Proteoform‐Resolved Interaction Studies of Plasminogen by CZE‐MS and SEC‐MS Under Near‐Native Conditions

TL;DR

This paper introduces a new method using CZE-MS and SEC-MS to study plasminogen proteoforms and their interactions under near-native conditions.

Contribution

The study demonstrates a novel combination of CZE-MS and SEC-MS for proteoform-resolved interaction analysis of plasminogen.

Findings

CZE-MS separates plasminogen proteoforms differing in phosphorylation and sialylation under near-native conditions.

The nanoCEasy interface allows ionization under native or denaturing conditions for detailed proteoform analysis.

SEC-MS detects new plasminogen proteoforms not observed in direct infusion experiments.

Abstract

Native mass spectrometry has become an important technique for studying proteins and protein complexes under physiologically similar conditions. However, the technique is still rarely coupled to separation techniques, as the widely used reversed‐phase chromatography mostly denatures proteins. Here we present a study combining both capillary zone electrophoresis and size exclusion chromatography, each coupled online to native electrospray ionization mass spectrometry for the analysis of proteoforms of plasminogen. Plasminogen, the zymogen form of plasmin, is an abundant plasma protein with multiple proteoforms whose activation via proteolytic cleavage is tightly regulated within the fibrinolytic system and is partly modulated by post‐translational modifications. Near‐native CZE conditions enable the separation of proteoforms differing in phosphorylation and glycosylation, that is, almost baseline separation of proteoforms differing in phosphorylation and sialylation. The nanoflow sheath liquid interface (nanoCEasy) enables efficient ionization as well as the flexibility to ionize under native or denaturing conditions. The CZE‐MS set‐up can be used to study proteoform‐resolved interaction directly in the capillary, as demonstrated for the activation of plasminogen by the co‐injection of tissue plasminogen activator (tPA). In these proof‐of‐concept experiments, various truncations of plasminogen were observed with a preferred cleavage for the N‐glycosylated and the phosphorylated proteoforms. SEC‐MS enables partial separation of the proteoforms, particularly between glycosylated and nonglycosylated proteoforms leading to the detection of new proteoforms not described by direct infusion experiments. Overall, the combination of near‐native CZE‐MS and SEC‐MS provides a detailed characterization of plasminogen and enables proteoforms‐resolved interaction studies.

Figures

Deconvoluted mass spectra of plasminogen acquired by near‐native CZE‐MS. Each deconvoluted mass spectrum on the left was averaged over a 1‐min interval of the CZE separation and is displayed from top to bottom, corresponding to 29.0–54.0 min. The Total Ion Electropherogram (TIE) of the near‐native CZE separation is shown on the right. Separation is performed in a BGE of 50 mM AmAc +20 mM HAc (pH = 5.0). See text for more details on the method.

Near‐native CZE‐MS EIEs of different plasminogen proteoforms. The following four major proteoforms can be identified: (i) type I—nonphosphorylated (T1‐A, T1‐C, T1‐E, and T1‐G), (ii) type II—nonphosphorylated (T2‐A, T2‐C, T2‐E, and T2‐G), (iii) type I—phosphorylated (T1‐B, T1‐D, T1‐F, and T1‐H), and (iv) type II—phosphorylated (T2‐B, T2‐D, T2‐F, and T2‐H). The top 4 charge states from each of the major forms have been used to create the EIEs. See text for details on the method.

Deconvoluted mass spectra of plasminogen acquired by near‐native SEC‐MS. Each deconvoluted mass spectrum on the left was averaged over a 15‐s interval of the SEC timescale and is displayed from top to bottom, corresponding to 12.75–17.00 min. The total ion chromatogram (TIC) of the near‐native SEC separation is shown on the right.

Near‐native SEC‐MS EICs of different plasminogen proteoforms. The following four major proteoforms can be identified: (i) type I—nonphosphorylated (T1‐A, T1‐C, T1‐E, and T1‐G), (ii) type II—nonphosphorylated (T2‐A, T2‐C, T2‐E, and T2‐G), (iii) type I—phosphorylated (T1‐B, T1‐D, T1‐F, and T1‐H) and (iv) type II—phosphorylated (T2‐B, T2‐D, T2‐F, and T2‐H). The top 4 charge states from each of the major forms have been used to create the EICs shown in this figure. See text for details on the method.

In‐capillary activation of plasminogen by tPA (1:10 tPA:plasminogen) in CZE‐MS. The concept of in‐capillary CZE‐MS for the activation study of plasminogen by tPA is shown in Figure 5A. Deconvoluted mass spectra of a CZE‐MS analysis of plasminogen with the co‐injection of tPA (1.1% of capillary volume of 0.25 μg/μL tPA solution) (Figure 5B). Each deconvoluted mass spectrum on the left was averaged over a 1‐min interval of the CZE separation and is displayed from top to bottom, corresponding to 30.0–50.0 min. The Total Ion Electropherogram (TIE) of the near‐native CZE separation is shown on the right. CZE‐MS conditions are the same as in Figure 1.

Proteoform distribution of plasminogen and truncated plasminogen after co‐injection of tPA in CZE‐MS experiments. All data are normalized to the proteoform type II, +H1N1S1+P (most abundant proteoforms of intact plasminogen). Blue bars represent the proteoforms distribution of the intact plasminogen, orange bars indicate the results of the co‐injection of tPA (1.1% of capillary volume of 0.25 μg/μL tPA solution); full orange: remaining plasminogen, striped: truncated forms with the loss of AA1‐68.

Tables

TABLE 1. Summary of identified plasminogen proteoforms by different techniques.

Full text

Introduction

1

Native mass spectrometry has become an important technique for studying proteins and protein complexes under near‐physiological conditions [1, 2, 3, 4, 5]. It enables the study of the biological functions of proteins, protein complexes, and proteoforms, which are highly associated with their conformation. In particular, interaction with drugs or drug candidates can be studied [3, 6]. Electrospray ionization has been shown to maintain the functional conformation of proteins and proteoforms when performed from (volatile) salt‐rich and pH‐buffering conditions and avoiding high contents of denaturing solvents [7]. For direct infusion ESI‐MS, samples typically need to be desalted and buffer exchanged to volatile salts such as ammonium acetate [8]. This can be omitted when chromatographic or electrophoretic separation techniques are applied prior to MS. More importantly, the separation of proteins and proteoforms allows for reduced ion suppression and, thus, better sensitivity and improved quantitation.

In contrast to the commonly used reversed phase chromatography, hydrophobic interaction chromatography, ion‐exchange chromatography, and size exclusion chromatography can be used to separate proteins under near‐native conditions [9, 10, 11, 12]. Especially SEC‐MS is relatively straightforward as no volatile salts are required. Thus, SEC‐MS has been used to study monoclonal antibodies under native conditions by the group of Guillarme [13, 14, 15]. SEC is used often for online buffer exchange but can also add another separation dimension as shown for various biological systems [16]. Native SEC‐MS has also been shown to be useful to study protein aggregation [17].

Capillary zone electrophoresis‐mass spectrometry is a powerful technique to characterize the proteoforms of complex proteins [18], such as glycosylated, phosphorylated, or acetylated proteins [19]. Moreover, CZE‐MS can be performed under near‐native conditions [20, 21, 22] enabling the separation of proteins of a proteome [23] or proteoforms such as charge variants of monoclonal antibodies [24, 25, 26, 27]. Nevertheless, there are still only a few studies applying native CZE‐MS for proteoform separation [28], protein complexes [29], or affinity studies [30, 31]. Capillary coatings are required for CZE analyses of proteins, as most proteins interact with the capillary wall, hampering the performance of the separation. For native CZE‐MS, static coatings are preferred not only to reduce interactions of the proteins but also to suppress the electroosmotic flow (EOF) to improve peak resolution. This EOF suppression is particularly important when a BGE at near neutral pH is applied. A hydroxypropyl methylcellulose (HPMC) capillary coating is relatively easy to produce and has been recently shown to be very effective for CZE‐MS of charge variants at near‐neutral pH [24]. The interfacing of CZE to mass spectrometry is of great importance in this context, as not all interfaces are equally effective for the study of proteins under near‐native conditions [21, 32]. NanoESI interfaces are especially useful because of increased sensitivity and reduced clustering.

Plasminogen is a 90 kDa glycoprotein that circulates in blood at approximately 180 mg/mL (around 2 μM). It serves as a central protease in the fibrinolytic system, playing a pivotal role in blood clot dissolution, extracellular matrix remodeling, and other physiological processes. Upon activation to plasmin, it degrades fibrin and other extracellular matrix components, contributing not only to hemostasis but also to wound healing, inflammation, and cancer metastasis [33]. Although plasminogen has long been studied as a single functional entity, plasminogen is now recognized as a diverse family of proteoforms. These distinct molecular forms arise from genetic variation, alternative splicing, and posttranslational modifications (PTMs), including N‐glycosylation, O‐glycosylation, and phosphorylation [34, 35]. By using the near‐native MS technique, Heck and coworkers identified more than 14 distinct proteoforms in plasma‐derived plasminogen [36].

Despite these findings, the biological significance of plasminogen proteoforms remains incompletely understood. Emerging evidence suggests that specific variants may differ in activation kinetics, binding affinities, or tissue distribution. For example, variations in N‐glycosylation influence plasminogen structure through the position of KR3 [37] and affect its binding and protease functions. Plasminogen type I (N‐glycosylated) exhibits approximately tenfold lower affinity for cells compared to type II [38], yet type I demonstrates greater protease activity [39]. In contrast, the structure–function relationships of O‐glycosylation and phosphorylation remain poorly characterized.

In this study, we employ near‐native CZE‐MS and SEC‐MS to achieve an in‐depth characterization of the proteoform landscape of human plasma‐derived plasminogen. By coupling MS with native‐condition separation, we expand the known proteoform profile beyond what near‐native MS alone can provide. Additional proteoforms were confirmed through peptide PTM mapping studies. Furthermore, we evaluated in a proof‐of‐concept study the proteoform‐resolved activation of plasminogen by co‐injecting tissue‐type plasminogen activator (tPA), one of the activators, in the native CZE‐MS approach. This work underscores the complementarity of CZE‐MS and SEC‐MS and lays the foundation for future investigations into protease‐substrate interactions in complex biological systems.

Materials and Methods

2

Chemicals and Reagents

2.1

Plasma‐derived human Glu‐plasminogen was purchased from Prolytix (Vermont, USA). Tissue plasminogen activator (tPA) was purchased from Merck (Darmstadt, Germany). Ammonium acetate (AmAc, 5.0 M in H_2_O, Molecular Biology grade), potassium phosphate monobasic (KH_2_PO_4_, 1.0 M in H_2_O, Reagent grade), urea (8.0 M in H_2_O, BioUltra grade), dithiotreitol (DTT, 1 M in H_2_O, Reagent grade), iodoacetamide (IAA, ≥ 99%), acetonitrile (≥ 99.9% gradient grade), isopropanol (IPA, ≥ 99.9% gradient grade), acetic acid (HAc, glacial, 100%), and trifluoroacetic acid (TFA) (≥ 99.0% LC–MS grade) were purchased from Merck (Darmstadt, Germany). Tris–HCl buffer (1.0 M solution, UltraPure grade) and formic acid (99+%, LC–MS grade) were purchased from Thermo Fischer Scientific (Massachusetts, USA). Trypsin Gold (Mass Spectrometry Grade) and Glu‐C (Sequencing Grade) were purchased from Promega (Wisconsin, USA). Titanium dioxide (TiO_2_)‐beads were purchased from GL Sciences (Tokyo, Japan). All chemicals and reagents were used as received.

Near‐Native CZE‐MS

2.2

CZE‐MS analysis was performed applying a 7100 CE System (Agilent Technologies GmbH, Waldbronn, Germany) coupled to either a Q‐Exactive Plus MS (Thermo Fisher Scientific, Bremen, Germany) or an Orbitrap Exploris 480 MS (Thermo Fisher Scientific, Bremen, Germany). A 65‐cm long HPMC coated capillary was used for all CZE‐MS experiments. HPMC coating was performed as described previously [24]. The capillaries were etched down to an outer diameter of < 100 μm (corresponding to a wall thickness of about 10–25 μm using a microscope for optical control) at a length of about 10–15 mm at one end. The capillary end, where the polyimide was removed was closed with standard hot glue and etched with HF (40% (v/v)) for 1 h at room temperature. After capillary etching, the glued end of the capillary was cut and the capillary shortened to a length of 65 cm at the not‐etched end. The BGE consisted of 50 mM AmAc and 20 mM HAc in ultrapure water. Plasminogen samples were injected hydrodynamically using 50 mbar pressure for 12 s. Plasminogen samples were buffer exchanged using ZipChip diluent (Repligen, Massachusetts, USA) to a final concentration of 5 mg/mL. For separation, a voltage of either 20 or 25 kV was applied to the inlet.

For hyphenation of CZE to MS, the homebuilt nanoCEasy interface was applied [40]. Emitters were obtained from BioMedical Instruments (Zoellnitz, Germany) and had an opening of 30 μm at the tip. The emitter was placed 3 mm in front of the MS orifice. The position of the emitter was controlled using a digital microscope (Dino‐Lite AM7515MZTL, Almere, the Netherlands). For native MS conditions, a sheath liquid (SL) containing IPA:H_2_O (30:70) and 20 mM AmAc was applied. For denaturing MS conditions, the SL contained IPA:H_2_O (50:50) and 5% HAc. The SL was delivered at a flow rate of around 10 μL/min using a syringe pump with the excess of the SL being drained backwards and the nanoES flow being determined primarily by applied voltage [41]. The Orbitrap Exploris 480 was applied in CZE‐MS measurements in high pressure mode with positive ESI with the following final conditions: Mass range 1500–8000 m/z; resolution = 30 000, RF lens = 200; in‐source CID = 120 eV, ion transfer tube: 250°C; 10 microscans, AGC target: 300% with a max injection time: 100 ms.

Near‐Native SEC‐MS

2.3

All SEC‐MS experiments were performed on an Ultimate 3000 RSLC HPLC (Thermo Scientific, Germering, Germany) coupled to a Q‐Exactive UHMR MS (Thermo Scientific, Bremen, Germany) using a Yarra SEC‐X150 column (1.8 μm, 4.6 × 150 mm, Phenomenex, California, USA). The mobile phase used was 100 mM AmAc, the flowrate was 100 μL/min (isocratic), column temperature was set to 25°C, and UV detection was performed at 214 and 280 nm. All samples were injected as is without buffer exchange at an injection volume of 6 μL. All MS measurements were performed in positive mode with typical conditions shown below but could be slightly adjusted to improve signal‐to‐noise ratio and ion desolvation: resolution = 25 000, in‐source CID = 10 eV, in‐source trapping on, desolvation voltage = −10 V, maximum IT = 200 ms, S‐lens RF level = 200, detector m/z optimization = high m/z, ion transfer target m/z = high m/z.

Data Analysis of Intact Plasminogen Proteoforms

2.4

Thermo Scientific FreeStyle software was used for qualitative and quantitative (peak area) analysis of total ion chromatogram (TIC), extracted ion chromatograms (EIC), and UV‐chromatograms. The theoretical m/z values used to evaluate the EIC of different plasminogen proteoforms were calculated using the top 4 charge states (19^+^ − 22^+^) of plasminogen protein sequence, taking into account the 24 disulfide bonds as well as different combinations of O‐glycosylation (types H1N1S1 and H1N1S2), N‐glycosylation (type A2), and phosphorylation.

Additionally, the Byosphere software platform as well as Intact Mass (Version: v3.8‐11) from ProteinMetrics was used to perform molecular weight deconvolution of CZE‐MS and SEC‐MS data and identify different plasminogen proteoforms. Sequence information for plasminogen (P00747) and tPA (P00750) was taken from Uniprot database (www.uniprot.org). The calculation of pI values was performed using the Expasy tool (https://web.expasy.org/cgi‐bin/compute_pi/pi_tool.cgi).

Results and Discussion

3

Separation of Plasminogen Proteoforms by CZE‐MS Under Near‐Native Conditions

3.1

Native capillary zone electrophoresis‐mass spectrometry is typically performed applying a background electrolyte (BGE) containing AmAc [20, 21]. Although high concentrations of AmAc are preferred to preserve the native form of proteins in direct infusion native ESI‐MS, the concentration in CZE is restricted to values of about 50 mM (when applying the typical 50‐μm ID capillaries) due to limitations of the current. Recently, we showed excellent proteoform separation applying a BGE of 50‐mM AmAc (neutral pH) in a HPMC coated capillary [24]. Here, we successfully applied this method, however, the pH of the BGE needed to be adapted to the pI of plasminogen, which has been determined to be in the range of 6.2–6.8 depending on the proteoforms [42]. Because the pH of the BGE needs to be below the pI, in order to ensure a net positive charge of the protein, we adjusted the pH of the BGE to 5.0 by adding 20 mM HAc to a 50 mM AmAc solution. This pH not only enables protein migration but is also close enough to the pI (i.e., about 1–2 pH units below the pI) to have a good selectivity for charge‐modified proteoforms (i.e., containing sialic acids or phosphorylation). By applying the near‐native CZE conditions using the nanoCEasy interface, several proteoforms of plasminogen were separated as illustrated for charge‐deconvoluted spectra in Figure 1. The spectra shown on the left were averaged over 1‐min intervals across the CZE timescale and arranged from top to bottom, corresponding to migration times between 29 and 54 min. The identity of these proteoforms was determined by comparing experimentally measured molecular weights with theoretical values calculated from the plasminogen sequence, incorporating various PTMs. These modifications include phosphorylation, O‐glycosylation (type H1N1S1 and H1N1S2), and N‐glycosylation (A2 type). The Total Ion Electropherogram (TIE) of the CZE separation is presented on the right, providing an overview of the migration profile under near‐native conditions.

Deconvoluted mass spectra of plasminogen acquired by near‐native CZE‐MS. Each deconvoluted mass spectrum on the left was averaged over a 1‐min interval of the CZE separation and is displayed from top to bottom, corresponding to 29.0–54.0 min. The Total Ion Electropherogram (TIE) of the near‐native CZE separation is shown on the right. Separation is performed in a BGE of 50 mM AmAc +20 mM HAc (pH = 5.0). See text for more details on the method.

At least 20 well defined proteoforms can be distinguished based on differences in glycosylation and phosphorylation. The separation according to changes in the charge‐to‐size ratio is clearly visible with the type I plasminogen (containing an additional N‐linked complex type glycan with a biantennary structure and two sialic acids [type A2]) which are migrating later than type II plasminogen. The separation of selected proteoforms due to changes in the phosphorylation and N‐glycosylation is shown in Figure 2.

Near‐native CZE‐MS EIEs of different plasminogen proteoforms. The following four major proteoforms can be identified: (i) type I—nonphosphorylated (T1‐A, T1‐C, T1‐E, and T1‐G), (ii) type II—nonphosphorylated (T2‐A, T2‐C, T2‐E, and T2‐G), (iii) type I—phosphorylated (T1‐B, T1‐D, T1‐F, and T1‐H), and (iv) type II—phosphorylated (T2‐B, T2‐D, T2‐F, and T2‐H). The top 4 charge states from each of the major forms have been used to create the EIEs. See text for details on the method.

CZE‐MS under the condition described above results in baseline separation of type I versus type II plasminogen, both in the nonphosphorylated and phosphorylated form as presented in Figure 2A using extracted ion electropherograms (EIEs) of the specific proteoform. In addition, phosphorylated forms are clearly separated from nonphosphorylated forms for type I and type II, respectively, as shown in Figure 2B. Interestingly, a reproducible split of type I peaks is observed for both the phosphorylated and nonphosphorylated form. The reason for the split peak remains unclear; however, it could be explained by the presence of differently folded isobaric isoforms.

All detected proteoforms of plasminogen along with static nanoESI data and results from the SEC‐MS are listed in Table 1. Despite the limitation of the absolute injection volume in the CZE (only 71 ng of total protein was injected), the CZE‐MS approach resulted in significantly more identified plasminogen proteoforms compared to the static nanoESI experiment. The nanoCEasy interface shows excellent performance even when using a sheath liquid containing 30% aqueous IPA and 20‐mM AmAc leading to a native charge envelope (m/z range 3500–6000 with a maximum intensity for charge state 20+) (Figure S1A in supporting information). In contrast, a charge envelope in the m/z range 2200–3200 (maximum intensity for charge state 35+) is obtained when a denaturing SL containing 50% aqueous IPA with 5% HAc is used. Obviously, the 30% IPA does not denature plasminogen, whereas the addition of 5% HAc does. This is in agreement with findings for monoclonal antibodies [21, 24]. Still, the addition of 30% IPA contributes to increased sensitivity when compared to spraying from pure aqueous solutions. This flexibility of the nanoSL interface allows the use of near‐native separation conditions in CZE combined with denaturing ionization conditions enabling the use of mass spectrometers with limited m/z range for this type of application. Additionally, the use of denaturing ionization conditions leads to an increase in charge (lower m/z), potentially resulting in higher sensitivity (less cluster‐prone) in mass spectrometric detection, as demonstrated previously [24].

Separation of Plasminogen Proteoforms by SEC‐MS Under Near‐Native Conditions

3.2

The various proteoforms of plasminogen were also characterized by SEC‐MS method under near‐native conditions using 100 mM AmAc as the mobile phase (see Method section for more details). Figure 3 illustrates the deconvoluted mass spectra of plasminogen obtained using a near‐native SEC‐MS approach. The spectra shown on the left were averaged over 15‐s intervals across the SEC timescale and arranged from top to bottom, corresponding to retention times between 12.75 and 17 min. The total ion chromatogram (TIC) of the SEC separation is presented on the right, providing an overview of the elution profile under near‐native conditions. This approach enables the characterization of intact proteoforms under size‐exclusion conditions while preserving noncovalent interactions, offering complementary insights to the above‐mentioned CZE‐MS method.

Deconvoluted mass spectra of plasminogen acquired by near‐native SEC‐MS. Each deconvoluted mass spectrum on the left was averaged over a 15‐s interval of the SEC timescale and is displayed from top to bottom, corresponding to 12.75–17.00 min. The total ion chromatogram (TIC) of the near‐native SEC separation is shown on the right.

Similar to observations from CZE‐MS methods, the SEC‐MS analysis revealed a high degree of heterogeneity in plasminogen, with multiple proteoforms detected across the chromatographic window. The detected proteoforms can be broadly classified into two categories based on their elution behavior. Early‐eluting species predominantly contained N‐glycosylation and correspond to type I plasminogen, whereas later‐eluting species lacked N‐glycosylation and represent type II plasminogen. This separation suggests that glycosylation significantly influences the hydrodynamic properties of plasminogen under near‐native conditions, affecting its SEC retention time. These findings not only confirm the structural diversity of plasminogen but also demonstrate the utility of SEC‐MS for probing glycoform‐dependent conformational differences in complex plasma proteins.

Figure 4 demonstrates the capability of near‐native SEC‐MS to resolve plasminogen proteoforms based on structural differences and PTMs. Extracted ion chromatograms (EICs) were generated for representative proteoforms grouped into the following four major categories: (i) type I—nonphosphorylated, (ii) type II—nonphosphorylated, (iii) type I—phosphorylated, and (iv) type II—phosphorylated. Within each category, the top four proteoforms were selected based on abundance and structural diversity, encompassing combinations of N‐glycosylation (A2), O‐glycosylation (H1N1S1 or H1N1S2), and phosphorylation. This targeted approach provides a clear visualization of how glycosylation and phosphorylation influence chromatographic behavior under near‐native conditions.

Near‐native SEC‐MS EICs of different plasminogen proteoforms. The following four major proteoforms can be identified: (i) type I—nonphosphorylated (T1‐A, T1‐C, T1‐E, and T1‐G), (ii) type II—nonphosphorylated (T2‐A, T2‐C, T2‐E, and T2‐G), (iii) type I—phosphorylated (T1‐B, T1‐D, T1‐F, and T1‐H) and (iv) type II—phosphorylated (T2‐B, T2‐D, T2‐F, and T2‐H). The top 4 charge states from each of the major forms have been used to create the EICs shown in this figure. See text for details on the method.

The top panel of Figure 4A highlights the separation between type I and type II plasminogen, both in phosphorylated and nonphosphorylated forms. Type I proteoforms, which contain N‐glycosylation (A2), elute earlier than type II proteoforms lacking N‐glycosylation, consistent with the increased hydrodynamic size imparted by N‐linked glycans. Interestingly, O‐glycosylation also influences retention time, as shown by the shift from H1N1S1 to H1N2S2. This suggests that even modest increases in glycan complexity can alter protein conformation or hydration shell, affecting SEC behavior. However, further additions of HexHexNAc and additional sialic acids did not produce significant retention time changes, indicating a threshold beyond which additional glycan complexity has minimal impact on hydrodynamic size under these conditions.

The bottom panel (Figure 4B) compares EICs for phosphorylated versus nonphosphorylated proteoforms within both type I and type II categories. While phosphorylation introduces only a small mass increment, subtle shifts in retention time were observed, suggesting that phosphorylation may influence protein conformation or charge distribution, even under near‐native conditions. Although the separation between phosphorylated and nonphosphorylated species was not complete, the observed trend demonstrates the sensitivity of SEC‐MS for detecting PTM‐induced structural changes.

The ability of near‐native SEC‐MS to resolve plasminogen proteoforms based on N‐glycosylation, O‐glycosylation, and phosphorylation underscores its utility for intact protein analysis in complex biological systems. Unlike denaturing LC–MS workflows, this approach preserves higher‐order structure, enabling differentiation of proteoforms that vary in hydrodynamic size and conformation. Similar to the CZE‐MS method described above, the SEC‐MS approach identified more plasminogen proteoforms compared to previously reported direct infusion nanoESI data (see Table 1). These findings provide valuable insights into the structural heterogeneity of plasminogen and highlight the potential of SEC‐MS for studying glycoform‐dependent functional properties and PTM‐driven structural modulation.

Comparison of CZE‐MS and SEC‐MS Methods Under Near‐Native Conditions

3.3

Both CZE‐MS and SEC‐MS reveal the characterization of intact proteoforms not described so far by direct infusion experiments (cp. [36]). Despite the different separation modes of the two techniques, the relative abundances of plasminogen proteoforms were largely comparable (see Figure S2 in supporting information for the top 20 proteoforms).

To further investigate and confirm the PTMs underlying the plasminogen proteoforms listed in Table 1, we performed a peptide‐based bottom‐up analysis to characterize both the types and locations of these PTMs. Details of the peptide‐based method are provided in the supporting information. Major results are summarized in Figure S3, and further elaborated in Figure S4 and Tables S1–S5 in the supporting information. This analysis confirmed that plasminogen primarily contains multiple PTMs, including N‐glycosylation (type A2 at Asn‐308), O‐glycosylation (types H1N1S1 and H1N1S2 at Ser‐268, Thr‐365, Thr‐475, Thr‐707, and Thr‐710), and phosphorylation (at Ser‐68, Ser‐358, and Ser‐741). For each of these PTM sites, the peptide without the respective PTM, including N308, was also observed.

As shown in Figures 1, 2, and S2, the most abundant proteoform detected by both CZE‐MS and SEC‐MS is T2‐B (type II phosphorylated, cp. Table 1 for nomenclature), which contains one O‐glycosylation (H1N1S1) and one phosphorylation. The proteoform abundance decreases progressively with increasing numbers of glycosylations. In SEC‐MS, even very minor proteoforms containing three different O‐glycans carrying mono‐ and doubly sialylated proteoforms (1× H1N1S1+ 2× H1N1S2) could be detected. These proteoforms could not be detected by CZE‐MS, most likely due to the three orders of magnitude lower injection volume and protein mass (injected protein mass: 71 ng in CZE‐MS and 59 μg in SEC‐MS), potentially also due to the different separation selectivity causing potentially co‐migration with other more abundant proteoforms—in the case of type II these highly sialylated proteoforms are expected to comigrate with the high abundant proteoforms of type I. In general, the resolution of proteoforms is better in CZE‐MS with close to baseline separation of charge variants, however, requiring much longer separation time (CZE‐MS: 60 min; SEC‐MS: 20 min).

Near‐Native CZE‐MS Method to Monitor the Activation of Plasminogen by tPA

3.4

Capillary electrophoresis is an excellent tool to perform online enzyme assays [43, 44], also known as electrophoretically mediated microanalysis (EMMA) [45]. In addition, “in‐capillary” reactions can be used to digest proteins by trypsin, pepsin, or IdeS prior to separation of the resulting peptides [46, 47, 48] or to cleave off glycans. However, very few “in‐capillary” protein reactions have been studied by CZE‐MS. Wang et al. studied the interaction of human recombinant MMP‐9 protein with its substrate peptide by CZE‐MS. Here, we examined how plasminogen is activated by tPA, which represents one of the two main activation pathways in humans. The approach involves co‐injecting plasminogen and tPA, allowing their differing electrophoretic mobilities to mix the proteins, and then analyzing the reaction products within the same analytical run. The concept of this experiment is shown in Figure 5A.

In‐capillary activation of plasminogen by tPA (1:10 tPA:plasminogen) in CZE‐MS. The concept of in‐capillary CZE‐MS for the activation study of plasminogen by tPA is shown in Figure 5A. Deconvoluted mass spectra of a CZE‐MS analysis of plasminogen with the co‐injection of tPA (1.1% of capillary volume of 0.25 μg/μL tPA solution) (Figure 5B). Each deconvoluted mass spectrum on the left was averaged over a 1‐min interval of the CZE separation and is displayed from top to bottom, corresponding to 30.0–50.0 min. The Total Ion Electropherogram (TIE) of the near‐native CZE separation is shown on the right. CZE‐MS conditions are the same as in Figure 1.

Because of its higher pI (7.6) and smaller size, tPA is expected to migrate faster at pH 5.0 compared to plasminogen (pI = 7.0). These values correspond to a relatively higher number of basic amino acid residues and a relatively lower number of acidic amino acid residues of tPA, respectively, when compared to plasminogen. Even though these pI values are calculated without taking the PTMs into considerations (experimental pI of plasminogen 6.2–6.8 [42]) we decided to inject tPA directly following the injection plug of plasminogen expecting a mixing due to faster migration of tPA. As anticipated, we observed plasminogen reacting with tPA. When a 12‐s, 50‐mbar, 0.5‐μg/μL plug of tPA was injected immediately after the same volume of a 5 mg/mL plasminogen solution, the resulting CZE‐MS data revealed both unreacted plasminogen and truncated plasminogen proteoforms within the 80‐ to 85‐kDa mass range (Figure 5B). These truncated forms can be attributed to the cleavage of the first 68 amino acids, which is in agreement with the data from Cramer et al. [36]. In addition, the cleaved fragment can be observed in the effluent of the CZE, when a pressure is applied. This is in perfect agreement to the pI of the cleaved peptide (AA1‐68: pI = 4.84), which is close to the pH of the BGE (5.0), leading to hardly any migration and, thus, appearing only after the application of pressure. In agreement, the truncated proteoforms are migrating slightly faster than plasminogen due to slightly higher basicity (truncated plasminogen (‐AA1‐68): pI = 7.7 in comparison to plasminogen with a pI of 7.0).

Furthermore, the truncated plasminogen shows separation of proteoforms of type I and type II, different degrees of glycosylation as well as phosphorylation similar to the intact plasminogen. In order to evaluate the selectivity of the tPA, we determined the ratios of all major proteoforms as shown in Figure 6.

Proteoform distribution of plasminogen and truncated plasminogen after co‐injection of tPA in CZE‐MS experiments. All data are normalized to the proteoform type II, +H1N1S1+P (most abundant proteoforms of intact plasminogen). Blue bars represent the proteoforms distribution of the intact plasminogen, orange bars indicate the results of the co‐injection of tPA (1.1% of capillary volume of 0.25 μg/μL tPA solution); full orange: remaining plasminogen, striped: truncated forms with the loss of AA1‐68.

Figure 6 shows the proteoform distribution of intact plasminogen in blue, and the experiment after treatment with tPA (orange), both of the remaining plasminogen (full orange) and the truncated plasminogen (striped orange). All intensities are normalized to the most abundant proteoform of intact plasminogen (1× H1N1S1+P). It is evident that type I (the N‐glycosylated form) reacts more quickly with tPA than type II (the non‐N‐glycosylated form), as shown by both a lower amount of remaining plasminogen and a higher amount of truncated plasminogen. Likewise, phosphorylated proteoforms undergo processing faster than their nonphosphorylated counterparts.

Up to this point, only data with a tPA:plasminogen ratio of 1:10 are shown. When injecting larger amounts of tPA (1:5 ratio), we observed a complete activation of plasminogen (hardly any intact plasminogen visible) and further processing to the hydrolyzed form of the AA(1‐68) truncation, as well as the truncation of the first 77AA (hydrolyzed) and additional so far unidentified fragments (Figure S5 in supporting information). In addition, in the case of a lower tPA:plasminogen ratio, type I plasminogen was activated faster, because only the type II remains. It is worth mentioning that the chosen experimental conditions in the CZE approach reassemble the activation conditions of free plasminogen in a “closed conformation.” The activation of plasminogen with tPA in complex with fibrin (“open conformation” [37]) can potentially lead to different effects of the PTMs [49].

This proof‐of‐concept study demonstrates that it is possible to perform proteoform‐resolved activation studies of plasminogen with very low amounts of proteins (mid‐ng range). The same concept could also be used to study the interaction of binding partners with plasminogen and other proteins.

Concluding Remarks

4

In this work, we have demonstrated that the combination of near‐native CZE‐MS and SEC‐MS enables a highly detailed and complementary characterization of the human plasma derived plasminogen proteoforms beyond what can be achieved by direct infusion native MS alone. Both separation techniques preserve native‐like conditions while introducing critical orthogonal selectivity, enabeling the resolution and confident assignment of plasminogen proteoforms differing by subtle PTMs like N‐glycosylation, O‐glycosylation, and phosphorylation.

Near‐native CZE‐MS provides superior resolution for charge modifying PTMs, achieving near baseline separation of phosphorylated and sialylated proteoforms at minimal injection amounts in the low nanogram range. The use of an HPMC coated capillary and a pH‐controlled AmAc BGE proved critical for maximizing selectivity close to the pI of plasminogen, while the nanoflow‐MS interface offered the flexibility to operate under native or denaturing ESI conditions without compromising separation performance. In contrast, near‐native SEC‐MS offers robust separation based on hydrodynamic size and glycosylation‐dependent conformational differences, enabling the detection of additional low‐abundance plasminogen proteoforms and providing complementary structural insight under conditions readily compatible with high mass range instrumentation.

Importantly, we also showed that the CZE‐MS platform further enables proteoform‐ resolved interaction studies directly within the separation capillary. Using plasminogen activation by tPA as a proof‐of‐concept, we demonstrate that in‐capillary mixing and reaction monitoring can adequately resolve proteoform‐specific processing kinetics. The observed preferential activation of N‐glycosylated and phosphorylated plasminogen proteoforms highlights the functional relevance of PTMs and illustrates the potential of native CZE‐MS for studying enzymatic selectivity under solution‐ phase and near physiological conditions using minimal sample amounts.

Taken together, this report underscores the strong complementarity of near‐native CZE‐MS and SEC‐MS for intact plasminogen proteoform analysis and interaction studies. Beyond understanding the complexity of plasminogen better, the workflows described in this report are broadly applicable to other heterogeneous plasma proteins and biotherapeutic proteins where subtle PTM differences may modulate structure, interaction, and function. The ability to couple high‐resolution proteoform separation with near‐native MS detection and in‐capillary reaction monitoring provides a powerful complementary analytical toolbox for advancing proteoform‐resolved structural and functional studies.

Supporting information

Figure S1: Exemplary mass spectrum from near‐native CZE‐MS (A) and SEC‐MS (B) showing the major proteoform of type II (T2‐A). Figure S2: Comparison of relative abundance (%) of different proteoforms measured by CZE‐MS and SEC‐MS (n = 3 injections). Figure S3. Plasminogen domain organization and post‐translational modifications Plasminogen comprises an N‐terminal Pan‐apple (PAp) domain, five kringle domains, and a C‐terminal serine protease domain. And can be classified by the presence (type I) or absence (type II) of N308 glycosylation. Phosphorylation (predominantly at S358) and O‐glycosylation at several other sites contribute to a complex proteoform profile of plasminogen. Novel sites of post‐translational modification and labelled in red. Dashed lines indicate sites of observed tPA‐induced cleavage. Figure S4: MS/MS spectra of phosphorylated peptides enriched from plasminogen. (A) MS/MS spectrum of tryptic peptide covering phosphorylated S358 and O‐glycosylated T365. (B) MS/MS spectrum of a Glu‐C generated peptide covering phosphorylated S358. (C) MS/MS spectrum of a Glu‐C generated peptide covering phosphorylated S741. (D) MS/MS spectrum of tryptic peptide covering phosphorylated S68. Table S1: Posttranslational modifications found on plasminogen. Glycosylation and phosphorylation contribute to a complex proteoform profile of plasminogen. Table S2: Position, composition and quantification of plasminogen O‐glycosylation All of the most abundant O‐glycosylation found on plasminogen comprises 1× HexNAc, 1× Hex and either 1× or 2× NeuAc, contributing to a mass shift of 656 and 947 Da, respectively. Table S3: Position, composition and quantification of plasminogen N‐glycosylation Plasminogen N308 glycosylation is primarily 4× HexNAc, 5× Hex and 2× or 1× NeuAc, contributing mass shifts of 2204.77 and 1913.68 Da respectively. Table S4: Identification and abundance of plasminogen phosphorylation sites using trypsin digestion followed by enrichment of phosphorylated peptides. Four unique phosphorylation sites were identified using trypsin to digest plasminogen before enrichment of phosphorylated peptides using TiO_2_. Of these sites, phosphorylated S358 was the most abundant. Table S5: Identification and abundance of plasminogen phosphorylation sites using GluC digestion followed by enrichment of phosphorylated peptides. Four unique phosphorylation sites were identified using GluC to digest plasminogen before enrichment of phosphorylated peptides using TiO_2_. Of these sites, phosphorylated S358 was the most abundant. Figure S5: In‐capillary activation of plasminogen by tPA (1:5 tPA:plasminogen) in CZE‐MS. Deconvoluted mass spectra of a CZE‐MS analysis of plasminogen with the co‐injection of tPA (1.1% of capillary volume of 0.5 μg/μL tPA solution). Each deconvoluted mass spectrum on the left was averaged over a 1‐min interval of the CZE separation and is displayed from top to bottom, corresponding to 23.0–43.0 min. The total ion electropherogram (TIE) of the near‐native CZE separation is shown on the right. CZE‐MS conditions are the same as in Figure 1.