The effect of anionic surfactant on poliovirus particles during capillary electrophoresis

Journal of Pharmaceutical and Biomedical Analysis 71 (2012) 79–88

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The effect of anionic surfactant on poliovirus particles during capillary electrophoresis Iuliana Oita a , Hadewych Halewyck b,c , Bert Thys b,c , Bart Rombaut b,c , Yvan Vander Heyden a,∗ a Vrije Universiteit Brussel-VUB, Center for Pharmaceutical Research-CePhaR, Department of Analytical Chemistry and Pharmaceutical Technology, Laarbeeklaan 103, B-1090, Brussels, Belgium b Vrije Universiteit Brussel-VUB, Center for Pharmaceutical Research-CePhaR, Department of Pharmaceutical Biotechnology & Molecular Biology, Laarbeeklaan 103, B-1090, Brussels, Belgium c Vrije Universiteit Brussel-VUB, Center for Pharmaceutical Research-CePhaR, Center for Neurosciences, Laarbeeklaan 103, B-1090, Brussels, Belgium

a r t i c l e

i n f o

Article history: Received 10 May 2012 Received in revised form 26 July 2012 Accepted 30 July 2012 Available online 4 August 2012 Keywords: Capillary electrophoresis Poliovirus SDS Capsid dissociation Stability

a b s t r a c t Because of its essential role in SDS-PAGE, sodium dodecylsulphate (SDS) is generally associated with protein denaturation. However, for SDS-PAGE, proteins are linearized in the presence of SDS, following the exposure to high temperatures and reducing agents. In comparison, the conditions employed during a capillary electrophoretic (CE) separation involve only a limited exposure to SDS, at much lower temperatures. As the outer surface of the non-enveloped viruses consists of proteins, virus interaction with SDS can be judged from the perspective of SDS–protein interaction. Several studies have indicated that proteins have a different susceptibility to SDS, depending on their secondary structure and number of subunits. Therefore it is not straightforward to estimate what should be expected when intact polioviruses and subviral particles obtained by thermal conversion of the poliovirions, are exposed to SDS during CE separation. In this study it is shown that, during CE separations, SDS has no effect on the integrity of the poliovirion, but the presence of SDS in the separation system influences the poliovirus peak height and shape. The implication of SDS in the CE separation of poliovirus is discussed in detail. On the contrary, the proteinaceous subviral particles, such as the empty capsids, are less stable in the presence of SDS during the CE separation, and aggregates between the individual poliovirus capsid proteins and SDS are formed. Finally, we have proposed an alternative separation approach, involving an SDS gradient, for an improved separation of the subviral particles. © 2012 Elsevier B.V. All rights reserved.

1. Introduction Sodium dodecylsulphate is a common chemical in most biochemistry, forensics and molecular biology labs. It is an essential reagent for SDS-PAGE, an electrophoretic technique used for protein separation according to size. The role in SDS-PAGE established a fame of protein denaturant for SDS. Besides this, SDS is also known to inactivate enzymes, to disrupt biological systems, such as cells, viruses or membranes, as it has powerful dissociation and solubilization properties arising from its amphiphilic structure [1]. The amphiphilic properties of SDS were successfully exploited in analytical sciences for selectivity manipulation in modern separation techniques, such as liquid chromatography or capillary electrophoresis [2]. When included above its critical micellar concentration in the composition of the CE buffers, the formed SDS micelles create a pseudostationary phase and the separation

∗ Corresponding author. E-mail address: [email protected] (Y.V. Heyden). 0731-7085/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jpba.2012.07.033

principles of both electrophoresis and chromatography are combined in a method called micellar electrokinetic chromatography [3]. In bioanalytical CE applications, SDS is a valuable additive to prevent or minimize protein adsorption to the capillary wall [4]. In well defined conditions, the inclusion of SDS in the CE separation system offers the prerequisites for intra-column signal enhancement. For instance, the involvement in an isotachophoretic process has been described for high SDS concentrations in the injection plug [5,6]. Sweeping, i.e. a signal enhancing mechanism involving the presence of a pseudostationary phase, consists in picking and accumulating analyte molecules by the SDS micelles that enter the sample zone [7]. A feature of the signal enhancing mechanisms, involving SDS, refers to the importance of the relative conductivities of sample and separation buffer, as described in detail elsewhere [6–10]. SDS was also found indispensable to obtain reproducible CE separation of human rhinovirus [11–14], poliovirus [15–17], bacteriophage T5 [18], or rotavirus-like particles [19]. The concentration of SDS in the separation buffer appears to be a compromise between the separation needs and the virus stability. However,

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from a virological point of view, a limitation of SDS concentration would be desired as the interaction between SDS and viruses might harm the integrity of the virus particle [20]. On the contrary, from a separation science point of view, larger SDS concentrations might increase the mobility of the virus particles, similarly to what was observed for CE separations of nanoparticles [22–25]. These contradictory facts indicate the need for a thorough study of the interactions between virus particles and SDS during CE separations. The virus particles mobility is mainly attributed to the ionization of outer chargeable groups of the proteinaceous capsid [26]. Therefore, the effect of SDS on the viruses during the CE separations can be better understood from the perspective of SDS–protein interactions. Although known as denaturant, it was showed that SDS does not completely unfold proteins in aqueous conditions, but it alters the helical structure of many proteins [1], or it forms rod-shape complexes with some proteins [21]. In SDS-PAGE, the proteins are intentionally completely denaturated and unfolded following boiling at slightly basic pH (∼8) in 2% (∼70 mM) SDS in the presence of a reducing agent [27]. Following this treatment, aggregates with similar charge/mass ratio are obtained [28]. Less is however known about the evolution of the protein structure after transient exposure to SDS, such as during a CE separation. The formation of non-native conformations of the proteins following temporary contact with aqueous SDS solutions, and a differentiation following limited SDS exposure was discussed in few reports [29,30]. SDS–protein interaction is a complex process and the mechanism is not completely elucidated yet. It seems to be influenced by pH, temperature and other experimental conditions. It appears that SDS induces and/or stabilizes secondary structure, possibly by hydrophobic interactions [31] or salt bridges between the sulphate group and the positively charged amino acids [32]. However, the strongest evidences point for an electrostatic interaction with positively charged residues of the protein, accompanied by binding to hydrophobic regions [28]. Additionally, Jones [28] indicated that the SDS presence at even sub-molar levels already initiates protein denaturation, while Gudiksen et al. [29] demonstrated SDS aggregate formations with several proteins, below critical micellar concentrations. A protein’s susceptibility to SDS is related to the number of positively charged amino acids on the protein surface and the conformation, i.e. the more relaxed the structure, the more susceptible [33]. Thus it seems that proteins packed mainly in a ␤-sheet structure are more resistant to denaturation. For instance, ferritin, a protein with a complex quaternary structure formed by the association of 24-mers, involved in multiple interactions, undergoes substantial denaturation only at higher SDS concentrations unlike simpler proteins [30]. Similarly, bovine superoxide dismutase (SOD), a protein containing close packing of hydrophobic interfaces in an 8-stranded ␤-vessel structure, thus with a structure of low flexibility, offers only a low SDS access to the binding sites. Consequently, SOD has excellent stability and higher resistance to SDS action [30]. Additionally, the lack of sulphide bridges increases the sensitivity to SDS, but packing seems more important for the protein susceptibility to SDS [28]. In viral capsids, the proteins are kept together by a combination of non-covalent interactions, thermodynamically extremely stable and comparable to subunit interfaces in protein–protein complexes and homodimers [34]. As the size of the virus particles, for which CE separations have already been developed, are within 30–100 nm range, additional insight to SDS–virus interactions can be also obtained extrapolating the interactions between SDS and spherical nanoparticles. In the latter case, SDS adsorbs onto the surface of the particles, imparts negative charges and improves the colloidal stability, inducing electrostatic repulsion between particles. The extent of adsorption depends on adsorption density, packing orientation and the nature

of the particle charges [22]. Recently, it was demonstrated that SDS adsorbs even on negatively charged silica particles despite the equal sign, creating supercharged systems [35]. Similar SDS properties are exploited for re-dispersion of pellets of viral particles [20]. However, there are a few major differences between the nanoparticles and viral particles regarding the surface exposure. While for nanoparticles the outer surface is generally smooth, the capsid of virus particles has spatial features, such as the receptor binding sites. Additionally, while most of the nanoparticles are generally impermeable to the liquid flow during separations, the viral capsid might be semipermeable or even exhibit pores [36]. These properties of the viral capsids influence the electrophoretic behavior of virus particles, as described in detail in [37,38]. Poliovirus, probably the most studied picornavirus, is extremely simple. The mature particle consists of a protein shell, with an outer diameter of 27–30 nm, surrounding the naked RNA of 7.5–8 kb [39,40]. The capsid is build up only from four different structural proteins, i.e. viral proteins 1–4 (VP1, VP2, VP3, and VP4). The native VP1, VP2 and VP3 have molecular weights of 33.5, 26.4 and 30.0 kDa, respectively, and share the same folding pattern, a “jelly-roll ␤-barrel”, described in detail in [41]. The particular folding and arrangement in the structural proteins allows forming a dense and rigid capsid that explains the low capsid permeability and the chemical stability of the poliovirus. The virus is insensitive to lipid solvents, and is relatively resistant to many common laboratory disinfectants. However, formaldehyde, glutaraldehyde, strong acids and bases, or sodium hypochlorite inactivate it. It is believed that inactivation occurs as a result of chemical modification of the virion [42]. Poliovirus particle intermediates have been observed in vivo during cell entry and morphogenesis [40–44]. These intermediates are generically referred to as subviral particles, i.e. “incomplete” viral particles or mixtures of components of the viral particles. As the isolation of these intermediates is not straightforward, heating the poliovirus suspensions between 50 and 56 ◦ C was found to be a reasonable alternative to obtain subviral particles in vitro. It was found that VP4 is externalized after heating, triggering a structural rearrangement of the viral capsids. Finally, RNA is released from the capsid and particles sedimenting at 80S are formed, corresponding to empty poliovirus capsids [40–44]. Literature indicated that the empty capsids obtained in vitro, are indistinguishable from the final product of infection and were used as models for cryo-electron microscopy studies of the RNA release [44]. Koch and Koch [41] have showed that the poliovirus is stable in SDS solutions up to 1% (about 35 mM). We believe that the poliovirus is still intact within our experimental conditions since the concentration of SDS in the separation buffers does not exceed 25 mM. Additionally, the integrity of the poliovirus was demonstrated based on an affinity reaction with N-neutralizing antibodies [16]. In this paper, the interaction occurring between SDS and both poliovirus and in vitro obtained subviral particles, during the CE separations, was studied. The need for SDS in the separation buffer and the SDS impact on the separation between poliovirus and subviral particles were considered.

2. Materials and methods 2.1. Chemicals and reagents Sodium dodecyl sulphate (SDS, 98.5%) was purchased from Sigma (Steinheim, Germany), o-phthalic acid (puriss, >99.5%) from Fluka (Steinheim, Germany), and sodium hydroxide (NaOH) 1 M from Fisher Scientific (Leicestershire, UK). All other chemicals were purchased from Merck (Darmstadt, Germany). All chemicals were used as bought.

I. Oita et al. / Journal of Pharmaceutical and Biomedical Analysis 71 (2012) 79–88

2.2. Solutions

0.40

Solutions were prepared using ultra pure water, produced in-house by a Nanopure Diamond water purification system (Barnstead, Dubuque, Iowa). Buffers were prepared by dissolving the necessary amounts of chemicals in water and adjusting the pH before bringing to volume. The pH measurements were performed using an Orion 520 A pH-meter (Orion Research, Boston, MA). Solutions were degassed by ultrasonication for 20 min in an ultrasonic bath (Branson Ultrasonic Corporation, CT) and filtered through a polypropylene membrane with 0.2 ␮m pore size (VWR, Leuven, Belgium) prior to CE analysis.

0.35

2.3. Capillary electrophoresis CE experiments were performed on a Beckman P/ACE MDQ CE system (Fullerton, CA). Fused-silica capillaries with a total length of 50.2 cm (effective length 40 cm) and inner diameters of 50 ␮m were purchased from Composite Metal Services (Ilkley, UK). A pressure pulse was applied to inject a sample plug, the latter expressed further as the percentage of the capillary length to the detection window. The necessary pressures and injection times needed for a given sample plug were estimated using Beckman “CE Expert” software. Separations were performed at 26 ◦ C in 100 mM borate buffer pH 8.3, containing 25 mM SDS, as separation buffer and applying 10 kV voltages. New capillaries were conditioned by flushing with 100 mM hydrochloric acid, followed by water, 1 M sodium hydroxide, and again water, each time for 10 min, using 20 psi pressure. Prior to each measurement the capillary was rinsed with 0.1 M NaOH, water, and separation buffer for 2 min each, applying 14 psi pressure. To all samples, 0.005% dimethylformamide, as EOF marker, and 100 ␮g/ml o-phthalic acid, as internal standard, were added [45]. 2.4. Samples Several batches of the Family Picornaviridae, genus Enterovirus, species Poliovirus, Sabin strain (type 1) (poliovirus) were grown, collected and purified as described in [43]. Compared to the Mahoney strain, regularly used for inactivated poliovirus vaccine (IPV) manufacturing, the Sabin strain has attenuated infective properties, and was used for biosafety considerations. Optionally, the virus can be purified by a sucrose gradient ultracentrifugation and samples with very high viral purity are then obtained. The viral concentration was estimated spectrophotometrically [46]. Two poliovirus batches were used: OC09 and OC10. Sample OC09 was used without sucrose gradient ultracentrifugation. The poliovirus concentration of sample OC09 was estimated about 530 ␮g/ml. Before injection, all samples containing OC09 were diluted 5 times with water. Sample OC10 was first purified by sucrose gradient ultracentrifugation and the poliovirus concentration was estimated to be about 245 ␮g/ml. Poliovirus samples were ultrafiltrated at a 30,000 Da molecular weight cut-off (UF 30.000) using Amicon Ultra-4 devices (Millipore, Bedford, MA, USA) with an Ultracel3 membrane against 5 mM borate buffer pH 8.3, for buffer exchange purposes. The upper ultrafiltrate was further concentrated using UF 300.000, Vivaspin 6 centrifugal concentrators with polyethersulphone membrane (Sartorius-Stedim Biotech, Goettigen, Germany) [47]. Ultrafiltration was performed using a Harrier 18/80 R refrigerated centrifuge (MSE, London, UK). After purification and concentration of sample OC10, a poliovirus concentration about 2500 ␮g/ml was obtained. Before CE analysis, the purified and concentrated poliovirus suspension was diluted 15 times using 20 mM borate buffer pH 8.3.

0.30

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Subviral parcles A

AU

0.25

0.20

A

0.15

0.10

B

(2)

B (3)

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6.5

7.0

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9.5 10.0 10.5 11.0 11.5 12.0 12.5 13.0

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Fig. 1. Poliovirus and subviral particles/contaminant electromigration. (1) Sample OC09 after 5 times dilution in water, poliovirus ∼100 ␮g/ml. (2) Poliovirions–empty capsids mixture, poliovirus ∼160 ␮g/ml and empty capsids ∼500 ␮g/ml. (3) Empty capsids obtained using a poliovirus suspension ∼500 ␮g/ml. (4) Highly purified poliovirus suspension, poliovirus ∼160 ␮g/ml. Separation conditions: uncoated fused silica capillary 40 cm effective length, 50 ␮m internal diameter; 100 mM borate buffer pH 8.3 with 25 mM SDS; 10 kV; detection wavelength 205 nm; sample injected as 12% plug.

Empty capsids were obtained in vitro by heating the purified and concentrated OC10 at 56 ◦ C for 10 min after diluting 5 times with 20 mM borate buffer pH 8.3. Empty capsids were mixed with purified poliovirions to obtain a final ratio of 3:1 (v/v) and a poliovirions–empty capsids mixture was obtained. The matrix of this mixture contained only about 20 mM borate buffer besides poliovirus and empty capsids. The empty capsids were further denaturated by exposing the purified and concentrated OC10 to 100 ◦ C for 10 min and by SDS denaturation. For the SDS denaturation, the poliovirions were incubated with 1% SDS and ␤-mercaptoethanol at 96 ◦ C. Further, buffer exchange was performed as described above to remove the excess of ␤-mercaptoethanol and the upper resulted from ultrafiltration was injected in the CE. 3. Results and discussions 3.1. Characterization of the poliovirus samples During earlier experiments, aiming developing and optimizing the analytical procedure, only a separation buffer containing SDS was able to separate properly the poliovirus from the EOF marker peak [15–17]. Recently, the impact of the sample composition on the CE separations of poliovirus was demonstrated [47] and the results of that study triggered a re-assessment of the need for SDS in the separation buffer. For the current study, two types of samples were investigated, with different purity levels according to their A260 /A280 ratio, a well established virological approach for fast purity estimation. Using this approach, a pure poliovirus suspension is considered to have an A260 /A280 ratio of 1.70 [46]. The electrophoretic profile of the two samples investigated in the current study can be compared in Fig. 1 (trace 1 vs. trace 2). The first sample, OC09, was not purified using sucrose gradient centrifugation and it was a relatively “dirty” sample according to its A260 /A280 ratio of 1.35. The low A260 /A280 ratio is explained

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by an excess of protein entities of unknown nature, occurring from growth and purification steps. Consequently, a group of 8–9 peaks corresponding to contaminants or subviral particles with a migration time range of 9–11 min can be observed on the electropherogram (Fig. 1-trace 1). The second sample was a poliovirions–empty capsids mixture suspended in borate buffer pH 8.3, free from growth and purification contaminants. This mixture was obtained in several steps. Initially, buffer exchange and concentration, as described in [47], were applied to a sucrose gradient purified poliovirus sample. A suspension containing about 2500 ␮g/ml poliovirions in 5 mM borate buffer pH 8.3 was obtained, with a A260 /A280 ratio of 1.67. Subsequently, the poliovirions were converted to empty capsids. Finally, these empty capsids and poliovirions were combined to obtain a mixture containing about 160 ␮g/ml poliovirions and 500 ␮g/ml empty capsids. The electropherogram of the mixture is presented in Fig. 1-trace 2, that of the empty capsids in Fig. 1trace 3 and of the poliovirions suspension in Fig. 1-trace 4. The poliovirus peak is well separated from the empty capsids peaks and the same peaks can be identified in the electropherogram of the mixture and those corresponding to poliovirus particles and empty capsids, respectively. The electrophoretic separation of the empty capsids was shown to be more complex than that of poliovirus [16]. It should be taken into account that cryo-electron microscopy studies characterizing empty particles (80S) demonstrated that at least three different kinds of particles, classified based on RNA density inside their capsid, are formed depending on the length of the exposure time to 56 ◦ C [44,48]. In general, it is assumed that the longer the thermal exposure time, the emptier the capsids [44]. Nevertheless, less than 10% difference in the sample profile was found when 20 min of thermal exposure were used instead of 4 min [44]. Still, when poliovirions were heated only 10 min the particles have been caught in the act of releasing their viral RNA [44]. Therefore, as the samples prepared for this study were exposed only 10 min to 56 ◦ C, prior to CE injection the sample contained empty viral capsids, but the RNA was probably still attached. Very small peaks compared to the poliovirus peak were obtained in Ref. [16]. Therefore, in this study, the empty capsids were injected undiluted and in 12% plugs in order to ensure the highest signal possible within the experimental conditions. In Fig. 1-trace 3, a group of peaks of nature discussed in detail further was observed around 9–11 min. The position of these peaks is in agreement to what was previously published by Halewyck et al. [16], but the shape and number is slightly different. However, there is a number of experimental differences between the current study and the one in [16], i.e. the empty capsids were now obtained after only 10 min of thermal exposure vs. 20 min, using a more concentrated sample with a matrix of 20 mM borate buffer pH 8.3 vs. 20% sucrose–20 mM phosphate buffer pH 7.2, and injecting samples as 12% plug vs. 5%. Sample OC09 and the poliovirions–empty capsids mixture were used to investigate poliovirus electromigration in a separation buffer containing SDS concentrations from 0 to 125 mM (Fig. 2). A successful separation of poliovirus in OC09 in the absence of SDS was not possible because of too strong interactions between the protein contaminants of the sample and the capillary wall. Even at SDS concentrations of 5 and 12.5 mM, repeatability problems were observed for sample OC09. On the contrary, no separation problems were observed for the poliovirions–empty capsids mixture. These findings suggest that neither poliovirus nor the empty capsids are prone to interactions with the capillary wall. Practically, this means that, for highly purified poliovirus suspensions, SDS is not necessary as buffer additive to prevent interactions with the capillary wall. This result is different from what was previously published for human rhinovirus, the first picornavirus studied using CE [11]. For rhinovirus it was found that detergents above their respective CMC

are essential for the reproducibility of the electrophoretic results [13].

3.2. SDS concentration in the separation buffer and poliovirus electromigration When SDS is present in the separation buffer, it becomes constantly available, at a given concentration, for further adsorption to the viral particle surface or to the peptide backbone of individual viral proteins. For nanoparticles, an increase of the migration time in the presence of detergents was shown [23]. Similar observations were described for proteins, explained as a possible increase in the negative net charge because of SDS attachment [30]. Additionally, for proteins, the increase of the absolute net mobility, relative to SDS-free buffers, was correlated with SDS–protein aggregate formation during CE separation [29]. However, it should also be considered that when the SDS concentration of the separation buffer increases, a consequent increase in the viscosity is produced, that has the tendency to decrease mobility [49]. Based on the data previously published for proteins, only massive shifts in migration times accompanied by important alterations in peak shape can be considered as an indication of an essential change in the protein structure in result to SDS–protein aggregate formation [29,30]. For poliovirus, independent of the investigated sample, we have observed a small increase in the migration time when [SDS] in the separation buffer increased (Fig. 2A). However, a similar increase was also observed for the migration time of the EOF marker. The latter observation indicates a decrease in the EOF magnitude, suggesting an overall effect of the buffer viscosity increase. Consequently, the net absolute mobility did not seem to be influenced by the [SDS] in the buffer for the range 12.5–125 mM (Fig. 2B). The net absolute mobility of poliovirus was slightly higher at SDS concentrations below 12.5 mM (Fig. 2B). For both investigated samples, the poliovirus peak width at half height was minimal at 12.5–25 mM SDS and increased at high [SDS], while the corrected area (observed area divided by migration time) was rather constant for [SDS] above 5 mM (data not shown). Accordingly, the resolution between poliovirus and the EOF marker peak decreased when the SDS concentration in the separation buffer increased (Fig. 2C), even though, theoretically, an increase was expected. The above observations indicated that SDS is actively involved in the electrophoretic separation. This hypothesis is also supported by the evolution of the poliovirus peak height when the SDS concentration increased (Fig. 2D). For concentrations above 5 mM, the height increases, reaching a maximum when [SDS] is 12.5 mM for the highly purified poliovirus sample and 25 mM for the OC09 sample. The height then decreased for higher SDS concentrations and remained rather constant above 50 mM. Considering the separation principle of CE, these observations are consistent with the presence of a preconcentration mechanism, responsible for sharpening poliovirus peaks when the separation buffer contains SDS in concentrations between 12.5 and 25 mM. However, for proteins, it was earlier indicated that peak sharpening is an indication of reduced differences in microheterogeneity because of the formation of a stable complex, thus the SDS aggregates approach the saturation point [29,30]. Accordingly, we have shown that the electrophoretic mobility of poliovirus is not influenced by an [SDS] increase in the separation buffer (Fig. 2B). This indicates that only a limited number of SDS molecules are binding to a low number of binding sites and no structural changes are observed following poliovirus–SDS interaction during the CE separation. This is not surprising considering the tight packing of the poliovirus capsid proteins, confirmed by the stability and limited capsid permeability of poliovirus [41,42]. Also for rhinoviruses, the published data indicate that the 10 mM SDS used in the separation buffer has no effect on the virus particle

I. Oita et al. / Journal of Pharmaceutical and Biomedical Analysis 71 (2012) 79–88

B.

9

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Fig. 2. Influence of SDS concentration in the separation buffer on the poliovirus CE separation. (A) Migration times of EOF and poliovirus; (B) poliovirus mobility; (C) resolution between poliovirus and EOF marker peak; (D) poliovirus peak height. spv ∼ 2500 ␮g/ml; OC09 ∼ 530 ␮g/ml.

integrity and the mobility of the virus does not depend on [SDS] [11–14]. However, SDS appears to be involved in sharpening the poliovirus peak, as its concentration influences the peak shape. Additionally, SDS is needed for a good repeatability when samples containing additional protein contaminants have to be separated. 3.3. Electromigration of poliovirus and empty capsids in the absence of SDS During the above experiments, in the electropherogram of the poliovirions–empty capsids mixture obtained using SDS depleted buffer, we observed that the poliovirus peak appeared to be different than expected (Fig. 3). The corrected area of the first major peak, expected to be poliovirus, was almost double compared to all other corrected poliovirus peak areas, observed for separation buffers containing 5–125 mM SDS. Additionally, the peak width at half height (0.39) in the absence of SDS was larger than for any poliovirus peak observed before (0.04–0.37) (Fig. 3). These observations suggest the existence of multiple species within the same peak and motivated us to perform additional tests. When empty capsids were injected separately, using a separation buffer without SDS, we observed that peaks A and B were perfectly separated (Table 1), while they were not in the electropherogram of the empty capsids (Fig. 1-trace 3). Nevertheless, the migration time of poliovirus in the absence of SDS equaled that observed for peak A in the electropherogram of the empty capsids (Table 1). When the electropherograms of poliovirus and empty capsids were observed at 260 nm, peak A almost disappeared, while peak B decreased considerably upon examination at 280 nm, in the same manner as presented in Fig. 3 for the poliovirions–empty capsids mixture. It seemed that peak A, observed in the electropherogram of the heat exposed poliovirus (Fig. 1-trace 3), corresponded to a protein assembly or mixture, possibly the empty capsids, while peak B seems to be RNA.

The former hypothesis was also consistent with the ratios between the peak heights or areas observed at 260 and 280 nm, computed for poliovirus and empty capsids (Table 1). Generally, A260 /A280 ratios smaller than 1.7–1.8 indicate the presence of additional proteins while larger values indicate the presence of additional nucleic acids. Similar ratios for peak heights and areas were computed for the main peaks observed in the electropherograms of viral RNA (vRNA) synthesized in vitro, and of ferritin, as references for nucleic acids and proteins, respectively (Table 1). The vRNA data from Table 1 were acquired during an earlier study as no vRNA was available at the moment this research was performed. The separation conditions were comparable, except that the voltage was 18 kV instead of 10, but this is not expected to have any influence on the computed ratios or on similarity factors. As a consequence, the migration time of vRNA was shorter than for the peak assumed to represent RNA in the electropherogram of the empty capsids. Similarly to what was presented in Table 1, the ratios between the peak heights or areas observed at 260 and 280 nm were computed for the poliovirions–empty capsids mixture peaks when separation buffer without SDS was used. On the electropherogram of the poliovirions–empty capsids mixture in Fig. 3, the ratios between the peak heights or areas observed at 260 and 280 nm for peak A (about 1.4) were intermediary between those of poliovirus (1.8) and empty capsids (0.6). The ratios for peak B from the same electropherogram (about 2.2) are consistent with those of vRNA. The small difference in migration times observed between the electropherograms of poliovirus, empty capsids and their mixture was probably caused by the well known day-to-day variation of the EOF, as their net absolute mobilities are extremely similar. The results indicated that the presence of SDS in the separation buffer triggers the disintegration of the protein assembly in the empty capsids sample and peak A (Fig. 1 traces 2 and 3) is obtained. Therefore, peak A corresponds probably to a mixture of viral proteins–SDS aggregates. In fact, the same observations

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No SDS in the separaon buffe r

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Fig. 3. CE separation of the poliovirions–empty capsids mixture with and without SDS measured at several wavelengths. Separation conditions: uncoated fused silica capillary 40 cm effective length, 50 ␮m internal diameter; separation buffer 100 mM borate buffer pH 8.3 with 25 mM SDS (right pane) and no SDS (left pane); 10 kV; detection wavelength see fig; sample injected as 12% plug. Sample: poliovirions–empty capsids mixture (see text for details).

Table 1 UV absorption ratio at 260 and 280 nm, migration time, PV similarity, and RNA similarity for poliovirus, poliovirus exposed to 56 ◦ C for 10 min (hPV), reference proteins and RNA. Poliovirus and heated poliovirus samples were separated using buffers without SDS. Separation conditions as in Fig. 1. Sample

Migration time

 = 260 Area

 = 280 Height

Area

Area260 /Area280

Height260 /Height280

PV similaritye

RNA similaritye

Height

Poliovirus hPVa

5.7 5.8c 9.3d

38,100 2100 64,100

2160 180 2000

20,400 4000 25,300

1200 250 840

1.87 0.53 2.53

1.80 0.72 2.38

1.000 0.978 0.870

0.887 0.771 0.998

Mixture of PV and hPV

5.1c 7.4d

89,000 422,600

4200 8900

61,500 189,300

3000 4000

1.45 2.23

1.40 2.23

0.993 0.884

0.995 0.999

RNAb Ferritin

6.9 7.5

200,500 97,900

7640 3040

87,200 88,200

3300 2690

2.30 1.11

2.32 1.13

0.883 0.994

0.999 0.873

a b c d e

Sample exposed to 56 ◦ C for 10 min for the thermal conversion of the poliovirions to empty capsid particles. Sample separated using 18 kV and injected as 10% plug. Corresponding to peak A. Corresponding to peak B. Factor calculated to compare UV spectra of the investigated peak and the reference peak; values larger than 0.97 indicate a match with the reference.

were also published for the empty capsids of the rhinovirus, where non-dissociated empty capsids were only observed when a milder detergent was used [13,14]. When SDS is absent in the separation system, the protein assembly is conserved and migrates with a velocity comparable to that of the poliovirus particles. Thus, in the absence of SDS, it was not possible to separate the empty capsids from the poliovirus particles within the employed separation conditions, i.e. 12% sample plug injection of heated or unheated purified poliovirus suspension in borate buffer 20 mM pH 8.3 (Fig. 3).

3.4. Integrity of poliovirus particles and SDS presence in the separation buffers Regardless the amount of SDS used, the poliovirus peak was always detected. The peak shape, i.e. height and width, was influenced by the amount of SDS in the buffer, while the corrected area was rather constant. However, in the presence of SDS, peak A, assumed to be a protein assembly, was separated from the poliovirus peak (Table 2), but not from peak B, the RNA peak. The shape of peak A was also highly influenced by the SDS concentration. Acceptable peaks, symmetric and almost baseline separated

were obtained only for the separation buffer containing 12.5 and 25 mM SDS (Fig. 1). When [SDS] increases from 0 to 125 mM the migration time of the peak A shifts from 5.1 to 15.3 min (Table 2). Additionally, its shape is severely altered, i.e. the peak became extremely broad and for [SDS] above 50 mM SDS included the peak B. When poliovirions are converted to empty capsids, RNA is released and the capsid proteins undergo structural rearrangement [44]. Compared to the virions, the empty capsids are Table 2 Migration times of poliovirus and subviral particles for different SDS concentrations in the separation buffer. SDS, mM

0 5 12.5 25 50 75 100 125

Migration time, min Poliovirus

Capsid proteins A

RNA B

5.1 5.3 6.1 6.3 6.9 7.0 7.1 7.4

5.1 6.6 8.7 9.4 13.2 14.8 15.4 15.3

7.5 8.1 10.0 10.4 14.7 15.8 15.4 18.1

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Fig. 4. CE separation of off-line dissociated poliovirus capsids. (1) Poliovirus incubated with 1% SDS and ␤-mercaptoethanol at 96 ◦ C; (2) reference protein mixture incubated with 1% SDS and ␤-mercaptoethanol at 96 ◦ C; (3) reference protein mixture before incubation with 1% SDS and ␤-mercaptoethanol at 96 ◦ C; (4) poliovirus exposed to 100 ◦ C (right side: comparison of electropherogram at 205, 260 and 280 nm). Separation conditions: uncoated fused silica capillary 40 cm effective length, 50 ␮m internal diameter; 100 mM borate buffer pH 8.3 with 25 mM SDS; 10 kV; detection wavelength 205 nm; sample injected as 12% plug.

thermodynamically less stable and, consequently, during CE, they are destabilized by SDS. The individual proteins are probably released, forming SDS-aggregates. As SDS is UV transparent and the detection wavelength at 205 nm is non-specific, in the electropherograms, the peak attributed to empty capsids corresponds to all forms of subviral proteins, i.e. native, non-native, and aggregates with SDS. Additionally, when saturated protein–SDS aggregates are formed, these cannot be CE separated as there is no charge-tomass difference, thus no mobility difference. The peak broadening is probably due to the microheterogeneity, i.e. the presence of multiple species, or to a rate of aggregates formation and unfolding comparable with the time necessary to reach the detector [25,29]. To estimate the impact of the empty capsids exposure to SDS during CE analysis, samples containing poliovirions pre-exposed to conditions known to alter the native conformation of the capsid were injected. Thus, before injecting in the CE system, poliovirus was exposed to SDS denaturation, i.e. incubation with 1% SDS and ␤-mercaptoethanol at 96 ◦ C, as well as the thermal degradation at 100 ◦ C for 10 min. All samples were separated using a buffer containing 25 mM SDS. After exposure to these conditions, the poliovirus peak disappeared (Fig. 1-trace 4 vs. Fig. 4-trace 4). In the SDS denaturated poliovirus, it was difficult to interpret the electropherogram because of large ␤-mercaptoethanol interference (data not shown). Buffer exchange was performed and the electropherograms are presented in Fig. 4-traces 1 and 2. Even after ultrafiltration, the ␤-mercaptoethanol peak was still visible. An important, broad and tailing peak was visible at about 9.9 min in the electropherogram of the SDS denaturated poliovirions (Fig. 4traces 1 and 2). The peak purity assessment of this peak indicated that it corresponds to a mixture of compounds with similar chargeto-mass ratios, probably RNA and SDS saturated viral proteins. The electropherograms of a protein mixture before and after SDS denaturation are presented in Fig. 4-traces 2 and 3. In trace 2, an important peak is visible at about 10 min. When measured at 260 nm, this peak decreased more than 50 times. Thus, no absorbtion was observed at 260 nm for the denaturated proteins. The corresponding peak in trace 1, i.e. SDS denaturated poliovirus, had ratios for peak heights or areas at 260 and 280 nm around 2, indicating also RNA presence. In trace 3, peaks consistent with protein-SDS

aggregates corresponding to non-native conformations, as also described in [29,30] were observed between 10 and 13 min. The exposure to 100 ◦ C for 10 min is expected to dissociate completely the virus particles. The electropherogram (Fig. 4-trace 4) is very similar to the one corresponding to the exposure of poliovirus to 56 ◦ C for 10 min (Fig. 1-trace 3). However, after the exposure to 100 ◦ C, the width of peak A became almost double and it overlapped consequently a larger part of peak B. Examined at 260 nm,

Fig. 5. Electropherograms related to SDS addition to the sample. (1) Poliovirus sample without SDS added; (2) poliovirus sample containing 25 mM SDS; (3) poliovirus sample containing 25 mM SDS separated with an SDS depleted buffer. Separation conditions: uncoated fused silica capillary 40 cm effective length, 50 ␮m internal diameter; 100 mM borate buffer pH 8.3; 25 mM SDS in the separation buffer for (1) and (2); 10 kV; detection wavelength 205 nm; sample injected as indicated; sample: sucrose gradient purified poliovirus, 4 times diluted with water –A260 /A280 = 1.73, 245 ␮g/ml poliovirus.

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peak A completely disappeared, but it was slightly visible at 280 nm (Fig. 4-right pane). This supported our hypothesis that the first peak corresponds to SDS–protein capsid aggregates, while the second corresponds to RNA. The injection of poliovirus incubated with 1% SDS and ␤mercaptoethanol at 96 ◦ C (trace 1), confirmed that peak A, supposed to have protein origin and observed in the electropherograms of the empty capsids (Fig. 1-trace 3) corresponds to aggregates between individual capsid proteins and SDS.

3.5. Electromigration of poliovirus when SDS is added to the sample The results presented until now were obtained using a discontinuous separation system, where SDS was present in the separation buffer, but not in the injected sample plug. When 25 mM SDS was added to the sample immediately before injection, to create a homogeneous separation system, the poliovirus peak shape was severely modified, i.e. the peak height decreased to less than half and the peak broadening increased visibly (Fig. 5-trace 1 vs 2). Attempts to use a reversed system, i.e. sample containing SDS and an SDS depleted separation buffer, compromised completely the separation between poliovirus and the EOF marker peak regardless the plug size (Fig. 5-trace 3). Additional peaks were visible (Fig. 5-trace 3), probably as a result of a limited dissociation of poliovirus particles induced by the SDS present in the sample plug.

Possibly, the presence of SDS in the sample plug revokes the preconcentration mechanism developed in the discontinuous separation system. We thus have shown that the addition of SDS in the sample plug is detrimental to poliovirus CE separation because of interference with the preconcentration mechanism.

3.6. Electromigration of poliovirus and empty capsids in SDS gradient We did not observe interactions between poliovirus or empty capsids and the capillary wall as long as the original sample matrix was removed and the sample was resuspended in diluted separation buffer without SDS (Fig. 1). The SDS presence in the separation buffer was shown necessary to separate poliovirus from the empty capsids, but detrimental to the empty capsids integrity. Therefore we decided to study the separation of the poliovirions–empty capsids mixture using an SDS gradient. The gradient was formed by (i) filling first the capillary with separation buffer containing SDS, followed by (ii) injection of the sample plug (12%) without SDS and a small plug of separation buffer (2%) without SDS, and finally (iii) performing the separation in buffers without SDS. Within this setup, the SDS micelles are positioned only at the cathodic side of the sample zone. The sample was prepared in borate buffer 20 mM pH 8.3, thus the conductivity of the sample zone was lower than that of the separation buffer. Fig. 6 shows the separations of the poliovirions–empty capsids mixture in SDS gradients, at two different temperatures. When the

Fig. 6. Separation of a mixture of highly purified poliovirus and subviral particles in SDS gradient. Below, the profile of the current with (a) the capillary is filled with the sample (12%) and SDS containing separation buffer and (b) the capillary is filled with separation buffer without SDS. Sample: poliovirus and subviral particles in borate buffer 20 mM pH 8.3. PV indicates poliovirus peak, A and B the subviral particle peaks and * spikes, i.e. artifactual peaks corresponding to aggregates. Separation conditions: uncoated fused silica capillary 40 cm effective length, 50 ␮m internal diameter; 10 kV; detection wavelength 205 nm; sample injected as 12% plug; separation buffer 100 mM borate buffer pH 8.3; between runs capillary was filled with separation buffer 100 mM borate buffer pH 8.3 and 25 mM SDS – (1) and (2) or 100 mM borate buffer pH 8.3 and 12.5 mM SDS – (3) and (4); separation performed at 25 ◦ C – (1) and (3) or 35 ◦ C – (2) and (4); (5) – control, separation conditions as in Fig. 1.

I. Oita et al. / Journal of Pharmaceutical and Biomedical Analysis 71 (2012) 79–88

voltage was applied, we observed a steep decrease of the current until the sample matrix and the SDS micelles leaved the capillary (a-lower pane) followed by a stable current (b-lower pane). The current stabilization indicates a capillary filled with separation buffer only, thus without SDS micelles. As SDS is an anionic surfactant, the natural migration tendency of the SDS micelles will be toward the anode. Since the sample conductivity is lower, the electrical field will be higher in the sample zone. Therefore when the micelles enter the sample zone they will stack at the opposite side of sample–separation buffer interface and sweep [7–9]. A slight sharpening of the poliovirus peaks, i.e. decrease of peak width and increase in height, was observed when the SDS gradient was used (Fig. 6 vs. Fig. 1). The effect was more pronounced for the buffer containing 12.5 mM SDS (Fig. 6-traces 3 and 4). Similar results were also obtained for higher EOF, obtained increasing the temperature by 10 ◦ C (Fig. 6-trace 1 vs. 2 and trace 3 vs 4). The separation of the empty capsids was much improved compared to the situation when separation buffer and filling buffer were identical (Fig. 6 vs. Fig. 1). Peak A obtained in the SDS gradient seems to correspond to the empty capsids, because based on the UV spectrum, it is a pure peak matching the empty capsids spectrum (99.94% peak purity) and which almost has no absorption at 260 nm. However, additional data are needed to confirm. Besides peaks A and B, already described in the previous sections on empty capsids, an additional peak at 9.5 min, which seems to be protein related, was observed for the experiments with 25 mM SDS in the filling buffer (Fig. 6-trace 1). Since this peak is not visible for the experiments using 12.5 mM SDS, it is likely to correspond to saturated aggregates between SDS and capsid proteins. Spikes, i.e. the very sharp peaks in Fig. 6, indicating the formation of aggregates [13,14], were sometimes observed. The incidence of the spikes was higher for the experiments with 12.5 mM SDS. Summarized, sharper peaks are obtained for separations in SDS gradient, as well as an improved separation between the peaks corresponding to empty capsids. Possibly the use of the SDS gradient conserved the empty capsids structure and provided the separation between the empty capsids and poliovirus, but additional tests are needed to confirm this hypothesis. 4. Concluding remarks Our results indicate that the electrophoretic mobility of poliovirus is not influenced by the increase of SDS concentration in the separation buffer. Therefore, no poliovirus–SDS aggregates are formed or no structural changes are observed during the CE separation when SDS-containing buffers are used for separation. Additionally, SDS appears to be involved in sharpening of poliovirus peak and it improves the overall repeatability when samples containing additional protein contaminants have to be separated. The SDS involved preconcentration mechanism needs a discontinuous system. However, in the SDS containing separation buffer, the empty capsids dissociate in the individual proteins during the CE separation. Furthermore, the individual proteins form aggregates with SDS and they migrate as single peaks. Sharper peaks and improved separation of the empty capsids were obtained when an SDS gradient was used. Possibly, within the SDS gradient, the peak corresponding to the empty capsids conserves its integrity and is separated from the poliovirus particles, but additional tests are needed to confirm this hypothesis. Acknowledgements This work was supported by a Horizontale Onderzoeksactie (HOA) of the Vrije Universiteit Brussel and a research grant

87

(G.0051.08) of the FWO. We thank Monique De Pelsmacker for the preparation of the viruses, and Katrien Decq and Frank Van der Kelen for the technical and logistic assistance.

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