Yihan Li

Antibody–siRNA conjugates (ARCs) are emerging as a promising approach for targeted delivery of small interfering RNA (siRNA) to specific cells outside the liver, harnessing the advantages of antibody-based delivery to overcome limitations inherent to siRNA molecules, including rapid degradation, limited tissue penetration, and poor specificity. To address the critical need for robust stability assessment of ARCs in biological matrices, we developed a novel in vitro serum stability assay featuring immunoaffinity purification that purified a tool molecule, ARC1, under neutral conditions from biological matrices to preserve the molecular integrity and prevent artificial degradation. The purified ARC1 was analyzed in three complementary liquid chromatography–high-resolution mass spectrometry (LC-HRMS) methods: native mass spectrometry, protein mass spectrometry, and oligonucleotide mass spectrometry. This assay enabled assessment of each individual ARC component, including the parental antibody, linker, sense strand, and antisense strand. The results demonstrated that the linker remained stable in serum for 4 days, while hydrolysis of the conjugated siRNA occurred in both sense and antisense strands. Various biotransformation products were identified by mapping the results from three LC-HRMS methods to visualize the degradation pathway of ARC1. This work represents the first in vitro serum stability analysis workflow of ARCs that can capture biotransformation of each ARC component using immunoaffinity purification. This method offers a valuable tool to proactively identify potential liabilities in ARC designs and inform ARC candidate selection, siRNA engineering, and strategic planning for bioanalytical, toxicity, and immunogenicity assessment during drug discovery and early development.

Rationale: A common strategy for antibody‐drug conjugate (ADC) quantitation from in vivo study samples involves measurement of total antibody, conjugated ADC, and free payload concentrations using multiple reaction monitoring (MRM) mass spectrometry. This not only provides a limited picture of biotransformation but can also involve lengthy method development. Quantitation of ADCs directly at the intact protein level in native conditions using high‐resolution mass spectrometers presents the advantage of measuring exposure readout as well as monitoring the change in average drug‐to‐antibody ratio (DAR) and in vivo stability of new linker payloads with minimal method development. Furthermore, site‐specific cysteine‐conjugated ADCs often rely on non‐covalent association to retain their quaternary structure, which highlights the unique capabilities of native mass spectrometry (nMS) for intact ADC quantitation. Methods: We developed an intact quantitation workflow involving three stages: automated affinity purification, nMS analysis, and data processing in batch fashion. The sample preparation method was modified to include only volatile ion‐pairing reagents in the buffer systems. A capillary size‐exclusion chromatography (SEC) column was coupled to a quadrupole time‐of‐flight high‐resolution mass spectrometer for high‐throughput nMS analysis. Samples from two mouse pharmacokinetic (PK) studies were analyzed using both intact quantitation workflow and the conventional MRM‐based approach. Results: A linear dynamic range of 5–100 μg/mL was achieved using 20 μL of serum sample volume. The results of mouse in vivo PK measurement using the intact quantitation workflow and the MRM‐based approach were compared, revealing excellent method agreement. Conclusions: We demonstrated the feasibility of utilizing nMS for the quantitation of ADCs at the intact protein level in preclinical PK studies. Our results indicate that this intact quantitation workflow can serve as an alternative generic method for high‐throughput analysis, enabling an in‐depth understanding of ADC stability and safety in vivo.

The direct analysis of intact proteins via MS offers compelling advantages in comparison to alternative methods due to the direct and unambiguous identification and characterization of protein sequences it provides. The inability to efficiently analyze proteins in the “middle mass range,” defined here as proteins from 30 to 80 kDa, in a robust fashion has limited the adoption of these “top‐down” methods. Largely, a result of poor liquid chromatographic performance, the limitations in this mass range may be addressed by alternative separations that replace chromatography. Herein, the short migration times of CZE‐ESI‐MS/MS have been extended to size‐sorted whole proteins in complex mixtures from Pseudomonas aeruginosa PA01. An electrokinetically pumped nanospray interface, a coated capillary, and a stacking method for on‐column sample concentration were developed to achieve high‐loading capacity and separation resolution. We achieved full width at half maximum of 8–16 s for model proteins up to 29 kDa and identified 30 proteins in the mass range of 30–80 kDa from P. aeruginosa PA01 whole cell lysate. These results suggest that CZE‐ESI‐MS/MS is capable of identifying proteins in the middle mass range in top‐down proteomics.

Capillary zone electrophoresis-multiple/single reaction monitoring (CZE-MRM/SRM), which employed an electrokinetically driven sheath-flow electrospray interface, was used for the rapid and highly sensitive detection of protein analytes in complex tryptic digests. MRM channels were developed against a commercial exponential mixture of bovine proteins. Five proteins spanning four orders of magnitude concentration range were confidently detected from only 2.5 ng of the digest mixture; the mass detection limits (S/N = 3) of two detected proteins, alpha-casein and glutamate dehydrogenase were about 600 zmol and 30 amol, respectively. This technique was then applied to a RAW 264.7 cell lysate digest. Three proteins were confidently and reproducibly detected from 100 pg of this digest. The sample amount corresponds to the approximate protein content from a single cell, which suggests that CZE-MRM may be a useful analytical tool in chemical cytometry. In addition to providing highly sensitive detection of proteins in complex mixtures, this system is highly rapid; migration time of the protein digests was less than 10 min.

We report the shotgun proteomic analysis of mammalian cell lysates that contain low nanogram amounts of protein. Proteins were denatured using methanol, digested using immobilized trypsin, and analyzed by UPLC-ESI-MS/MS. The approach generated more peptides and higher sequence coverage for a mixture of three standard proteins than the use of free trypsin solution digestion of heat- or urea-denatured proteins. We prepared triplicate RAW 264.7 cell lysates that contained 6, 30, 120, and 300 ng of protein. An average of 2 ± 1, 23 ± 2, 134 ± 11, and 218 ± 26 proteins were detected for each sample size, respectively. The numbers of both protein and peptide IDs scaled linearly with the amount of sample taken for analysis. Our approach also outperformed traditional methods (free trypsin digestion of heat- or urea-denatured proteins) for 6–300 ng RAW 264.7 cell protein analysis in terms of number of peptides and proteins identified. The use of accurate mass and time (AMT) tags resulted in the identification of an additional 16 proteins based on 20 peptides from the 6 ng cell lysate prepared with our approach. When AMT analysis was performed for the 6 ng cell lysate prepared with traditional methods, no reasonable peptide signal could be obtained. In all cases, roughly ∼30% of the digested sample was taken for analysis, corresponding to the analysis of a 2 ng aliquot of homogenate from the 6 ng cell lysate.

We report the performance of capillary zone electrophoresis coupled with an electrokinetically pumped electrospray interface and an Orbitrap‐Velos mass spectrometer for high sensitivity protein analysis. We first investigated the system for quantitation of the tryptic digest of BSA. The system produced outstanding linearity with respect to peak height, number of peptide IDs, and spectral counts across the range of 12 nM to 750 nM (60 amol to 3.5 fmol) of BSA injected. One peptide produced a detection limit of 0.3 nM (1.5 amol) injected. We also analyzed 700 pg of a tryptic digest prepared from a RAW264.7 cell lysate; ten proteins were identified in triplicate analyses after filtering the data with peptide confidence value as high. This sample size corresponds to the protein content of approximately ten eukaryotic cells.

We demonstrate the use of capillary zone electrophoresis with an electrokinetic sheath-flow electrospray interface coupled to a triple-quadrupole mass spectrometer for the accurate and precise quantification of Leu-enkephalin in a complex mixture using multiple-reaction monitoring (MRM). Assay time is <6 min, with no re-equilibration required between runs. A standard curve of Leu-enkephalin was performed in the presence of a background tryptic digest of bovine albumin. We demonstrate reasonably reproducible peak heights (21% relative standard deviation), retention times (better than 1% relative standard deviation), and robust electrospray quality. Our limit of detection (3σ) was 60 pM, which corresponds to the injection of 335 zmol of peptide. This is a 10–20-fold improvement in mass sensitivity than we have obtained by nano HPLC/MRM and substantially better than reported for LC/MS/MS. Further quantification was performed in the presence of stable-isotope-labeled versions of the peptides; under these conditions, linearity was observed across nearly 4 orders of magnitude. The concentration detection limit was 240 pM for the stable-isotope-labeled quantification.

We demonstrate the use of capillary zone electrophoresis with an electrokinetically pumped sheath-flow electrospray interface for the analysis of a tryptic digest of a sample of intermediate protein complexity, the secreted protein fraction of Mycobacterium marinum. For electrophoretic analysis, 11 fractions were generated from the sample using reverse-phase liquid chromatography; each fraction was analyzed by CZE-ESI-MS/MS, and 334 peptides corresponding to 140 proteins were identified in 165 min of mass spectrometer time at 95% confidence (FDR < 0.15%). In comparison, 388 peptides corresponding to 134 proteins were identified in 180 min of mass spectrometer time by triplicate UPLC-ESI-MS/MS analyses, each using 250 ng of the unfractionated peptide mixture, at 95% confidence (FDR < 0.15%). Overall, 62% of peptides identified in CZE-ESI-MS/MS and 67% in UPLC-ESI-MS/MS were unique. CZE-ESI-MS/MS favored basic and hydrophilic peptides with low molecular masses. Combining the two data sets increased the number of unique peptides by 53%. Our approach identified more than twice as many proteins as the previous record for capillary electrophoresis proteome analysis. CE-ESI-MS/MS is a useful tool for the analysis of proteome samples of intermediate complexity.