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. 2024 Aug 30;35(10):2490–2498. doi: 10.1021/jasms.4c00281

Sequencing of Phosphorodiamidate Morpholino Oligomers by Hydrophilic Interaction Chromatography Coupled to Tandem Mass Spectrometry

Mingming Wang 1,*, Brian O’Day 1, Brian Michaels 1, Jurjus Jurayj 1, Bao Zhong Cai 1, Tao Wei 1,*
PMCID: PMC11450806  PMID: 39213635

Abstract

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Sequencing of phosphorodiamidate morpholino oligomers (PMOs) by hydrophilic interaction chromatography (HILIC) coupled to tandem mass spectrometry (MS/MS) is reported. The MS/MS analysis was performed using a quadrupole/time-of-flight (Q-ToF) mass analyzer and collision induced dissociation (CID) in negative ion mode. To improve MS sensitivity in negative ion mode, HILIC conditions, including the separation column, mobile phases, and MS parameters, were optimized. Using the developed HILIC–CID-MS/MS method, 100% sequence coverage was achieved for PMOs ranging from 18-mer to 25-mer. Additionally, the method was successfully applied to identifying positional isomers of n – 1 deletion impurities present in PMO drug substances.

Keywords: tandem mass spectrometry (MS/MS), collision induced dissociation (CID), hydrophilic interaction chromatography (HILIC), phosphorodiamidate morpholino oligomer (PMO), sequencing, impurity characterization

Introduction

Phosphorodiamidate morpholino oligomers (PMOs) are short single stranded oligonucleotides consisting of morpholine rings connected by uncharged phosphorodiamidate linkages. The unique design of PMOs imparts several useful therapeutic and physical properties including resistance to enzymatic degradation, specificity due to tight binding with complementary strands, low toxicity, and high water solubility.1−4 PMOs have been recognized as an increasingly important class of antisense therapeutics for the treatment of various diseases,5−12 most notably for Duchenne muscular dystrophy (DMD). To date, four PMO drugs including eteplirsen, golodirsen, casimersen (Sarepta Therapeutics Inc.), and viltolarsen (NS Pharma) have been approved by the U.S. Food and Drug Administration for the treatment of DMD.13

Driven by the drug development need and regulatory requirements, critical quality attributes (CQAs) of oligonucleotide therapeutics, such as purity, impurity profile, and sequence, must be characterized to ensure process consistency, product quality, safety, and efficacy. It is challenging to characterize oligonucleotides and their impurities due to their relatively large molecular weights, complexity, and diastereomeric nature (where applicable). Consequently, the coelution of impurities with their parent oligonucleotides makes it unfeasible to achieve complete separation by traditional liquid chromatography (LC) approaches such as anion-exchange chromatography (AEX) and ion-pairing reversed phase (IP-RP) chromatography.14,15 To overcome the chromatographic resolution limitation, mass spectrometry (MS) is coupled to LC as it allows identification of coeluting impurities with specificity.14 Recently, IP-RP LC coupled with mass spectrometry (IP-RP LC/MS) has been established as the method of choice for the analysis of oligonucleotides.16 Advances such as the addition of fluoroalcohols in the mobile phase with IP reagents like alkylamines have led to increased MS sensitivity in negative ion mode.17 However, IP reagents can cause ion suppression in positive ion mode when subsequent mass spectral analysis is performed. Dedicated systems are commonly implemented for IP-RP LC/MS analysis.15,16,18,19 Therefore, hydrophilic interaction chromatography (HILIC) has emerged as an attractive alternative for oligonucleotide analysis as it addresses the concern of ion suppression in IP-RP.16,20,21 When coupled to electrospray ionization MS, HILIC may afford higher MS sensitivity because the utilization of high-organic mobile phases leads to more efficient desolvation and thus improved MS sensitivity.22 So far, HILIC–MS has not been previously explored for the characterization of PMOs.

The utility of LC–MS has also been extended to the sequence confirmation of oligonucleotides. A typical MS-based approach to sequencing oligonucleotides involves backbone cleavages followed by identification of the resulting fragments by MS. Oligonucleotide backbones are typically cleaved enzymatically,23−27 chemically,28−31 or via gas-phase dissociation.32−35 Enzymatic digestion of phosphodiester and phosphorothioate oligonucleotide backbones is well documented,23−26 where exonucleases sequentially cleave individual nucleotides from the 3′ or 5′ terminus of the oligonucleotide. However, this approach does not apply to PMOs as their backbones are resistant to enzymatic degradation.1−3

Therefore, PMO sequencing was achieved by controlled acid hydrolysis followed by analysis of resultant fragments via matrix-assisted laser desorption/ionization (MALDI) ToF MS.36 This method requires time-consuming acid incubation, and MALDI ToF MS suffers from relatively low mass accuracy and resolution. In contrast, gas-phase dissociation such as collision-induced dissociation (CID) requires little sample pretreatment and has thus been widely used for sequencing of oligonucleotides with phosphodiester and phosphorothioate backbones.35,37−40 However, it has not been employed to sequence PMOs. Thus, CID-MS/MS for the sequencing of PMOs was evaluated in this work. Unlike the negatively charged oligonucleotides such as phosphodiesters and phosphorothioates, PMOs have neutral backbones, possibly leading to a distinctive fragmentation pathway (Figure 1). In the course of our investigation, Karasawa et al.41 reported an MS/MS method for sequencing PMOs from 18-mer to 25-mer for the first time, where electron capture dissociation (ECD) in positive ion mode was used to induce backbone cleavage. ECD generated mainly d and z ions (Figure 1) that were used to confirm the sequence of the PMOs.

Figure 1.

Figure 1

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Nomenclature of PMO fragmentation and representative fragments that were detected by CID-MS/MS. Note: d – N(CH3)2 fragments result in a mass difference of 45 amu from the corresponding d ion, whereas z – N(CH3)2 fragments lead to a mass difference of 43 amu from the respective z ion.

In addition to sequence confirmation, MS/MS can be applied to impurity characterization of PMOs, which share similar analytical challenges with other types of oligonucleotides. However, compared to negatively charged oligonucleotides, the distinct nature of neutral PMO backbones adds more challenges in chromatographic separation, as their separation mostly relies on the charges of nucleobases under different pH conditions. As a result, it is particularly challenging to resolve the full-length PMO products (FLP) from their n – 1 deletions, which are defined as impurities structurally related to the parent PMO by deletion of a single nucleotide. Arising from incomplete detritylation or coupling during solid-phase oligomer synthesis (SPOS), n – 1 deletions constitute the majority of PMO impurities and exist as mixtures of components differentiated by the deletion location (termed positional isomers). Considering that n – 1 deletions are usually the predominant PMO impurities, it is necessary to identify the problematic synthetic cycles to facilitate upstream process development and optimization. However, different positional n – 1 isomers have identical molecular weights and closely elute. Consequently, they cannot be distinguished and characterized by LC–MS alone. Therefore, sequencing via MS/MS can aid in the identification of different n – 1 positional isomers by pinpointing the exact deletion location.

Herein, HILIC coupled to CID-MS/MS was evaluated for sequencing PMOs from 18-mer to 25-mer. In this study, CID-MS/MS in both positive and negative ion modes were initially explored with negative ion mode yielding more useful sequence information. To improve the MS sensitivity in negative ion mode, the HILIC conditions and MS parameters were optimized. To the best of our knowledge, this is the first report on sequencing of PMOs in negative ion mode by CID-MS/MS. Additionally, this method was applied to identifying positional isomers of n – 1 deletion impurities present in PMO drug substance (DS).

Experimental Section

Materials and Reagents

PMO-1, PMO-2, and PMO-3 (Table 1) were synthesized via SPOS, purified, and isolated as lyophilized solids by Sarepta Therapeutics. LC/MS grade water, acetonitrile (ACN), acetic acid, and ammonium hydroxide were purchased from Fisher Scientific (Waltham, MA). LC/MS grade ammonium acetate was purchased from Honeywell (Charlotte, NC).

Table 1. List of PMOs.

PMO length (bases) base sequence (5′ → 3′)
PMO-1 18 GCTATTACCTTAACCCAG
PMO-2 22 CAATGCCATCCTGGAGTTCCTG
PMO-3 25 GTTGCCTCCGGTTCTGAAGGTGTTC
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Sample Preparation

Stock solutions of PMOs were prepared in LC/MS grade water at 10 mg/mL and stored at 4 °C. Each PMO was diluted 10-fold by mixing 100 μL of each stock solution with 200 μL of water and 700 μL of ACN prior to LC/MS analysis.

HILIC–MS/MS Method

HILIC separations were performed on a 1290 Infinity II HPLC system (Agilent, USA) using a Shodex HILICpak VN-50 (2.0 × 150 mm, 5 μm) column at 40 °C. Mobile phase A was 1 mM ammonium acetate in water, and mobile phase B was ACN. Gradient elution at 0.2 mL/min started with 70% B (held for 2 min), gradually decreased to 50% B at 5 min, held at 50% B for 2 min, immediately returned to the initial condition (70% B) at 7.1 min, and then held for 3 min at 70% B. The detailed HILIC conditions are listed in Table S1.

A 6545 XT Q-ToF mass spectrometer (Agilent, USA) was used to acquire the MS data. For MS analysis, a full scan (MS mode) was performed in negative ion mode to find the precursor ions. For the MS/MS analysis, targeted MS/MS was conducted to obtain fragment ions. The detailed MS parameters and MS/MS parameters are listed in Table S2. MS and MS/MS data were processed by an Agilent MassHunter Qualitative Analysis 10.0. Bioconfirm 12 software was used for deconvolution. MS/MS spectra were interpreted manually. Fragments obtained from multiple collision energies (CEs) and multiple precursor ions were used to improve the sequence coverage.

Results and Discussion

HILIC–MS Method Development

Achieving sufficient sequence coverage for PMOs requires a highly sensitive LC–MS/MS method. To this end, both HILIC conditions and CID-MS/MS conditions were optimized.

As CID-MS/MS in positive ion mode failed to provide fragments that convey sequence information (vide infra), we explored MS/MS in negative ion mode instead. MS sensitivity for PMOs in negative ion mode was observed to be generally lower than that in positive ion mode. To achieve satisfactory MS sensitivity in negative ion mode, initial investigations were conducted using columns with various stationary phase surface chemistries including diol (Shodex HILICpak VN-50, 2.0 × 150 mm, 5 μm), penta-hydroxyl (BIOshell Glycan, 2.1 × 50 mm, 2.7 μm), zwitterionic (ZIC-HILIC, 2.1 × 50 mm, 3.5 μm, and ZIC-cHILIC, 2.1 × 50 mm, 3 μm), and amide (BEH Amide, 3.0 × 50 mm, 1.7 μm). As shown in Figure 2A,B, a diol column (HILICpak VN-50, 2.0 × 150 mm) afforded the highest MS sensitivity manifested by the extracted ion chromatogram (EIC) peak area and was chosen as the separation column. To improve the MS sensitivity, ammonium acetate as the mobile phase modifier was evaluated at different concentrations ranging from 0.25 to 10 mM. As shown in Figure 2C, the MS sensitivity was suppressed when the concentration of ammonium acetate increased. Additionally, a higher ammonium acetate concentration led to an earlier retention time (Figure S1A). The MS sensitivity was comparably high when 0.25 and 1 mM ammonium acetate was used; however, 1 mM ammonium acetate was selected as the mobile phase modifier due to less sodium adduct formation (Figure S1B). The pH of the mobile phase could affect the ionization efficiency and thus the MS sensitivity. Therefore, the MS sensitivity using 1 mM ammonium acetate with 0.02% (v/v) acetic acid, or 0.02% (v/v) ammonium hydroxide, was evaluated. As shown in Figure 2D, addition of 0.02% ammonium hydroxide greatly suppressed the MS signal, whereas introduction of 0.02% (v/v) acetic acid did not improve the MS sensitivity but did cause peak broadening (Figure S1C). Notably, the pH of the mobile phase affected the charge state distribution, as well as the retention time. Under basic conditions, the −3 charge state and −2 charge state were comparable in abundance, whereas the −3 charge state was predominant under acidic conditions (Figure S1D). Additionally, the retention time shifted earlier under basic conditions (Figure S1C). Collectively, 1 mM ammonium acetate was finally chosen as the mobile phase modifier.

Figure 2.

Figure 2

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Optimization of HILIC conditions: (A) HILIC chromatograms using different columns; (B) MS sensitivity (expressed by the EIC peak area) using different HILIC columns; (C) effect of concentration of ammonium acetate on MS sensitivity; (D) effect of pH of mobile phase on MS sensitivity.

To further improve the MS sensitivity, the MS parameters were optimized. Gas temperatures ranging from 250 to 325 °C were tested. As shown in Figure S2A, the gas temperature had a marginal effect on the MS sensitivity. A gas temperature of 275 °C was chosen. Nozzle voltages ranging from 0 to 2000 V were also optimized. As shown in Figure S2B, the EIC peak area increased when the nozzle voltage increased; 2000 V was thus chosen as the nozzle voltage. Fragmentor voltages from 175 to 325 V were also evaluated. As shown in Figure S2C, the fragmentor voltage of 275 V afforded the highest MS sensitivity.

Sequencing

Our initial attempt at sequencing the PMOs was made using targeted MS/MS in positive ion mode since PMOs have been observed empirically to ionize more efficiently in positive ion mode, possibly due to their neutral backbones. In positive ion mode, multiple charge states (+4 to +7) were observed in PMO-1 (Figure 3A). With the use of the most abundant charge state (+5) as the precursor ion for subsequent CID fragmentation, the predominant fragmentation pathway was base loss (Figure 3C,D). This observation was consistent with previous results.41 Unfortunately, such base loss does not provide adequate sequence information. Changing the CEs did not lead to appreciable backbone fragmentation that would generate fragments useful for sequence confirmation (data not shown). Additionally, different charge states (+4 and +6) as the precursor ions were also subjected to CID fragmentation; however, base loss remained the predominant fragmentation pathway (data not shown). Notably, the preference of base loss seems to follow the order G > C > A ≫ T (Figure 3D), which is likely due to the different gas-phase proton affinities of nucleobases.42 Such charged base loss (i.e., protonated form) occurs to primarily relieve Coulombic repulsion.43

Figure 3.

Figure 3

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Initial attempt to sequence PMO-1 in positive ion mode: (A) MS spectrum; (B) deconvoluted mass spectrum. The relative intensities of n – C (deletion of one morpholino cytidine from FLP) and n – A (deletion of one morpholino adenosine from FLP) are 11 and 4%, respectively. (C) MS/MS spectrum using +5 charge state as the precursor ion; (D) expanded view of (C).

Alternatively, the sequencing of PMOs in negative ion mode was evaluated by utilizing PMO-1. Compared to positive ion mode, PMOs exhibited fewer charge states in negative ion mode. As shown in Figure 4A, the most abundant charge state was −3 for PMO-1. Therefore, the ion at m/z 2069.7261 was selected as the precursor ion. The MS/MS spectra are shown in Figure 4B–G. Despite minor base loss (Figure 4C), the negative ion mode afforded more pronounced backbone fragmentation. Interestingly, d ions were the most abundant species detected among other ions, including b ions, d – N(CH3)2 ions, z ions, and z – N(CH3)2 ions. The proposed fragmentation pathway of PMO and structures of the resultingfragments are presented in Figure 1. This indicates PMO fragments differently from negatively charged oligonucleotides such as phosphodiesters where a-B and w ions are commonly observed (Figure S3).33 The detected fragments of PMO-1 are summarized in Table 2. As shown in Figure 5, 100% sequence coverage of PMO-1 was achieved, thus confirming the sequence of PMO-1. Sequencing of longer PMOs (PMO-2 and PMO-3) in negative ion mode was also assessed. Similarly, d ions were the most abundant fragments observed besides b ions, d – N(CH3)2 ions, z ions, and z – N(CH3)2 ions. As shown in Figure S4 and Tables S3 and S4, 100% sequence coverage was also achieved for PMO-2 and PMO-3.

Figure 4.

Figure 4

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Mass spectra of PMO-1 in negative ion mode: (A) MS spectrum; (B) MS/MS spectrum; (C–G) expanded views of (B). Precursor ion ([M – 3H]3–) at m/z 2069.7261 was selected for targeted MS/MS. Multiple CEs including 60, 63, 66, and 69 V were applied to improve sequence coverage.

Table 2. Theoretical and Detected Fragments of PMO-1a.

position (5′) theoretical m/z (charge) detected m/z mass error (ppm) position (3′) theoretical m/z (charge) detected m/z mass error (ppm)
0 368.1598 (−1) 368.1597 –0.3 18 2919.0061 (−2) 2919.0332 9.3
1 723.2755 (−1) 723.2754 –0.1 17 2741.4482 (−2) 2741.4404 –2.8
2 1038.3852 (−1) 1038.3845 –0.7 16 2583.8934 (−2) 2583.8934 0.0
3 1368.4945 (−1) 1368.4937 –0.6 15 2418.8387 (−2) 2418.8314 –3.0
4 1707.6154 (−1) 1707.6132 –1.3 14 2249.2783 (−2) 2249.2756 –1.2
5 2037.7247 (−1) 2037.7212 –1.7 13 2084.2236 (−2) 2084.2224 –0.6
6 2367.8340 (−1) 2367.8347 0.3 12 1919.1690 (−2) 1919.1685 –0.2
7 2706.9549 (−1) 2706.9488 –2.3 11 1749.6086 (−2) 1749.6109 1.3
8 1510.5284 (−2) 1510.5289 0.4 10 1592.0537 (−2) 1592.0565 1.8
9 1668.0832 (−2) 1668.0826 –0.3 9 2870.0051 (−1) 2869.9984 –2.3
10 1833.1378 (−2) 1833.1382 0.2 8 2539.8958 (−1) 2539.8886 –2.8
11 1998.1925 (−2) 1998.1887 –1.9 7 2209.7864 (−1) 2209.7833 –1.4
12 2167.7529 (−2) 2167.7694 7.6 6 1870.6656 (−1) 1870.6634 –1.2
13 2337.3134 (−2) 2337.3112 –0.9 5 1531.5447 (−1) 1531.5442 –0.3
14 2494.8682 (−2) 2494.8714 1.3 4 1216.4351 (−1) 1216.4350 –0.1
15 2652.4230 (−2) 2652.4136 –3.5 3 901.3254 (−1) 901.3241 –1.4
16 2809.9778 (−2) N/D N/A 2 586.2158 (−1) 586.2088 –11.9
17 1986.0231 (−3) 1986.0288 2.9 1 247.0949 (−1) 247.0951 0.8
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a

d ions are listed for 5′-fragments, and z ions are listed for 3′-fragments. N/D and N/A denote “not detected” and “not applicable”, respectively.

Figure 5.

Figure 5

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Sequence coverage of PMO-1.

Characterization of Impurities

Impurity profile is a CQA required for oligonucleotide therapeutics by regulatory agencies. PMO drug substances are manufactured first by SPOS to yield oligomers on resin, which are subsequently cleaved and deprotected to generate the crude PMO materials. Several impurities could arise during this process. Common PMO impurities generated during SPOS include deletion, addition, backbone modification, and fragmentation. Common impurities generated during cleavage and deprotection include loss of the triethylene glycol tail at the 5′-end and base modifications such as C to U conversion, M + 106, etc. Among these common impurities, deletions such as n – 1 are the predominant impurities observed in PMOs. An n – 1 deletion often exists as a mixture of different positional isomers. In the case of PMO-1, despite the presence of six cytosine subunits (Table 1), there are three positional isomers that have n – C deletions, namely n – C2, n – C8/9, and n – C14/15/16, because deletion of any repeating bases adjacent to each other in the sequence leads to the same deletion sequence. Thus, n – C8/9 and n – C14/15/16 are simplified as n – C8 and n – C14, respectively, using the first deletion site. Due to chromatographic coelution and identical molecular weights, it is not possible to differentiate n – 1 positional isomers by LC–MS alone. However, sequence analysis via MS/MS could potentially address this challenge by identifying the “fingerprint” fragments of each positional isomer. Therefore, HILIC MS/MS in negative ion mode was utilized here to elucidate the deletion site of the n – 1 impurities present in PMO DS.

The initial analysis of PMO-1 DS (Figure 3B, performed in positive ion mode) indicated n – C and n – A are the most abundant n – 1 deletions present, with relative intensities of approximately 11 and 4%, respectively. Therefore, these two n – 1 deletions were selected for subsequent MS/MS sequencing in the negative ion mode for elucidation of the deletion site. The precursor ion of m/z 1964.3541 (n – C) and the precursor ion of m/z 1956.3503 (n – A) were subjected to targeted MS/MS analysis. Of the three types of n – C deletions (e.g., n – C2, n – C8, and n – C14) in PMO-1, the sequencing result (Figure 6A and Table S6) confirmed the C-deletion mainly occurred at 5′-14. Deletions at 5′-8 were also detected but at a lower level than 5′-14 (data not shown). Similarly, of the four possible n – A deletions in PMO-1, consisting of n – A4, n – A7, n – A12, and n – A17, sequencing results confirmed that the A-deletion site occurred primarily at 5′-12 (Figure 6B and Table S5). Deletions at 5′-4 and 5′-7 were also observed but at lower levels (data not shown). In summary, this HILIC–CID-MS/MS method was able to locate the exact deletion site in n – 1 deletion impurities present in PMO DS at an intensity as low as 4% (relative to FLP). Elucidating the deletion site provides opportunities for troubleshooting during process development. For instance, a longer coupling time and/or detritylation time can be implemented at a certain cycle to minimize n – 1 deletion impurities. Furthermore, this method was simple and high-throughput and could achieve PMO impurity identification without the need for offline impurity enrichment, greatly reducing the development timeline and increasing development efficiency.

Figure 6.

Figure 6

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Sequence coverage of n – 1 impurities in PMO-1: (A) n – C; (B) n – A.

Conclusion

To the best of our knowledge, here we report the first sequencing of PMOs using HILIC–CID-MS/MS in negative ion mode. This method afforded 100% sequence coverage for PMOs ranging from 18-mer to 25-mer. Additionally, we successfully applied this method to identifying n – 1 positional isomers present in the PMO drug substance. This workflow was capable of differentiating n – 1 positional isomers at low levels, and thus, no additional offline impurity enrichment was needed. Our approach is simple and high-throughput and can be easily adopted in most laboratories.

Acknowledgments

The authors would like to thank Shashank Jain and Guannan Li from Agilent Technologies for kindly providing software support and discussion to enable data analysis.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/jasms.4c00281.

  • Additional data for HILIC–MS/MS method optimization; sequence confirmation of PMO-2 and PMO-3; theoretical/observed fragmentation patterns of n – A and n – C deletions present in PMO-1 (PDF)

Author Contributions

T.W. and M.W. conceptualized the presented ideas, designed, and planned the experiments. B.Z.C. and T.W. supervised the project. M.W. performed the experiments with help from B.M. and B.O. M.W. collected and analyzed the data and prepared the manuscript with supervision and support from B.Z.C., J.J., and T.W. B.M. verified the analytical methods and results. All authors discussed the results and contributed to the final manuscript.

The authors declare no competing financial interest.

Supplementary Material

js4c00281_si_001.pdf (574.8KB, pdf)

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