Circulating Truncated Alpha-1 Antitrypsin Glycoprotein in Patient Plasma Retains Anti-Inflammatory Capacity

Abstract

Alpha-1 antitrypsin (AAT) is an acute phase protein that possesses immune-regulatory and anti-inflammatory functions independent of antiprotease activity. AAT deficiency (AATD) is associated with early-onset emphysema and chronic obstructive pulmonary disease. Of interest are the AATD nonsense mutations (termed null or Q0), the majority of which arise from premature termination codons in the mRNA coding region. We have recently demonstrated that plasma from an AATD patient homozygous for the Null Bolton allele (Q0_bolton) contains AAT protein of truncated size. Although the potential to alleviate the phenotypic consequences of AATD by increasing levels of truncated protein holds therapeutic promise, protein functionality is key. The goal of this study was to evaluate the structural features and anti-inflammatory capacity of Q0_bolton-AAT. A low-abundance, truncated AAT protein was confirmed in plasma of a Q0_bolton-AATD patient and was secreted by patient-derived induced pluripotent stem cell–hepatic cells. Functional assays confirmed the ability of purified Q0_bolton-AAT protein to bind neutrophil elastase and to inhibit protease activity. Q0_bolton-AAT bound IL-8 and leukotriene B₄, comparable to healthy control M-AAT, and significantly decreased leukotriene B₄–induced neutrophil adhesion (p = 0.04). Through a mechanism involving increased mRNA stability (p = 0.007), ataluren treatment of HEK-293 significantly increased Q0_bolton-AAT mRNA expression (p = 0.03) and Q0_bolton-AAT truncated protein secretion (p = 0.04). Results support the rationale for treatment with pharmacological agents that augment levels of functional Q0_bolton-AAT protein, thus offering a potential therapeutic option for AATD patients with rare mutations of similar theratype.

Introduction

Alpha-1 antitrypsin (AAT) is the archetypal member of the serpin superfamily produced mainly by hepatocytes and is the most abundant endogenous serine protease inhibitor in the blood. The predominant role of AAT is as a serine protease inhibitor, primarily inhibiting neutrophil elastase (NE) but also other proteases, including cathepsin G and proteinase 3 (1). The structure of the AAT molecule is critical for its antiprotease activity and is comprised of three β sheets (A, B, and C), nine α helices, and a reactive center loop (RCL) at the C-terminal end. Additionally, AAT undergoes posttranslational modifications with the addition of N-linked oligosaccharides at asparagines 46, 83, and 247. AAT is also an acute phase protein, the levels of which become elevated within hours of developing inflammation or postinfection (2), and is known to be raised in a number of conditions, ranging from acute community-acquired pneumonia to postsurgery (3, 4). Indeed, during the resolving phase of community-acquired pneumonia, there is a significant increase in circulating levels of sialylated negatively charged glycoforms of AAT complexed to potent chemokines, including IL-8: a regulatory binding event that negates CXCR1 engagement (2). Moreover, AAT has been shown to possess substantial anti-inflammatory properties independent of its antiprotease activity, affecting many cell types, including pancreatic islet β cells (5), B cells (6), neutrophils (7, 8), and macrophages (9), and has been implicated in modulating cellular processes as diverse as endothelial cell apoptosis (10) and fibroblast-mediated cytokine expression (11).

AAT deficiency (AATD) is a hereditary disorder characterized by low circulating levels of AAT and is associated with the development of chronic obstructive pulmonary disease often by the third or fourth decade, liver disease, and in rare cases, skin panniculitis (12). Studies indicate that neutrophilic inflammation plays a major role in the pathogenesis of AATD-associated emphysema, with increased lung neutrophil burden described in AATD subjects even with mild functional lung impairment (13). Infusion of plasma-purified AAT protein (augmentation therapy) has proven therapeutic benefit in AATD (14, 15) and has been explored in a number of disease models, including diabetes (16, 17) and arthritis (6, 11) and airway diseases, including bronchiectasis/chronic obstructive pulmonary disease and cystic fibrosis (18, 19). Most recently, the Randomized, Placebo-controlled Trial of Augmentation Therapy in α-1 Proteinase Inhibitor Deficiency trial demonstrated the benefit of AAT augmentation therapy as compared with placebo (14), and the Randomized, Placebo-controlled Trial of Augmentation Therapy in α-1 Proteinase Inhibitor Deficiency open-label extension trial further supported the continued efficacy of AAT in decelerating the progression of AATD lung disease over 4 y (15). However, plasma-purified AAT is not a limitless resource, and alternative sources of augmentation therapy, in addition to novel therapeutics, are being sought.

The common normal AAT allele is M, which accounts for 95% of alleles, and in healthy individuals leads to AAT plasma levels >1.04 g/L or 20 μM. The AATD Z allele accounts for 1–2% of alleles in the population and is caused by the substitution of glutamic acid for lysine at position 342. AATD associated with the ZZ phenotype is well described and understood; conversely, little is known about the family of SERPINA1 mutations termed null or Q0, such as the Q0_bolton and Q0_hongkong mutations. Null mutations arise from a number of different types of mutations, including nonsense and frameshift mutations (20), resulting in a premature termination codon (PTC) in the mRNA coding region and undetectable serum levels of AAT by routine nephelometry and isoelectric focusing (IEF) methods. Recently, we identified the presence of truncated AAT protein in plasma of a patient homozygous for the Q0_bolton mutation (21), which represents an attractive therapeutic target. Drugs such as geneticin (G418), amikacin, and ataluren (PTC124) force the ribosome to read through early stop codons (22) and can suppress disease-causing PTCs in mammalian cells in vitro and in vivo (23–25), with therapeutic potential demonstrated in Duchenne muscular dystrophy (DMD) (26) and cystic fibrosis (27–29). Alternatively, several pharmacological agents inhibit nonsense-mediated mRNA decay (NMD), and this is particularly beneficial when the truncated protein encoded by PTC mRNAs retain normal function (30, 31). NMD inhibition (referred to as PTC suppression therapy) has the potential to alleviate the phenotypic consequences of a wide range of genetic diseases by increasing levels of truncated, yet functional, protein to the protective threshold level (32), with associated potential benefit for a range of disorders, including haemophilia and several cancers (25). In this article, we extend these concepts to AATD and characterize the structural and anti-inflammatory properties of the circulating truncated AAT protein arising because of the Q0_bolton mutation. Moreover, we extend our tests on the circumvention of translation-dependent mRNA surveillance to enhance Q0_bolton-AAT protein production.

Materials and Methods

Study design

Ethical approval was obtained from Beaumont Hospital institutional review board, and written informed consent was obtained from all study participants. Healthy control individuals (n = 10, mean age = 31.55 ± 7.05 y) had a mean forced expiratory volume in 1 s (FEV₁) of 103 ± 14.6% predicted, showed no evidence of any disease, were all nonsmokers, and none were taking medication. This group were defined as having an MM phenotype by IEF, with serum AAT concentrations within the normal range (20–50 μM). AAT measurements were performed by a rate immune nephelometric method (Array 360 System; Beckman Coulter) or by immune turbidimetry (AU5400; Beckman Coulter). IEF for phenotyping AAT from plasma was performed using the HYDRASYS platform (Sebia). A nonsmoking ZZ-AATD patient who was not receiving augmentation therapy (mean FEV₁ of 73% predicted) and a heterozygous MZ-AATD individual (mean FEV₁ of 103% predicted) were recruited. A patient (never smoker, mean FEV₁ of 59% predicted) homozygous for the Q0_bolton mutation was recruited, as determined by sequencing all coding exons (II-V) of the AAT gene (SERPINA1, RefSeq: NG_008290) as previously described (33), using the CEQ 8800 Genetic Analysis System (Beckman Coulter) or the Big Dye Terminator Cycle Sequencing Kit 3.1 (Applied Biosystem) with the 3130 Genetic Analyzer. Following sequencing of all coding exons (II-V) of the SERPINA1 gene, the patient was identified as homozygous for the Q0_bolton mutation, with two PTCs at aa 373 and 374 on exon V.

Isolation of plasma and purification of active AAT

Blood was obtained from consenting volunteers in 7.5-ml heparinized S-Monovette tubes (10 U/ml; Sarstedt, Germany). Plasma was immediately isolated by centrifugation of the blood (1000 × g, 10 min at room temperature [RT]) and was then aliquoted and stored at −80°C until required. AAT was isolated from human plasma by use of Alpha-1 Antitrypsin Select Resin packed into a Tricorn column (GE Healthcare Life Sciences, Buckinghamshire, U.K.) and chromatographed by fast protein liquid chromatography on an ӒKTAprime Plus (GE Healthcare Life Sciences). Plasma was diluted with Buffer A (20 mM Tris, 150 mM NaCl, pH 7.4) at a ratio of 1:3 and then loaded onto the column at a flow rate of 0.5 ml/min. Bound AAT was eluted from the resin with a gradient of 0–100% Buffer B (2 M MgCl₂) over 20 ml. Fractions containing AAT were concentrated using Amicon Ultra centrifugal filters (Millipore) and desalted into PBS using NAP-10 desalting columns (GE Healthcare Life Sciences). AAT was quantified by ELISA using the SERPINA1 DuoSet (R&D Systems) according to manufacturer’s instructions, and the plate was recorded using a Spectra Max M3 (Molecular Devices).

Following this, the ability of purified AAT to inhibit NE (Athens Research) activity was evaluated by an NE-specific fluorescence resonance energy transfer (FRET; substrate Abz-APEEIMRRQ-EDDnp; assay buffer [0.5 M NaCl, 0.1% (v/v) Brij-35, 0.1 M HEPES, pH 7.6]) assay. Fixed volumes of NE (containing 1 × 10⁻⁶ M active NE) were incubated with a range of concentrations of AAT (0–5 nM) at 37°C for 30 min. NE with no AAT acted as a control for each experiment. Postincubation, each sample was mixed with FRET substrate in NE assay buffer, and fluorescence was recorded at excitation 320 nm and emission 420 nm at 20 s intervals at 28°C. The percentage of remaining NE was calculated by converting the values into percentages and taking the value obtained for the AAT (2 nM) samples at 5.4 min away from the value obtained from the control NE (4 nM) samples. IC₅₀ measurements for AAT were calculated from the plot of the percentage of remaining activity versus the AAT concentrations, using GraphPad Software (Prism 5.0), as was the amount of AAT required to completely inhibit NE activity. The IC₅₀ for AAT against 1 × 10⁻⁶ M NE activity was 5.41 × 10⁻⁷ M. To completely inhibit 1 × 10⁻⁶ M active NE, 1 × 1.09⁻⁶ M-AAT was required. Reactions were run on SDS-PAGE and immunoblotted for AAT (34).

Protein electrophoresis and Western blot analyses

For one-dimensional gel electrophoresis, SDS-PAGE sample buffer (10× containing 0.2% (w/v) bromophenol blue, 50% (w/v) sucrose, 1% (w/v) SDS, 1% (w/v) DTT, 200 mM EDTA, 3 M Tris-HCL) was added to protein samples, boiled for 3 min, and then subjected to 12.5% (w/v) SDS-PAGE under denaturing conditions. Alternatively, samples were subjected to NuPAGE Novex 4–12% Bis-Tris gel electrophoresis (Bio-Sciences Limited). Two-dimensional (2D) PAGE was performed as previously described (34). Gels were stained with Coomassie Blue stain for visualization of protein banding patterns, or, alternatively, proteins were transferred onto polyvinylidene difluoride membrane for Western blotting. Polyvinylidene difluoride membranes were incubated with 1 μg/ml polyclonal goat AAT Ab or 1 μg/ml polyclonal rabbit AAT Ab (Abcam). The secondary Abs were HRP-linked anti-rabbit (Cell Signaling) or HRP-linked anti-goat (Santa Cruz). Visualization of immune-reactive protein bands was achieved using Immobilon Western Chemiluminescent HRP Substrate (Millipore) and the Syngene G:Box Chemi XL gel documentation system. Densitometry was performed using the GeneSnap SynGene program (Synoptics).

AAT leukotriene B₄ and IL-8 binding and activity assays

UV spectra were recorded on a Jenway 6405 for the in vitro binding of leukotriene B₄ (LTB₄) to AAT or human serum albumin (HSA; control plasma protein) as previously reported (8). The ratio of ligand/protein was 5 μM:1 μM, and UV spectra values were recorded between 230- and 330-nm wavelengths (5-nm intervals) in a quartz cuvette. The molar absorptivity (ε) at 270 nm was calculated as described by Gill and von Hippel (35), with a path length of 1 cm and represented as M⁻¹cm⁻¹. A molar absorptivity of 4.3 × 10⁴ M⁻¹cm⁻¹ at 270 nm (in PBS) was adopted for LTB₄. The interaction of LTB₄ with fast protein liquid chromatography–purified M-AAT or Q0_bolton-AAT was analyzed with solutions prepared in PBS (8). Neutrophil adhesion assays in response to LTB₄ (100 nM) were performed by measuring the number of calcein AM (5 μM; Life Technologies)–loaded neutrophils adhering to a fibronectin-coated plate in the presence or absence of AAT, HSA (control plasma protein, 27.5 μM), or the BLT1 antagonist U-75302 (1 μM; Enzo Life Sciences, Exeter, U.K.) as previously described (8).

The Whatman Minifold I Slot-Blot System (GE Healthcare) was employed to assess binding of AAT to IL-8 using 0.4 μm of nitrocellulose. AAT or HSA (protein control, 5 μg) was added to each well in a volume of 200 μl of PBS under vacuum. Human recombinant carrier-free IL-8 (0.1 μg/ml; Cambridge Bioscience) or PBS was incubated for 1 h at RT, without vacuum. The unbound IL-8 was removed by washing with PBS (3 times), and then each well was blocked with 1% (w/v) gelatin in PBS for 1 h. The wells were then incubated with 1 μg/ml IL-8 mAb (R&D Systems) for 1 h, followed by horse anti-mouse secondary HRP-linked Ab (Cell Signaling). Controls for this experiment excluded IL-8 but included primary and secondary Abs. A final wash step was carried out prior to visualization of the nitrocellulose using Immobilon Western Chemiluminescent HRP Substrate (Millipore).

Preparation of AAT glycans for ultra-performance liquid chromatography

For removal of N-linked glycans, all AAT gel spots were excised from Coomassie Blue–stained 2D gels. Each gel spot was cut into 1-mm² pieces and placed into a polypropylene 96-well microplate. The gels were first washed with acetonitrile (ACN) for 10 min followed by 20 mM NaHCO₃ (pH 7.2)_; this was repeated three times_, with the buffers vacuumed to waste each time. N-glycans were removed by employing the high-throughput method described by Royle et al. (36). The N-linked glycans were released using peptide N-glycanase F (PNGase F), which was prepared using 50 μl of 100 mU/ml PNGase F (GKE-5006D; Prozyme) in 20 mM NaHCO₃. Each gel piece was allowed to soak for 5 min and was then further covered with 50 μl of 20 mM NaHCO₃ buffer alone and transferred to a 2-aminobenzamide (2AB) collection block. The plate was sealed with adhesive film and incubated overnight at 37°C. The following day, the released glycans were collected. Distilled H₂O (dH₂O; 200 μl) was added to each well and placed on the plate mixer for 10 min. This was then vacuumed to the 2AB collection block. This was followed by addition of 200 μl of ACN for 10 min and subsequent vacuuming to the collection block. This cycle was repeated 4 times. Once the samples were dry, 20 μl of 1% (v/v) formic acid was added to the samples and then incubated at RT for 40 min. The samples were dried overnight in a vacuum centrifuge (SpeedVac).

Glycans were fluorescently labeled with 2AB by a reductive amination reaction. In brief, 2AB labeling mixture (LudgerTag kit, U.K.) was added to the samples and incubated at 65°C for 2 h with gentle agitation. Excess 2AB reagent was removed on Whatman 3-mm paper (Clifton, NJ) in ACN. Pieces of Whatman paper were cut (1 cm³) and washed in a beaker of dH₂O 3 times. They were then placed in aluminum foil and dried at 65°C. The pieces were folded into quarters and placed in the wells of a Whatman protein precipitation plate (prewashed with ACN followed by dH₂O). The samples in 5 μl of 2AB labeling solution were then added to the center of the folded paper in the wells and allowed to dry for 15 min. The excess 2AB was removed using ACN with agitation for 15 min; this was then vacuumed to waste and repeated 4 times. The 2AB-labeled glycans were then eluted from the paper by adding 2 × 900 μl of dH₂O and agitating for 30 min and were then collected by vacuum in a 96-well plate. Finally, this was dried overnight in a vacuum centrifuge. The dried glycans were then redissolved in a known volume of dH₂O for analysis by ultra-performance liquid chromatography (UPLC). The glycans were rehydrated in 9 μl of H₂O, and 21 μl of ACN was additionally added and agitated for 1 min. The 30-μl sample was then transferred to an analysis plate. UPLC was performed using a BEH Glycan 1.7 μm, 2.1 × 150 mm column (Waters, Milford, MA) on an Acquity UPLC (Waters). Solvent A was 50 mM formic acid adjusted to pH 4.4 with ammonia solution. Solvent B was ACN. The column temperature was set as 30°C. Fluorescence was measured at 420 nm with excitation at 330 nm. The system was calibrated using an external standard of hydrolyzed and 2AB-labeled glucose oligomers to create a dextran ladder, as previously described (37).

For exoglycosidase digestions, all enzymes were purchased from Prozyme (San Leandro, CA). The labeled glycans were digested in a volume of 10 μl for 16 h at 37°C in 50 mM sodium acetate buffer, pH 5.5 (except in the case of jack bean α-mannosidase, in which the buffer was 100 mM sodium acetate, 2 mM Zn²⁺, pH 5), using the following enzymes: 0.5 U/ml Arthrobacter ureafaciens sialidase (EC 3.2.1.18), 1 U/ml Streptococcus pneumonia sialidase (NAN1, EC 3.2.1.18), 1 U/ml bovine testes β-galactosidase (EC 3.2.1.23), 1 U/ml bovine kidney α-fucosidase (EC 3.2.1.51), 8 U/ml β-N-acetylglucosaminidase cloned from S. pneumonia expressed in Escheria coli (EC 3.2.1.30), 60 U/ml jack bean α-mannosidase (EC 3.2.1.24), and 0.4 mU/ml almond meal α-fucosidase (EC 3.2.1.111). After incubation, enzymes were inactivated by incubation at 65°C for 15 min. The enzymes were then removed by filtration through a 10-kDa protein-binding EZ filter (Millipore). N-glycans were then dried by vacuum centrifuge and analyzed by UPLC (38).

Liquid chromatography tandem mass spectrometry was performed on individual gel pieces so as to confirm the identity of AAT, using Proteome Discoverer Software (v1.4; Thermo Fisher Scientific) using a complementary two stream algorithm search with SEQUEST and MASCOT against a human subset from the UniProtKB/Swiss-Prot database. Carbamidomethylation of cysteine residues was selected as a fixed modification, and oxidation of methionine was considered as a variable modification also allowing for two missed cleavages. The following SEQUEST filters were applied: for charge state 1, X_corr > 1.9; 2, X_corr > 2.2; 3, X_corr > 3.75 and peptide Delta correlation (maximum delta Cn0.1). The following MASCOT filters were applied: MASCOT threshold score of 40 and MASCOT significance threshold of 0.05.

Generation and differentiation of induced pluripotent stem cell–derived hepatic cells

To isolate human dermal fibroblasts for reprogramming, 6-mm full-thickness skin biopsies were performed and fibroblasts were expanded using previously described methods (Supplemental Fig. 1) (39). Fibroblasts (1 × 10⁵) were transduced with the excisable hSTEMCCA reprogramming vector, and emerging colonies were picked and characterized to confirm normal karyotype and expression of markers of pluripotency before proceeding to additional experiments (Supplemental Fig. 1) (40). Wild-type control induced pluripotent stem cell (iPSC) experiments were performed using the previously published human iPSC WT lines, BU-1 and BU-3 (41). Human iPSCs were adapted to feeder-free conditions and cultured on matrigel in mTeSR1 media (StemCell Technologies) prior to initiation of directed differentiation experiments. Directed hepatic differentiation was performed using a previously described differentiation protocol with slight modifications. Briefly, undifferentiated iPSCs were dissociated with Gentle Cell Dissociation reagent (StemCell Technologies) and 1 × 10⁶ cells plated per well of a matrigel-coated six-well plate. Twenty-four hours later, culture media was aspirated and cells were cultured using the STEMdiff Definitive Endoderm Kit according to the manufacturer’s instructions. After 6 d of culture in these media conditions, cells were differentiated according to a previously published protocol (40).

Lentiviral production and cell culture of HEK-293 cells

Q0_bolton-AAT cDNA was amplified by PCR from differentiated Q0_bolton iPSC-hepatic cells using primers including Not I and Bgl II restriction sequences at the 5′ and 3′ ends, respectively. Amplified cDNA was digested and cloned into the first position of the previously generated pHAGE lentiviral backbone, lenti CMV-AAT-UBC-GFP (42). To induce expression of transgenes, HEK-293 cells were transduced using the aforementioned lentiviral vectors to coexpress the enhanced GFP together with either M or Q0_bolton-AAT cDNA under control of the CMV or UBC promoter element. GFP⁺ cells were sorted using a MoFlo cell sorter (DakoCytomation) to 98.5% purity for use in further experiments. HEK-293 cells were cultured in DMEM (Life Technologies) supplemented with 10% (v/v) FBS (Biosciences), 1% (v/v) L-Glutamine (Life Technologies), and 0.2% (v/v) primocin (Invivogen). Once the cells were 80% confluent, media was removed and discarded, and wells were washed with Dulbecco's PBS before detaching the adherent cells using 0.05% (w/v) trypsin-EDTA (Life Technologies) at 37°C for 2–3 min or until cellular detachment occurred. After this period, 7 ml of growth medium was added to inhibit the activity of trypsin, and the cells were centrifuged at 300 × g for 5 min. Cells at a density of 1 × 10⁵ were seeded in 12-well plates and allowed to adhere overnight before culturing in serum-free media containing either gentamicin (0.5 mg/ml) or ataluren (0.1, 0.5, 2.5, 12.5, or 62.5 μg/ml). The employed doses were based on that of a study with gentamicin and ataluren (PTC124) on fibroblast cells (43). The supernatants were collected after this time, and RNA was collected by subjecting the cells to TRI Reagent. Protein was isolated from the TRI Reagent sample using four times the volume of 100% (v/v) acetone with incubation on ice for 30 min. The resultant pellet was washed briefly with acetone and solubilized with 2× SDS-PAGE sample buffer and then analyzed by Western blotting.

Throughout the experiments, cell viability was assessed as per manufacturer’s instructions using the CellTiter 96 AQueous One Solution Cell Proliferation Assay (Promega) by pipetting 20 μl of CellTiter96 reagent into wells of a clear 96-well plate containing 100 μl of samples in culture medium. This was incubated at 37°C for 1–4 h in a humidified 5% CO₂ atmosphere. The absorbance was recorded at 490 nm on a Spectra Max M3 (Molecular Devices, U.K.). Toxicity was determined by comparing the viability of treated cells to untreated control cells.

RNA isolation and quantitative real-time PCR

Total RNA was extracted from HEK-293 cells using TRI Reagent as per the manufacturer’s instructions. In brief, 500 μl of TRI Reagent was added to cell pellets and subsequently, 100 μl of chloroform was added, vortexed briefly, and incubated at RT for 10 min. Centrifugation at 12,000 × g at 4°C separated the sample into three phases. The aqueous RNA fraction at the top was taken off and transferred into a fresh tube with 250 μl of isopropanol and incubated for 5 min at RT. A pellet was formed by centrifugation at 12,000 × g at 4°C for 10 min. The supernatant was discarded, and the pellet was washed with 700 μl of 75% (v/v) ethanol and centrifuged for 7500 × g at 4°C for 5 min. The RNA pellet was subsequently air dried, and 25 μl of diethyl pyrocarbonate dH₂O was added. The RNA was stored at −80°C until ready to use. The RNA content was quantified using a Nanodrop 8000 Spectrophotometer (Thermo Scientific, Ireland). The isolated RNA was considered free of contaminants when the A260/280 was >1.7.

Prior to cDNA synthesis, contaminating genomic DNA was removed from 500 ng of RNA in a total volume of 14 μl using genomic DNA Wipeout Buffer from the QuantiTect Reverse Transcription Kit (Qiagen). This was heated at 42°C for 2 min. The RNA was then reverse transcribed to cDNA with 1 μl of RTase, 4 μl of 5× reverse transcriptase buffer, and 1 μl of reverse transcriptase primer mix per 2-μl sample. An RTase-free sample was prepared by supplementing RNase-free H₂O for RTase. cDNA was prepared on a PTC-200 Thermo Cycler (MJ Research, MN) with the following cycles: 30 min at 42°C, 3 min at 95°C, and at 4°C for infinity. For quantitative real-time PCR (qRT-PCR) analyses, the Lightcycler 480 (Roche Diagnostics) was employed, and the following program was used: preincubation (95°C, 3 min); amplification, 40 cycles consisting of denaturation (10 s, 95°C), annealing (with an optimized annealing temperature of 56°C for 10 s), and elongation (72°C for 10 s); melting curve analysis (95°C, 5 s; 65°C, 1 min; and 97°C continuous acquisition); and final cooling step at 4°C. The expression of target genes relative to GAPDH was determined using the 2^−∆∆cycle threshold method (44). AAT forward primer was 5′-3′:5′-TGGATTTGGTCAAGGAGCTT-3′, and reverse primer was 5′-3′:5′-CATGCCTAAACGCTT CATCA-3′. GAPDH forward primer was 5′-3′:5′-CATGAGAAGTATGACAACAGCCT-3′, and reverse primer was 5′-3′:5′-AGTCCTTCCACGATACCAAAGT-3′.

For mRNA stability assays, experiments were carried out as previously described (45). In brief, HEK-293 cells at a density of 1 × 10⁵ were seeded in 12-well plates, allowed to adhere overnight, and then were serum starved for 16 h. After this time, the media was removed, and the wells were washed with Dulbecco's PBS before the addition of serum-free media with or without ataluren (62.5 μg/ml) for 30 min, followed by actinomycin D (10 μg/ml) for 30 min. After 1, 2, or 3 h, the cells were washed with PBS before being lysed in TRIzol reagent. Following this, total RNA was extracted from cells as per the manufacturer’s instructions. RNA was quantified using the Nanodrop 8000 Spectrophotometer (Fisher Scientific, Ireland). RNA (100 ng) was reverse transcribed to cDNA using the QuantiTect Reverse Transcription Kit (Qiagen, U.K.) as per manufacturer’s instructions. qRT-PCR was performed using a Lightcycler 480. Q0_bolton-AAT was detected using the PCR primers 5′-TGGATTTGGTCAAGGAGCTT-3′ and 5′-CATGCCTAAACGCTTCATCA-3′, and RPLPO were detected using the 5′-GGCAGCATCTACAACCCTGA-3′ and 5′-GGCAGCATCTACAACCCTGA-3′ primers (MWG Eurofins). qRT-PCR was performed using the following protocol: preincubation (95°C for 3 min); amplification (50 cycles consisting of denaturation, annealing, elongation [10 s at 95°C, 10 s at 55°C, and 10 s at 72°C]); melting curve analysis (95°C for 5 s, 65°C for 1 min, and 97°C for five continuous acquisitions); and a final cooling step to 4°C. Expression of AAT relative to RPLPO was determined using the 2^−ΔΔ cycle threshold method. The t_1/2 was determined by comparing the percentage of Q0_bolton-AAT mRNA remaining at each time point with that at time 0.

Confocal microscopy

Cells were fixed with 4% (w/v) paraformaldehyde on a polysine glass slide, and cell membranes were permeabilized using 0.2% (v/v) Triton x-100 in PBS for 5 min at RT. Nonspecific binding was prevented by blocking with 4% (w/v) BSA in PBS. To determine protein colocalization, cells were incubated with 1 μg/ml FITC-labeled goat polyclonal anti-AAT (Abcam, Cambridge, U.K.). Cells were mounted using Vectashield mounting medium with DAPI for nuclear staining (Vectashield Lab, Burlingame, CA). The controls for these experiments included cells alone and nonspecific isotype control IgG (Santa Cruz, TX). All immunofluorescence was viewed and images were acquired using a Zeiss LSM 710 confocal immunofluorescence microscope (Zeiss, Germany). Images were captured with excitation wavelengths for DAPI and FITC of 364 and 488 nm, respectively. The images were represented as two-and-a-half–dimensional reconstructions using Zen software (2011 Edition; Zeiss).

Statistical analysis

Data sets were analyzed for statistical significance using GraphPad Prism 5.0 software package (GraphPad Software) with significance determined at p < 0.05. For data sets with three or more paired groups, a repeated measures ANOVA was employed followed by post hoc Bonferroni multiple comparisons test. To determine mRNA stability experiments, a nonlinear fit one-phase exponential decay curve was employed. For the glycosylation data set, the 95% confidence intervals (CI) were calculated. Results are expressed as mean ± SEM of biological replicates or independent experiments, as stated in the figure legends.

Results

Q0_bolton-AAT mutation is associated with truncated AAT protein

A patient was identified as homozygous for the Q0_bolton mutation as a result of two PTCs at aa 373 and 374 on exon V (21). High-resolution computerized tomography scan of the thorax demonstrated panlobular emphysema with bibasal predominance. Pulmonary function testing subsequently confirmed an obstructive pattern of moderate severity (FEV₁ = 59% predicted; FEV₁/forced vital capacity (FVC) = 0.55) with a minimal bronchodilator response and impaired diffusion capacity of the lung for carbon monoxide (56% predicted; Fig. 1A). AAT phenotype testing by IEF revealed no discernable AAT protein banding pattern in plasma of the patient homozygous for the Q0_bolton mutation (Fig. 1B). Of note, IEF used for AAT phenotyping has a lower limit of sensitivity of 0.05 g/L (46). Controls for the latter assay include a sample from an individual homozygous for the common normal AAT M allele. The resultant M-AAT protein has five distinct bands on IEF gels termed M2, M4, M6, M7, and M8. The ZZ-AATD protein demonstrates three IEF bands, Z2, Z4, and Z6, as well as the classic cathodal shift (Fig. 1B). To understand potential conformational changes to the tertiary structure of the Q0_bolton-AAT molecule, molecular modeling was performed (Fig. 1C). The native AAT structure is in a kinetically trapped meta-stable state, with the rapid folding of residues 383–400 (18 residues, strands 1c and 4b) responsible for kinetic trapping (47). Of these 18 residues, 14 remain in the Q0_bolton-AAT molecule; however, it is also missing a further 18 residues (strand 5b). Molecular dynamics simulations of Q0_bolton-AAT based on the meta-stable AAT form (Fig. 1C, missing residues in orange) did not show any significant changes to residues 383–396, suggesting that there are sufficient interactions involving these residues to stabilize a native-like structure should it form during the early steps of protein folding, and thus these residues may be sufficient to kinetically trap a meta-stable form. Ensuing experiments used conventional column chromatography to purify Q0_bolton-AAT.

FIGURE 1.

PTC in the SERPINA1 gene encoding truncated AAT protein. (A) Chest high-resolution computerized tomography scan demonstrating panacinar emphysema and bronchiectatic changes of the lower lobes. (B) IEF patterns of AAT phenotypes. MM-AAT glycoforms (M2–M8) are denoted on the left and ZZ (Z2, Z4, and Z6) are shown by the broken arrow. Plasma from an individual homozygous for the Q0_bolton mutation is presented on the right. Levels of plasma AAT (grams per liter) for each phenotype are indicated. (C) Molecular model of glycosylated Q0_bolton-AAT. Green, peptide; blue, glycans; red, RCL (residues M382–S383); pink, altered amino acid sequence (residues 386–396); orange, deleted amino acid sequence (residues 396–418).

Q0_bolton-AAT is produced by, and secreted from, hepatocytes

The reported amount of circulating AAT in the proband was <0.1 g/L as measured by immune nephelometry, a commonly used standard clinical assay for serum protein determinations. This suggests Q0_bolton-AAT is present in extremely low concentrations that are below the lower limit of detection for this standard clinical assay. In turn, however, truncated Q0_bolton-AAT protein was purified from the patient’s plasma by use of Alpha-1 Antitrypsin Select, a resin with high selectivity for AAT. From whole plasma (200 μl), we purified ∼190 μg of AAT from the MM plasma and 2 μg (0.01 g/L) of protein from the Q0_bolton plasma, as determined by BCA assay. Coomassie-stained one-dimensional SDS-PAGE bands from the peak fraction containing the AAT protein is shown in Fig. 2A. 2D-PAGE staining revealed eight glycoforms of M-AAT and seven glycoforms of the Q0_bolton-AAT protein run over a pH range of 4–7 (Fig. 2B), with Q0_bolton-AAT demonstrating an anodal shift compared with M-AAT. To further confirm the identity of the protein that was extracted from whole plasma, both the spots representing purified M-AAT and the Q0_bolton-AAT protein were excised from the gel and assessed by LC-MS/MS. Mass spectrometry confirmed each of the protein spots as AAT with a high MASCOT score achieved for both samples (mean 63.64 and 54.50% coverage for M-AAT and Q0_bolton-AAT, respectively). Western blot analysis with a polyclonal goat AAT Ab (Fig. 2A, middle panel) confirmed the presence of AAT and a size shift for Q0_bolton-AAT. We further investigated the reduced molecular mass of Q0_bolton-AAT by running 8% (w/v) SDS-PAGE and visualized the 49-kDa truncated protein on a Western blot using a rabbit Ab for AAT (Fig. 2A, bottom panel). Glycosylation of proteins can result in a heterogeneous migratory pattern on gels; thus, to confirm the reduced molecular mass, we treated purified M-AAT and the Q0_bolton protein with PNGase F to remove the entire N-glycan structure and then subjected the treated protein to Western blot analysis. All of the 52-kDa M-AAT was reduced to 44 kDa, and the 49-kDa Q0_bolton-AAT protein was reduced to 41 kDa upon PNGase F treatment (Fig. 2C). These results confirm the successful isolation of Q0_bolton-AAT as a low-m.w. truncated glycosylated protein.

FIGURE 2.

Isolation of truncated AAT protein from Q0_bolton plasma. (A) Coomassie Blue–stained SDS-PAGE showing AAT purified from MM and Q0_bolton plasma at 1 μg (lane 1 and 3) and 0.1 μg (lane 2 and 4) loadings. Purified MM and Q0_bolton-AAT ran at 52 and 49 kDa, respectively (top panel). To confirm identity of AAT, purified protein was subjected to Western blot analyses with goat polyclonal (Gt anti-AAT, middle panel) or rabbit polyclonal Ab to AAT (Rb anti-AAT, bottom panel). Molecular mass markers are indicated at the left margin in kilodaltons. (B) Coomassie Blue–stained 2D-PAGE run over a linear pH range of 4–7. AAT glycoforms are numbered. (C) Western blot of purified AAT protein was treated with PNGase, and electrophoretic mobility of glycosylated 52-kDa M-AAT and 49-kDa truncated Q0_bolton-AAT compared with deglycosylated (degly) protein (44 and 41 kDa, respectively) is demonstrated. (D) MM control and Q0_bolton patient-derived iPSC-hepatic cells exhibit intracellular AAT. By confocal microscopy, the distribution of AAT (green) in MM and Q0_bolton patient-derived iPSC-hepatic cells presented as punctuate staining throughout the cell. Cell nuclei are stained blue with DAPI. (E) MM control and Q0_bolton patient-derived iPSC-hepatic cells exhibit extracellular AAT. ELISA demonstrated that MM and Q0_bolton iPSC-hepatic cells secrete ∼8500 and 600 ng/ml of AAT, respectively (ANOVA, p < 0.0001; n = 3 technical repeats). Experiments illustrated in (A)–(C) are each representative gels and blots of three separate experiments. Confocal analysis in (D) is a representative result of three experiments.

Next, we explored whether iPSC-derived hepatic cells could serve as an appropriate cellular model to study the Q0_bolton-AAT protein (Supplemental Fig. 1) and to confirm that the vascular origin of truncated Q0_bolton-AAT was via hepatic secretion. Confocal imaging of both healthy MM and Q0_bolton iPSC-hepatic cells stained with anti-human AAT Abs revealed a punctuate pattern throughout the cell, confirming the production of AAT protein by hepatic cells despite the nonsense mutation, albeit with considerably reduced intensity (Fig. 2D). In accordance with low plasma levels of Q0_bolton-AAT in vivo, we found secreted levels of Q0_bolton-AAT from patient-derived iPSC-hepatic cells to be ∼7% of controls (581 ± 40 ng/ml versus 8522 ± 674 ng/ml; mean ± SD; p < 0.0001) after 25 d in culture (Fig. 2E). Collectively, these results confirm the hepatic production and secretion of Q0_bolton-AAT and its presence in the plasma of a null AATD homozygote.

Q0_bolton-AAT maintains antiprotease and anti-inflammatory capacity

The main function of AAT is to act as an antiprotease, and previously we demonstrated the ability of Q0_bolton-AAT to inhibit NE (21), a result confirmed in this study. AAT readily forms a covalent complex with NE at approximately a 1:1 molar ratio, yielding a protein complex of a combined molecular mass of 81 kDa (48). The Western blot in Fig. 3A depicts the AAT complex that occurs when M-AAT (52 kDa) and NE (29 kDa) interact at a 1:1 ratio for 30 min, with minimal levels of free AAT (52 kDa) remaining. Although less active than M-AAT, Q0_bolton-AAT successfully reacted with the protease, as indicated by an increase in the intensity of an immunoband of ∼78 kDa (Fig. 3A, lower panel). We extended this set of experiments and employed FRET analysis to demonstrate that both M-AAT and Q0_bolton-AAT can inhibit NE activity (Fig. 3B). The percentage of NE (4 nM) remaining after treatment with AAT (2 nM) for 5.4 min was recorded at excitation 320 nm and emission 420 nm. M-AAT was shown to reduce NE activity by 58%, and although significantly less, Q0_bolton-AAT reduced NE activity by 13% (n = 3, p = 0.04) (Fig. 3B), suggesting that it may provide lung tissue protection if present in sufficient levels.

FIGURE 3.

Q0_bolton-AAT maintains antiprotease and anti-inflammatory properties. (A) Formation of AAT/NE inhibitory complexes. M-AAT (top panel) was incubated with NE at an AAT/NE molar ratio of 1:1 for 10 min (M-AAT). Reaction products were subjected to Western blot analysis employing a goat anti-human AAT Ab for AAT or AAT/NE complexes. M-AAT incubated with NE resulted in the formation of an 81-kDa AAT/NE complex. Q0_bolton-AAT incubated with NE for 0, 5, 10, or 20 min resulted in the formation of a 78-kDa AAT/NE complex. Experiments illustrated are a representative of three separate experiments. (B) FRET analyses of the percentage of NE (4 nM) activity remaining after treatment with AAT (2 nM) for 5.4 min. M-AAT significantly reduced the level of active NE compared with Q0_bolton-AAT (n = 3, two-tailed Student t test). (C) Q0_bolton-AAT binds IL-8. Slot blot of 5 μg of M-AAT, Q0_bolton-AAT, or HSA (positive control) for IL-8 binding. Blots were incubated with carrier-free IL-8, and the binding event was detected by employing an IL-8 mAb. Control (Con) for nonspecific binding excluded IL-8 but included primary and secondary Ab. Total immobilized protein was visualized by Ponceau S Stain (bottom panel). Densitometry values of IL-8 binding to both AAT forms were normalized to the HSA control and demonstrate the ability of M-AAT (p = 0.005) and Q0_bolton-AAT (p = 0.007) to bind IL-8 (Student t test; n = 3 independent experiments). (D) Glycoanalysis of AAT protein. Q0_bolton-AAT and M-AAT were excised from Coomassie Blue–stained gels and analyzed by UPLC. Values indicate individual biological replicates (n = 3) of the M-AAT.

In subsequent experiments, we investigated the anti-inflammatory capacity of Q0_bolton-AAT. It has previously been shown that M-AAT binds IL-8 via glycan sites on the molecule, thereby modulating neutrophil migration (34). To confirm the competence of Q0_bolton-AAT as an immunoregulatory molecule, its ability to bind IL-8 was assessed in vitro by use of slot blot analysis. To confirm that the effects shown in this study do not simply reflect the nonspecific effect of a protein, in the assay, we extended this experiment to include HSA as a control plasma protein. Results revealed that, compared with HSA, Q0_bolton-AAT bound significantly higher levels of IL-8 (p = 0.007), equivalent to that of M-AAT (Fig. 3C). In a subset of control experiments, IL-8 was omitted from reactions, with no nonspecific interaction between AAT, or HSA, and the anti–IL-8 Abs detected.

As the glycan moieties of AAT are essential for not only the serum t_1/2 of AAT but also its capacity to bind IL-8, it was essential to investigate the glycosylation pattern of Q0_bolton-AAT. Consequently, glycoanalysis was performed on plasma-purified M-AAT and Q0_bolton-AAT cut from Coomassie Blue–stained 2D gels. The N-glycans of purified M-AAT and Q0_bolton-AAT were analyzed by UPLC in combination with exoglycosidase digestions and structural assignments. Typical chromatograms of hydrophilic interaction liquid chromatography columns of undigested AAT from a healthy MM individual and the Q0_bolton AATD patient are shown in Supplemental Fig. 2, and the list of identified Q0_bolton-AAT glycans is presented in Supplemental Table I. The core (α1-6 linked), outer-arm (α1-3 linked), and total fucosylation as well as amounts of biantennary, triantennary, and tetra-antennary glycans were calculated in all samples from the percentages of peak areas after A. ureafaciens sialidase digest, as presented in Supplemental Table II. The 95% CI were calculated for each of the M-AAT samples. For all except biantennary, the percentage of glycans in the Q0_bolton-AAT sample was greater than the 95% CI (Fig. 3D). Core fucosylation for M-AAT was calculated to be 5.35% (95% CI: 4.69–6.02) compared with 8.3% for the Q0_bolton-AAT sample, and outer-arm fucosylation for M-AAT was reported to be 9.51% (95% CI: 6.43–12.60) compared with 16.32% for the Q0_bolton-AAT. Total fucosylation was demonstrated to be 14.36% (95% CI: 11.11–17.61) in M-AAT samples, whereas it was found to be increased to 23.86% in the Q0_bolton-AAT samples. Furthermore, there appears to be a difference in the percentage of branched glycans between the two sample types, with M-AAT reporting a higher percentage of biantennary glycans (77.62%; 95% CI: 71.67–83.56) compared with the 62.81% reported in the Q0_bolton-AAT samples. Additionally, there was a decrease in the percentage of both tri- (19.84%; 95% CI: 15.20–24.48) and tetra-antennary (2.54%; 95% CI: 0.63–4.45) glycans in M-AAT samples compared with Q0_bolton-AAT samples (32.01 and 5.16%, respectively) (Fig. 3D). This trend toward increased core and outer-arm fucosylation differentiates Q0_bolton-AAT from M-AAT and is consistent with persistent inflammation (49), although these differences do not appear to impact the binding capacity of Q0_bolton-AAT for IL-8.

Moreover, the capacity of AAT to disable the stimulating effect of LTB₄ has been previously shown, with the mechanisms of inhibition involving direct binding of LTB₄ via a hydrophobic pocket on the surface of the molecule (8). Additionally, a previous study has detailed the ability of LTB₄ to interact with HSA (50). For this analysis, the structure of LTB₄ is key, as it comprises a triene moiety that can be recorded by UV absorption at 262, 270, and 282 nm (50), and when protein conjugated, the triene structure of LTB₄ becomes planar. In control experiments of the current study, the UV spectrum of free LTB₄ gave rise to the characteristic vibrational structure in PBS solution (Fig. 4A) (51). By use of a 5:1 mixed ratio of LTB₄ to HSA or M-AAT, it was confirmed that LTB₄ interacts with HSA and M-AAT, demonstrating a reduction in ε_max at 270 nm compared with that of LTB₄ alone (Fig. 4A). Moreover, an LTB₄/AAT molar ratio of 5:1 resulted in an ∼45% decrease compared with LTB₄ alone, with area under the curve analysis of UV profiles of three technical replicates performed on three separate days revealing a significant decrease in the presence of M-AAT when compared with the control unbound free LTB₄ or HSA bound LTB₄ (p = 0.006 and p = 0.001, respectively) (Fig. 4A). Subsequent experiments explored the ability of Q0_bolton-AAT to bind LTB₄. In the current study, Q0_bolton-AAT protein retained the ability to bind LTB₄ as evaluated by a reduction in ε_max at 270 nm, similar to that of M-AAT_. An LTB₄/Q0_bolton-AAT molar ratio of 5:1 resulted in a significant 40% decrease compared with the LTB₄ profile alone (p = 0.004), as deduced by area under the curve analysis (Fig. 4C). Q0_bolton-AAT bound to LTB₄ to the same extent as M-AAT at equivalent concentration.

FIGURE 4.

Q0_bolton-AAT binds LTB₄. (A) The triene chromophore structure of LTB₄ (solid line) with the addition of M-AAT (dashed line) or HSA (dotted line). The molar ratio of ligand to protein was 5:1. UV spectra were recorded between 230 and 320 nm. The three arrows indicate covalently bound atoms in the LTB₄ structure at 262, 270, and 283 nm. The molar absorptivity (ε) at 270 nm was calculated and represented as M⁻¹cm⁻¹ with a path length of 1 cm. Area under the curve analysis demonstrates the ability of M-AAT (p = 0.006) to bind LTB₄ (Student t test; n = 3 independent experiments). (B) The triene chromophore structure of LTB₄ (solid line) with the addition of M-AAT or Q0_bolton-AAT (broken line). Area under the curve analysis demonstrates the ability of M-AAT (p = 0.0006) and Q0_bolton-AAT (p = 0.004) to bind LTB₄ compared with uncomplexed LTB₄ (Student t test; n = 3 independent experiments). (C) Neutrophil adhesion. Neutrophils (5 × 10⁶ cell/ml) were loaded with calcein AM dye and stimulated with LTB₄ (100 nM) in the presence or absence of AAT (27.5 μM) or HSA (27.5 μM). AAT treatment significantly reduces neutrophil adhesion compared with HSA-treated cells (p = 0.01). (Student t test; n = 5 technical repeats). (D) Q0_bolton-AAT significantly reduced cell adhesion (p = 0.04). The BLT1 antagonist U-75302 served as a positive control (Student t test, n = 3 independent experiments).

A number of studies have documented the activating effects of LTB₄ on neutrophil adhesion (52), and exposure of cells to LTB₄ in the presence of AAT has been shown to reduce neutrophil adhesion (8). Accordingly, ensuing experiments examined the effect of Q0_bolton-AAT on LTB₄-induced neutrophil adhesion. In control experiments, calcein AM–loaded neutrophils were stimulated with LTB₄ (100 nM) in the presence and absence of AAT or HSA (control plasma protein, 27.5 μM), and adherence to fibronectin-coated surfaces was assessed after 30 min incubation at 37°C (Fig. 4C). LTB₄ significantly increased neutrophil adhesion compared with unstimulated control cells (p = 0.01). A physiological concentration of 27.5 μM AAT significantly reduced the LTB₄ response to that of unstimulated control cells (p = 0.02), and AAT significantly reduced cell adhesion compared with the effect of HSA (p = 0.01) (Fig. 4C). Moreover, as illustrated in Fig. 4D, 27.5 μM Q0_bolton-AAT (p = 0.04) significantly inhibited LTB₄-induced cell adhesion, comparable to the effect of M-AAT. U-75302 (1 μM) was employed as a positive control (p = 0.04, Student t test, n = 3). Collectively, these results demonstrate that Q0_bolton-AAT, despite its truncated size, maintains antiprotease activity and normal ability to interact with the proinflammatory molecules IL-8 and LTB₄, thereby modulating cell adhesion.

AAT production is increased in response to readthrough compounds

The fundamental objective of theratyping is to partner medications to specific mutations. Therapies for AATD individuals with nonsense mutations could include readthrough compounds, leading to translation of the full-length AAT protein. iPSC-derived Q0_bolton hepatic cells treated with single-dose ataluren demonstrated increased AAT protein secretion (21). With this in mind, our aim was to evaluate additional readthrough compounds, over a range of concentrations, for their ability to overcome the PTC in the SERPINA1 gene. To achieve this, HEK-293 cells transduced with lentiviral vectors containing M-AAT or Q0_bolton-AAT were generated (Fig. 5A). A 2-fold reduction in basal mRNA expression was apparent in the PTC containing Q0_bolton-AAT mRNA compared with M-AAT mRNA (p = 0.01) (Fig. 5B), and the level of truncated AAT secreted from Q0_bolton-AAT–expressing cells was of lower abundance and molecular mass compared with M-AAT (p = 0.0001) (Fig. 5C). The in vitro data indicate that the protein levels detected are reduced and out of proportion to the mRNA levels. To understand why this was the case, we assessed whether the Q0_bolton-AAT protein was susceptible to degradation. To analyze this, M-AAT and Q0_bolton-AAT protein was subjected to two freeze-thaw cycles. Robustness of a protein against freezing and thawing depends on its conformational stability, and results of the current study demonstrate that although M-AAT remained intact after two cycles of freeze-thaw (−80°C), the level of Q0_bolton-AAT protein detected by Western blot analysis was reduced (Fig. 5D).

FIGURE 5.

HEK-293 M-AAT and Q0_bolton-AAT protein production. (A) Schematic of lenti CMV-AAT-UBC-GFP. The enhanced GFP reporter gene is constitutively expressed in the second position together with either M-AAT or Q0_bolton-AAT under control of the human UBC promoter. (B) Q0_bolton-AAT HEK-293 cells express significantly less AAT mRNA than the cells expressing M-AAT (p = 0.011, Student t test; n = 3). Cells not transduced with AAT acted as a negative control for AAT mRNA expression (Con). (C) AAT protein produced by M-AAT– or Q0_bolton-AAT–expressing HEK-293 cells. Cell culture supernatants were harvested and the cells lysed for protein analysis. Supernatants and cell lysates were run on an SDS-PAGE gel (12% [w/v]) and Western blotted for AAT. Densitometry analysis was performed, and the bar graph represents AAT relative expression. M-AAT is secreted to a greater extent than Q0_bolton-AAT (p = 0.0001, Student t test; n = 3). (D) M-AAT and Q0_bolton-AAT protein was subjected to two freeze-thaw cycles (cycle 1 = F/T 1, cycle 2 = F/T 2) and detected by Western blot analysis (n = 3 technical repeats).

Given that endogenous levels of mRNA are reduced in the Q0_bolton-AAT–expressing HEK-293 cells, we next assessed the ability of readthrough compounds to augment the levels of protein. We first investigated the effect of two readthrough compounds, using two previously described drug concentrations (43). Neither gentamicin (0.5 and 1 μg/ml) nor ataluren (30 and 62.5 μg/ml) had a significant impact on the viability of HEK-293 cells expressing M-AAT (Fig. 6A). On examination of mRNA levels in response to readthrough compounds, no significant change in mRNA levels in either the M-AAT or Q0_bolton-AAT expressing cells was observed in response to treatment with gentamicin (0.5 mg/ml) (Fig. 6B). In contrast, however, treatment of Q0_bolton-AAT expressing cells with ataluren (62.5 μg/ml) resulted in a modest but significant increase in mRNA levels (p = 0.03) (Fig. 6B). Moreover, by Western blot analyses, it was observed that ataluren treatment induced a significant 1.5-fold increase in AAT protein secretion by both M-AAT (p = 0.03) and Q0_bolton-AAT treated cells (p = 0.04) (Fig. 6C). As Western blot analysis is only semiquantitative, we performed an ELISA for secreted AAT in response to ataluren. For this experiment, Q0_bolton-AAT or M-AAT cells were treated with ataluren (62.5 μg/ml) for 48 h. After this time, secreted AAT levels were measured by ELISA. At baseline, Q0_bolton-AAT untreated cells secreted 2.8-fold less AAT (p < 0.001) compared with M-AAT cells. Both cell lines treated with ataluren (62.5 μg/ml) secreted significantly higher levels of AAT compared with respective untreated cells (p < 0.001); however, 1.85-fold less Q0_bolton-AAT was detected compared with M-AAT (p < 0.001) (Fig. 6D).

FIGURE 6.

Increased M-AAT and Q0_bolton-AAT production in response to ataluren. (A) Viability of HEK-293 cells expressing M-AAT exposed to either gentamicin (GT, 0.5 or 1 mg/ml) or ataluren (Atal, 30 or 62.5 μg/ml) for 48 h. Results revealed no significant impact on cell viability (n = 3, Student t test). (B) AAT gene expression in HEK-293 cells expressing M-AAT or Q0_bolton-AAT. HEK-293 cells overexpressing M-AAT or Q0_bolton-AAT were exposed to GT (0.5 mg/ml), Atal (62.5 μg/ml), or vehicle control (Con) for 24 h. M-AAT and Q0_bolton-AAT mRNA expression is significantly increased in the presence of Atal compared with Con cells. (C) AAT protein expression in response to readthrough compounds. Culture supernatants of HEK-293 exposed to GT (0.5 mg/ml) or Atal (62.5 μg/ml) for 48 h were Western blotted for secreted AAT. Protein levels were normalized to that of respective Con. An increase in AAT protein secretion post-Atal treatment was observed for HEK-293 cells expressing M-AAT (p = 0.03) and Q0_bolton-AAT (p = 0.04, Student t test, three independent experiments). (D) Q0_bolton-AAT or M-AAT cells were treated with Atal (62.5 μg/ml) for 48 h. Secreted AAT levels were measured by ELISA. An increase in AAT protein secretion post-Atal treatment was observed for M-AAT cells and Q0_bolton-AAT cells (p < 0.001, n = 4 independent experiments, ANOVA).

An ataluren dose response was next studied, with a significant increase in Q0_bolton-AAT secretion apparent at low concentrations of ataluren employed at 0.1 and 0.5 μg/ml (p = 0.001 and p = 0.002, respectively) and up to 62.5 μg/ml (p = 0.04) (Fig. 7A). We further investigated by Western blot analysis whether ataluren treatment of Q0_bolton-AAT–expressing HEK-293 cells led to translation of the full-length AAT protein. By Western blot analyses, it was confirmed that ataluren employed at a concentration of 62.5 μg/ml increased levels of secreted Q0_bolton-AAT protein compared with 12.5 μg/ml treatment, although the reduced molecular mass of Q0_bolton-AAT (49 kDa) compared with M-AAT (52 kDa) was still apparent (Fig. 7B).

FIGURE 7.

Stabilization of mRNA leads to increased Q0_bolton-AAT truncated protein (A) AAT protein production by Q0_bolton-expressing cells in response to increasing concentrations of ataluren (Atal). Atal employed at 0.1, 0.5, 2.5, 12.5, and 62.5 μg/ml induced a significant increase in levels of secreted AAT protein compared with untreated cells (control [Con]) (Student t test, three independent experiments). (B) AAT protein size in response to readthrough compounds. Electrophoretic mobility of AAT secreted by Q0_bolton-expressing HEK-293 cells in response to Atal (12.5 and 62.5 μg/ml) was reduced (49 kDa) compared with AAT produced by HEK-293 cells expressing M-AAT (42 kDa), indicating that Atal caused increased expression but not an increase in size. Representative image of three separate experiments. In (A) and (B), postnuclear supernatants were prepared of untreated and treated cells, and Western blots for actin demonstrated equal protein loading, supporting the use of equal cell numbers per reaction. (C) Effect of Atal (62.5 μg/ml) on Q0_bolton-AAT mRNA stability over 3 h. HEK cells were cultured in the presence or absence of Atal for 30 min and actinomycin D (10 μg/ml) for 30 min. mRNA levels were normalized to RPLP0 mRNA levels and expressed as a percentage of AAT/RPLP0 mRNA levels at time 0. Pretreatment with Atal resulted in an increased t_1/2 (1.3 h) of Q0_bolton-AAT mRNA compared with mRNA from untreated cells (40 min) (p = 0.007, n = 3, nonlinear-fit one-phase exponential decay curve).

One explanation for the increased protein expression in response to ataluren could be through stabilization of Q0_bolton-AAT mRNA levels. To analyze mRNA stability, we blocked cellular transcription with the inhibitor actinomycin D, which interferes with transcription (Fig. 7C). HEK-293 cells expressing Q0_bolton-AAT were treated with ataluren (62.5 μg/ml) for 30 min, followed by actinomycin D (10 μg/ml) for 30 min, and the amount of Q0_bolton-AAT mRNA remaining at 1 h of treatment was calculated. The t_1/2 of Q0_bolton-AAT mRNA in cells that had not received ataluren treatment was determined to be 40 min. This was significantly increased to 1.3 h in the presence of ataluren, accounting for a 1.95-fold increase in message stability (p = 0.007) (Fig. 7C). This result may account for the heightened expression of AAT following ataluren treatment, amplified translation of Q0_bolton-AAT, and increased protein secretion. Collectively, these data demonstrate increased AAT protein production in the presence of ataluren, and this is particularly useful in this disorder as the truncated protein encoded by Q0_bolton mRNAs retains anti-inflammatory function.

Discussion

The first AATD case study due to a null mutation was described in a 24-y-old patient with undetectable serum levels of AAT (53). Currently, at least 32 AATD null mutations have been reported (54). Historically, it has been thought that no AAT protein arises from null mutations, but as part of this study, we purified truncated AAT protein because of the Q0_bolton null mutation from human plasma and evaluated its capacity as an anti-inflammatory mediator. Furthermore, we used transduced HEK-293 cells expressing Q0_bolton-AAT to study the efficacy of readthrough compounds on successfully augmenting secreted protein levels.

The discovery of the truncated Q0_bolton-AAT protein of 49 kDa is expected due to the position of the PTC on the mRNA sequence. However, why we detected Q0_bolton-AAT protein within the circulation, whereas studies of other null variants have not, is of interest. One possible explanation for this may be the unfolded protein response, a response observed associated with the null variant Q0_hongkong, which leads to degradation of the protein before it can enter the circulation (55). However, the fact that we identified circulating truncated Q0_bolton-AAT may be due to the different locations of the PTC; Q0_hongkong is located 39 aa upstream of the Q0_bolton PTC, before the RCL, which is in contrast to the Q0_bolton mutation transcripts, which retain the active site of the molecule. Consequently, in contrast to Q0_hongkong, which is characterized by a complete absence of secreted protein, the concentration of Q0_bolton-AAT protein purified was ∼1% of the healthy MM control level. From confocal microscopy and Western blot analyses of HEK-293 cells, it would appear that Q0_bolton-AAT is not retained intracellularly and that the Q0_bolton-AAT protein that we have purified from plasma represents protein that was not targeted for destruction. It is noteworthy to mention that research developments in this area suggest that NMD can further contribute to disease by preventing the translation of truncated proteins that may retain function (56).

Proteomic and glycomic analyses were performed to characterize the plasma-purified Q0_bolton-AAT protein. Using 2D-PAGE, a significant anodal shift of the Q0_bolton-AAT protein compared with the native M-AAT form was observed. This is in contrast to the cathodal shift that occurs as a result of the point mutation (Glu342Lys) in the SERPINA1 gene that results in the ZZ-AAT protein (38). The results of this study also indicate that the glycosylation of Q0_bolton-AAT protein is altered, comprising a trend toward increased branching of glycans with a decrease of glycans containing biantennary branching and a concomitant increase in those containing tri- and tetra-antennary branching. A similar degree of altered branching has been reported for the acute phase protein α-1 acid glycoprotein in asthmatic patients and this correlates with FEV₁ and eosinophil numbers (57). Results of the current study also demonstrate increased fucosylation of Q0_bolton-AAT protein, a phenomenon previously shown for AAT in patients with rheumatoid arthritis and chronic joint inflammation (58). Of particular interest, the low level of plasma Q0_bolton-AAT detected may indicate that the mutant protein is more susceptible to clearance (or alternatively, degradation) compared with M-AAT. Indeed, glycans on AAT are important not only for conformational stability of AAT by decreasing the energy level of the native state protein but also for the plasma t_1/2. The plasma t_1/2 of M-AAT is 4–5 d, which is in contrast to nonglycosylated AAT that is rapidly eliminated from plasma (59). Furthermore, AAT can be removed from the circulation via galactose residues that, when exposed, trigger removal of AAT through endocytosis-mediated uptake by asialoglycoprotein receptors on hepatocytes (60, 61). Although the results of this study indicate that the glycosylation of Q0_bolton-AAT protein is altered compared with M-AAT, the recorded changes are most likely not the main cause of the low levels of plasma Q0_bolton-AAT, but may possibly be due to the observed reduced protein stability.

Crucially, AAT glycans are important for controlling IL-8 (2). Indeed, a recent study demonstrated the enhanced anti-inflammatory quality of sialylated AAT involving superior inhibitory influence on neutrophil IL-8–induced chemotaxis. During the resolving phase of infection, there was a significant increase in circulating levels of sialylated AAT/chemokine complexes, in keeping with a pivotal role of AAT in modulation of cell activity, facilitating resolution of inflammation (2). In contrast, glycosylation does not affect binding of AAT to LTB₄, but instead, the lipid locates to a hydrophobic pocket along the protein surface against the s6A and s5A strands (8). AAT-LTB₄ complex formation modulates BLT1 engagement and neutrophil adherence (8). Results of the current study revealed that Q0_bolton-AAT bound IL-8 and LTB₄ to the same extent as M-AAT, confirming the potential immune-regulatory ability of this truncated protein. Glycosylation is also unrelated to the antielastase capacity of the protein, as demonstrated by the retained antiprotease function in a recombinant, aglycosylated AAT molecule (62). In the current study, the Q0_bolton-AAT protein retained anti-NE capacity at a lower level than M-AAT but nevertheless may provide antiprotease protection if present in sufficient levels. Epidemiological studies have revealed that although normal circulating levels of M-AAT are ∼27.5 μM, a concentration of ∼11 μM may be sufficient to protect the lung from an increased risk of emphysema (63). As such, if adequate amounts of Q0_bolton-AAT were released from hepatocytes and other AAT-producing cells, the resulting protein in its truncated form may provide a layer of protection against neutrophil-mediated damage by, first, preventing neutrophil migratory responses to LTB₄ and IL-8 and, second, as an antielastase screen to protect the lungs. Supportive studies in line with this theory include the W1282× CFTR PTC, a C-terminal CFTR mutation that exhibits partial channel activity and is moderately affected via ivacaftor therapy (64).

We next assessed the ability of readthrough compounds to augment the levels of AAT protein, by treating HEK-293 M-AAT– and Q0_bolton-AAT–expressing cells with gentamicin and ataluren. Gentamicin, as the progenitor drug in this category, has been shown to cause suppression of PTC in a number of in vitro cell models (32). Ataluren has the ability to induce the readthrough of stop codons, with readthrough highest in the genetic code UGA, followed by UAG and then UAA. The mutation to the SERPINA1 gene resulting in the Q0_bolton phenotype causes UAA in the coding region, suggesting that ataluren may potentially trigger readthrough and production of full-length protein. Results of the current study demonstrate that AAT protein levels were boosted in HEK-293 M-AAT– and Q0_bolton-AAT–expressing cells treated with ataluren, with a dose between 0.1 and 62.5 μg/ml causing significant increase in protein levels. The highest concentration of ataluren employed minimally affected cell viability. In vivo, two separate studies have explored the effect of ataluren in patients with DMD. The first recruited patients who received ataluren orally three times daily (10–40 mg/kg) (65), and the second involved a multicenter, randomized, double-blind, placebo-controlled, phase-3 trial in patients (40 mg/kg per day) (26). In both cases, the dose of ataluren was generally well tolerated and within the range used in the current study. Moreover, our data demonstrate variations in the ataluren effect by dose, and in line with this observation, a bell-shaped dose response with the use of ataluren has been documented. In this regard, by use of cultured myotubes from a murine model of DMD, and also patients with DMD, a bell-shaped response curve for dystrophin production following ataluren treatment was observed (25, 66). In addition to these findings, a bell-shaped dose response relationship to ataluren was apparent in a zebra fish DMD model (67). It was clear from our data that ataluren did not cause readthrough to full-length protein but augmented levels of both M-AAT and truncated Q0_bolton-AAT protein, a result confirming data obtained with the use of iPSC-hepatic cells (21). Although the main function performed by readthrough compounds is suppression of PTC, recent data suggest that some of the compounds may additionally act as NMD inhibitors (68). Moreover, it has been proposed that ataluren can affect premature translation termination by altering the competition between regulatory factors and near-cognate tRNAs for binding to the ribosomal A site (69). Results of the current study indicate an alternative explanation for the increased protein expression in response to ataluren, involving stabilization of AAT mRNA levels, a documented effect of agents that induce readthrough of PTC (70).

In conclusion, results of this study demonstrate the possible use of ataluren to induce production and secretion of augmented levels of Q0_bolton-AAT protein in vivo. This approach has the potential to alleviate the phenotypic consequences of AATD by increasing levels of truncated, yet functional, protein. Certainly, the theratypes theme that now features in our understanding of CFTR variants reported in CF (and that guides the type of therapy patients may respond to best) can apply to AATD. By this approach, grouping AATD mutations according to theratype could provide a new definition for targeting AATD. However, an important target that must be considered is the threshold and distribution of restored protein level required to provide lung protection in AATD; future studies testing the ability of ataluren treatment to exceed this threshold could support its use as a viable treatment option in this disease setting.

Disclosures

The authors have no financial conflicts of interest.