For successful vector-based gene therapy manufacturing, the selected adeno-associated virus (AAV) vector production system must produce vector at sufficient scale. However, concerns have arisen regarding the quality of vector produced using different systems. In this study, we compared AAV serotypes 1, 8, and 9 produced by two different systems (Sf9/baculovirus and HEK293/transfection) and purified by two separate processes. We evaluated capsid properties including protein composition, post-translational modification, particle content profiles, and in vitro and in vivo vector potency. Vectors produced in the Sf9/baculovirus system displayed reduced incorporation of viral protein 1 and 2 into the capsid, increased capsid protein deamidation, increased empty and partially packaged particles in vector preparations, and an overall reduced potency. The differences observed were largely independent of the harvest method and purification process. These findings illustrate the need for careful consideration when choosing an AAV vector production system for clinical production.

Studies were performed as previously described. Briefly, ammonium bicarbonate, DTT, and iodoacetamide (IAM) were purchased from Sigma Aldrich, and acetonitrile, formic acid, trifluoroacetic acid (TFA), 8 M guanidine hydrochloride (GndHCl), and trypsin from ThermoFisher Scientific. Stock solutions of 1 M DTT and 1.0 M IAM were prepared. Capsid proteins were denatured and reduced at 90°C for 10 min in the presence of 10 mM DTT and 2 M GndHCl. Samples were allowed to cool to room temperature and then alkylated with 30 mM IAM at room temperature for 30 min in the dark. The alkylation reaction was quenched with the addition of 1 mL of DTT. 20 mM ammonium bicarbonate (pH 7.5–8) was added to the denatured protein solution at a volume that diluted the final GndHCl concentration to 200 mM. Trypsin solution was added for a 1:20 trypsin-to-protein ratio and incubated at 37°C overnight. After digestion, TFA was added to a final concentration of 0.5% to quench the digestion reaction.

Online chromatography was performed with an Acclaim PepMap column (15 cm long, 300 μm inner diameter) and a Thermo UltiMate 3000 RSLC system (ThermoFisher Scientific) coupled to a Q Exactive HF with a NanoFlex source (ThermoFisher Scientific). During online analysis, the column temperature was maintained at a temperature of 35°C. Peptides were separated with a gradient of mobile phase A (MilliQ water with 0.1% formic acid) and mobile phase B (acetonitrile with 0.1% formic acid). The gradient was run from 4% B to 6% B over 15 min, to 10% B for 25 min (40 min total), and then to 30% B for 46 min (86 min total).  Samples were loaded directly to the column. The column size was 75 cm × 15 µm identifier (I.D.) and was packed with C18 media (Acclaim PepMap, Thermofisher Scientific). Due to the loading, lead-in, and washing steps, the total time for each liquid chromatography tandem-mass spectrometry run was ~ 2 hr.

Mass spectrometry data was acquired using a data-dependent top-20 method on a Q Exactive HF mass spectrometer, dynamically choosing the most abundant not-yet-sequenced precursor ions from the survey scans (200–2,000 m/z). Sequencing was performed via higher-energy collisional dissociation fragmentation with a target value of 1e5 ions determined with predictive automatic gain control; isolation of precursors was performed with a window of 4 m/z. Survey scans were acquired at a resolution of 120,000 at 200 m/z.  The resolution was set for higher-energy collisional dissociation (HCD) spectra to 30,000 at 200 m/z with a maximum ion injection time of 50 ms and a normalized collision energy of 30. The S-lens radio frequency (RF) level was set to 50, which gave optimal transmission of the m/z region occupied by the peptides from our digest. Precursor ions were excluded with single, unassigned, or six and higher charge states from fragmentation selection. BioPharma Finder 1.0 software (ThermoFisher Scientific) was used to analyze all data acquired. For peptide mapping, searches were performed using a single-entry protein Federal Acquisition Streamlining Act (FASTA) database with carbamidomethylation set as a fixed modification while oxidation, deamidation, and phosphorylation were set as variable modifications. A 10-ppm mass accuracy, a high protease specificity, and a confidence level of 0.8 for tandem-mass spectrometry spectra were used.

Mass spectrometric identification of deamidated peptides was relatively straightforward; deamidation adds to the mass of intact molecule +0.984 Da (the mass difference between -OH and -NH₂ groups). The percent deamidation of a particular peptide was determined by dividing the mass area of the deamidated peptide by the sum of the area of the deamidated and native peptides. Considering the number of possible deamidation sites, isobaric species that are deamidated at different sites may co-migrate at a single peak. Consequently, fragment ions originating from peptides with multiple potential deamidation sites can be used to locate or differentiate multiple sites of deamidation. In these cases, the relative intensities within the observed isotope patterns can be used to specifically determine the relative abundance of the different deamidated peptide isomers. This method assumes that the fragmentation efficiency for all isomeric species was the same and independent of the site of deamidation. This approach allows for the definition of specific sites involved in deamidation and the potential combinations involved in deamidation.

These are .raw files from Thermo Orbitrap.