Skip to main content
NCBI home page
ACS AuthorChoice logoLink to ACS AuthorChoice
. 2024 Jun 10;96(24):9994–10002. doi: 10.1021/acs.analchem.4c01384

Unravelling the Link between Oligonucleotide Structure and Diastereomer Separation in Hydrophilic Interaction Chromatography

Honorine Lardeux †,‡, Kathrin Stavenhagen §, Clément Paris §, Rikke Dueholm §, Camille Kurek §, Leonardo De Maria §, Felix Gnerlich §, Tomas Leek §, Werngard Czechtizky §, Davy Guillarme †,‡, Manasses Jora §,*
PMCID: PMC11190878  PMID: 38855895

Abstract

graphic file with name ac4c01384_0006.jpg

Therapeutic oligonucleotides (ONs) commonly incorporate phosphorothioate (PS) modifications. These introduce chiral centers and generate ON diastereomers. The increasing number of ONs undergoing clinical trials and reaching the market has led to a growing interest to better characterize the ON diastereomer composition, especially for small interfering ribonucleic acids (siRNAs). In this study, and for the first time, we identify higher-order structures as the major cause of ON diastereomer separation in hydrophilic interaction chromatography (HILIC). We have used conformational predictions and melting profiles of several representative full-length ONs to first analyze ON folding and then run mass spectrometry and HILIC to underpin the link between their folding and diastereomer separation. On top, we show how one can either enhance or suppress diastereomer separation depending on chromatographic settings, such as column temperature, pore size, stationary phase, mobile-phase ionic strength, and organic modifier. This work will significantly facilitate future HILIC-based characterization of PS-containing ONs; e.g., enabling monitoring of batch-to-batch diastereomer distributions in full-length siRNAs, a complex task that is now for the first time shown as possible on this delicate class of therapeutic double-stranded ONs.

Therapeutic oligonucleotides (ONs) have proven effective as a pharmacological strategy toward previously “undruggable” targets by regulating gene expression.1 Single-stranded antisense ONs (ASOs) and double-stranded small interfering ribonucleic acids (siRNAs) are currently the main ON-based therapeutics undergoing clinical trials.2 Unmodified nucleic acids often suffer from enzymatic degradation, immune activation, low target affinity, and limited cellular uptake.3 Therefore, chemical modifications are essential for more effective, stable, and specific ON therapeutics.4

Among the 20 FDA-approved ON therapeutics, six are fully phosphorothioated ASOs, two are partially phosphorothioate (PS)-modified ASOs (i.e., 13 and 15 PS modifications), and five are double-stranded siRNAs incorporating six PS modifications (i.e., two PS located on one strand and four PS located on the other).2,5 Originally introduced to enhance nuclease resistance, PS modifications also confer improved pharmacokinetic properties and target affinity to the ON.6 In PS-modified ONs, one (or several) nonbridging oxygen atoms in the phosphodiester backbone are replaced by sulfur. Each PS modification adds chirality to a phosphorus atom leading to either the Rp or Sp configuration. As a result, PS-containing ONs are a mixture of 2n diastereomers, where n equals the number of PS linkages.

Despite the growing interest in stereocontrolled ON synthesis,7,8 stereopure PS-modified compounds remain a minority among ONs in clinical trials.9 Hence, the pharmacological and physicochemical profiles of PS-modified drugs are the cumulative result of the individual properties of their diastereomer population. It has been demonstrated that a single change in the PS configuration can result in a significant change in the ON topology, leading to differences in lipophilicity, ionic character, and molecular interactions.10−12

In this context, monitoring the diastereomeric distribution resulting from PS modifications and batch-to-batch fingerprinting are critical to avoid unpredictable variations in product safety and efficacy.13,14 This is particularly crucial during drug manufacturing, where small modifications to the process can lead to significant changes in the diastereomeric population. Equally, the location of fractionation points during purification can also affect the diastereomeric composition.15,16

Among the liquid chromatography (LC)-based techniques used for characterizing PS-modified ONs,17 anion-exchange chromatography and ion-pairing reversed-phase LC (IP-RPLC) have demonstrated significant potential for (partially) separating ON diastereomers.18−20 Additional complementary methods for studying diastereomers in PS-modified ONs are capillary electrophoresis using chiral selectors, ion mobility spectrometry, and spectroscopic methods [e.g., 31P nuclear magnetic resonance (NMR) and circular dichroism].21−23

Due to the significant number of diastereomers in full-length therapeutic ONs, even their partial separation is usually not achievable. Therefore, more commonly than not, the batch-to-batch diastereomeric distribution of ONs remains uncharacterized. Instead, changes in the ON synthesis and purification processes are kept to a minimum to reduce batch-to-batch disparities. In addition, purity assessments and impurity profiling are usually performed using strategies that suppress diastereomer separation.24 More specifically, bulky and highly hydrophobic IP reagents,25 highly hydrophobic stationary phases,26 and elevated column temperature27 are used for standard ON analysis.

Recently, hydrophilic interaction chromatography (HILIC), increasingly recognized for its applicability to ON analysis,28 has shown promising ON diastereomer separation. Goyon et al.23 have demonstrated that ONs modified with methoxy and fluoro substituents at position 2′ or locked nucleic acid moieties (a methylene bridge between 2′-O and 4′-C) show improved diastereomeric separation due to enhanced structural rigidity.

Here, we confirm that HILIC is indeed a valuable chromatographic technique for the partial separation of diastereomers in therapeutic full-length, single- and double-stranded PS-modified ONs. Using a combination of conformational predictions, melting profiles, mass spectrometry (MS) data, and diverse HILIC settings, we demonstrate that intrinsic higher-order structures (HOSs) of ONs play a pivotal role for (partial) separation of ON diastereomers. Through a better understanding of the parameters (such as column stationary phase, pore size and temperature, mobile-phase ionic strength, and organic modifier), we provide insights into modulating (including suppressing) ON diastereomer separation in HILIC.

Experimental Section

Materials

The ON sequences employed in this study and details of their secondary structure predictions are provided in Tables S1 and S2. For the sake of clarity, names are written as follows: as and ss refer to antisense and sense strands, respectively, followed by the number of PS modifications. Purified as-202PS and ss-202PS were purchased from Axolabs (Kulmbach, Germany) and resuspended in deionized water at 100 μM prior to analysis. All additional single- and double-stranded ONs were synthesized in-house. Additional details on ON synthesis are available in the Supporting Information. Deionized water (H2O) was obtained from an Elix Technology Inside purification system (Merck Millipore, Darmstadt, Germany). Optima LC–MS-grade acetonitrile (ACN) and methanol (MeOH) were purchased from Fisher Scientific (Loughborough, UK). Ammonium acetate (AA, LiChropur LC–MS) was purchased from Sigma-Aldrich.

LC–MS Analysis

LC–MS experiments were carried out using a Premier LC system (Waters, Milford, MA, USA) equipped with a binary solvent delivery pump coupled to a photodiode array detector (Waters) for UV detection at 260 nm and a high-resolution RDa mass spectrometer (benchtop TOF, with about 10,000 FWHM mass resolution, Waters). Unless stated otherwise, the ONs were mainly analyzed on a Premier Glycoprotein BEH Amide (100 × 2.1 mm, 300 Å, 1.7 μm) column, but other columns were also tested (Figure S8 only): Premier BEH Amide (100 × 2.1 mm, 130 Å, 1.7 μm), Triart Diol-HILIC (100 × 2.1 mm, 120 Å, 1.9 μm), BEH HILIC (100 × 2.1 mm, 130 Å, 1.7 μm), and Poroshell 120 HILIC-Z (100 × 2.1 mm, 120 Å, 1.9 μm). Injection volumes of 0.1 μL (for single strands) and 0.2 μL (for duplexes)—10 pmol were loaded on the column. Mobile phases were composed of AA in 30:70 H2O/ACN (mobile phase A, MPA) and AA in 70:30 H2O/ACN (mobile phase B, MPB). The flow rate was set to 0.4 mL/min (unless stated otherwise). Specific AA concentrations in the mobile phases, organic modifier, column temperature, and gradient programs for each experiment are described in the main text, as needed.

A mass spectrometer was used in electrospray negative mode with an acquisition range of 400 to 5000 m/z. The desolvation temperature was set at 450 °C, source temperature was set at 120 °C, the cone voltage was set to 40 V, and the capillary voltage was set to 0.8 kV. These conditions were derived from preliminary optimization studies (data not shown). UNIFI (Waters) was used as the data acquisition and processing software.

Melting Temperature Curves

A Lambda 365 UV/vis spectrophotometer equipped with a thermostated multicell Peltier holder and a Peltier temperature controller (PerkinElmer, Upplands Väsby, Sweden) was used to acquire melting temperatures of purified ONs. UV absorbance was measured at 5 μM ON concentration in different buffers from 70 to 10 °C. Experiments were performed using the following parameters: 260 nm (wavelength), 0.5 min (holding time), 0.5 °C (data acquisition interval), 1 °C/min (heating rate), 0.1 °C (error range). The experimental melting temperatures were calculated by using a first derivative curve. UV data was acquired by UV Express software (PerkinElmer).

Results and Discussion

Column Temperature and Mobile-Phase Ionic Strength Play a Critical Role in Diastereomer Separation in HILIC

Figure 1 shows extracted ion chromatograms (EICs) obtained for as-202PS, a model single-stranded siRNA sequence containing four PS modifications, two at each sequence end, i.e., 16 diastereomers. This sequence is predicted to adopt a hairpin conformation with a melting temperature of 40 °C (Table S2). HILIC analyses were performed with varying AA concentration in the mobile phases (i.e., 10 or 100 mM) and different column temperatures (i.e., 90, 40, and 5 °C). A generic gradient profile of 35–50% H2O (12.5–50% MPB) in 5 min, derived from preliminary optimization studies, was used. Similar experiments were performed using as-1PS (same sequence as as-202PS but one PS-modified linkage, thus two diastereomers) and as-0PS (same sequence as as-202PS but only phosphodiester linkages, thus no diastereomers).

Figure 1.

Figure 1

Open in a new tab

Effect of column temperature and mobile-phase ionic strength on HILIC analysis of as-202PS, that possess four PS modifications, indicated by a purple V. EICs showing the comparison of 90 (red), 40 (orange), and 5 °C (blue) as column temperature and 10 mM (i) and 100 mM (ii) as AA concentration in the mobile phases. Circled nucleotides indicate sugar modifications (2′-OMe indicated by a solid line and 2′-F by a dotted line).

As depicted in Figure 1, a single chromatographic peak was observed at 90 °C for as-202PS, independent of the AA concentration in the mobile phase. This is likely resulting from the absence of the ON HOS at temperatures above the melting temperature of as-202PS.29 Temperature-induced denaturation of the ON HOS exposes more hydrophilic sites,30 thus enhancing ON interactions with the amide stationary phase. This very likely explains the increase in the retention time with temperature. Lowering the temperature to 40 °C and further to 5 °C triggers the chromatographic peak to split into three chromatographic peaks at 10 mM AA and 5 °C.

As for as-202PS, analysis of as-1PS (Figure S1A) showed the same trends in terms of relative retention time vs column temperature and mobile-phase ionic strength. However, two resolved isobaric peaks for the two diastereomers were detected at 10 mM AA and 5 °C.

As illustrated in Figure 1, increasing the salt concentration to 100 mM AA resulted in nine partially separated chromatographic peaks at 5 °C for as-202PS and thus enhanced ON retention and diastereomer resolution at higher AA concentrations in the mobile phases. This phenomenon can be partially explained by minimized electrostatic repulsion between the residual silanols of the stationary phase31 and the negatively charged ONs, as well as higher ON solvation.32 It is worth mentioning that AA-containing solutions of different concentrations, including 100 mM, are known to stabilize the HOS of proteins due to AA’s kosmotropic nature,33 allowing its use to effectively mimic physiological conditions in native MS-based analysis of proteins and nucleic acids.34

Another important factor that helps explaining the enhanced ON retention and diastereomer resolution at high AA concentrations is the increase of polarity of the immobilized water layer associated with the stationary phase under high ionic strength,32 especially in combination with low column temperatures. Within the gradient profile of 35–50% H2O, the formation of a salt-enriched water layer on the surface of the stationary phase is favored at higher mobile-phase ionic strength, since AA is insoluble in ACN. Therefore, in addition to surface adsorption, one can expect partitioning to play an important role in the HILIC separation at high AA concentrations and low column temperatures.35 This is in contrast to ON separations in HILIC at low ionic strength, where the immobilized water layer on the surface of the stationary phase is destabilized due to the relatively high water content (i.e., > 30%) in the mobile phases.36

Additional experiments were performed to confirm whether the ON isobaric species separated by HILIC are indeed diastereomers. Figure S1B shows the corresponding chromatograms for as-0PS, which has the same sequence as the other two compounds (i.e., as-1PS and as-202PS), but no diastereomer. Regardless of the column temperature or mobile-phase ionic strength, only a single chromatographic peak was observed for as-0PS. This confirms that the partial separation patterns observed for as-1PS and as-202PS can be attributed to ON diastereomers.

To better understand the pivotal role of column temperature on the ON HOS, we investigated the charge state distribution (CSD) of as-202PS during electrospray ionization MS (ESI-MS) analysis. Figure S2 shows the mass spectra acquired for as-202PS at column temperatures of 90, 40, and 5 °C, employing mobile phases containing 100 mM AA. At 40 and 5 °C, the mass spectra are characterized by the prevalence of the ON at low charge states (z), ranging from z = −3 to z = −5. The two most abundant ions are [M – 4H]4– and [M – 5H]5–, with a subtle shift toward higher mass-to-charge (m/z) values (i.e., lower charge states) at lower temperatures. Lowering the temperature to 5 °C results in [M – 4H]4– being more populated (69% of the total MS signal) when compared to 40 °C (48%). Such a limited number of charge states in the mass spectra strongly indicate a more compact ON structure37 and are consistent with the predicted HOS of as-202PS under these conditions (Table S2).

This is further explained by the high mobile-phase ionic strength that masks a high proportion of the phosphate groups, favoring a narrower CSD at lower charge states.37 Conversely, at a temperature of 90 °C, ions shift toward lower m/z values (i.e., higher charge states), displaying a broader CSD from z = −3 to z = −9. This distinctive pattern is characteristic of unfolded ONs, in which more sites are available for deprotonation. Despite the relatively high ionic strength of the eluent, the elevated column temperature effectively disrupts intramolecular hydrogen bonding, leading to the adoption of an unfolded ON conformation.37

Together, these results highlight the critical role of both column temperature and mobile-phase ionic strength in driving the folding and unfolding dynamics of ONs and therefore enabling the (at least partial) separation of diastereomer species in HILIC. Moreover, they suggest that despite the high levels of organic solvents in HILIC mobile phases, the HOSs adopted by nucleic acids are preserved (at least to a high level) at low column temperature and high mobile-phase ionic strength.

ON Sequences Favoring Rigid HOSs Provide Better Chromatographic Resolution of ON Diastereomers

While a comparison of MS spectra at different temperatures strongly suggests the presence of distinct temperature-dependent ON conformations (i.e., folded and unfolded), it is important to acknowledge the possibility of conformational changes occurring during the ESI-MS process. Therefore, we decided to characterize the structure of the ON in solution to confirm our MS findings.

UV absorbance vs temperature profiles are valuable tools for evaluating the conformation of nucleic acids in solution (and their propensity to fold in a given environment), especially for monitoring conformational changes with temperature and ionic strength (due to changes in UV absorbance). Due to the hypochromic effect, folded ONs present a decreased absorbance of UV light when compared to unfolded species.38

Figure 2A shows melting curves measured for as-202PS in various media: pure H2O, HILIC diluent, and phosphate-buffered saline solution (1× PBS). In H2O, as-202PS shows no/minimal melting characteristics. This result aligns with expectations, as the negatively charged phosphate groups, being less shielded by counterions along the ON, lead to a reduced folding propensity.39 However, when diluted in 100 mM AA in 35:65 H2O/ACN, simulating the initial conditions of HILIC analysis, a clear transition in absorbance occurs with a temperature decrease from 70 to 10 °C. This transition manifests as a sigmoidal curve, characteristic of RNA melting, and confirms the presence of a HOS at temperatures below the melting temperature. This observation is supported by computational predictions (Table S1). A similar melting profile was obtained for as-202PS diluted in 1× PBS. The local maxima of the first derivatives of the melting curves indicate a melting temperature for as-202PS of 38 °C in HILIC diluent vs 41 °C in 1× PBS, in close agreement with the theoretical value of 40 °C. This observation is congruent with our assumption that the chromatographic conditions used in our study are unlikely to disrupt the HOS of ONs below their melting temperature. Through their ability to indicate conformational changes of ONs in solution, UV melting curves enable the differentiation between sequences prone to intramolecular Watson–Crick base pairing (e.g., as-202PS) and unstructured sequences.40 While sigmoidal melting curves are expected for ONs with HOSs, a more linear response with no pronounced transition is expected for unstructured ONs.40 In this context, we further examined the melting curve of the complementary strand of as-202PS and ss-202PS. ss-202PS was chosen as the computational model that did not predict any HOS for this sequence (Table S2). Not surprisingly, even in the presence of a highly ionic environment represented by the HILIC eluent, no classical melting behavior was observed in the UV spectrum of ss-202PS (Figure 2A). This experimentally confirms the absence of substantial intramolecular base pairing, thus HOS, for ss-202PS, under the HILIC conditions employed in this study.

Figure 2.

Figure 2

Open in a new tab

(A) Melting curves of as-202PS (blue traces) and ss-202PS (green traces) measured over the temperature range of 70–10 °C and using either 1× PBS, H2O, or 100 mM AA in 35:65 H2O/ACN as the diluent. (B) Corresponding EICs of as-202PS and ss-202PS using the generic gradient and 100 mM as AA concentration in the mobile phases and 5 °C as column temperature.

Figure 2B displays the EICs for as-202PS and ss-202PS. These results were obtained using the generic gradient profile of 35–50% H2O in 5 min, 100 mM AA in the HILIC mobile phases, and column temperature set to 5 °C. Chromatographic analysis of as-202PS shows a partial separation of its 16 diastereomers, resulting in about nine observed chromatographic peaks (Figures 1 and 2B). In contrast, the 16 diastereomers of the complementary unstructured strand ss-202PS yield two barely separated chromatographic peaks with a much narrower elution window (Figure 2B). This observation supports the correlation between the ON’s capacity to adopt a HOS and the achieved diastereomer resolution under favorable HILIC conditions.

It is worth noting that as-202PS and ss-202PS have different base compositions and that the difference in overall hydrophilicity may also impact the ability of HILIC to separate their diastereomers.

To mitigate the impact of the base composition, six additional PS-modified ONs were synthesized (Table S3). All these sequences shared the same molecular mass (6646.8 Da), 2′-fluoro (2′-F) and 2′-methoxy (2′-OMe) modifications located at the same alternating positions, the same number and location of PS modifications, and the same proportion of each nucleobase as the model sequence as-202PS. According to RNAfold, the permutations in the base sequence resulted in either compounds with a more rigid HOS than as-202PS (three of them) or unstructured compounds (another three of them) (Table S2). Figure 3 illustrates the EICs corresponding to two of these sequences, analyzed using the generic gradient profile of 35–50% H2O in 5 min, 100 mM AA HILIC mobile phases, and a column temperature of 5 °C. EICs of the other sequences are shown in Figure S3.

Figure 3.

Figure 3

Open in a new tab

EICs of highly structured (HS1) and unstructured (L1) ONs using 100 mM as the AA concentration in mobile phases and 5 °C as the column temperature.

The highly structured ONs, namely, HS1, HS2, and HS3, resulted in three to five narrowed chromatographic peak clusters for a total of five to eight closely separated diastereomers. Unsurprisingly, HS1, which has the lowest minimum free energy (MFE, −8.0 kcal/mol) and is therefore predicted to be the most rigid HOS among the compounds tested here, exhibited the highest diastereomer resolution with five baseline-resolved chromatographic peak clusters compared to (i) HS3 (MFE −5.9 kcal/mol, highest MFE among the three highly structured sequences), which shows only three peak clusters, (ii) HS2 (MFE −6.8 kcal/mol), which presents an in-between HS1 and HS3 behavior with about six partially resolved chromatographic peaks, and (iii) as-202PS (MFE −0.2 kcal/mol), which gives nine observable chromatographic peaks, but with the poorest resolution. The relatively high number of observable chromatographic peaks and poor resolution for as-202PS as opposed to more structured compounds like HS1, HS2, and HS3 may be attributed to its lower frequency of the MFE structure (31.5% compared to > 96.5% for HS1, HS2, and HS3). For as-202PS, the structure represented in this article becomes one among a larger number of ordered structures, as illustrated with the dot plot containing the base pair probabilities (Figure S4). On the contrary, a broad chromatographic peak was obtained for the analysis of unstructured ONs (namely, L1, L2, and L3), whose MFE is predicted to be 0 kcal/mol and whose frequency of the MFE structure is >95%. This suggests that the inherent flexibility of unstacked single-stranded ONs impedes diastereomer resolution in HILIC. These findings point at intrinsic HOSs of ONs contributing to improved chromatographic resolution of diastereomers—in contrast with previous studies claiming that the formation of a secondary structure might be an obstacle for diastereomer resolution.23

They further offer a deeper understanding of the chromatographic behavior of PS-modified ONs. It has been previously reported that a single change in PS configuration can significantly affect the electronic structure of PS-modified ONs.11 Previous structural studies indicated a correlation between the Rp/Sp configuration and the position of the sulfur within the PS group in the case of a double-stranded ON.10 More specifically, it was found that when both strands exhibit the Sp configuration, the sulfur tends to be positioned on the outside of the helix, making it more exposed to interactions, whereas the Rp configuration results in the sulfur being buried inside the structure. This suggests that the diastereomer resolution observed here is derived from each (or clusters of) individual diastereomer species having slightly different available groups for hydrogen-bonding interactions with the stationary phase. Differences in the partitioning behavior of these isobaric species are also expected to play a role in separation, as these species present (at least slight) differences in hydrophilicity due to differences in solvent accessibility. Together, this may explain the enhanced diastereomer separation, as the HILIC conditions employed here (i.e., mobile phases with high ionic strength and low column temperature) favor rigid and compact ON HOSs (which favor hydrogen-bonding differentiation) as well as the water layer stabilization on the surface of the stationary phase (which favors partitioning).

Sugar Modifications, PS Number and Location, and Duplexes Also Modulate ON Diastereomer Separation in HILIC

2′-Substituent of the Ribose Sugar

In addition to exploring the impact of structural rigidity on the chromatographic resolution of diastereomers, we also investigated the influence of ribose 2′ substituents. Given that predictions using RNAfold are based on unmodified ribose sugars, and our model siRNA sequences are entirely substituted with alternating 2′-F and 2′-OMe groups, we decided to further assess the contribution of sugar modifications to diastereomer resolution.

Figure S5A shows the EICs obtained using 100 mM AA, 5 °C HILIC conditions for ONs with different ribose substituents at the 2′ position: m(as-202PS), f(as-202PS), and r(as-202PS) feature full 2′-OMe, 2′-F, or 2′-hydroxy (2′–OH) modification, respectively. First of all, it is important to note that the gradient window was adjusted to 55–70% H2O in 5 min for r(as-202PS) (vs the generic 35–50% H2O in 5 min for the other ONs), as the presence of 2’–OH sugars made r(as-202PS) too retained under the generic gradient conditions used here (data not shown). While about nine chromatographic peaks were partially separated for as-202PS with a selectivity of 1.3 between the first and last detected chromatographic peaks, m(as-202PS) and f(as-202PS) presented a profile with three major chromatographic peak clusters partially separated and a selectivity of 1.2 and 1.3, respectively. Surprisingly, only one sharp chromatographic peak was observed for r(as-202PS), suggesting that the presence of 2′–OH sugars interferes with the retention mechanism driving diastereomeric separation. Notably, diastereomer separation is facilitated when ONs are fully sugar-modified by either 2′-F or 2′-OMe groups (or combinations of both). As all these 2′ groups favor sugar pucker at the C3′-endo configuration,23 it is likely that the difference in diastereomer selectivity is related to the enhanced hydrogen-bonding capabilities and/or hydrophilicity of the 2′–OH-containing ON. Likely, r(as-202PS) gets heavily solvated in the immobilized water layer and/or forms excessive hydrogen-bonding interactions with the stationary phase, thus making separation of its diastereomers more challenging (if at all possible).

Number and Location of PS-Modified Linkages

Another variable influencing diastereomer separation is the number and location of the PS-modified linkages. The generic HILIC gradient was maintained (i.e., 35–50% H2O in 5 min), using mobile phases containing 100 mM AA and a column temperature of 5 °C. Figure S5B provides insights into the chromatographic separation of diastereomers of single-stranded ONs with varying numbers of PS modifications: two for as-2PS and as-101PS, six for as-6PS, and 19 for as-19PS.

First, the chromatograms of as-2PS and as-101PS were compared. The sequences differ only in the positioning of their two PS-modified linkages; as-2PS presents two PS modifications at the 5′ end of the sequence, whereas as-101PS presents one PS linkage at each sequence end. Baseline resolution of the four expected diastereomer peaks was not achieved for any of these compounds. However, the analysis of as-2PS showed a baseline separation of three distinct chromatographic peaks with the second eluting chromatographic peak showing a shoulder, likely attributed to the presence of two closely unresolved diastereomers. In the case of as-101PS, two major chromatographic peaks and a shoulder could be observed but with reduced selectivity between the first and last eluting peaks (1.1 as opposed to 1.2 for as-2PS). Enhanced diastereomer separation was observed when PS modifications were located at the same end of the sequence. Despite not being fully investigated here, and considering only the hairpin representation illustrated in Figure S5, it seems that as-2PS presents more accessible PS-modified sites (both PS modifications are located in the overhanging linear portion of the structure) when compared to as-101PS (one of the PS modifications is buried within the helix portion of the structure). These differences suggest that ONs with HOSs presenting more accessible groups for interaction are more likely to return higher diastereomeric separation. However, further experiments using, for instance, NMR are needed to confirm this speculation.

The complexity of the chromatographic profile increases when analyzing as-6PS, which possesses two PS-modified linkages at each end of the sequence and additional two internal nonconsecutive PS-modified linkages. Within this mixture of sixty-four diastereomers, we identified more than ten chromatographic peaks, resulting in a selectivity of 1.4 calculated between the first and last observed peaks. The number of detectable chromatographic peaks is limited by the increasing number of diastereomers, which exhibit more overlapping properties in terms of hydrophilicity and therefore the ability of the amide stationary phase/immobilized water layer to differentiate them. Analytical complexity is maximized with as-19PS, a fully phosphorothioated sequence mimicking a therapeutic ASO with regard to the phosphate backbone. Such a sequence presents more than 500,000 possible diastereomers. Unsurprisingly, a broad chromatographic peak, due to the high number of diastereomers, is observed for as-19PS, highlighting the limitations of current analytical techniques, including HILIC, for assessing diastereomeric distribution in full-length fully PS-modified ONs such as therapeutic ASOs.

Double-Stranded siRNAs with a Varying Number of PS Modifications

We also explored HILIC at low temperature (i.e., 5 °C) and high ionic strength (i.e., 100 mM AA) using the generic gradient profile (i.e., 35–50% H2O in 5 min) for the analytical characterization of double-stranded siRNAs, which typically incorporate four to eight PS-modified linkages.2 Two model siRNAs, namely, ds-4PS featuring four PS-modified linkages on one strand (two PS on each sequence end) and ds-8PS with four PS modifications on each strand (again, two PS on each sequence end), were studied here. Four and eight partially resolved chromatographic peaks were observed for ds-4PS and ds-8PS, respectively (Figure 4). As for single-stranded ONs with HOSs, double-stranded ONs are also highly ordered species in solution. Therefore, these results illustrate that HILIC is also able to preserve HOSs of double-stranded siRNAs, and as a consequence (at least partially), it resolves diastereomers of these complex species. Although far from the 16 and 256 expected chromatographic peaks for the diastereomers of these double-stranded ONs, it is important to highlight that HILIC can deliver partial diastereomer resolution of highly representative full-length modified double-stranded siRNAs of approximately 12 kDa. Such an outcome paves the way for monitoring batch-to-batch changes in the diastereomeric populations of these delicate species during siRNA development.

Figure 4.

Figure 4

Open in a new tab

Analysis of intact double-stranded siRNAs. (A) EICs of ds-4PS and ds-8PS using 100 mM as AA concentration in mobile phases and 5 °C as column temperature. Selectivities were calculated between the first and last detectable peaks (marked by an orange dot) in the apparent cluster of chromatographic peaks. For more information about the sequences, please refer to Table S1.

It is important to mention here that in addition to not being disrupted in the HILIC conditions employed in our study, the duplexed nature of siRNAs was also kept intact during both the ionization and ion-transfer processes under the ESI-MS parameters employed in our analyses. As seen in Figure S6A, only m/z values corresponding to the duplexes (ranging from [M – 7H]7– to [M – 4H]4–) are observed in the mass spectra extracted underneath the chromatographic peaks corresponding to the duplex species. Figure S6B highlights that the main charge states of the individual sense and antisense strands were not observed in these mass spectra. Additional tests revealed that this is true even when more traditional HILIC conditions are employed (e.g., 25 mM AA in the mobile phases and column temperature set to 30 °C; data not shown). Therefore, this highlights that the mass spectrometer employed here (i.e., Waters RDa) is capable of delivering the soft ionization and ion-transfer conditions needed for the intact characterization of double-stranded siRNAs. As described elsewhere,41 duplexes are commonly at least partially denatured during ESI-MS analysis. As a consequence, MS-based impurity profiling of nondenatured siRNAs is usually not feasible. Outcomes illustrated in Figure S6 suggest that the Waters RDa mass spectrometer (and instruments with similar features) is suitable for such a task. However, such a study is beyond the scope of this manuscript, thus not followed on.

Diastereomer Separation May be Suppressed under Alternative HILIC Conditions

HILIC conditions can be tuned to enhance the separation of diastereomers in ONs with HOSs. This is especially true for siRNAs due to the limited number of PS-modified linkages. However, challenges emerge when dealing with highly PS-modified ONs, such as ASOs, as chromatographic peak broadening becomes a complicating factor (Figure S5B). Therefore, as also implemented for IP-RPLC-based methods,17 strategies to suppress the diastereomer selectivity of ONs in HILIC are needed, particularly because studies using this chromatographic mode for ON analysis are barely emerging.

ACN is commonly used as the organic modifier in HILIC. It is generally preferred over MeOH because of its aprotic nature, facilitating the formation of a stable immobilized water layer on the stationary-phase surface. This water layer indeed plays a key role for hydrophilic partitioning in HILIC mode.42 However, previous reports on HILIC of ONs suggest that ionic interactions and hydrogen bonding with the ligand at the surface of the stationary phase play a more substantial role in the HILIC retention mechanism when more traditional HILIC conditions are employed in the analysis (e.g., 15–25 mM AA in the mobile phases; 30–40 °C for column temperature).36 Therefore, we explored the impact of MeOH as a mobile-phase organic modifier during HILIC analysis of ON diastereomers.

The EICs of as-202PS under HILIC conditions that enhance diastereomer separation (100 mM AA, 5 °C) using MeOH as the organic modifier are presented in Figure 5 (vs with ACN in Figure 2B). The gradient profile with ACN involved the generic 35–50% H2O range in 5 min at a flow rate of 0.4 mL/min, while the MeOH gradient profile was adjusted to 29–44% H2O over the same duration at a flow rate of 0.3 mL/min. Interestingly, the use of MeOH in the mobile phase resulted in a single narrowed chromatographic peak for as-202PS, contrary to about nine chromatographic peaks observed with ACN. Such an outcome may be attributed to the destabilization of the water layer on the stationary-phase surface in the presence of protic MeOH molecules. The protic nature of MeOH promotes hydrogen bonding among the solvent molecules, the ONs, and the amide stationary phase,43 thus suppressing molecular interactions that drive ON diastereomer separation. Another potential explanation for such a MeOH-driven diastereomer separation suppression is that the presence of MeOH may disrupt the ON HOS in solution. Therefore, to verify the latter, we compared melting curves for as-202PS, the hairpin-like sequence, diluted in the HILIC diluents composed of either 100 mM AA in 35:65 H2O/ACN or 100 mM AA in 35:65 H2O/MeOH (Figure S7). Very similar melting profiles were obtained for as-202PS in diluents composed of either ACN or MeOH as an organic modifier. Thus, the apparent suppression of diastereomer separation can likely not be attributed to MeOH-induced ON HOS denaturation. Rather, it is due to perturbation in the hydrogen-bonding interactions between ONs and the amide stationary phase and/or the immobilized water layer.

Figure 5.

Figure 5

Open in a new tab

EIC of as-202PS under favorable chromatographic conditions (100 mM AA, 5 °C) using mobile phases containing MeOH as the organic modifier.

It was beyond the scope of this study to evaluate the impact of MeOH-based HILIC mobile phases on the separation of ON full-length products and their related impurities (e.g., shortmers, longmers, oxidation products, etc.). Nonetheless, the previous observations underscore the significant contribution of hydrogen-bonding interactions between ONs and the stationary phase and immobilized water layer in the HILIC separation mechanism of ONs. More importantly, it reveals that diastereomer separation in HILIC is a result of diastereomers possessing distinct hydrogen-bonding capabilities with both stationary and mobile phases.

As shown earlier (Figures 1 and S1), parameters such as the temperature and ionic strength can further tune the diastereomer selectivity. Reducing the AA content and working at elevated column temperatures are straightforward approaches to disrupt HOSs and thus minimize/suppress diastereomer separation. Variation of the column pore size allows for further modulation of diastereomer separation. As shown in Figure S8A, under the same generic HILIC conditions (i.e., 100 mM AA, 5 °C, and 35–50% H2O in 5 min), diastereomer separation decreased when analyzing the as-202PS diastereomers using an amide phase with 130 Å rather than 300 Å pores. The observed decrease in selectivity may be attributed to the decreased accessible surface area with decreasing pore size, as well as the increased saturation capacity of the stationary phase, as explained elsewhere.44 In the case of the separation of closely related negatively charged compounds such as ON diastereomers, smaller pore sizes restrict ONs from accessing inner pore volumes, thus decreasing accessible surface areas for each diastereomer, compromising their separation.

We further suspected that the nature of the stationary phase and especially its chemistry would also significantly affect the diastereomer resolution. Outcomes of three alternative HILIC stationary phases were tested: a silica (BEH HILIC, 130 Å) column, a diol (Triart Diol-HILIC, 120 Å) column, and a zwitterionic (Poroshell HILIC-Z, 120 Å) column.

Again, the same generic HILIC conditions were employed (i.e., 100 mM AA, 5 °C, and 35–50% H2O in 5 min). As shown in Figure S8B, the amide stationary phase exhibited the highest diastereomer separation (which may partially be due to larger pore sizes), followed by the diol column with partial diastereomer separation, while the silica and zwitterionic columns seem to suppress diastereomer separation. This observation aligns with the notion that different phases possess varying hydrogen-bonding capacities.36 Overall, by combining the above different strategies, carefully choosing the appropriate column stationary phase and pore size, and tuning HILIC chromatographic conditions, such as mobile-phase ionic strength, column temperature, and organic modifier, one can suppress diastereomer separation when needed.

Conclusions

Our results underscored the mobile-phase ionic strength and column temperature to enhance the separation of diastereomers in HILIC. By elevating the ionic strength to 100 mM AA and reducing the column temperature to 5 °C, the separation of diastereomers was enhanced. Such conditions preserved the HOS of ONs, facilitating diastereomer separation. Confirmatory experiments using ESI-MS revealed a temperature-dependent CSD of ONs, with lower column temperatures favoring ON HOSs. Temperature-dependent findings were corroborated by UV melting curves, which confirmed that mobile phases with high AA levels and low column temperature maintain ON HOSs in solution.

As important as identifying HILIC conditions that favor diastereomer separation, this investigation marks a pioneering exploration into diastereomer separation of full-length ONs, unravelling the pivotal role of ON HOSs in diastereomer separation in HILIC. Under the evaluated conditions, compounds with a more rigid and limited number of HOSs returned higher diastereomer selectivity. Insights into the mechanisms driving the diastereomer separation were also gathered. Mobile phases with high ionic strength in combination with low column temperature seem to stabilize the immobilized water layer on the amide stationary-phase surface and minimize ionic interactions, suggesting that the diastereomer separation in HILIC is mostly driven by hydrogen-bonding interactions, through either adsorption or partitioning.

Additionally, we explored the behavior of various ON designs, including ONs with different 2′ modifications, different numbers and locations of PS-modified linkages, and single- and double-stranded siRNAs. While 2′-F and 2′-OMe modifications favored diastereomer separation both individually and in combination, 2′–OH groups disrupted it, likely because of enhanced hydrogen-bonding capabilities of the 2′–OH-containing ONs. Enhanced diastereomer separation was observed when PS modifications were located at more accessible/exposed locations within the ON structure, strongly suggesting that diastereomer separation is correlated to the exposure of PS-modified groups to molecular interactions with the stationary phase and/or immobilized water layer. However, further experiments are needed to better elucidate this. Importantly, HILIC was proven to be effective for partially separating diastereomers in double-stranded siRNAs, paving the way for monitoring batch-to-batch changes in diastereomer population during double-stranded siRNA development.

Apart from this, we propose strategies to suppress diastereomer separation, including using MeOH instead of ACN as the organic modifier, considering different HILIC column chemistries and/or pore sizes, and fine-tuning column temperature and mobile-phase ionic strength. Combining these strategies allowed total suppression of diastereomer separation through modulation of the HILIC mechanism and/or ON conformation, offering a new degree of flexibility regarding specific application requirements. Analysis of fully PS-modified ONs, which possess hundreds of thousands of diastereomers, or impurity analysis of PS-modified ONs would benefit from the suppression of diastereomer resolution, which often hampers the analysis.

Acknowledgments

The authors thank Waters corporation, in especial, Jakob Rajgard (Principal Chemistry Account Manager), for sponsoring this project with some of the chromatographic columns evaluated here.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.analchem.4c01384.

  • Information about ON sequences, synthesis, and secondary structure predictions; effect of column temperature and mobile-phase ionic strength on HILIC analysis and effect of column temperature on CSD; base pair probabilities of as-202PS; impact of HOSs on diastereomer separation; base pair probabilities of as-202PS; impact of the 2′-substituent of the sugar ring and number and position of PS linkages on diastereomer separation; ESI-MS spectra for double-stranded siRNAs and their corresponding single-stranded sequences; and impact of column pore size and alternative column stationary-phase chemistry on diastereomer separation (DOCX)

Author Contributions

All authors have given approval to the final version of the manuscript.

The authors declare no competing financial interest.

Supplementary Material

ac4c01384_si_001.docx (22.6MB, docx)

References

  1. Crooke S. T.; Witztum J. L.; Bennett C. F.; Baker B. F. RNA-Targeted Therapeutics. Cell Metab. 2018, 27 (4), 714–739. 10.1016/j.cmet.2018.03.004. [DOI] [PubMed] [Google Scholar]
  2. Egli M.; Manoharan M. Chemistry, Structure and Function of Approved Oligonucleotide Therapeutics. Nucleic Acids Res. 2023, 51 (6), 2529–2573. 10.1093/nar/gkad067. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Dowdy S. F. Overcoming Cellular Barriers for RNA Therapeutics. Nat. Biotechnol. 2017, 35 (3), 222–229. 10.1038/nbt.3802. [DOI] [PubMed] [Google Scholar]
  4. Khvorova A.; Watts J. K. The Chemical Evolution of Oligonucleotide Therapies of Clinical Utility. Nat. Biotechnol. 2017, 35 (3), 238–248. 10.1038/nbt.3765. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Mullard A. 2023 FDA Approvals. Nat. Rev. Drug Discovery 2024, 23, 88–95. 10.1038/d41573-024-00001-x. [DOI] [PubMed] [Google Scholar]
  6. Hyjek-Składanowska M.; Vickers T. A.; Napiórkowska A.; Anderson B. A.; Tanowitz M.; Crooke S. T.; Liang X.; Seth P. P.; Nowotny M. Origins of the Increased Affinity of Phosphorothioate-Modified Therapeutic Nucleic Acids for Proteins. J. Am. Chem. Soc. 2020, 142 (16), 7456–7468. 10.1021/jacs.9b13524. [DOI] [PubMed] [Google Scholar]
  7. Li M.; Lightfoot H. L.; Halloy F.; Malinowska A. L.; Berk C.; Behera A.; Schümperli D.; Hall J. Synthesis and Cellular Activity of Stereochemically-Pure 2′-O-(2-Methoxyethyl)-Phosphorothioate Oligonucleotides. Chem. Commun. 2017, 53 (3), 541–544. 10.1039/C6CC08473G. [DOI] [PubMed] [Google Scholar]
  8. Wei X. Coupling Activators for the Oligonucleotide Synthesis via Phosphoramidite Approach. Tetrahedron 2013, 69 (18), 3615–3637. 10.1016/j.tet.2013.03.001. [DOI] [Google Scholar]
  9. Hall J. Future Directions for Medicinal Chemistry in the Field of Oligonucleotide Therapeutics. RNA 2023, 29 (4), 423–433. 10.1261/rna.079511.122. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Lan W.; Hu Z.; Shen J.; Wang C.; Jiang F.; Liu H.; Long D.; Liu M.; Cao C. Structural Investigation into Physiological DNA Phosphorothioate Modification. Sci. Rep. 2016, 6 (1), 25737. 10.1038/srep25737. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Bohr H. G.; Shim I.; Stein C.; Ørum H.; Hansen H. F.; Koch T. Electronic Structures of LNA Phosphorothioate Oligonucleotides. Mol. Ther. Nucleic Acids 2017, 8, 428–441. 10.1016/j.omtn.2017.05.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Frey P. A.; Sammons R. D. Bond Order and Charge Localization in Nucleoside Phosphorothioates. Science 1985, 228 (4699), 541–545. 10.1126/science.2984773. [DOI] [PubMed] [Google Scholar]
  13. Arrico L.; Stolfi C.; Marafini I.; Monteleone G.; Demartis S.; Bellinvia S.; Viti F.; McNulty M.; Cabani I.; Falezza A.; Di Bari L. Inhomogeneous Diastereomeric Composition of Mongersen Antisense Phosphorothioate Oligonucleotide Preparations and Related Pharmacological Activity Impairment. Nucleic Acid Ther. 2022, 32 (4), 312–320. 10.1089/nat.2021.0089. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Capaldi D.; Akhtar N.; Atherton T.; Benstead D.; Charaf A.; De Vijlder T.; Heatherington C.; Hoernschemeyer J.; Jiang H.; Rieder U.; Ring F.; Peter R.; Stolee J. A.; Wechselberger R. Strategies for Identity Testing of Therapeutic Oligonucleotide Drug Substances and Drug Products. Nucleic Acid Ther. 2020, 30 (5), 249–264. 10.1089/nat.2020.0878. [DOI] [PubMed] [Google Scholar]
  15. Enmark M.; Rova M.; Samuelsson J.; Örnskov E.; Schweikart F.; Fornstedt T. Investigation of Factors Influencing the Separation of Diastereomers of Phosphorothioated Oligonucleotides. Anal. Bioanal. Chem. 2019, 411 (15), 3383–3394. 10.1007/s00216-019-01813-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Roussis S. G.; Cedillo I.; Rentel C. Characterizing the Diastereoisomeric Distribution of Phosphorothioate Oligonucleotides by Metal Ion Complexation Chromatography, In-Series Reversed Phase-Strong Anion Exchange Chromatography, and 31P NMR. Anal. Chem. 2021, 93 (48), 16035–16042. 10.1021/acs.analchem.1c03593. [DOI] [PubMed] [Google Scholar]
  17. Chen T.; Tang S.; Fu Y.; Napolitano J. G.; Zhang K. Analytical Techniques for Characterizing Diastereomers of Phosphorothioated Oligonucleotides. J. Chromatogr. A 2022, 1678, 463349. 10.1016/j.chroma.2022.463349. [DOI] [PubMed] [Google Scholar]
  18. Stec W. J.; Zon G.; Uznaǹski B. Reversed-Phase High-Performance Liquid Chromatographic Separation of Diastereomeric Phosphorothioate Analogues of Oligodeoxyribonucleotides and Other Backbone-Modified Congeners of Dna. J. Chromatogr. A 1985, 326, 263–280. 10.1016/S0021-9673(01)87452-5. [DOI] [PubMed] [Google Scholar]
  19. Li L.; Leone T.; Foley J. P.; Welch C. J. Separation of Small Interfering RNA Stereoisomers Using Reversed-Phase Ion-Pairing Chromatography. J. Chromatogr. A 2017, 1500, 84–88. 10.1016/j.chroma.2017.04.008. [DOI] [PubMed] [Google Scholar]
  20. Enmark M.; Harun S.; Samuelsson J.; Örnskov E.; Thunberg L.; Dahlén A.; Fornstedt T. Selectivity Limits of and Opportunities for Ion Pair Chromatographic Separation of Oligonucleotides. J. Chromatogr. A 2021, 1651, 462269. 10.1016/j.chroma.2021.462269. [DOI] [PubMed] [Google Scholar]
  21. Demelenne A.; Nys G.; Nix C.; Fjeldsted J. C.; Crommen J.; Fillet M. Separation of Phosphorothioated Oligonucleotide Diastereomers Using Multiplexed Drift Tube Ion Mobility Mass Spectrometry. Anal. Chim. Acta 2022, 1191, 339297. 10.1016/j.aca.2021.339297. [DOI] [PubMed] [Google Scholar]
  22. Ghassemi K M.; Demelenne A.; Crommen J.; Servais A.-C.; Fillet M. Improvement of Chemo- and Stereoselectivity for Phosphorothioate Oligonucleotides in Capillary Electrophoresis by Addition of Cyclodextrins. J. Chromatogr. A 2022, 1676, 463270. 10.1016/j.chroma.2022.463270. [DOI] [PubMed] [Google Scholar]
  23. Goyon A.; Blevins M. S.; Napolitano J. G.; Nguyen D.; Goel M.; Scott B.; Wang J.; Koenig S. G.; Chen T.; Zhang K. Characterization of Antisense Oligonucleotide and Guide Ribonucleic Acid Diastereomers by Hydrophilic Interaction Liquid Chromatography Coupled to Mass Spectrometry. J. Chromatogr. A 2023, 1708, 464327. 10.1016/j.chroma.2023.464327. [DOI] [PubMed] [Google Scholar]
  24. Kanavarioti A. HPLC Methods for Purity Evaluation of Man-Made Single-Stranded RNAs. Sci. Rep. 2019, 9 (1), 1019. 10.1038/s41598-018-37642-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Donegan M.; Nguyen J. M.; Gilar M. Effect of Ion-Pairing Reagent Hydrophobicity on Liquid Chromatography and Mass Spectrometry Analysis of Oligonucleotides. J. Chromatogr. A 2022, 1666, 462860. 10.1016/j.chroma.2022.462860. [DOI] [PubMed] [Google Scholar]
  26. Enmark M.; Bagge J.; Samuelsson J.; Thunberg L.; Örnskov E.; Leek H.; Limé F.; Fornstedt T. Analytical and Preparative Separation of Phosphorothioated Oligonucleotides: Columns and Ion-Pair Reagents. Anal. Bioanal. Chem. 2020, 412, 299–309. 10.1007/s00216-019-02236-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Kadlecová Z.; Kalíková K.; Tesařová E.; Gilar M. Phosphorothioate Oligonucleotides Separation in Ion-Pairing Reversed-Phase Liquid Chromatography: Effect of Temperature. J. Chromatogr. A 2022, 1681, 463473. 10.1016/j.chroma.2022.463473. [DOI] [PubMed] [Google Scholar]
  28. Liu A.; Cheng M.; Zhou Y.; Deng P. Bioanalysis of Oligonucleotide by LC-MS: Effects of Ion Pairing Regents and Recent Advances in Ion-Pairing-Free Analytical Strategies. IJMS 2022, 23 (24), 15474. 10.3390/ijms232415474. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Kadlecová Z.; Kalíková K.; Tesařová E.; Gilar M. Phosphorothioate Oligonucleotides Separation in Ion-Pairing Reversed-Phase Liquid Chromatography: Effect of Ion-Pairing System. J. Chromatogr. A 2022, 1676, 463201. 10.1016/j.chroma.2022.463201. [DOI] [PubMed] [Google Scholar]
  30. Goyon A.; Zhang K. Characterization of Antisense Oligonucleotide Impurities by Ion-Pairing Reversed-Phase and Anion Exchange Chromatography Coupled to Hydrophilic Interaction Liquid Chromatography/Mass Spectrometry Using a Versatile Two-Dimensional Liquid Chromatography Setup. Anal. Chem. 2020, 92 (8), 5944–5951. 10.1021/acs.analchem.0c00114. [DOI] [PubMed] [Google Scholar]
  31. Alpert A. J. Electrostatic Repulsion Hydrophilic Interaction Chromatography for Isocratic Separation of Charged Solutes and Selective Isolation of Phosphopeptides. Anal. Chem. 2008, 80 (1), 62–76. 10.1021/ac070997p. [DOI] [PubMed] [Google Scholar]
  32. Alpert A. J. Effect of Salts on Retention in Hydrophilic Interaction Chromatography. J. Chromatogr. A 2018, 1538, 45–53. 10.1016/j.chroma.2018.01.038. [DOI] [PubMed] [Google Scholar]
  33. Hofmeister F. On the understanding of the effects of salts. Arch. Exp. Pathol. Pharmakol. 1888, 24 (4–5), 247–260. 10.1007/BF01918191. [DOI] [Google Scholar]
  34. Xia Z.; DeGrandchamp J. B.; Williams E. R. Native Mass Spectrometry beyond Ammonium Acetate: Effects of Nonvolatile Salts on Protein Stability and Structure. Analyst 2019, 144 (8), 2565–2573. 10.1039/C9AN00266A. [DOI] [PubMed] [Google Scholar]
  35. McCalley D. V. Understanding and Manipulating the Separation in Hydrophilic Interaction Liquid Chromatography. J. Chromatogr. A 2017, 1523, 49–71. 10.1016/j.chroma.2017.06.026. [DOI] [PubMed] [Google Scholar]
  36. Lardeux H.; Guillarme D.; D’Atri V. Comprehensive Evaluation of Zwitterionic Hydrophilic Liquid Chromatography Stationary Phases for Oligonucleotide Characterization. J. Chromatogr. A 2023, 1690, 463785. 10.1016/j.chroma.2023.463785. [DOI] [PubMed] [Google Scholar]
  37. Largy E.; König A.; Ghosh A.; Ghosh D.; Benabou S.; Rosu F.; Gabelica V. Mass Spectrometry of Nucleic Acid Noncovalent Complexes. Chem. Rev. 2022, 122 (8), 7720–7839. 10.1021/acs.chemrev.1c00386. [DOI] [PubMed] [Google Scholar]
  38. D’Abramo M.; Castellazzi C. L.; Orozco M.; Amadei A. On the Nature of DNA Hyperchromic Effect. J. Phys. Chem. B 2013, 117 (29), 8697–8704. 10.1021/jp403369k. [DOI] [PubMed] [Google Scholar]
  39. Gruenwedel D. W.; Hsu C. Salt Effects on the Denaturation of DNA. Biopolymers 1969, 7 (4), 557–570. 10.1002/bip.1969.360070412. [DOI] [PubMed] [Google Scholar]
  40. Guo X. Secondary Structural Characterization of Oligonucleotide Strands Using Electrospray Ionization Mass Spectrometry. Nucleic Acids Res. 2005, 33 (11), 3659–3666. 10.1093/nar/gki671. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Huang M.; Xu X.; Qiu H.; Li N. Analytical Characterization of DNA and RNA Oligonucleotides by Hydrophilic Interaction Liquid Chromatography-Tandem Mass Spectrometry. J. Chromatogr. A 2021, 1648, 462184. 10.1016/j.chroma.2021.462184. [DOI] [PubMed] [Google Scholar]
  42. Dinh N. P.; Jonsson T.; Irgum K. Water Uptake on Polar Stationary Phases under Conditions for Hydrophilic Interaction Chromatography and Its Relation to Solute Retention. J. Chromatogr. A 2013, 1320, 33–47. 10.1016/j.chroma.2013.09.061. [DOI] [PubMed] [Google Scholar]
  43. Subirats X.; Casanovas L.; Redón L.; Rosés M. Effect of the Solvent on the Chromatographic Selectivity in Reversed-Phase and HILIC. Adv. Sample Prep. 2023, 6, 100063. 10.1016/j.sampre.2023.100063. [DOI] [Google Scholar]
  44. Bagge J.; Enmark M.; Leśko M.; Limé F.; Fornstedt T.; Samuelsson J. Impact of Stationary-Phase Pore Size on Chromatographic Performance Using Oligonucleotide Separation as a Model. J. Chromatogr. A 2020, 1634, 461653. 10.1016/j.chroma.2020.461653. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

ac4c01384_si_001.docx (22.6MB, docx)

Articles from Analytical Chemistry are provided here courtesy of American Chemical Society

RESOURCES