems, Minneapolis, MN), followed by immunoblotting with anti-CFTR antibody (kindly provided by J. Riordan, University of North Carolina School of Medicine, Chapel Hill, NC) (Mall et al., 2004). The blots were incubated with horseradish peroxidase (HRP)-conjugated anti-mouse IgG (Santa Cruz Biotechnology, Santa Cruz, CA). Immmunoreactive bands were visualized with an enhanced chemiluminescence (ECL) detection kit (Pierce Biotechnology, Rockford, IL).
FRT-YFP cells were harvested and seeded onto a 35-mm plate containing a sterile, 15-mm diameter, round glass coverslip. The plasmids were suspended at 300μg/ml in injection buffer (140mM KCl, 10mM HEPES; pH 7.4). The injection solution contained Alexa Fluor 647 (0.04mM; Invitrogen) to permit identification of the microinjected cells. Microinjection (~2pl, corresponding to 0.6pg of DNA) was performed on an Olympus IX71 inverted microscope (Olympus America, Center Valley, PA) equipped with an injector (Pico Injector model PLI100; Harvard Apparatus, Holliston, MA), micromanipulator (Narashige, East Meadow, NY), and glass pipettes pulled with a Flaming/Brown micropipette puller (model P-87; Sutter Instrument, Novato, CA). Twenty-four hours after injection, CFTR activity of injected cells was measured by evaluation of YFP fluorescence intensity (Jayaraman et al., 2000; Galietta et al., 2001a,b). Cells plated on coverslips were mounted in a low-volume, heated chamber (model RC-20 with a TC-324B thermostatic controller to regulate stage temperature; Warner Instruments, Hamden, CT), using a multireservoir flow system (model VC-6; Warner Instruments), with an in-line glass heating coil to prewarm the buffers before entering the flow chamber (model 158830; Radnoti Glass Technology, Monrovia, CA) connected to a thermostatically controlled water-circulating pump (model 1104; VWR, West Chester, PA). All buffers also included a final concentration of 0.5% dimethyl sulfoxide (DMSO) to eliminate the potential for changes in membrane permeability as conditions were changed. The cells were initially perfused with Cl−–phosphate-buffered saline (PBS) (137mM NaCl, 2.7mM KCl, 0.7mM CaCl2, 1.1mM MgCl2, 1.5mM KH2PO4, 8.1mM Na2HPO4; pH 7.4) at 1.8ml/min at 37°C. In the first cycle of buffer exchanges, no cAMP activators were included. After equilibration in Cl−–PBS, cells were switched to I−–PBS in which NaCl was replaced with NaI (137mM). After equilibration in I−–PBS, cells were returned to Cl−–PBS and reequilibrated. In a second cycle of buffer exchanges, cells were equilibrated in I−–PBS and then switched to I−–PBS in the presence of cAMP activators (5μM forskolin, 50μM isobutylmethylxanthine). Cell fluorescence was measured every 5sec on an inverted Olympus IX70 microscope equipped with a cooled charge-coupled device (CCD) camera (Quantix camera operating at 1MHz; Photometrix, Tucson, AZ) and the fluorescence intensity of individual cells was assessed with MetaMorph image analysis software (Molecular Devices/Universal Imaging, Downingtown, PA). To assess the relative amount of CFTR activity, the rate of iodide influx (decreasing YFPH148Q fluorescence intensity) was assessed in the presence of cAMP activation and the maximal point-to-point slope was calculated. Background measured in the absence of cells was subtracted.
Cells were defined as being CFTR+ if they met three criteria, including (1) slope of YFP fluorescence that exceeded the mean slope of negative control cells (FRT-YFP) by >2 standard deviations (i.e., a change in slope >0.03×10−3/sec), (2) a total decrease in fluorescence intensity of >25% compared with the initial fluorescence intensity before I−, and (3) a recovery of fluorescence intensity >15% compared with the initial fluorescence intensity.
The AAV6.2 serotype was chosen for expression of the CFTR segmental trans-splicing donor and acceptor constructs on the basis of data showing that this capsid conferred high-efficiency gene transfer to airway epithelial cells in rodents, humans, and nonhuman primates (Limberis et al., 2009). The CFTR segmental trans-splicing donor or acceptor constructs were cloned into an AAV packaging plasmid containing a cytomegalovirus (CMV) promoter and a simian virus 40 (SV40) polyadenylation site. AAV6.2 vectors packaging CFTR donor or acceptor constructs were produced by a three-plasmid transfection strategy, with the following three plasmids: (1) pAAVCFTR Donor or Acceptor genome plasmid; (2) pAAV6.2 plasmid, which provides Rep proteins and Cap proteins; and (3) pAdΔF6, an adenoviral helper plasmid that provides adenoviral helper functions of E2, E4, and VA RNAs. To initiate a round of vector production, pAAVCFTR genome plasmid (donor or acceptor, 600μg each), pAAV6.2 (800μg), and pAdΔF6 (1.2mg) were cotransfected into HEK-293 cells (which contain an integrated copy of the Ad E1 region) with PolyFect (Qiagen) according to the manufacturer's instructions. At 72hr posttransfection, cells were harvested, and AAV particles were purified as previously described (De et al., 2008). Briefly, a crude viral lysate was prepared by three cycles of freeze–thaw, and clarified by centrifugation. AAV6.2CFTR Donor and AAV6.2CFTR Acceptor were purified by iodixanol gradient and HiTrap Q HP Sepharose anion-exchange chromatography (Amersham/GE Healthcare Life Sciences, Piscataway, NJ) as previously described (De et al., 2008). The purified AAV6.2 vectors were concentrated with a Biomax 100 membrane concentrator (Millipore, Billerica, MA) and stored in PBS (pH 7.4) at −80°C until use. Vector genome titers were determined by TaqMan real-time PCR, using a CMV enhancer-specific primer–probe set (Applied Biosystems, Foster City, CA) (De et al., 2008).
293orf6 cells seeded in 60- or 100-mm dishes were infected with AAV6.2CFTR Donor, AAV6.2CFTR Acceptor, or a combination of AAV6.2CFTR Donor and AAV6.2CFTR Acceptor for 90min at a dose of 2×104 genome copies/cell for each vector. Virus infection was performed with serum-free DMEM. After infection, cells were cultured in DMEM containing 100μM ZnCl2 and 10% FBS. After 72hr, infected cells were lysed and total RNA and protein were isolated to evaluate the expression of CFTR mRNA and protein, using the assays described previously. As a positive control, HEK-293 cells were transfected with a plasmid expressing full-length CFTR cDNA (pCFTR), described previously, according to the manufacturer's instructions. RNA levels were assessed by TaqMan real-time PCR with relative quantitation, using rRNA as reference and reagents from Applied Biosystems. The primer and probes were also from Applied Biosystems, and consisted of a CFTR Donor-specific set (Hs00357004_m1), spanning the junction of exons 9 and 10; a CFTR Acceptor specific set (Hs00357011_m1), spanning the junction of exons 21 and 22; and a segmental trans-splicing specific set (Hs01565534_m1), spanning the junction of exons 14a and 14b. Linearity of response was determined by ensuring that the relative amounts were unaffected by dilution of RNA by a factor of 10.
FRT-YFP cells (1.5×105) seeded on 15-mm round, glass coverslips were infected with AAV6.2CFTR Donor, AAV6.2CFTR Acceptor, or a combination of AAV6.2CFTR Donor and AAV6.2CFTR Acceptor in a volume of 30μl of OptiMEM medium (Invitrogen) at a concentration of 5×104 genome copies/cell. Virus infection was performed for 90min followed by addition of 2ml of fresh Coon's modified F-12 medium containing 5% FBS, penicillin (100U/ml), and streptomycin (100μg/ml). After 72hr, the CFTR functional assay was performed as described previously. FRT-YFP cells stably transfected with CFTR served as the positive control (Galietta et al., 2001a).
To create an adenoviral gene transfer vector to modify cells to express YFP, the pH-sensitive YFP-H148Q gene was amplified from FRT-YFP cells. It was sequenced and subcloned into pShuttleCMV. This plasmid was recombined with AdLacZ pK7 adenovirus (Wickham et al., 1996), and the resulting plasmid was transformed in HEK-293 cells, expanded, and purified as described previously; the addition of the pK7 motif (which includes a stretch of seven consecutive lysine residues in the knob of the fiber protein) is intended to enhance entry into cells not expressing high levels of the adenovirus receptor (Wickham et al., 1996, 1997; Hidaka et al., 1999). To confirm the presence of both YFP and the pK7 fiber construct, the infectivity of the virus was compared with that of Ad5YFP (prepared as described previously, using a conventional E1−, E3− adenovirus serotype 5 backbone) for infection of human dermal fibroblasts, known to be deficient in the high-affinity coxsackievirus–adenovirus receptor (Hidaka et al., 1999). AdpK7YFP-H148Q infectivity was >2 logs higher than that of Ad5YFP-H148Q (the same vector without the seven lysine residues) in these cells (data not shown).
IB3-1 cells grown on 15-mm round, glass coverslips were infected with 30μl of rAAV-containing DMEM. IB3 cells were treated with 40μM N-acetyl-l-leucyl-l-leucyl-l-norleucine (LLnL; Boston Biochemical, Boston, MA), a proteasome inhibitor that enhances AAV-mediated gene expression (Duan et al., 2000b). In separate experiments, the ability of LLnL to enhance transgene expression by a factor of >10-fold after AAV infection of IB3-1 cells with AAV6.2LacZ vector was confirmed (data not shown). IB3 cells were transferred to fresh medium containing LLnL (40μM) and AAV6.2CFTR Donor, AAV6.2CFTR Acceptor, or a combination of AAV6.2CFTR Donor and AAV6.2CFTR Acceptor under the infection conditions described previously for FRT cells. After 90min, cells received fresh medium containing 40μM LLnL for a period of 24hr. Cells were then transferred to medium without LLnL for 24hr followed by infection with AdpK7.YFPH148Q (3000particles/cell) for 60min. The cells were then trypsinized, replated on 18-mm round coverslips, and incubated overnight. Coverslips were used to evaluate CFTR function assay as described previously. As a positive control, naive IB3 cells were infected with both AdCFTR (5000particles/cell) (Harvey et al., 1999) and AdpK7YFP-H148Q (3000particles/cell) for 60min and incubated for 24hr before assessment of CFTR activity.
Data are presented as means±standard error (SE), where mean values for a condition were calculated on the basis of mean values from each coverslip tested for that condition. Tests for significance of difference between groups employed the Student t test, where p<0.05 was taken as an indication of a significant difference between groups.
To test the feasibility of CFTR segmental trans-splicing in vitro, 293 cells were transfected with the plasmids containing either 5′ sequences (CFTR exons 1–14a followed by an intron with a splice donor, “CFTR Donor”), 3′ sequences (an intron with a splice acceptor sequence followed by CFTR exons 14b–24, “CFTR Acceptor”), or both constructs. CFTR gene expression was evaluated at the mRNA and protein levels. Intact CFTR message produced by segmental trans-splicing was assessed by collecting total RNA from transfected cells and conducting RT-PCR with primers that bridged the splice junction. The presence of a single 623-bp product was diagnostic for the presence of the upstream forward primer-binding site in the donor sequence (CFTR nucleotides 2349–2370) and the downstream reverse primer-binding site in the acceptor sequence (CFTR nucleotides 2949–2971) on a contiguous stretch of nucleic acids that could only result from a splicing event (Fig. 2A). The presence of the 623-bp band required reverse transcriptase activity as well as the presence of both the donor and acceptor plasmids, indicating that the splicing event was a result of a rearrangement at the RNA level rather than at the DNA level. Although the amplified product size matched a PCR product generated from intact CFTR message (pCFTR plasmid), further confirmation of correct splicing was provided by sequencing the PCR product, which showed that the mRNA contained the correct CFTR mRNA sequence through the splice junction (Fig. 2B). To determine whether cells transfected with pCFTR Donor plus pCFTR Acceptor produced CFTR protein, CFTR protein expression was evaluated by Western analysis after immunoprecipitation. In agreement with the RT-PCR data, the 170-kDa CFTR protein was detected in 293 cells transfected with a plasmid containing full-length CFTR cDNA (pCFTR) or with a pair of plasmids containing the segmental trans-splicing donor and acceptor pair (pCFTR Donor plus pCFTR Acceptor; Fig. 2C). The 170-kDa band was absent from naive cells (data not shown) and from cells transfected with either the donor or acceptor construct alone. Two additional bands at 55 and 25kDa resulted from immunoreactivity of the immunoglobulin used during immunoprecipitation; these bands were less prominent in the pCFTR sample, where less total protein was loaded. The banding pattern and size of immunoreactive protein products were similar to those of the product produced from 293 cells transfected with full-length CFTR cDNA. As expected, CFTR protein were not found in 293 cells transfected with the control plasmids pcDNA3, pCFTR Donor plus pcDNA3, or pCFTR Acceptor plus pcDNA3.
A fluorescence-based assay was used to assess functional expression of CFTR (Galietta et al., 2001a,b). To evaluate CFTR function after segmental trans-splicing, microinjection (~0.6pg/cell) was used to deliver CFTR segmental trans-splicing plasmids into the nuclei of Fischer rat thyroid cells (FRT; known to be CFTR negative) that stably express a Cl−-sensitive yellow fluorescent protein (YFP-H148Q). Microinjected cells were identified by codelivery of a soluble red fluorophore (Alexa Fluor 647). After microinjection, CFTR function was assessed in naive and microinjected cells by measuring YFP fluorescence as extracellular Cl− was replaced with I−. Channel activity was stimulated by addition of the adenylate cyclase activator, forskolin, and the phosphodiesterase inhibitor, isobutylmethylxanthine (IBMX). Although all cells contained yellow fluorescent protein due to stable expression of YFP-H148Q (displayed in the green channel), only microinjected cells contained red fluorescence (displayed in the red channel). Viable injected cells were identified by the presence of both YFP and Alexa Fluor 647 (colocalization of digital green and red signals appears yellow/orange; Fig. 3A, white arrowheads). Some microinjected cells became nonviable; these cells appear red in digital images because of a lack of detectable YFP-H148Q (Fig. 3A, black arrowheads).
One day after injection, CFTR function was evaluated through the change in YFP-H148Q fluorescence during exchange of chloride and iodide followed by addition of cAMP activators (Galietta et al., 2001a,b). In the absence of CFTR, the change in YFP fluorescence was negligible in FRT-YFP cells after the addition of cAMP activators (Fig. 3B, black triangles). These data confirmed earlier reports that FRT cells have low basal halide permeability (Galietta et al., 2001a). A significant decrease in YFP fluorescence was observed in FRT-YFP cells microinjected with a plasmid containing CFTR cDNA (Fig. 3B, green triangles) after addition of cAMP activators (IBMX/forskolin), whereas YFP fluorescence increased rapidly when the cell medium was returned to a buffer containing Cl− (Fig. 3B); some of the microinjected cells were negative for CFTR activity. In FRT-YFP cells injected with pCFTR Donor or pCFTR Acceptor individually, YFP fluorescence failed to show that same dramatic decrease after addition of iodide with cAMP activators (data not shown). As expected, microinjection of both pCFTR Donor and pCFTR Acceptor plasmids produced detectable CFTR-mediated chloride conductance (Fig. 3B, yellow triangles) that was significantly increased compared with microinjection of pcDNA3, a negative control plasmid that lacks an expression cassette (Fig. 3B, gray triangles).
Cells were defined as being “positive” for CFTR (CFTR+) if they met three criteria including a significant increase in the slope of fluorescence intensity compared with control cells, a large decrease in fluorescence intensity on exchange of I− for Cl−, and a large increase in fluorescence intensity on return to Cl− (see Materials and Methods). Cells that met all three criteria were considered CFTR+ whereas cells that failed to meet all three criteria were considered CFTR−. A significant fraction of microinjected cells acquired a phenotypic response to the chloride/iodide exchange (~60% of pCFTR-injected cells and ~40% of pCFTR Donor/pCFTR Acceptor-coinjected cells). To assess CFTR activity quantitatively, the maximal slope per 5-sec interval during iodide influx in the presence of forskolin/IBMX was calculated; this parameter has previously been established as the best index of CFTR activity (Galietta et al., 2001b). Multiple cells from at least three independent experiments were examined for each condition (see the caption to Fig. 3 concerning panel C). Whereas microinjection of either the pCFTR Donor or pCFTR Acceptor plasmid alone failed to change the slope of the change in YFP fluorescence, microinjection of either pCFTR or the combination of pCFTR Donor and pCFTR Acceptor led to significant increases in the slope of the change in YFP fluorescence (p<0.001 for both conditions compared with uninjected cells; Fig. 3C). The mean slopes for the pCFTR and pCFTR Donor plus pCFTR Acceptor groups were the same (p>0.1).
Among the AAV serotypes that have been used for gene transfer to the airway epithelium, evidence suggests that a novel capsid designated AAV6.2 is highly efficient (Limberis et al., 2009). After cloning the CFTR Donor and CFTR Acceptor expression cassettes into an AAV packaging plasmid, each plasmid was cotransfected with a plasmid containing AAV6.2 cap genes, resulting in production of AAV6.2CFTR Donor and AAV6.2CFTR Acceptor.
To test the ability of AAV6.2CFTR Donor and AAV6.2CFTR Acceptor to produce CFTR through segmental trans-splicing, the two vectors were used to infect 293orf6 cells alone or in combination. Infection of 293 cells by this serotype was confirmed with AAV6.2LacZ under the same conditions, with >80% of the cells staining positive with 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside (X-Gal). As a positive control, 293orf6 cells were transfected with pCFTR as described previously. The presence of an intact 14a/b junction at the mRNA level was confirmed by RT-PCR (Fig. 4A). In the absence of reverse transcriptase, no PCR fragments were generated. In the presence of reverse transcriptase, a 623-bp fragment was generated in the positive control and in the culture that was infected with both AAV6.2CFTR Donor and AAV6.2CFTR Acceptor. Cells infected with either AAV6.2CFTR Donor or AAV6.2CFTR Acceptor alone did not exhibit an intact 14a/b junction. Sequencing of the PCR fragment from cells infected with AAV6.2CFTR Donor and AAV6.2CFTR Acceptor confirmed that the junction formed correctly (Fig. 4B). Western analysis for CFTR demonstrated the presence of a 170-kDa band in cells infected with AAV6.2CFTR Donor and AAV6.2CFTR Acceptor together, but not in cells infected with AAV6.2CFTR Donor or AAV6.2CFTR Acceptor alone (Fig. 4C). These data confirmed that RNA transcripts derived from AAV vectors were able to participate in segmental trans-splicing leading to production of CFTR protein.
To estimate the efficiency of segmental trans-splicing, TaqMan real-time PCR with relative quantitation was used. RNA from 293orf6 cells infected with AAV6.2CFTR Donor, AAV6.2CFTR Acceptor, or both was analyzed, using rRNA as a relative reference and human tracheal epithelium RNA as a calibrator. Three primers and probe combinations were compared: the first within the CFTR Donor; the second within the CFTR Acceptor, and the third spanning the STS junction (Fig. 4D). As expected, the TaqMan primer and probe combination specific to the CFTR Donor detected mRNA only in the samples in which AAV6.2CFTR Donor was used for infection. Similarly, the TaqMan primer and probe combination specific to the CFTR Acceptor detected mRNA only in the samples in which AAV6.2CFTR Acceptor was used for infection. In the doubly infected cells, the apparent levels for the AAV6.2CFTR Donor-specific mRNA and AAV6.2CFTR Acceptor-specific mRNA were comparable, showing both vectors had comparable transduction efficiencies. The TaqMan primer and probe combination spanning the segmental trans-splicing junction showed amplification only in the RNA from cells that received both vectors. The amount was 4.7% of the estimated amount of RNA from the AAV6.2CFTR Donor and 12.1% of the amount from the AAV6.2CFTR Acceptor. Moreover, the overall amount of CFTR mRNA was greater than was seen in human tracheal epithelium, suggesting that physiological levels of CFTR may be obtained after in vivo gene transfer.
To determine whether CFTR protein produced by AAV-mediated segmental trans-splicing was functional, we evaluated CFTR activity in FRT-YFP cells infected with AAV6.2CFTR Donor and AAV6.2CFTR Acceptor individually or in combination. Infection of FRT-YFP cells with this serotype was confirmed with AAV6.2LacZ under the same conditions, with 18±1% of the cells staining positive with X-Gal. CFTR function was assessed by analysis of the fluorescence intensity of the YFP-H148Q mutant as described previously. Fluorescence intensity profiles of individual cells were evaluated, and the slope of the decrease in fluorescence intensity was calculated after the replacement of extracellular chloride with iodide in the presence of cAMP activators (Fig. 5A). Whereas naive FRT-YFP cells showed little variation in fluorescence intensity after addition of iodide and cAMP activators, approximately 50% of the FRT-YFP cells expressing CFTR exhibited a decrease in fluorescence on addition of the cAMP activators. In cultures that received either AAV6.2CFTR Donor or AAV6.2CFTR Acceptor, cells never exhibited a decrease in YFP fluorescence intensity, but cells that received both AAV6.2CFTR Donor and AAV6.2CFTR Acceptor had clear decreases in YFP fluorescence intensity on addition of cAMP activators to iodide-containing medium. Cells were designated as CFTR+ when they met the three criteria described in Materials and Methods and the slope of fluorescent change was assessed from multiple cells per group from at least three independent experiments (see Fig. 5B caption). Consistent with the fraction of cells infected with AAV6.2LacZ under these conditions, 19% of the cells infected with AAV6.2CFTR Donor plus AAV6.2CFTR Acceptor vectors were functionally CFTR positive. The CFTR-negative cells were assumed to be uninfected or to have been infected with only AAV62.CFTR Donor or AAV6.2CFTR Acceptor, but not both. The rate of change of fluorescence intensity in cells infected with AAV vectors carrying the CFTR donor/acceptor pair was significantly less than in cells with endogenous expression of CFTR (p<0.01; Fig. 5B), but based on the knowledge that 5–10% of CFTR activity is necessary to correct CFTR mutations (Trapnell et al., 1991; Johnson et al., 1992; Farmen et al., 2005), the function mediated by the donor/acceptor vector is likely sufficient.
Expression of functional CFTR in FRT cells after infection with AAV6.2CFTR Donor and AAV6.2CFTR Acceptor demonstrated the principle that segmental trans-splicing was successful in a cell that was CFTR negative. We sought to determine whether the process would work in cells containing mRNAs encoding mutant forms of CFTR. To answer this question, IB3-1 cells, SV40-transformed airway epithelial cells derived from an individual with CF (Zeitlin et al., 1991), were used as the model cell line for the study. Unlike FRT-YFP cells, IB3-1 cells do not constitutively express the chloride-sensitive YFP-H148Q. For the purpose of detecting CFTR expression, an adenoviral vector expressing the YFP-H148Q gene was created. Although the vector was based on adenovirus serotype 5, the construct employed a modified form of the adenoviral fiber protein with a stretch of seven lysine residues in the globular head domain (AdpK7). The pK7 fiber was designed to confer efficient gene transfer with broad tropism conveyed by the polylysine motif (Wickham et al., 1996). The resulting adenoviral vector (AdpK7-YFPH148Q) efficiently expressed chloride-sensitive YFP in a manner independent of the expression of the coxsackievirus–adenovirus receptor, indicating that the pK7 fiber was functioning appropriately (data not shown). To enhance the efficiency of AAV-mediated gene expression in IB3-1 cells, the cells were cultured with LLnL, a proteasome inhibitor, previously shown to enhance expression of transgenes from AAV serotype 2 vectors in vitro in human polarized epithelia (Duan et al., 2000b). We tested the ability of LLnL to enhance AAV6.2-mediated gene expression in IB3-1 cells, using both LacZ and GFP as marker genes. In all cases, cells treated with LLnL showed superior expression compared with expression without LLnL treatment (data not shown).
To evaluate gene expression, IB3-1 cells were infected with AAV6.2CFTR Donor and AAV6.2CFTR Acceptor individually or together, treated with LLnL-containing medium, infected with AdpK7-YFPH148Q, and assessed for YFP fluorescence intensity after exchange of iodide for chloride in extracellular medium (Fig. 6A). Infection of IB3 cells with this serotype was confirmed, using AAV6.2LacZ under identical conditions, with 39±5% of the cells staining positive with X-Gal. The results in IB3-1 cells mirrored those in FRT-YFP cells. Naive IB3-1 cells showed no chloride conductance. A large proportion of IB3-1 cells infected with AdCFTR demonstrated clear chloride conductance. Cells infected with either AAV6.2CFTR Donor or AAV6.2CFTR Acceptor had no chloride conductance, whereas a small percentage of cells infected with both AAV6.2CFTR Donor and AAV6.2CFTR Acceptor were positive for chloride conductance. The percentage of doubly infected cells that were functionally CFTR positive was lower (11%) than expected on the basis of the efficiency of infection with AAV6.2LacZ. When evaluating only cells that were considered positive for CFTR conductance by the criteria described in Materials and Methods, positive cells were found only among cells that received AAV6.2CFTR Donor and AAV6.2CFTR Acceptor, whereas cells that received only one vector were never positive. Among positive cells, the slope of the conductance in CFTR+ cells was significantly greater than the slope observed in control cells (Fig. 6B). The rate of change of fluorescence intensity in cells infected with AAV vectors carrying the CFTR donor/acceptor pair was less than for cells infected with AdCFTR (p<0.01), but the level was likely sufficient to correct the Cl− conductance defect.
Cystic fibrosis, a common lethal hereditary disorder, is caused by mutations in the cystic fibrosis transmembrane conductance regulator gene (Riordan et al., 1989; Rommens et al., 1989; Welsh and Smith, 1993; Riordan, 2008). Although multiple organs are affected by the CFTR mutation, the lung disease is responsible for most of the morbidity and mortality in individuals with CF. Hence, the major focus of gene therapy for CF is to correct CFTR function in the airway epithelium. Among the candidate gene therapy vectors for CF, AAV appears to be a good choice because of its ability to confer persistent, high-level gene expression in immunocompetent animal models of genetic diseases (Warrington and Herzog, 2006; Wu et al., 2006). However, for application to CF, the AAV vector is faced with the challenge that the packing capacity of AAV cannot accommodate the full-length CFTR cDNA in combination with an active promoter and relevant 3′ signals (Flotte et al., 1993a,b; Zhang et al., 1998; Sirninger et al., 2004; Ostedgaard et al., 2005). In this study, segmental trans-splicing was used to solve this challenge. This strategy relies on delivery of two halves of the CFTR cDNA in different vectors (Pergolizzi et al., 2003; Pergolizzi and Crystal, 2004; Liu et al., 2005; Nakayama et al., 2005). When the two vectors infect the same cells, the two pre-mRNAs come together by virtue of complementary sequences inserted into the 3′ and 5′ noncoding regions of the upstream and downstream vectors, respectively. By including relevant splicing signals near the complementary hybridization sequences, the nuclear splicing machinery joins the two half-transcripts into an intact mRNA. In this study, the CFTR cDNA was divided between exon 14a and exon 14b. RT-PCR and Western analysis results after plasmid transfection in 293 cells demonstrated expression of correctly trans-spliced CFTR mRNA and full-length protein. A CFTR functional assay after microinjection confirmed the presence of functional CFTR after segmental splicing.
After demonstrating the feasibility of the strategy, the constructs were packaged into AAV capsids to determine whether AAV-mediated expression of the constructs was consistent with segmental trans-splicing. A novel AAV capsid, AAV6.2, was chosen for the vector because of its highly efficient in vitro infection of human polarized airway epithelial cultures and highly efficient in vivo infection of mouse airway (Limberis et al., 2009). Coinfection of 293orf6 cells with AAV6.2CFTR Donor and AAV6.2CFTR Acceptor vectors led to the production of correctly spliced CFTR RNA and full-length protein. After coinfection with the same pair of vectors, chloride conductance was observed in FRT-YFP cells as well as in IB3-1 cells, an airway epithelial cell line derived from an individual with CF (Zeitlin et al., 1991).
The efficiency of segmental trans-splicing in this study, with no optimization of splicing signals, was measured as 4.7–12.1%. However, several previous studies involving dual-AAV vector gene delivery strategies that rely on genome concatenation have been published (Halbert et al., 2002; Reich et al., 2003; Lai et al., 2006; Ghosh et al., 2007; Yan et al., 2007). Although these studies demonstrate in vivo efficacy, there are several limitations that prevent efficiency comparisons with the RNA-based trans-splicing approach presented here. These include the use of different promoters controlling gene expression, the use of reporter gene expression as readout as opposed to the percentage of donor and acceptor incorporated into mature mRNA, and the use of coding sequences with differing gene-splitting sites, which affects the efficiency of the process. Several studies compare the efficiency of the dual-gene delivery strategy with small reporter genes with that of a single vector containing the full-size coding sequence. In the case of CFTR, these types of comparisons are not possible because the CFTR coding sequence cannot be packaged into a single AAV vector (Halbert et al., 2002; Reich et al., 2003; Ghosh et al., 2007). The most analogous measurement of splicing efficiency to that presented in our current study involved quantitation of RNA accumulation by RNase protection assay after dual-vector delivery of a minidystrophin gene (Lai et al., 2005, 2006). These studies measured the level of donor cleaved at the splice donor site as a measure of successful trans-splicing; however, it is possible that all donor cleaved at the splice site may not be correctly spliced into the intended acceptor and, therefore, efficiency comparisons with our study, which measures the amount of product formed from successful splicing of donor to acceptor, cannot be made.
Although the efficiency of the segmental trans-splicing process needs to be improved, together, the data reveal that segmental trans-splicing may be a feasible strategy to use in conjunction with AAV6.2 to accomplish sustained expression of CFTR in the airway of individuals with CF.
A variety of other strategies have been used in attempts to overcome the packaging limit of AAV vectors as it relates to CFTR gene therapy. In some studies, investigators dispensed with the promoter region, relying instead on latent promoter activity in the inverted terminal repeat (ITR) of the AAV genome (Flotte et al., 1993a,b). The low efficiency of the ITR as a promoter may have led to reduced levels of AAV-mediated gene expression, using this strategy. In other studies, a truncated form of an active promoter was used with a full-length CFTR cDNA (Zhang et al., 1998; Sirninger et al., 2004). Yet another strategy employed a truncated, functional form of the CFTR gene with an active promoter (Ostedgaard et al., 2005). Finally, a novel strategy used the ability of AAV genomes to undergo intermolecular concatamerization between two independent AAV vectors, resulting in end-to-end continuous DNA segments (Duan et al., 2000b, 2001). If this strategy was applied to two vectors expressing two halves of the CFTR gene, then this mechanism would have the potential to create a complete gene encoding full-length CFTR.
Finally, conventional (as opposed to segmental) trans-splicing has been used to correct endogenous CFTR mRNA (Mansfield et al., 2000; Liu et al., 2002, 2005). In this strategy, a single downstream construct was delivered to cells and trans-splicing was accomplished by using a hybridization sequence that was complementary to an intron sequence upstream of the exon containing the common ΔF508 mutation. Cis-splicing (joining of exons in the original, endogenous CFTR pre-mRNA) occurred preferentially over trans-splicing (joining of the upstream endogenous CFTR pre-mRNA exons to the downstream exogenous, corrected CFTR pre-mRNA exons). Rare, but productive, trans-splicing resulting in CFTR activity was observed in some treated cells. The potential advantages of classical trans-splicing are that only one gene transfer vector is needed and that the production of protein is limited by the amount of endogenous CFTR message and, thus, inherently relies on endogenous regulatory mechanisms that act on the CFTR promoter. However, CFTR exhibits low endogenous transcription rates, and thus classical trans-splicing efficiency is limited by the concentration of one of the two partners in the bimolecular hybridization reaction.
The present study uses segmental trans-splicing, a method developed, in part, to increase the efficiency of trans-splicing by expressing high levels of both of the hybridization partners in the bimolecular hybridization reaction (Pergolizzi et al., 2003; Pergolizzi and Crystal, 2004; Nakayama et al., 2005). In prior studies using segmental trans-splicing, sufficient segmental trans-splicing product was used to mediate phenotypes and correct a genetic defect. Any increase in efficiency of CFTR production through the high expression of both upstream and downstream splicing partners, using the segmental trans-splicing strategy, will have to be balanced against any reduction in efficiency that may occur given the requirement to have a single cell infected by two different AAV vectors. This question will be answered only by further studies using complex in vitro models (e.g., polarized human epithelium) or in vivo models (e.g., CF knockout mice).
The authors thank R. Pergolizzi, J. Schwartz, L. Xu, J. Ang, B. De, and R.B. Silver for help with the studies; and N. Mohamed for help in preparing this manuscript. These studies were supported, in part, by P01 HL51746, Cystic Fibrosis Foundation RDP R881, and the Will Rogers Memorial Fund (Los Angeles, CA).
No competing financial interests exist.