On Behalf of the UK Cystic Fibrosis Gene Therapy Consortium

A Phase I/IIa safety and efficacy study of nebulised liposome-mediated gene therapy for cystic fibrosis supports a multidose trial

Eric WFW Alton*, A Christopher Boyd*, Gwyneth Davies*, Jane C Davies*, Deborah R Gill*, Uta Griesenbach*, Tracy E Higgins*, Stephen C Hyde*, J Alastair Innes*, David J Porteous*,

*Senior Authors

On behalf of the UK Cystic Fibrosis Gene Therapy Consortium

Eric WFW Alton*, Katie J Bayfield, Nicholas Bell, A Christopher Boyd*, June Brand, Andrea Brum, Tabinda Burney, Roberto Calcedo, Paula Carvelli, Faye Chalmers, Mario Chan, Seng H Cheng, D David S Collie, Steve Cunningham, Heather Davidson, Gwyneth Davies*, Jane C Davies*, Lee A Davies, Sarah Davis, Maria Dewar, Ann Doherty, Jackie Donovan, Natalie S Dwyer, Mathew Embley, Duncan M Geddes , James SR Gibson, Deborah R Gill*, Andrew P Greening, Uta Griesenbach*, David M Hansell, Luci Hellings, Tracy E Higgins*, Stephen C Hyde*, Laura Hyndman, J Alastair Innes*, Joseph Jacob, Nancy Jones, Brian F Keogh, Belinda Lees, Maria P Limberis, Paul Lloyd-Evans, Michelle C Manvell, Dominique McCormick, Gerry McLachlan, Fiona McLean, Cuixiang Meng, Gordon D Murray, Nikki Newman, Andrew G Nicholson, Javier Parra-Leiton, David J Porteous*, Ian A Pringle, Phil Reid, Gina Rivellini, Clare J Saunders, Ronald K Scheule, Sarah Sheard, Helen Sheridan, Andrew Simpson, Keith Smith, Stephen N Smith, Samia Soussi, Barbara Stevenson, Stephanie G Sumner-Jones, Nia Voase, Marguerite Y Wasowicz, Abigail Wilson, James M Wilson, Paul Wolstenholme-Hogg, Monica Yanez-Lopez.

Supplementary data

Reconstitution and preparation of pGM169/GL67A

Immediately prior to use, pGM169 and GL67A were thawed to room temperature and GL67A hydrated in water for injection. pGM169/GL67A complexes were prepared by rapidly mixing 5 ml pGM169 and 5 ml GL67A using a disposable syringe-based static mixer device (1). The final dosage form of pGM169/GL67A contains nominally 2.6 mg/ml pGM169, 3.7 mg/ml GL67, 8.9 mg/ml DOPE and 1.7 mg/ml DMPE-PEG5000. The shelf-life of the final formulation was 6 hr.

Dose delivery

Patients were pre-treated with 200-400 g salbutamol (albuterol) via a metered dose inhaler and spacer device approximately 20 minutes prior to receiving the trial treatment to prevent bronchospasm in response to the hypotonic nature of the trial preparation. Nebulization took place in sealed negative-pressure cubicles with external venting to limit spread within the hospital setting; patients were observed and communicated with through a glass window by the trial team. Following weighing, each 5 ml aliquot was nebulized during cycles of tidal breathing whilst patients were wearing a nose clip for 3 minutes, following which the nebulizer air supply (8 L/min) was turned off for a 2 minute rest period. Nebulization continued until delivery was complete, following which devices were re- weighed. Patients in the nasal subgroup administered one actuation of the nasal spray device to each nostril during the 2 minute ‘off-nebulizer’ period, which resulted in an approximate 1 ml dose to each nostril. After dosing, patients either remained in the cubicle for 40 minutes, or were able to leave wearing a mask to limit spread of exhaled trial product in the unit.

Study design and dosing cohorts

This trial was designed as an iterative study, data from dosed patients being presented to the independent Scientific Advisory Commiteee (SAC) and Data Safety & Monitoring Board (DSMB) and used to inform both dosing and outcome measures in future participants; this led to uneven numbers in the three dosing groups and subgroups of patients undergoing CT scans and bronchoscopies. There was no randomisation involved. The first cohort of 3 patients received 10 ml, half the dose administered in our previous trial (Samia, please add ref for Lancet paper- numbers for others will need changing). Once this was determined safe, subsequent patients received 20 ml, some of whom underwent bronchoscopic measurements. Dose-related adverse events led to protocol amendments to explore adjuvant therapies (ibuprofen/ prednisone) and slow delivery rates, following which the 10 ml cohort was increased in numbers and the 5 ml cohort added (both of the latter included some patients receiving adjuvant therapy with paracetamol). On the advice of SAC, imaging was excluded from the protocol once subjects in the 20 ml dosing cohort had shown little change in CT scans, as these carry a significant radiation burden and it was considered unlikely that this safety measure would be useful in the lower dosing cohorts. For similar reasons, bronchoscopies, with the associated significant burden for the patients and trial team, were omitted once the relatively low yield of bronchial transgene-specific mRNA was observed in the 20 ml group.

Lung clearance index

LCI is a measure derived from a multiple breath washout (MBW) which provides a global measurement of ventilation inhomogeneity. It can be performed either with inhalation of an inert tracer gas such as sulphur hexafluoride (SF6) or by using 100% oxygen to wash out resident nitrogen. In the case of an exogenous tracer, the gas is inspired until equilibrium is reached between the inhaled and exhaled air at which point the gas source is removed. The number of lung volume turnovers required until the expired tracer gas concentration falls to an arbitrary 1/40th of its starting value is used to calculate the LCI. Individuals with lung disease and greater ventilation inhomogeneity require longer to clear the tracer gas and therefore will have a higher (more abnormal) LCI. The method adopted in this trial used the tracer gas, SF6 detected by the Innocor photoacoustic gas analyser (2). In brief, the procedure is performed with a mouthpiece during tidal breathing whilst patients wear a nose clip. Triplicate measures are undertaken, each test taking approximately 15 minutes. Data are subsequently analysed off-line according to strict Standard Operating Procedures for quality control using software developed within the Consortium.

Gas transfer

Transfer factor of the lung for carbon monoxide (TLCO, DLCO) was measured using a single breath technique (London site: Jaeger Masterscreen, CareFusion, Germany; Edinburgh site: CPL, nSpire Health, England. Subjects, who were seated and wearing a nose clip, were asked to exhale to residual volume, then take a maximal inspiration (required to be >90% vital capacity) of the test gas (medical air with 0.28% carbon monoxide and 9% helium) prior to a breath hold of 10 secs duration and a smooth exhalation. This procedure was repeated at least twice on each test day (minimal interval between tests 4 mins). Duplicate estimates of TLCO were required to be within 10% or 1.0 mmol.min-1.kPa-1 (whichever was greater) to meet requirements for inclusion. At least 2 technically acceptable tests from up to 10 attempts were obtained and the mean values used for analysis. The following values were recorded for each test: Single Breath TLCO, Alveolar Volume (VA), Transfer Coefficient (KCO, derived from TLCO/AV) and Inspiratory Vital Capacity (IVC). TLCO and KCO were corrected for most recent haemoglobin (Hb, usually obtained on the same day) and expressed as TLCOc and KCOc using standard formulae.

Assessment of immune responses

Anti-ds (double stranded) DNA antibodies (IgG) were processed by the clinical laboratories using a commercial ELiA dsDNA assay (Phadia 250, ThermoScientific) in accordance with routine hospital practice. Peripheral blood mononuclear cells were extracted and human interferon-γ enzyme-linked immunospot assay for the analysis of CFTR-specific T cells was performed as previously described (3).

Nasal potential difference (PD) measurements

Subjects underwent up to 3 pre-dosing nasal PD measurements and post-dosing measurements at various time points as illustrated in Fig 1c. At the time of the first measurement in any subject, single use, disposable, dual-lumen catheters, fashioned from Foley urinary catheters (4) were fixed in place at the site of maximal unstimulated PD in one nostril; for all subsequent measurements, the same nostril and the same distance into the nasal cavity was used. Solutions were manufactured by the Pharmacy at St George’s Hospital, UK according to standard protocols (5) and were perfused at room temperature a rate of 4 ml/min in the following sequence: Ringers, Ringers plus amiloride ( 100 µM), Zero chloride Ringers plus amiloride and Zero chloride Ringers plus amiloride containing isoproterenol ( 10 µM). Traces were obtained using a high-impedance, low resistance voltmeter (LR-4, Logan Research UK) onto a laptop computer and were scored using standard criteria.

Nasal brushings

Nasal brushings were performed in subjects in the nasal subgroup pre-dosing and at time intervals thereafter from under the middle or inferior turbinate of both nostrils using interdental brushes (Dent-o-care, London, UK). Cells were suspended in 800 µl of PBS and processed for transgene DNA/mRNA analysis (see below).

Bronchoscopy: Lower airway PD, brushings and endobronchial biopsy

Bronchoscopy was undertaken in a subgroup of patients for efficacy (gene transfer assessed by mRNA on brushings and PD) and safety (remodelling in endobronchial biopsies). All procedures were performed under general anaesthesia to overcome the challenges of interference from patient movement or coughing, and to avoid local anaesthetic agents which affect ion channel function. In the current trial, all bronchoscopies and associated procedures were performed by a single operator (JCD) with the same senior anaesthetist (BK). Solutions (Ringers and zero chloride) were formulated as for nasal measurements with the exception that amiloride was omitted from the latter. They were warmed prior to use to reduce the formation of microbubbles, but were used at room temperature and perfused at 100 ml/hour. Basal (non-stimulated) PD was measured on each wall of the distal trachea with sterile Ringer’s solution. Where possible, a stable period (<1mV change over 30 seconds) was recorded. Subsequently, at three sites more distally (approximately 5th generation airways; all in one lung), following a stable 30 second period with Ringer’s solution, the solution was switched to a zero-chloride equivalent containing isoproterenol (10 µM) for 5 min. The catheter was removed and re-primed between measurements. Hardware and software were as described for nasal PD. Measurements were completed before the samples described below were obtained.

For analysis, recordings (1-3) from the same patient at the same time point were pre- grouped and scored blinded both for acceptability and response. Recordings were only accepted for scoring if a) the time of any inflexion due to the zero chloride/isoproterenol solution was approximately 50-80 seconds after onset of perfusion, this duration having previously been shown to represent clearing of the dead space within the catheter (~50 secs) and allowing ~30 secs for the initiation of any isoproterenol response, b) the response fell into one of three categories namely no response, continuous upward (hyperpolarization) or continuous downward (depolarization) deflection. Recordings were excluded if a) the response at the 50-80 sec time point was unstable (rising or falling), or b) the deflection was characterised by both a depolarization and a hyperpolarization. In acceptable cases, response was scored at 300 seconds after the switch of solutions.

Ten bronchial brushing samples were obtained from throughout the airways of the same lung using disposable cytology brushes (BC-202D-5010, Olympus UK, Southend-on-Sea, UK); to prevent cell loss which occurs whilst re-sheathing the brush, the entire bronchoscope was removed each time and the brush cut whilst still exposed at the distal end of the scope. Brushed cells were placed into two aliquots of PBS (5 brushings in each) and processed for DNA/ mRNA analysis.

Two endobronchial biopsies were obtained from approximately 5th generation subcarinae with disposable forceps (BC-202D-5010, Olympus UK, Southend-on-Sea, UK) and processed using standard clinical protocols for haematoxylin/ eosin staining.

Molecular analysis of CFTR gene expression

DNA and RNA were simultaneously isolated from nasal and bronchial brushing samples using AllPrep (QIAGEN, Manchester, UK). Levels of pGM169-specific and endogenous CFTR DNA and mRNA were quantified using TaqMan (Life Technologies, Paisley, UK) real-time quantitative PCR instruments and supplies, following reverse transcription (RT) where appropriate, essentially as described (6). Absolute DNA and in vitro transcribed RNA standards allowed precise copy number quantification. pGM169 DNA levels were determined using a qPCR primer set specific for the soCFTR2 cDNA:

50 nM forward primer: 5’-GGAACAGCTCCAAGTGCAAGA-3’

900 nM reverse primer: 5’-CCTGGTGTCCTGCACTTCCT-3’

100 nM probe: 5’-FAM-CAAGCCCCAGATTGCTGCCCTG-TAMRA-3’.

Levels were normalised to total genomic DNA as determined using a qPCR primer set specific for the endogenous CFTR gene:

300 nM forward primer 5’- CTTCCCCCATCTTGGTTGTTC-3’

300 nM reverse primer: 5’-TGACAGTTGACAATGAAGATAAAGATGA-3’

100 nM probe: 5’-VIC-TGTCCCCATTCCAGCCATTTGTATCCT-TAMRA-3’. pGM169-derived mRNA was determined using qRT-PCR. Reverse transcription reactions (20µl) contained ≤5 µl (≤500ng) total RNA, 1.25 units of MultiScribe Reverse Transcriptase (LifeTechnologies) and 0.4 Units RNAse inhibitor (LifeTechnologies) and 400nM of the appropriate primer: soCFTR2 mRNA: 5’-CCAGCTGAAGAACAGCTTGCT-3’ hCFTR mRNA: 5’-CCAGCTGAAGAACAGCTTGCT-3’. pGM169-derived cDNA levels were determined using a qPCR primer set specific for the correctly spliced exon 1-2 soCFTR2 mRNA:

300 nM forward primer: 5’TCTCCCTCCTGTGAGTTTGGTT3’

300 nM reverse primer: 5’GCTCACCACAGAGGCCTTCT3’

100 nM probe: 5’FAMCTAGCCACCATGCAGAGAAGCCCTCTGTAMRA3’.

Levels were normalised to endogenous hCFTR mRNA as determined using a qPCR primer set specific for the correctly spliced exon 1-2 endogenous CFTR mRNA:

300 nM forward primer: 5’GGAAAAGGCCAGCGTTGTC3’

300 nM reverse primer: 5’CCAGGCGCTGTCTGTATCCT3’

100 nM probe: 5’VICCCAAACTTTTTTTCAGCTGGACCAGACCAATAMRA3’. qPCR reactions (10 µl) contained 0.6-6 ng total DNA or 2 µl cDNA and 1xTaqMan Universal MasterMix (LifeTechnologies). Thermocycling was performed in 384-well plates, on a 7900HT cycler, with SDS 2.2 software, and the following cycling conditions: 50°C 2 min, 95°C 10 min, then 40 cycles of 95°C 15 sec, 60°C 1 min. Data were calculated as the percentage of pGM169-specific to endogenous CFTR copy number. Where samples were positive for pGM169-specific signal, but either quantification fell below the linear range of the standards (LOQ - Limit Of Quantification) or quantification of endogenous CFTR signal was negative, they were scored as Positive But Not Quantifiable (PBNQ). Remaining samples that were negative for endogenous CFTR signal were scored as Not Determined (ND). Samples negative for pGM169-specific signal but positive for endogenous CFTR signal were scored as Zero. Where data are presented as post-pre treatment difference, a conservative approach was adopted for samples scored with the nominal value of PBNQ by substituting a value of either 0 or the LOQ as appropriate to maximise the post-pre difference for placebo samples and minimise the post-pre difference for active samples. For bronchial brushings, two nominally identical samples were typically available (each pools of five independent bronchial brushings) and the maximum score was reported.

Acknowledgments We would like to thank the patients and their families, the CF Trust for funding and members of the Scientific Advisory Committee (Stuart Elborn, Pierre Lehn, Brandon Wainwright, Gerry McElvaney, Nikki Samsa, Donna Harcombe) and DSMB (Prof Duncan Empey ([email protected]), Prof David Bennett ([email protected]), Prof Francesco Muntoni ([email protected])).

The research was supported by the National Institute for Health Research (NIHR) Respiratory Biomedical Research Unit at the Royal Brompton and Harefield NHS Foundation Trust and Imperial College London and the Biomedical Research Centre based at Imperial College Healthcare NHS Trust and Imperial College London.

References

1. 1. Davies LA, Nunez-Alonso GA, Hebel HL, Scheule RK, Cheng SH, Hyde SC, Gill DR. A novel mixing device for the reproducible generation of nonviral gene therapy formulations. Biotechniques. 2010;49:666-8.

2. Davies J, Sheridan H, Bell N, Cunningham S, Davis SD, Elborn JS, Milla CE,

Starner TD, Weiner DJ, Lee PS, Ratjen F. Assessment of clinical response to ivacaftor with lung clearance index in cystic fibrosis patients with a G551D-CFTR mutation and preserved spirometry: a randomised controlled trial. Lancet Respir

Med. 2013;1:630-8.

3. Calcedo R, Griesenbach U, Dorgan DJ, Soussi S, Boyd AC, Davies JC, Higgins TE, Hyde SC, Gill DR, Innes JA, Porteous DJ, Alton EW, Wilson JM, Limberis MP. Self-reactive CFTR T cells in humans: implications for gene therapy. Hum Gene Ther Clin Dev. 2013;24:108-15.

4. Middleton PG, Geddes DM, Alton EW. Protocols for in vivo measurement of the ion transport defects in cystic fibrosis nasal epithelium. Eur Respir J. 1994;7:2050-6.

5. Rowe SM, Clancy JP, Wilschanski M. Nasal potential difference measurements to assess CFTR ion channel activity. Methods Mol Biol. 2011;741:69-86.

6. Rose AC, Goddard CA, Colledge WH, Cheng SH, Gill DR, Hyde SC. Optimisation of real-time quantitative RT-PCR for the evaluation of non-viral medicated gene transfer to the airways. Gne Ther. 2002;9:1312-20.