RESEARCH ARTICLE – Pharmaceutical Biotechnology
Preformulation Study of Highly Purified Inactivated Polio Vaccine, Serotype 3 WEI QI,1 YUHONG ZENG,2 SCOTT ORGEL,1 ALAIN FRANCON,3 JAE HYUN KIM,2 THEODORE W. RANDOLPH,4 JOHN F. CARPENTER,1 C. RUSSELL MIDDAUGH2 1
Department of Pharmaceutical Sciences, University of Colorado Anschutz Medical Campus, Aurora, Colorado University of Kansas, Macromolecule and Vaccine Stabilization Center, Lawrence, Kansas 3 Sanofi Pasteur, March l’Etotile, France 4 Department of Biological and Chemical Engineering, University of Colorado, Boulder, Colorado 2
Received 14 September 2013; revised 23 October 2013; accepted 6 November 2013 Published online 26 November 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI 10.1002/jps.23801 ABSTRACT: To improve the effectiveness of the polio vaccination campaign, improvements in the thermal stability of the vaccine are being investigated. Here, inactivated polio vaccine, serotype 3 (IPV3) was characterized via a number of biophysical techniques. The size was characterized by transmission electronic microscopy and light scattering. The capsid protein conformation was evaluated by intrinsic fluorescence and circular dichroism (CD), and the D-antigen content by enzyme-linked immunosorbent assay (ELISA). The pH thermal stability of IPV3 (pH 3.0–8.0; 10◦ C–87.5◦ C) was evaluated by fluorescence, CD, and static light scattering. The transition temperatures reflect the responses, respectively, of tertiary structure, secondary structure, and size to applied thermal stress. The data were summarized as empirical phase diagrams, and the most stable conditions were found to be pH 7.0 with temperature lower than 40◦ C. CD detected a higher transition temperature for capsid protein than that for RNA. The effects of certain excipients on IPV3 thermal stability and antigen content were evaluated. The results of their effects, based on intrinsic fluorescence and ELISA, were in good agreement, suggesting the feasibility of applying intrinsic fluorescence as a high-throughput tool for formulation development. The study improves the understanding C 2013 Wiley of IPV3 thermal stability, and provides a starting point for future formulation development of IPV3 and other serotypes. Periodicals, Inc. and the American Pharmacists Association J Pharm Sci 103:140–151, 2014 Keywords: stability; excipients; vaccines; physical characterization; pre-formulation; stabilization
INTRODUCTION Since 1998, the World Health Organization (WHO) has been campaigning for the global eradication of polio.1 In 2009, over 361 million children were immunized.2 Disappointingly, in resource-poor regions, the most commonly used oral polio vaccine (OPV) formulation had reduced potency and generated live vaccine-derived poliovirus and rare vaccine-associated paralytic poliomyelitis (VAPP).3–7 Additionally, previously poliofree countries could be reinfected because of imported virus from polio-endemic areas, that is, the 2010 polio outbreak in Tajikistan.8 Therefore, a safer vaccine is desired to eliminate VAPP and vaccine-derived poliovirus circulation. In the USA and UK, inactivated polio vaccine (IPV) replaced OPV in 2000 and 2004, respectively.9 IPV is currently formulated as a liquid (medium 199) and is stable for over a year at 4◦ C, but loses potency in only weeks at 25◦ C or 37◦ C.10 To preserve vaccine potency during shipping, storage, and distribution, an effective cold chain is necessary but practically difficult in the resource-poor areas where the vaccine is most needed. Hence, a thermally stable formulation is strongly desired to maintain potency before ultimate administration of the vaccine.
Correspondence to: C. Russell Middaugh (Telephone: +785-864-5813; Fax: +785-864-5814; E-mail: [email protected]) This article contains supplementary material available from the authors upon request or via the Internet at http://onlinelibrary.wiley.com/. Journal of Pharmaceutical Sciences, Vol. 103, 140–151 (2014)
C 2013 Wiley Periodicals, Inc. and the American Pharmacists Association
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To develop new poliovirus vaccine formulations, it is critical to determine the effect of potential formulation additives on the structural stability of IPV. High-resolution structural information concerning poliovirus has been provided by EM11 and X-ray12 analysis. The process by which poliovirus enters cells was recently characterized by fluorescence microscopy.13 For rapid development of new formulations, however, more readily detected biophysical properties, rather than high-resolution structures, are needed. Tryptophan (Trp) fluorescence intensity was previously used to study the thermal stability of poliovirus in the presence of excipients14 and under different pH conditions.15 Circular dichroism (CD) spectroscopy and differential scanning microcalorimetry were also used in attempts to monitor poliovirus thermal stability,14 but the results were compromised by extremely low sample concentration. Currently, changes in the physical properties of IPVs upon thermal stress are poorly understood, with only limited data available at temperatures higher than 45◦ C.10 In addition, the routine active antigen (D-antigen) content test was not standardized, and inconsistencies among manufacturers and laboratories exist. These probably result from the variation in antibodies and procedures used in the D-antigen test.16 Furthermore, many studies14–18 have investigated either vaccine structure or potency individually, but the combined results are rarely reported. Finally, a more rapidly measured biophysical property might be useful as a complementary measurement of potency. A correlation between such a physical property and IPV potency, if it exists, would greatly facilitate rational and effective IPV formulation development.
RESEARCH ARTICLE – Pharmaceutical Biotechnology
In this work, we employed highly purified inactivated polio vaccine, serotype 3 (IPV3) samples from a commercial manufacturer to: (1) characterize IPV3 by different biophysical techniques, focusing on size and capsid protein structure; (2) establish a pH thermal stability profile of IPV3 employing empirical phase diagrams (EPDs); (3) evaluate IPV3 active antigen content with and without excipients upon thermal stress; and (4) correlate certain biophysical properties with antigen content.
MATERIALS AND METHODS Materials Inactivated polio vaccine, serotype 3 was shipped from Sanofi Pasteur (Lyon, France) in medium 199 without phenol red (see Sigma #M9163 for detailed composition). During shipping and storage, the temperature was kept between 2◦ C and 8◦ C. The IPV3 concentration in terms of protein mass was approximately 20 :g/mL and the D-antigen content was approximately 1128 UD/mL. The coating monoclonal antibody, mAb #520, was from NIBSC (National Institute for Biological Standards and Control). The secondary antibody, mAb #4B7-1H8-2E10, was obtained from Sanofi Pasteur and kept at −80◦ C until use. Anti-mouse IgG peroxidase antibody from goat (A3673) was purchased from Sigma (St. Louis, Missouri). Nonfat milk powder was purchased from Becton Dickson (Franklin Lakes, New Jersey). Ultrapure urea and deuterium oxide (D2 O, 99.9%) were from MP Biomedicals LLC (Aurora, Ohio). Sucrose and trehalose were obtained from J.T. Baker (Phillipsburg, New Jersey). Mannitol U.S.P. was from Spectrum (New Brunswick, New Jersey). Sorbitol was purchased from Calbiochem (Merck KGaA, Darmstadt, Germany). Glycine and anhydrous glycerol were purchased from Fisher (Fair Lawn, New Jersey). SYPRO Orange was from Invitrogen (Eugene, Oregon). All other chemicals were obtained from Sigma. Sample Preparation Inactivated polio vaccine, serotype 3 samples were first dialyzed R Dialysis Cassettes with a 10 kDa molecular in Slide-A-Lyzer weight (MW) cutoff (Pierce, Rockford, Illinois) against 20 mM phosphate-citrate buffer with a fixed ionic strength of 0.15M by adding an appropriate amount of NaCl, pH 7.0 at 4◦ C overnight before any other specified treatment. Dynamic Light Scattering Backscattering from solutions containing IPV3 in approximately 150 :L samples in a low-volume quartz cuvette with 1 cm path length was measured at 173◦ with a Malvern Zetasizer (Worcestershire, UK). The temperature was set at either 25◦ C or ramped from 25◦ C to 90◦ C with 5◦ C steps. Samples were held at each temperature for 2–3 min before data acquisition. Transmission Electronic Microscopy Three microliter IPV3 samples was placed onto a carbon-coated grid and allowed to adsorb for 3 min. The grid then was washed twice with ddH2 O. The sample was stained with 2% uranyl acetate solution for 2 min, air dried and then imaged by a model Technai G2 transmission electronic microscopy (TEM) equipped with a Gatan Ultrascan digital camera from FEI (Hillsboro, Oregon) at a magnification of 60,000×. DOI 10.1002/jps.23801
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CD Spectroscopy Dialyzed IPV3 samples were placed in Millipore Centricon tubes (MW cutoff 10 kDa) and centrifuged at approximately 2500 × g in a Sorvall Centrifuges RT 6000D (Wilmington, Delaware) at a temperature between 2◦ C and 6◦ C. The concentration of virus was determined by UV absorption spectroscopy. The UV absorbance at 280 nm in a 1 cm path length cuvette was at least 0.2 units. Approximately 400 :L of sample were then transferred into a 1 mm path length cuvette and the CD spectrum of the sample was measured from 190 to 300 nm with a scan step of 1 nm by a Chirascan Plus spectropolarimeter from Applied Photophysics (Leatherhead, UK). The CD spectra were analyzed by the CDNN software package to obtain the percentage of various secondary structure types. Fluorescence Spectroscopy Seven hundred microliter samples of dialyzed IPV3 was added to a 1 cm path length cuvette. Intrinsic Trp fluorescence spectra were acquired with a Photon Technology International spectrofluorometer (Lawrenceville, New Jersey) equipped with a Peltier thermostatted cuvette holder. The samples were excited at 295 nm (>95% Trp emission) and the emission spectra were collected from 300 to 400 nm, with a 1 nm step and 1 s integration time. Static light scattering (SLS) was collected at 295 nm by a second photomultiplier with the same instrument located 180◦ to the fluorescence emission detector. For stability tests with excipients, dialyzed IPV3 samples were concentrated approximately 20-fold by centrifugation at approximately 2500 × g in Millipore Centricon tubes with a 10kDa cutoff (2◦ C–6◦ C). Concentrated samples were mixed with specific excipient solutions in equal volumes to a final IPV3 concentration that was approximately 10-fold that of the freshly dialyzed sample. To prepare IPV3 samples in D2 O, IPV3 in medium 199 was centrifuged in Millipore Centricon tubes with 10-kDa cutoffs to approximately 5% or less of the initial volume, and then resuspended in D2 O to the initial volume. This was repeated three times to ensure that the D2 O content was greater than 95%. All the stability test experiments with excipients were carried out in an Optim 1000 spectrofluorimeter from Avacta (Wetherby, UK). Approximately 10 :L of IPV3 mixed with individual excipients was directly loaded into the micro cuvette arrays. The intrinsic fluorescence was excited at 266 nm with a laser source, and the emission spectra were collected from 250 to 500 nm with default settings. SLS data were collected simultaneously at 473 nm using a separate laser. For extrinsic fluorescence experiments, SYPRO Orange (Invitrogen) was mixed with IPV3 samples to a final dye concentration of 5× dilution from the original stock. The samples were excited at 473 nm and the spectra were collected from 450 to 700 nm. For isothermal incubation, the spectra were collected at specific temperatures (40◦ C, 50◦ C, 60◦ C, and 70◦ C) every 5 min during 3 h incubations. For temperature ramping experiments, the temperature was varied from 20◦ C to 90◦ C at a 1◦ C per intervals, and spectra were collected at each temperature after a hold time of 2–3 min. Temperature interval was set at 1◦ C to capture smoother transitions in the presence of different excipients. The raw spectra were processed with the embedded Optim1000 software (Avacta) using default settings. The ratio of the intensity at 350 to 330 nm was used as an indication of the relative abundance of unfolded to native protein. This Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
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intensity ratio provided similar transition information to that of the fitted peak position (see Suppl. Material, Suppl. Fig. S1). For CD and fluorescence thermal ramping experiments, the intensity or peak position was fitted to a standard two-state protein unfolding model, with the transition temperature, Tm , the mid-point of the transition. pH and Thermal Stability For pH and thermal stability studies, IPV3 was first dialyzed into 20 mM phosphate-citrate buffer with a fixed ionic strength of 0.15 M by adding an appropriate amount of NaCl. The pH was varied from 3.0 to 8.0 at 1.0 unit intervals. Samples were analyzed by CD, Trp fluorescence, and SLS, with temperature ramping from 10◦ C to 87.5◦ C at 2.5◦ C intervals. Samples were equilibrated at each temperature for 2–3 min before data acquisition. EPDs for IPV3 were generated as described previously.19–21 Temperature interval was set up at 2.5◦ C to be compatible with the MatLab code for EPD at that time. Five sets of data at each pH was used to construct EPDs, including SLS, fluorescence peak position, fluorescence intensity, and CD values at 205 and 220 nm. As described later in the text, these diagrams reveal different physical states of the vaccine under the specified pH and temperature conditions. Enzyme-Linked Immunosorbent Assay Indirect enzyme-linked immunosorbent assays (ELISAs) were performed to characterize the virus content of IPV3 in terms of D-antigen content. The measurement of D-antigen content has been routinely used to correlate with the protective immunogenicity of IPV.16,22 Briefly, 50 :L of coating antibody, mAb #520, were added to wells in a 96-well plate (PI-15041, Thermal Scientific) and incubated at 4◦ C overnight. mAb #520 was reported to bind to the active antigen site 1 for IPV3.22,23 Plates were washed at least three times with washing buffer [1× PBS (phosphate-buffered saline) with 0.025%, v/v, Tween 20 and 0.21% skim milk]. Sixteen microliter IPV3 samples was first diluted 10-fold into the washing buffer to a final volume of 160 :L. Eighty microliter out of the 160 :L was then added to the first well on the plate. The remaining 80 :L was diluted by washing buffer to 160 :L and another 80 :L was loaded to the second well, and so on. At each dilution, 80 :L sample was transferred into the corresponding well on the plate in sequence for that sample. Plates were then incubated at 37◦ C for 2 h. After another washing step, 50 :L of secondary antibody solution were added and incubated at 37◦ C for 1 h. Fifty microliter of tertiary antibody solution was added after washing and incubated at 37◦ C for another hour. Finally, 50 :L of TMB (3, 3 , 5, 5 tetramethylbenzidine) was added to each well, and the plate was held at room temperature for 10–15 min). HCl (1 N) was added to stop the reaction and the absorbance was read immediately at 450 nm with 630 nm as a reference, employing a VERSMax plate reader (Molecular Devices, Sunnyvale, California). Inactivated polio vaccine, serotype 3 samples in PBS after dialysis were used to determine assay reproducibility. Titer values were reproducible with a standard deviation within 15% for seven tested samples on different days (data not shown). The typical loss of the D-antigen from the medium 199 during dialysis was between 10% and 20% (data not shown). Incubation at 50◦ C for 1 h was chosen as an accelerated degradation condition to screen for the effects of excipients on Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
IPV3 stability. In preliminary tests, IVP3 stability was tested after incubation at 50◦ C and 60◦ C, with incubation times of 15, 30, and 60 min. Degradation at 60◦ C was rapid, leading to a complete loss of antigen detectable in the ELISA assay after only 30 min, whereas 10%–15% of the original antigen content remained after incubation at 50◦ C for 1 h. This level of thermal stress permitted an adequate window to detect effects of the excipients. A standard 4-parameter fit was performed for all ELISA data using Origin software, with EC50 values fit for each sample. The EC50 value for IPV3 samples immediately after dialysis was taken as 100%, and that for IPV3 samples after 1 h incubation at 50◦ C as 0%. To evaluate the effects of different excipients upon thermal stress, the EC50 values of those samples were normalized to the difference between 0% and 100% controls. Such values were used to represent the ELISA antigen content of the IPV3, which is referred to as percent of antigen content in the text. Sample Replicates and Error For all experiments, at least triplicates were studied unless otherwise specified. The error bars in the figures reflect the standard deviation of the specific measurement. For clarity and to avoid confusion, certain figures did not include the error bars.
RESULTS Biophysical Characterization of IPV3 Transmission electronic microscopy (TEM) and dynamic light scattering (DLS) were first applied to characterize the size and morphology of IPV3. As shown in Fig. 1a, IPV3 in freshly dialyzed samples appeared as spherical particles with diameters of approximately 30 nm, which is close to previous literature reports.11,24 No empty particles were observed, suggesting that the single-stranded RNA had not leaked out of the vaccine capsid. Although TEM directly represents IPV3 morphology and size, it is neither convenient nor practical for obtaining realtime images of IPV3 under thermal stress. In addition, formulations containing high concentrations of disaccharides may not be suitable for TEM measurements. Furthermore, the sampling area of a TEM image represents only a small fraction of the total sample, and the measurement might not be accurate because of sample preparation effects. Therefore, DLS was used as an alternative approach to measure virus particle size and size distributions. In Fig. 1b, for a freshly dialyzed IPV3 sample, a single peak dominated the volume distribution with a mid-point diameter of 33.9 ± 1.9 nm. The diameter determined from a corresponding intensity measurement was slightly larger than the reported value of approximately 30 nm, presumably because light scattering provides a hydrodynamic radius, which is indicative of the apparent size of the hydrated particle and could be influenced by even very small amount of larger particles. The intrinsic fluorescence of Trp residues has been routinely used to probe the tertiary structure of proteins in different solvent conditions. A fluorescence emission maximum from 320 to 330 nm suggests that on average Trp residues are located in a relatively apolar environment as seen typically in native states, whereas a wavelength maximum from 350 to 355 nm suggests Trp residues are more exposed to the aqueous solvent as seen in more structurally altered states.25 In Fig. 1c, we observed an emission peak for IPV3 with a maximum near 335 nm, DOI 10.1002/jps.23801
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Figure 1. Characterization of IPV3 in PBS buffer after dialysis and before any thermal treatment or exposure to excipients. (a) TEM images of IPV3. The black scale bar at the left corner is 50nm in length. (b) DLS of IPV3 in terms of volume percentage. (c) Intrinsic Trp fluorescence of IPV3 excited at 295nm. (d) CD spectrum of IPV3 from 195 to 300nm.
suggesting that the Trp residues in the virus proteins are on average found in a relatively apolar environment. The IPV3 sequence (http://www.ncbi.nlm.nih.gov/protein/AAA19636), contains 780 Trp residues located in a single virion (13 Trp residues present in single copies of the capsid proteins VP1 to VP4, with 60 copies of VP1 to VP4 in each virion). Thus, the signal seen reflects the averaged status of a large number of Trp residues, negating any high-resolution structural interpretation of spectral changes. Circular dichroism spectroscopy was used to examine the secondary structure of the capsid proteins. In Fig. 1d, a strong negative band around 205 nm is seen, together with a positive band from 260 to 280 nm. A minimum at 205 nm usually reflects the presence of crossed $-structure. The positive ellipticity near 265 nm may arise from the RNA component, as previously reported for hepatitis A virus.26 To further probe the nature of the secondary structure, the spectrum was deconvoluted to compare the secondary structure components reported for type 3 poliovirus (with PDB ID 2plv).27 In the spctra of IPV3 obtained between 190 and 260 nm, approximately 10% helix, 33% $-sheet, 25% $-turn, and 33% random coil were estimated. For type 3 poliovirus, 9% helix and 29% $-sheet were reported for VP 1 and 3, with 12% helix and 37% $-sheet for VP 2, and 10% helix and 14% $-sheet for VP 4, with the remaining structures being turns and/or unspecified structures (http://www.pdb.org/pdb/ explore/remediatedSequence.do?structureId=2PLV). Thus, the secondary structural content from our measurements were similar to previous estimates, even though the RNA component may interfere with the signal below 240 nm.26 Again, as in DOI 10.1002/jps.23801
the fluorescence results, we emphasize that CD spectroscopy measured the combined structural properties of all the capsid proteins. Thermal Stability of IPV3 at pH 7 as Assessed by Intrinsic Fluorescence Spectroscopy, CD Spectroscopy, SLS, and DLS Inactivated polio vaccine, serotype 3 is currently formulated at pH 7.0–7.4. To provide reference data to compare against results for IPV3 in other solution conditions, analyses of IPV3 at pH 7 were performed first using Trp and SYPRO Orange fluorescence, CD spectroscopy, SLS, and DLS. Intrinsic Trp fluorescence emission spectra collected at different temperatures are shown in Fig. 2a. As the temperature increased, the fluorescence signal gradually decreased, and the spectra broadened. Concomitantly, the central peak position red shifted from approximately 335 nm to 346 nm. Temperature-dependent transition curves of maximum intensity and peak position are shown in Figures 2b and 2c, respectively. The intensity curve showed a gradual decrease with increasing temperature, with an inflection point of approximately 50◦ C. In contrast, a more abrupt transition was observed with the curve for peak position, indicating increasing exposure of the Trp residues to a more hydrophilic environment with heating. The Tm obtained from the fluorescence peak position data was 57.5 ± 1.8◦ C. Intrinsic fluorescence was also used to probe vaccine structural changes for IPV3 during isothermal incubation, at 40◦ C, 50◦ C, 60◦ C, and 70◦ C. As shown in Fig. 2d, no significant peak position change was observed as a function of incubation time for IPV3 at 40◦ C. During incubation at 50◦ C, a gradual but Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
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Figure 2. Intrinsic Trp fluorescence spectra of IPV3 upon thermal stress. (a) Emission spectra of intrinsic fluorescence collected from 300 to 400nm with excitation at 295nm. The arrow direction indicates increased 5◦ C temperature steps, illustrated by alternative dark and gray lines. (b) Normalized intrinsic Trp fluorescence intensity as a function of temperature. The peak position of the maximum intensity for intrinsic Trp fluorescence as a function of increasing temperature (c) and isothermal incubation (d) at indicated temperatures. Triplicates were performed to the above experiments and the mean values were plotted with the standard deviation as the error bar in panel b and c.
subtle transition took place with the peak position red-shifting approximately 2 nm. When IPV3 was incubated at 60◦ C (close to the Tm ), the peak position red-shifted about 5 nm within 2000 s. After longer incubation times, the peak position slightly decreased, which may reflect the formation of large aggregates (see Suppl. Material, Supp. Fig. S2). At an even higher temperature (70◦ C), the peak position red-shifted up to approximately 10 nm within a shorter time period (∼1000 s), suggesting major structural disruption. Similar to the behavior at 60◦ C, there was a gradual blueshift in peak position at longer incubation times. Changes in SYPRO Orange fluorescence have recently been used to study thermal stability of monoclonal antibodies and adjuvanted pneumococcal proteins,28 as well as used for excipient/stabilizer screening.28,29 This dye is nonfluorescent in hydrophilic environments, but exhibits a substantial increase in fluorescence when it is exposed to apolar regions of proteins.28,29 Therefore, SYPRO Orange fluorescence intensity can be used as an indicator of protein structural changes, such as those occurring under thermal stress. In the case of IPV3, the extrinsic fluorescence from SYPRO Orange was detected with an intensity maximum near 600 nm and a slight shoulder at approximately 650 nm. As the temperature increased from 20◦ C to 90◦ C, the SYPRO Orange fluorescence intensity gradually decreased with no obvious cooperative transition, similar to the intrinsic fluorescence intensity trend. Furthermore, no significant shift was found for the wavelength maximum of SYPRO Orange fluorescence (see Suppl. Material, Suppl. Fig. S3). Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
Circular dichroism spectroscopic studies conducted as a function of temperature produced distinctive results (Fig. 3). Upon heating from 20◦ C to 90 ◦ C, the CD signal at approximately 205 nm for IPV3 decreased from −9.0 to −6.5 millidegrees, the wavelength minimum shifted from approximately 205 to 215 nm and there was a broadening of the spectra. For the RNA peak (positive ellipticity at ∼265nm), the signal decreased monotonically with increasing temperature, reaching a plateau close to 80◦ C. The apparent thermal transition temperatures for the protein and RNA portions of the spectra were 58.1 ± 1.4◦ C and 49.0 ± 2.0◦ C, respectively (Figs. 3c and 3d). This result suggests that the capsid protein is more thermally stable than the RNA component. When the IPV3 sample was cooled back to room temperature after thermal stress, the CD signal was regained to some extent; approximately 70% of the original RNA signal and approximately 50% of the protein signal were recovered within the region 200–220 nm (Fig. 3b). Simultaneously, the minimum wavelength shifted back to approximately 209 nm compared with the initial value of 205 nm. Thus, the loss of secondary structure of the capsid proteins at high temperature appeared to be at least partially reversible. As temperature was increased, the SLS intensity of IPV3 remained fairly constant until 60◦ C and then sharply increased to its maximum value at approximately 70◦ C (Fig. 4 a). The intensity decrease above 70◦ C was probably because of the precipitation of large aggregates. The large error bars at elevated temperature also reflect this phenomenon. DOI 10.1002/jps.23801
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Figure 3. Representative CD spectra of IPV3 upon thermal stress. (a) CD spectra of IPV3 at different temperatures 20◦ C–90◦ C at 10◦ C intervals. (b) CD spectra of IPV3 at 90◦ C, and at 20◦ C before and after the thermal treatment. (c) Wavelength shift of the minimum intensity between 200 and 230nm upon thermal stress. (d) The normalized intensity at 260nm upon thermal stress. Three replicates were performed and only representative results are shown for clarity. The Tm values for c and d were fitted as specified in Material and Methods.
Figure 4. The light scattering of the IPV3 at pH 7 upon thermal stress. (a) SLS. (b) DLS. The diameter of the two peaks in 90◦ C in panel b was measured at 25◦ C after thermal ramping to 90◦ C. Triplicates were performed to the above-mentioned experiments, and the mean values were plotted with the standard deviation as the error bar in panel a.
Dynamic light scattering results during heating of IPV3 (Fig. 4b) followed a similar trend to that observed with SLS. Below 70◦ C, a single peak gradually shifted from a position for approximately 30 nm to that of 70 nm. At 75◦ C, two peaks were detected, with one centered at 50 nm and the other at 400 nm, the latter clearly indicating the presence of aggregated virion particles. At even higher temperature (80◦ C to 90◦ C), very large aggregates appeared, resulting in unreliable data because of the high degree of polydispersity and the inability of DLS to accurately determine the size above 1 :m. After cooling samples to room temperature, two populations of approximately 80 nm and greater than 1 :m were seen with a volume ratio DOI 10.1002/jps.23801
close to 1:1 (Fig. 4b, 90◦ C panel). The formation of aggregates at high temperature was also confirmed by TEM, where disrupted virions and aggregated material with dimensions larger than a few hundred nanometers were observed (see Suppl. Material, Suppl. Fig. S4). pH and Thermal Stability—EPDs To explore the combined effects of pH and temperature, the methods described above were used to examine IPV3 at pH 3–8 at one unit intervals as the temperature was increased from 10◦ C to 90◦ C. Representative data are shown in Fig. 5. It is clear that the response of the virus to temperature was Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
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Figure 5. The representative experimental data for EPDs. (a) SLS. (b) Intrinsic fluorescence peak position calculated with the center of spectra mass method. (c) Normalized maximum fluorescence intensity. (d) CD intensity transition data at 220nm, with pH 3 (closed square), 4 (open circle), 5 (closed triangel), 6 (open triangel), 7(closed diamond), and 8 (open star).
highly pH dependent. Transitions (Tm values) observed in all methods generally follow the pattern pH 3 < 4 < 5 < 6 < 8 < 7, with clear evidence of multiple transitions, especially at low pH. To provide a more convenient and comprehensible view of this complex data, they were summarized in the form of several EPDs, Fig.6. In this method, the data are normalized and presented in the form of changes in color, facial features, or distorted polygons. Detailed descriptions of these visualization methods are presented elsewhere.19 Using a clustering technique to define apparent phase boundaries, all methods of presentation resolved five distinct phases. Using the three-index EPD, the nature of each phase can be at least partially assigned. This can be supported by direct analysis of the data in Fig. 5 that was used to construct the EPDs, as shown in Fig. 6. In summary, the native state of the viral particle appeared to be present at pH 6–7 below 45◦ C. Alteration in the tertiary structure of the viral proteins was evident at pH 5 and 8 with further changes at lower pH (3–4) with reduced thermal stability (25◦ C–32◦ C). Major changes in protein secondary structure (CD spectra) were most evident at pH 5–7 above 50◦ C. Aggregation appeared above 60◦ C at both high (8) and low (3–5) pH and appeared at lower temperature (95%), there is almost no detectable signal increase even up to 90◦ C, indicating little or no formation of large aggregates. Trehalose (25%) was almost as effective as D2 O at inhibiting aggregation. Sucrose and sorbitol produced similar results to those for trehalose. IPV3 samples in mannitol and glycerol have similar onset temperatures of aggregation to that of the control. IPV3 samples in urea (5%) aggregated only at much higher temperatures than those noted for samples with other excipients, indicating that urea might solubilize the virions to some extent DOI 10.1002/jps.23801
and thereby reduce aggregation even at temperatures above the Tm . Samples in sodium citrate generated significant noise so that a clear transition in SLS signal during heating could not be discerned. Because D2 O has previously been reported to be highly effective at increasing polio virus thermal stability,17,18 IPV3 samples in D2 O were further studied by DLS and CD spectroscopies. With DLS, there was almost no size change observed even up to 90◦ C (Fig. 7a). A single peak with a diameter of approximately 30 nm dominated the DLS results without appearance of a significant amount of larger species. By comparison, at the same solution temperatures, aggregates of approximately 1 :m with at a volume percentage of approximately 50% appeared for IPV3 samples in PBS buffer alone. The thermal transition temperature observed with CD spectroscopy for IPV3 in D2 O (open circle) was approximately 5◦ C higher than samples in PBS buffer (closed square), which is quite close to that observed by fluorescence (Fig. 7b). Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
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The use of the extrinsic fluorescence of SYPRO Orange was unsuccessful in terms of distinguishing excipient effects either by isothermal incubation or thermal ramping stress (see Suppl. Material). For all samples examined for 1 h incubation at 50◦ C (Suppl. Fig. S6), the fluorescence intensity dropped to approximately 50% of the original signal at 20◦ C without peak shape change. During thermal ramping (Suppl. Fig. S3D), the SYPRO Orange intensity for IPV3 samples in the absence and presence of the tested excipients were fairly close to each other. After normalization, all intensity trend lines were overlapping and could not be distinguished from one another. The effect of pH on IPV3 ELISA antigen content was also examined immediately after dialysis against the phosphatecitrate buffer at pH values from 3.0 to 8.0. As shown in Fig. 9a, IPV3 only maintained approximately 20% and 60% of the ELISA antigen content at pH 3 and 4, respectively. In contrast, the ELISA antigen contents were not significantly different from each other at pH 5, 6, 7, and 8, with more than 80% of the content remaining. The losses of ELISA antigen content at low pH are readily explained by the results shown in Fig. 5. At acidic pH, IPV3 samples formed large virion aggregates and lost both tertiary and secondary structure compared with results at higher pH. The fluorescence Tm of IPV 3 was also obtained in an attempt to correlate it with the ELISA antigen content (Fig. 9b). At pH 3, no clear transition could be seen because of the extensive unfolding of the capsid proteins. IPV3 manifested lower Tm values at pH 4 (∼44◦ C) and pH 5 (∼50◦ C) than at pH 6–8 (∼56◦ C–57 ◦ C). Thus, the thermal stability of IPV3 under different pH condition also correlated well with the remaining ELISA antigen content. All of the biophysical characterizations of IPV3 in the above pH conditions were included in the EPD (Fig. 6). For clarity and direct comparison, fluorescence, CD, SLS, and DLS data of IPV3 at different pH values at 25◦ C are summarized in Suppl. Fig. S7 and could be referred from Fig. 5.
DISCUSSION
Figure 7. The effects of selected excipients on IPV3 thermal stability and potency. (a) Remaining titer of IPV3 samples incubated at 50◦ C for 1h (open bar). The remaining percentage of titer was calculated based on the fresh dialyzed and 1h incubated 50◦ C PBS samples as 100% and 0%, respectively. Tm was determined by subtracting the Tm of IPV3 samples in PBS from that of IPV3 in the presence of the excipients. All the Tm values were fitted with a two-state model of the intensity ratio of 350–330nm of Trp intrinsic fluorescence. (b) The correlation between the Tm and remaining relative titer. (c) SLS of IPV3 with selected excipients. Triplicates were performed to the above experiments and the mean values were plotted with the standard deviation as the error bar for panel a and b. Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
We have characterized IPV3 in terms of size, morphology, tertiary, and secondary structure and ELISA antigen content, and defined the most stable conditions as pH 7 at less than 40◦ C. Because of the complex structure of the virion and low resolution of the techniques used here, we are not able to provide detailed structural information. Rather, to facilitate formulation development, our goal is to identify physical methods that can measure IPV thermal stability and provide a good correlation between stability and ELISA antigen content. Intrinsic fluorescence was identified as the most sensitive technique among those examined, with fluorescence peak position the most useful property for the desired studies. To achieve high throughput, the fluorescence/light scattering-based Avacta system was selected for excipient formulation studies. This work is not intended to discern the protective mechanisms of effective excipients, but aspects of potential mechanisms can be considered. Saccharides and polyols probably exert their protective effects through the preferential exclusion mechanism. This mechanism has been thoroughly described in the literature.30,31 Briefly, because of the preferential exclusion of the stabilizer from the surface of protein molecules, the free energy of the protein is increased. The magnitude of the free energy increase varies directly with the protein surface area DOI 10.1002/jps.23801
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Figure 8. The effect of high-concentration deuterium on the size (a) and CD (b) of IPV3 upon thermal ramping. (a) DLS of IPV3 in D2 O (>95%) at different temperatures. (b) Wavelength of minimum intensity of IPV3 in D2 O (>95%) (open circle) upon thermal ramping, as compared with IPV3 in PBS (closed square).
Figure 9. (a) The relative titer remaining of IPV3 samples dialyzed at varied pH. (b) The fitted fluorescence peak position transition temperature of IPV3 samples at different pH. The star notified that no clear transition temperature could be fitted for IPV3 samples at pH 3. Triplicates were performed in the above experiments and the mean values were plotted with the standard deviation as the error bars.
exposed to solvent, and thus is much larger for the unfolded than for the native state of the protein. As a result, in the presence of a preferentially excluded excipient, the native state is stabilized and the free energy of unfolding is increased.31,32 Stabilization through the preferential exclusion mechanism requires relatively high concentrations of solutes. This requirement is consistent with our observation of protective effects of trehalose, sucrose, and sorbitol at higher concentration (25%, w/w) but almost no increase in IPV3 stability with relatively low concentration (5%). Glycine, mannitol, and glycerol might also exert their effects on IPV3 through preferential exclusion,30,33,34 but are typically less effective than disaccharides. Mannitol and glycerol did not inhibit aggregation, whereas glycine actually promoted self-association at earlier times (see Suppl. Fig. S5). Sodium citrate greatly enhanced IPV3 thermal stability as detected by fluorescence and ELISA antigen content. This may also be attributed to the preferential exclusion mechanism but binding mechanisms are also possible. As expected, urea decreased the Tm of IPV3 but it also delayed the onset of aggregation. Urea is well known to unfold proteins as well as increase protein solubility.35–37 The precise mechanism of this activity is still under investigation, with either preferential binding to the unfolded state or altering of its solvent environment as commonly suggested explanations.36
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The stabilizing effects of D2 O on proteins and vaccines have been widely reported.38–41 This phenomenon has generally been attributed to enhanced hydrophobic interactions within or between proteins and/or the presence of stronger hydrogen bonding in D2 O.39,40 In the case of polio vaccine, the protective effect of D2 O was discovered more than a decade ago.18,42,43 In a previous study, Trp fluorescence polarization indicated that poliovirus existed in a more rigid environment in D2 O than H2 O.14 In addition, the amount of water penetrating into the polio virus virion was substantially reduced by D2 O with such an effect attributed to a reduction in the swelling of the virus in response to temperature increases.14 In the same study, however, the observed CD signals were low and noisy because of low virus concentration, and the effects of the D2 O were not clearly demonstrated.14 In the current study of IPV3, we further provide clear evidence of stabilization by D2 O. The Tm of IPV3 from fluorescence and CD measurements increased by approximately 5◦ C, with no significant size changes seen even at very high temperature (∼90◦ C) based on both SLS and DLS studies. This suggests that IPV3 did not swell or aggregate appreciably during heating, perhaps because of an increase in virion rigidity in D2 O. We speculate that CD spectra were better resolved than in earlier studies because of the high purity and concentration of the IPV3 sample used in this study.
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The capsid proteins of IPV3 responded to pH and thermal stress somewhat differently. For instance, both an acidic environment and 50◦ C isothermal incubation significantly reduced the ELISA antigen content. The size distribution and biophysical properties under the two treatments were completely different. At pH 3, with DLS analysis, a peak over 1000 nm in diameter appeared, along with low Trp fluorescence intensify and diminished CD signal (Suppl. Fig. S7). These changes account for the loss of antigen content. In contrast, with isothermal incubation at 50◦ C, we did not see significant changes in IPV3 size and fluorescence peak position (Fig. 2 and Suppl. Fig. S2). This observation may be because of the fact that all of the physical data are of lower resolution and reflects averaged signal over many viral components. Thus, the antigen binding site may have gone through local environment changes that cannot be detected with our methods, such as fluorescent spectroscopy, because of the large number of Trp residues. Nevertheless, we were able to use fluorescence-based Tm values to compare the IPV3 thermal stability under various solution conditions. The relative IPV3 stabilities measured with fluorescence spectroscopy matched well with those obtained with ELISA antigen content. Thus, biophysical assessment of thermal stability with fluorescence spectroscopy potentially provides a valuable tool for formulation development of IPV3 and perhaps other virus-based vaccines as well.
ACKNOWLEDGMENTS The authors thank Dr. Sangeeta Joshi for her kind help on the pH and thermal stability screening study, Dr. Hardeep Samra in MedImmune for his advice on SYPRO Orange dye application, and Dorothy Dill for her patient help on TEM. We are grateful to the Pall Corporation (Port Washington, New York) for the loan of the Optim 1000 and accessories. We also thank Sanofi Pastuer for funding and the supplies of the IPV3 and antibodies for the ELISA.
REFERENCES 1. WHO. 2003. Global Polio eradication initiative strategic plan 2004–2008. 1–40 http://www.polioeradication.org/content/publications/ 2004stratplan.pdf last accessed Nov. 21, 2013. 2. WHO. 2011. Pliomyelities fact sheet http://www.poliofree.org/ polio-fact-sheet.htm last accessed Nov. 21, 2013 . 3. Dowdle WR, De Gourville E, Kew OM, Pallansch MA, Wood DJ. 2003. Polio eradication: The OPV paradox. Rev Med Virol 13(5):277– 291. 4. Friedrich F. 1997. Rare adverse events associated with oral poliovirus vaccine in Brazil. Braz J Med Biol Res 30(6):695–703. 5. Kew OM, Sutter RW, de Gourville EM, Dowdle WR, Pallansch MA. 2005. Vaccine-derived polioviruses and the endgame strategy for global polio eradication. Annu Rev Microbiol 59:587–635. 6. Minor P. 2009. Vaccine-derived poliovirus (VDPV): Impact on poliomyelitis eradication. Vaccine 27(20):2649–2652. 7. Salas-Peraza D, Avila-Aguero ML, Morice-Trejos A. 2010. Switching from OPV to IPV: Are we behind the schedule in Latin America? Expert Rev Vaccines 9(5):475–483. 8. Tajikis WHOCO, Europe WRO, Contro ECDP. 2010. Outbreak of poliomyelitis in Tajikistan in 2010: Risk for importation and impact on polio surveillance in Europe? Eurosurveillance 15(17):5–7. 9. Atkinson W, Hamborsky J, McIntyre L, Wolfe S. 2008. Epidemiology and prevention of vaccine-preventable diseases. 10th ed. Department of Health and Human Services, Washington D.C. Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
10. Chen DX, Kristensen D. 2009. Opportunities and challenges of developing thermostable vaccines. Expert Rev Vaccines 8(5):547–557. 11. Hogle JM, Chow M, Filman DJ. 1985. Three-dimensional structure of poliovirus at 2.9A resolution. Science 229(4720):1358–1365. 12. Bubeck D, Filman DJ, Cheng N, Steven AC, Hogle JM, Belnap DM. 2005. The structure of the poliovirus 135S cell entry intermediate at 10angstrom resolution reveals the location of an externalized polypeptide that binds to membranes. J Virol 79(12):7745–7755. 13. Brandenburg B, Lee LY, Lakadamyali M, Rust MJ, Zhuang X, Hogle JM. 2007. Imaging poliovirus entry in live cells. PLoS Biol 5(7):e183. 14. Chen CH, Wu R, Roth LG, Guillot S, Crainic R. 1997. Elucidating mechanisms of thermostabilization of poliovirus by D2 O and MgCl2 . Arch Biochem Biophys 342(1):108–116. 15. Grimmel M, Zibirre R, Koch G. 1983. Fluorescence spectrophotometric study of structural alterations in the capsid of poliovirus. Arch Virol 78(3–4):191–201. 16. Westdijk J, Brugmans D, Martin J, van’t Oever A, Bakker WAM, Levels L, Kersten G. 2011. Characterization and standardization of Sabin based inactivated polio vaccine: Proposal for a new antigen unit for inactivated polio vaccines. Vaccine 29(18):3390–3397. 17. Milstien JB, Lemon SM, Wright PF. 1997. Development of a more thermostable poliovirus vaccine. J Infect Dis 175:S247–S253. 18. Verheyden B, Andries K, Rombaut B. 2001. Capsid and RNA stabilisation of the oral polio vaccine. Vaccine 19(15–16):1899–1905. 19. Kim JH, Iyer V, Joshi SB, Volkin DB, Middaugh CR. 2012. Improved data visualization techniques for analyzing macromolecule structural changes. Protein Sci 21(10):1540–1553. 20. Kueltzo LA, Ersoy B, Ralston JP, Middaugh CR. 2003. Derivative absorbance spectroscopy and protein phase diagrams as tools for comprehensive protein characterization: A bGCSF case study. J Pharm Sci 92(9):1805–1820. 21. Maddux NR, Joshi SB, Volkin DB, Ralston JP, Middaugh CR. 2011. Multidimensional methods for the formulation of biopharmaceuticals and vaccines. J Pharm Sci 100(10):4171–4197. 22. Rezapkin G, Martin J, Chumakov K. 2005. Analysis of antigenic profiles of inactivated poliovirus vaccine and vaccine-derived polioviruses by block-ELISA method. Biologicals 33(1):29–39. 23. Ferguson M, Wood DJ, Minor PD. 1993. Antigenic structure of poliovirus in inactivated vaccines. J Gen Virol 74(4):685–690. 24. Boublik M, Drzeniek R. 1977. Structural subunits of poliovirus particles by electron microscopy. J Gen Virol 37(1):127– 134. 25. Eftink MR. 1991. Fluorescence techniques for studying protein structure. Methods Biochem Anal 35:127–205. 26. Volkin DB, Burke CJ, Marfia KE, Oswald CB, Wolanski B, Middaugh CR. 1997. Size and conformational stability of the hepatitis A virus used to prepare VAQTA, a highly purified inactivated vaccine. J Pharm Sci 86(6):666–673. 27. Filman DJ, Syed R, Chow M, Macadam AJ, Minor PD, Hogle JM. 1989. Structural factors that control conformational transitions and serotype specificity in type-3 poliovirus. Embo J 8(5):1567–1579. 28. Goldberg DS, Bishop SM, Shah AU, Sathish HA. 2011. Formulation development of therapeutic monoclonal antibodies using highthroughput fluorescence and static light scattering techniques: Role of conformational and colloidal stability. J Pharm Sci 100(4):1306– 1315. 29. Ausar SF, Chan JD, Hoque W, James O, Jayasundara K, Harper K. 2011. Application of extrinsic fluorescence spectroscopy for the high throughput formulation screening of aluminum-adjuvanted vaccines. J Pharm Sci 100(2):431–440. 30. Arakawa T, Timasheff SN. 1982. Mechanism of stabilization of proteins by glycerol and sucrose. Seikagaku 54(11):1255–1259. 31. Lee JC, Timasheff SN. 1981. The stabilization of proteins by sucrose. J Biol Chem 256(14):7193–7201. 32. Foster TM, Dormish JJ, Narahari U, Meyer JD, Vrkljan M, Henkin J, Porter WR, Staack H, Carpenter JF, Manning MC. 1996. Thermal stability of low molecular weight urokinase during heat treatment. 3. Effect of salts, sugars and Tween 80. Int J Pharm 134(1–2):193–201. DOI 10.1002/jps.23801
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33. Haque I, Singh R, Moosavi-Movahedi AA, Ahmad F. 2005. Effect of polyol osmolytes on DeltaG(D), the Gibbs energy of stabilisation of proteins at different pH values. Biophys Chem 117(1):1– 12. 34. Santoro MM, Liu YF, Khan SMA, Hou LX, Bolen DW. 1992. Increased thermal-stability of proteins in the presence of naturallyoccurring osmolytes. Biochemistry 31(23):5278–5283. 35. Timasheff SN. 2002. Protein-solvent preferential interactions, protein hydration, and the modulation of biochemical reactions by solvent components. Proc Natl Acad Sci USA 99(15):9721–9726. 36. Bennion BJ, Daggett V. 2003. The molecular basis for the chemical denaturation of proteins by urea. Proc Natl Acad Sci USA 100(9):5142– 5147. 37. Chen J, Liu Y, Li X, Wang Y, Ding H, Ma G, Su Z. 2009. Cooperative effects of urea and L-arginine on protein refolding. Protein Expr Purif 66(1):82–90.
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38. Sen A, Balamurugan V, Rajak KK, Chakravarti S, Bhanuprakash V, Singh RK. 2009. Role of heavy water in biological sciences with an emphasis on thermostabilization of vaccines. Expert Rev Vaccines 8(11):1587–1602. 39. Cioni P, Strambini GB. 2002. Effect of heavy water on protein flexibility. Biophys J 82(6):3246–3253. 40. Efimova YM, Haemers S, Wierczinski B, Norde W, van Well AA. 2007. Stability of globular proteins in H2 O and D2 O. Biopolymers 85(3):264–273. 41. Verheul M, Roefs SP, de Kruif KG. 1998. Aggregation of betalactoglobulin and influence of D2 O. FEBS Lett 421(3):273–276. 42. Milstien JB, Lemon SM, Wright PF. 1997. Development of a more thermostable poliovirus vaccine. J Infect Dis 175 Suppl 1:S247–S253. 43. Wu R, Georgescu MM, Delpeyroux F, Guillot S, Balanant J, Simpson K, Crainic R. 1995. Thermostabilization of live virus vaccines by heavy water (D2 O). Vaccine 13(12):1058–1063.
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Preformulation Study of Highly Purified Inactivated Polio Vaccine, Serotype 3 WEI QI,1 YUHONG ZENG,2 SCOTT ORGEL,1 ALAIN FRANCON,3 JAE HYUN KIM,2 THEODORE W. RANDOLPH,4 JOHN F. CARPENTER,1 C. RUSSELL MIDDAUGH2 1
Department of Pharmaceutical Sciences, University of Colorado Anschutz Medical Campus, Aurora, Colorado University of Kansas, Macromolecule and Vaccine Stabilization Center, Lawrence, Kansas 3 Sanofi Pasteur, March l’Etotile, France 4 Department of Biological and Chemical Engineering, University of Colorado, Boulder, Colorado 2
Received 14 September 2013; revised 23 October 2013; accepted 6 November 2013 Published online 26 November 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI 10.1002/jps.23801 ABSTRACT: To improve the effectiveness of the polio vaccination campaign, improvements in the thermal stability of the vaccine are being investigated. Here, inactivated polio vaccine, serotype 3 (IPV3) was characterized via a number of biophysical techniques. The size was characterized by transmission electronic microscopy and light scattering. The capsid protein conformation was evaluated by intrinsic fluorescence and circular dichroism (CD), and the D-antigen content by enzyme-linked immunosorbent assay (ELISA). The pH thermal stability of IPV3 (pH 3.0–8.0; 10◦ C–87.5◦ C) was evaluated by fluorescence, CD, and static light scattering. The transition temperatures reflect the responses, respectively, of tertiary structure, secondary structure, and size to applied thermal stress. The data were summarized as empirical phase diagrams, and the most stable conditions were found to be pH 7.0 with temperature lower than 40◦ C. CD detected a higher transition temperature for capsid protein than that for RNA. The effects of certain excipients on IPV3 thermal stability and antigen content were evaluated. The results of their effects, based on intrinsic fluorescence and ELISA, were in good agreement, suggesting the feasibility of applying intrinsic fluorescence as a high-throughput tool for formulation development. The study improves the understanding C 2013 Wiley of IPV3 thermal stability, and provides a starting point for future formulation development of IPV3 and other serotypes. Periodicals, Inc. and the American Pharmacists Association J Pharm Sci 103:140–151, 2014 Keywords: stability; excipients; vaccines; physical characterization; pre-formulation; stabilization
INTRODUCTION Since 1998, the World Health Organization (WHO) has been campaigning for the global eradication of polio.1 In 2009, over 361 million children were immunized.2 Disappointingly, in resource-poor regions, the most commonly used oral polio vaccine (OPV) formulation had reduced potency and generated live vaccine-derived poliovirus and rare vaccine-associated paralytic poliomyelitis (VAPP).3–7 Additionally, previously poliofree countries could be reinfected because of imported virus from polio-endemic areas, that is, the 2010 polio outbreak in Tajikistan.8 Therefore, a safer vaccine is desired to eliminate VAPP and vaccine-derived poliovirus circulation. In the USA and UK, inactivated polio vaccine (IPV) replaced OPV in 2000 and 2004, respectively.9 IPV is currently formulated as a liquid (medium 199) and is stable for over a year at 4◦ C, but loses potency in only weeks at 25◦ C or 37◦ C.10 To preserve vaccine potency during shipping, storage, and distribution, an effective cold chain is necessary but practically difficult in the resource-poor areas where the vaccine is most needed. Hence, a thermally stable formulation is strongly desired to maintain potency before ultimate administration of the vaccine.
Correspondence to: C. Russell Middaugh (Telephone: +785-864-5813; Fax: +785-864-5814; E-mail: [email protected]) This article contains supplementary material available from the authors upon request or via the Internet at http://onlinelibrary.wiley.com/. Journal of Pharmaceutical Sciences, Vol. 103, 140–151 (2014)
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To develop new poliovirus vaccine formulations, it is critical to determine the effect of potential formulation additives on the structural stability of IPV. High-resolution structural information concerning poliovirus has been provided by EM11 and X-ray12 analysis. The process by which poliovirus enters cells was recently characterized by fluorescence microscopy.13 For rapid development of new formulations, however, more readily detected biophysical properties, rather than high-resolution structures, are needed. Tryptophan (Trp) fluorescence intensity was previously used to study the thermal stability of poliovirus in the presence of excipients14 and under different pH conditions.15 Circular dichroism (CD) spectroscopy and differential scanning microcalorimetry were also used in attempts to monitor poliovirus thermal stability,14 but the results were compromised by extremely low sample concentration. Currently, changes in the physical properties of IPVs upon thermal stress are poorly understood, with only limited data available at temperatures higher than 45◦ C.10 In addition, the routine active antigen (D-antigen) content test was not standardized, and inconsistencies among manufacturers and laboratories exist. These probably result from the variation in antibodies and procedures used in the D-antigen test.16 Furthermore, many studies14–18 have investigated either vaccine structure or potency individually, but the combined results are rarely reported. Finally, a more rapidly measured biophysical property might be useful as a complementary measurement of potency. A correlation between such a physical property and IPV potency, if it exists, would greatly facilitate rational and effective IPV formulation development.
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In this work, we employed highly purified inactivated polio vaccine, serotype 3 (IPV3) samples from a commercial manufacturer to: (1) characterize IPV3 by different biophysical techniques, focusing on size and capsid protein structure; (2) establish a pH thermal stability profile of IPV3 employing empirical phase diagrams (EPDs); (3) evaluate IPV3 active antigen content with and without excipients upon thermal stress; and (4) correlate certain biophysical properties with antigen content.
MATERIALS AND METHODS Materials Inactivated polio vaccine, serotype 3 was shipped from Sanofi Pasteur (Lyon, France) in medium 199 without phenol red (see Sigma #M9163 for detailed composition). During shipping and storage, the temperature was kept between 2◦ C and 8◦ C. The IPV3 concentration in terms of protein mass was approximately 20 :g/mL and the D-antigen content was approximately 1128 UD/mL. The coating monoclonal antibody, mAb #520, was from NIBSC (National Institute for Biological Standards and Control). The secondary antibody, mAb #4B7-1H8-2E10, was obtained from Sanofi Pasteur and kept at −80◦ C until use. Anti-mouse IgG peroxidase antibody from goat (A3673) was purchased from Sigma (St. Louis, Missouri). Nonfat milk powder was purchased from Becton Dickson (Franklin Lakes, New Jersey). Ultrapure urea and deuterium oxide (D2 O, 99.9%) were from MP Biomedicals LLC (Aurora, Ohio). Sucrose and trehalose were obtained from J.T. Baker (Phillipsburg, New Jersey). Mannitol U.S.P. was from Spectrum (New Brunswick, New Jersey). Sorbitol was purchased from Calbiochem (Merck KGaA, Darmstadt, Germany). Glycine and anhydrous glycerol were purchased from Fisher (Fair Lawn, New Jersey). SYPRO Orange was from Invitrogen (Eugene, Oregon). All other chemicals were obtained from Sigma. Sample Preparation Inactivated polio vaccine, serotype 3 samples were first dialyzed R Dialysis Cassettes with a 10 kDa molecular in Slide-A-Lyzer weight (MW) cutoff (Pierce, Rockford, Illinois) against 20 mM phosphate-citrate buffer with a fixed ionic strength of 0.15M by adding an appropriate amount of NaCl, pH 7.0 at 4◦ C overnight before any other specified treatment. Dynamic Light Scattering Backscattering from solutions containing IPV3 in approximately 150 :L samples in a low-volume quartz cuvette with 1 cm path length was measured at 173◦ with a Malvern Zetasizer (Worcestershire, UK). The temperature was set at either 25◦ C or ramped from 25◦ C to 90◦ C with 5◦ C steps. Samples were held at each temperature for 2–3 min before data acquisition. Transmission Electronic Microscopy Three microliter IPV3 samples was placed onto a carbon-coated grid and allowed to adsorb for 3 min. The grid then was washed twice with ddH2 O. The sample was stained with 2% uranyl acetate solution for 2 min, air dried and then imaged by a model Technai G2 transmission electronic microscopy (TEM) equipped with a Gatan Ultrascan digital camera from FEI (Hillsboro, Oregon) at a magnification of 60,000×. DOI 10.1002/jps.23801
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CD Spectroscopy Dialyzed IPV3 samples were placed in Millipore Centricon tubes (MW cutoff 10 kDa) and centrifuged at approximately 2500 × g in a Sorvall Centrifuges RT 6000D (Wilmington, Delaware) at a temperature between 2◦ C and 6◦ C. The concentration of virus was determined by UV absorption spectroscopy. The UV absorbance at 280 nm in a 1 cm path length cuvette was at least 0.2 units. Approximately 400 :L of sample were then transferred into a 1 mm path length cuvette and the CD spectrum of the sample was measured from 190 to 300 nm with a scan step of 1 nm by a Chirascan Plus spectropolarimeter from Applied Photophysics (Leatherhead, UK). The CD spectra were analyzed by the CDNN software package to obtain the percentage of various secondary structure types. Fluorescence Spectroscopy Seven hundred microliter samples of dialyzed IPV3 was added to a 1 cm path length cuvette. Intrinsic Trp fluorescence spectra were acquired with a Photon Technology International spectrofluorometer (Lawrenceville, New Jersey) equipped with a Peltier thermostatted cuvette holder. The samples were excited at 295 nm (>95% Trp emission) and the emission spectra were collected from 300 to 400 nm, with a 1 nm step and 1 s integration time. Static light scattering (SLS) was collected at 295 nm by a second photomultiplier with the same instrument located 180◦ to the fluorescence emission detector. For stability tests with excipients, dialyzed IPV3 samples were concentrated approximately 20-fold by centrifugation at approximately 2500 × g in Millipore Centricon tubes with a 10kDa cutoff (2◦ C–6◦ C). Concentrated samples were mixed with specific excipient solutions in equal volumes to a final IPV3 concentration that was approximately 10-fold that of the freshly dialyzed sample. To prepare IPV3 samples in D2 O, IPV3 in medium 199 was centrifuged in Millipore Centricon tubes with 10-kDa cutoffs to approximately 5% or less of the initial volume, and then resuspended in D2 O to the initial volume. This was repeated three times to ensure that the D2 O content was greater than 95%. All the stability test experiments with excipients were carried out in an Optim 1000 spectrofluorimeter from Avacta (Wetherby, UK). Approximately 10 :L of IPV3 mixed with individual excipients was directly loaded into the micro cuvette arrays. The intrinsic fluorescence was excited at 266 nm with a laser source, and the emission spectra were collected from 250 to 500 nm with default settings. SLS data were collected simultaneously at 473 nm using a separate laser. For extrinsic fluorescence experiments, SYPRO Orange (Invitrogen) was mixed with IPV3 samples to a final dye concentration of 5× dilution from the original stock. The samples were excited at 473 nm and the spectra were collected from 450 to 700 nm. For isothermal incubation, the spectra were collected at specific temperatures (40◦ C, 50◦ C, 60◦ C, and 70◦ C) every 5 min during 3 h incubations. For temperature ramping experiments, the temperature was varied from 20◦ C to 90◦ C at a 1◦ C per intervals, and spectra were collected at each temperature after a hold time of 2–3 min. Temperature interval was set at 1◦ C to capture smoother transitions in the presence of different excipients. The raw spectra were processed with the embedded Optim1000 software (Avacta) using default settings. The ratio of the intensity at 350 to 330 nm was used as an indication of the relative abundance of unfolded to native protein. This Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
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intensity ratio provided similar transition information to that of the fitted peak position (see Suppl. Material, Suppl. Fig. S1). For CD and fluorescence thermal ramping experiments, the intensity or peak position was fitted to a standard two-state protein unfolding model, with the transition temperature, Tm , the mid-point of the transition. pH and Thermal Stability For pH and thermal stability studies, IPV3 was first dialyzed into 20 mM phosphate-citrate buffer with a fixed ionic strength of 0.15 M by adding an appropriate amount of NaCl. The pH was varied from 3.0 to 8.0 at 1.0 unit intervals. Samples were analyzed by CD, Trp fluorescence, and SLS, with temperature ramping from 10◦ C to 87.5◦ C at 2.5◦ C intervals. Samples were equilibrated at each temperature for 2–3 min before data acquisition. EPDs for IPV3 were generated as described previously.19–21 Temperature interval was set up at 2.5◦ C to be compatible with the MatLab code for EPD at that time. Five sets of data at each pH was used to construct EPDs, including SLS, fluorescence peak position, fluorescence intensity, and CD values at 205 and 220 nm. As described later in the text, these diagrams reveal different physical states of the vaccine under the specified pH and temperature conditions. Enzyme-Linked Immunosorbent Assay Indirect enzyme-linked immunosorbent assays (ELISAs) were performed to characterize the virus content of IPV3 in terms of D-antigen content. The measurement of D-antigen content has been routinely used to correlate with the protective immunogenicity of IPV.16,22 Briefly, 50 :L of coating antibody, mAb #520, were added to wells in a 96-well plate (PI-15041, Thermal Scientific) and incubated at 4◦ C overnight. mAb #520 was reported to bind to the active antigen site 1 for IPV3.22,23 Plates were washed at least three times with washing buffer [1× PBS (phosphate-buffered saline) with 0.025%, v/v, Tween 20 and 0.21% skim milk]. Sixteen microliter IPV3 samples was first diluted 10-fold into the washing buffer to a final volume of 160 :L. Eighty microliter out of the 160 :L was then added to the first well on the plate. The remaining 80 :L was diluted by washing buffer to 160 :L and another 80 :L was loaded to the second well, and so on. At each dilution, 80 :L sample was transferred into the corresponding well on the plate in sequence for that sample. Plates were then incubated at 37◦ C for 2 h. After another washing step, 50 :L of secondary antibody solution were added and incubated at 37◦ C for 1 h. Fifty microliter of tertiary antibody solution was added after washing and incubated at 37◦ C for another hour. Finally, 50 :L of TMB (3, 3 , 5, 5 tetramethylbenzidine) was added to each well, and the plate was held at room temperature for 10–15 min). HCl (1 N) was added to stop the reaction and the absorbance was read immediately at 450 nm with 630 nm as a reference, employing a VERSMax plate reader (Molecular Devices, Sunnyvale, California). Inactivated polio vaccine, serotype 3 samples in PBS after dialysis were used to determine assay reproducibility. Titer values were reproducible with a standard deviation within 15% for seven tested samples on different days (data not shown). The typical loss of the D-antigen from the medium 199 during dialysis was between 10% and 20% (data not shown). Incubation at 50◦ C for 1 h was chosen as an accelerated degradation condition to screen for the effects of excipients on Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
IPV3 stability. In preliminary tests, IVP3 stability was tested after incubation at 50◦ C and 60◦ C, with incubation times of 15, 30, and 60 min. Degradation at 60◦ C was rapid, leading to a complete loss of antigen detectable in the ELISA assay after only 30 min, whereas 10%–15% of the original antigen content remained after incubation at 50◦ C for 1 h. This level of thermal stress permitted an adequate window to detect effects of the excipients. A standard 4-parameter fit was performed for all ELISA data using Origin software, with EC50 values fit for each sample. The EC50 value for IPV3 samples immediately after dialysis was taken as 100%, and that for IPV3 samples after 1 h incubation at 50◦ C as 0%. To evaluate the effects of different excipients upon thermal stress, the EC50 values of those samples were normalized to the difference between 0% and 100% controls. Such values were used to represent the ELISA antigen content of the IPV3, which is referred to as percent of antigen content in the text. Sample Replicates and Error For all experiments, at least triplicates were studied unless otherwise specified. The error bars in the figures reflect the standard deviation of the specific measurement. For clarity and to avoid confusion, certain figures did not include the error bars.
RESULTS Biophysical Characterization of IPV3 Transmission electronic microscopy (TEM) and dynamic light scattering (DLS) were first applied to characterize the size and morphology of IPV3. As shown in Fig. 1a, IPV3 in freshly dialyzed samples appeared as spherical particles with diameters of approximately 30 nm, which is close to previous literature reports.11,24 No empty particles were observed, suggesting that the single-stranded RNA had not leaked out of the vaccine capsid. Although TEM directly represents IPV3 morphology and size, it is neither convenient nor practical for obtaining realtime images of IPV3 under thermal stress. In addition, formulations containing high concentrations of disaccharides may not be suitable for TEM measurements. Furthermore, the sampling area of a TEM image represents only a small fraction of the total sample, and the measurement might not be accurate because of sample preparation effects. Therefore, DLS was used as an alternative approach to measure virus particle size and size distributions. In Fig. 1b, for a freshly dialyzed IPV3 sample, a single peak dominated the volume distribution with a mid-point diameter of 33.9 ± 1.9 nm. The diameter determined from a corresponding intensity measurement was slightly larger than the reported value of approximately 30 nm, presumably because light scattering provides a hydrodynamic radius, which is indicative of the apparent size of the hydrated particle and could be influenced by even very small amount of larger particles. The intrinsic fluorescence of Trp residues has been routinely used to probe the tertiary structure of proteins in different solvent conditions. A fluorescence emission maximum from 320 to 330 nm suggests that on average Trp residues are located in a relatively apolar environment as seen typically in native states, whereas a wavelength maximum from 350 to 355 nm suggests Trp residues are more exposed to the aqueous solvent as seen in more structurally altered states.25 In Fig. 1c, we observed an emission peak for IPV3 with a maximum near 335 nm, DOI 10.1002/jps.23801
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Figure 1. Characterization of IPV3 in PBS buffer after dialysis and before any thermal treatment or exposure to excipients. (a) TEM images of IPV3. The black scale bar at the left corner is 50nm in length. (b) DLS of IPV3 in terms of volume percentage. (c) Intrinsic Trp fluorescence of IPV3 excited at 295nm. (d) CD spectrum of IPV3 from 195 to 300nm.
suggesting that the Trp residues in the virus proteins are on average found in a relatively apolar environment. The IPV3 sequence (http://www.ncbi.nlm.nih.gov/protein/AAA19636), contains 780 Trp residues located in a single virion (13 Trp residues present in single copies of the capsid proteins VP1 to VP4, with 60 copies of VP1 to VP4 in each virion). Thus, the signal seen reflects the averaged status of a large number of Trp residues, negating any high-resolution structural interpretation of spectral changes. Circular dichroism spectroscopy was used to examine the secondary structure of the capsid proteins. In Fig. 1d, a strong negative band around 205 nm is seen, together with a positive band from 260 to 280 nm. A minimum at 205 nm usually reflects the presence of crossed $-structure. The positive ellipticity near 265 nm may arise from the RNA component, as previously reported for hepatitis A virus.26 To further probe the nature of the secondary structure, the spectrum was deconvoluted to compare the secondary structure components reported for type 3 poliovirus (with PDB ID 2plv).27 In the spctra of IPV3 obtained between 190 and 260 nm, approximately 10% helix, 33% $-sheet, 25% $-turn, and 33% random coil were estimated. For type 3 poliovirus, 9% helix and 29% $-sheet were reported for VP 1 and 3, with 12% helix and 37% $-sheet for VP 2, and 10% helix and 14% $-sheet for VP 4, with the remaining structures being turns and/or unspecified structures (http://www.pdb.org/pdb/ explore/remediatedSequence.do?structureId=2PLV). Thus, the secondary structural content from our measurements were similar to previous estimates, even though the RNA component may interfere with the signal below 240 nm.26 Again, as in DOI 10.1002/jps.23801
the fluorescence results, we emphasize that CD spectroscopy measured the combined structural properties of all the capsid proteins. Thermal Stability of IPV3 at pH 7 as Assessed by Intrinsic Fluorescence Spectroscopy, CD Spectroscopy, SLS, and DLS Inactivated polio vaccine, serotype 3 is currently formulated at pH 7.0–7.4. To provide reference data to compare against results for IPV3 in other solution conditions, analyses of IPV3 at pH 7 were performed first using Trp and SYPRO Orange fluorescence, CD spectroscopy, SLS, and DLS. Intrinsic Trp fluorescence emission spectra collected at different temperatures are shown in Fig. 2a. As the temperature increased, the fluorescence signal gradually decreased, and the spectra broadened. Concomitantly, the central peak position red shifted from approximately 335 nm to 346 nm. Temperature-dependent transition curves of maximum intensity and peak position are shown in Figures 2b and 2c, respectively. The intensity curve showed a gradual decrease with increasing temperature, with an inflection point of approximately 50◦ C. In contrast, a more abrupt transition was observed with the curve for peak position, indicating increasing exposure of the Trp residues to a more hydrophilic environment with heating. The Tm obtained from the fluorescence peak position data was 57.5 ± 1.8◦ C. Intrinsic fluorescence was also used to probe vaccine structural changes for IPV3 during isothermal incubation, at 40◦ C, 50◦ C, 60◦ C, and 70◦ C. As shown in Fig. 2d, no significant peak position change was observed as a function of incubation time for IPV3 at 40◦ C. During incubation at 50◦ C, a gradual but Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
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Figure 2. Intrinsic Trp fluorescence spectra of IPV3 upon thermal stress. (a) Emission spectra of intrinsic fluorescence collected from 300 to 400nm with excitation at 295nm. The arrow direction indicates increased 5◦ C temperature steps, illustrated by alternative dark and gray lines. (b) Normalized intrinsic Trp fluorescence intensity as a function of temperature. The peak position of the maximum intensity for intrinsic Trp fluorescence as a function of increasing temperature (c) and isothermal incubation (d) at indicated temperatures. Triplicates were performed to the above experiments and the mean values were plotted with the standard deviation as the error bar in panel b and c.
subtle transition took place with the peak position red-shifting approximately 2 nm. When IPV3 was incubated at 60◦ C (close to the Tm ), the peak position red-shifted about 5 nm within 2000 s. After longer incubation times, the peak position slightly decreased, which may reflect the formation of large aggregates (see Suppl. Material, Supp. Fig. S2). At an even higher temperature (70◦ C), the peak position red-shifted up to approximately 10 nm within a shorter time period (∼1000 s), suggesting major structural disruption. Similar to the behavior at 60◦ C, there was a gradual blueshift in peak position at longer incubation times. Changes in SYPRO Orange fluorescence have recently been used to study thermal stability of monoclonal antibodies and adjuvanted pneumococcal proteins,28 as well as used for excipient/stabilizer screening.28,29 This dye is nonfluorescent in hydrophilic environments, but exhibits a substantial increase in fluorescence when it is exposed to apolar regions of proteins.28,29 Therefore, SYPRO Orange fluorescence intensity can be used as an indicator of protein structural changes, such as those occurring under thermal stress. In the case of IPV3, the extrinsic fluorescence from SYPRO Orange was detected with an intensity maximum near 600 nm and a slight shoulder at approximately 650 nm. As the temperature increased from 20◦ C to 90◦ C, the SYPRO Orange fluorescence intensity gradually decreased with no obvious cooperative transition, similar to the intrinsic fluorescence intensity trend. Furthermore, no significant shift was found for the wavelength maximum of SYPRO Orange fluorescence (see Suppl. Material, Suppl. Fig. S3). Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
Circular dichroism spectroscopic studies conducted as a function of temperature produced distinctive results (Fig. 3). Upon heating from 20◦ C to 90 ◦ C, the CD signal at approximately 205 nm for IPV3 decreased from −9.0 to −6.5 millidegrees, the wavelength minimum shifted from approximately 205 to 215 nm and there was a broadening of the spectra. For the RNA peak (positive ellipticity at ∼265nm), the signal decreased monotonically with increasing temperature, reaching a plateau close to 80◦ C. The apparent thermal transition temperatures for the protein and RNA portions of the spectra were 58.1 ± 1.4◦ C and 49.0 ± 2.0◦ C, respectively (Figs. 3c and 3d). This result suggests that the capsid protein is more thermally stable than the RNA component. When the IPV3 sample was cooled back to room temperature after thermal stress, the CD signal was regained to some extent; approximately 70% of the original RNA signal and approximately 50% of the protein signal were recovered within the region 200–220 nm (Fig. 3b). Simultaneously, the minimum wavelength shifted back to approximately 209 nm compared with the initial value of 205 nm. Thus, the loss of secondary structure of the capsid proteins at high temperature appeared to be at least partially reversible. As temperature was increased, the SLS intensity of IPV3 remained fairly constant until 60◦ C and then sharply increased to its maximum value at approximately 70◦ C (Fig. 4 a). The intensity decrease above 70◦ C was probably because of the precipitation of large aggregates. The large error bars at elevated temperature also reflect this phenomenon. DOI 10.1002/jps.23801
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Figure 3. Representative CD spectra of IPV3 upon thermal stress. (a) CD spectra of IPV3 at different temperatures 20◦ C–90◦ C at 10◦ C intervals. (b) CD spectra of IPV3 at 90◦ C, and at 20◦ C before and after the thermal treatment. (c) Wavelength shift of the minimum intensity between 200 and 230nm upon thermal stress. (d) The normalized intensity at 260nm upon thermal stress. Three replicates were performed and only representative results are shown for clarity. The Tm values for c and d were fitted as specified in Material and Methods.
Figure 4. The light scattering of the IPV3 at pH 7 upon thermal stress. (a) SLS. (b) DLS. The diameter of the two peaks in 90◦ C in panel b was measured at 25◦ C after thermal ramping to 90◦ C. Triplicates were performed to the above-mentioned experiments, and the mean values were plotted with the standard deviation as the error bar in panel a.
Dynamic light scattering results during heating of IPV3 (Fig. 4b) followed a similar trend to that observed with SLS. Below 70◦ C, a single peak gradually shifted from a position for approximately 30 nm to that of 70 nm. At 75◦ C, two peaks were detected, with one centered at 50 nm and the other at 400 nm, the latter clearly indicating the presence of aggregated virion particles. At even higher temperature (80◦ C to 90◦ C), very large aggregates appeared, resulting in unreliable data because of the high degree of polydispersity and the inability of DLS to accurately determine the size above 1 :m. After cooling samples to room temperature, two populations of approximately 80 nm and greater than 1 :m were seen with a volume ratio DOI 10.1002/jps.23801
close to 1:1 (Fig. 4b, 90◦ C panel). The formation of aggregates at high temperature was also confirmed by TEM, where disrupted virions and aggregated material with dimensions larger than a few hundred nanometers were observed (see Suppl. Material, Suppl. Fig. S4). pH and Thermal Stability—EPDs To explore the combined effects of pH and temperature, the methods described above were used to examine IPV3 at pH 3–8 at one unit intervals as the temperature was increased from 10◦ C to 90◦ C. Representative data are shown in Fig. 5. It is clear that the response of the virus to temperature was Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
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Figure 5. The representative experimental data for EPDs. (a) SLS. (b) Intrinsic fluorescence peak position calculated with the center of spectra mass method. (c) Normalized maximum fluorescence intensity. (d) CD intensity transition data at 220nm, with pH 3 (closed square), 4 (open circle), 5 (closed triangel), 6 (open triangel), 7(closed diamond), and 8 (open star).
highly pH dependent. Transitions (Tm values) observed in all methods generally follow the pattern pH 3 < 4 < 5 < 6 < 8 < 7, with clear evidence of multiple transitions, especially at low pH. To provide a more convenient and comprehensible view of this complex data, they were summarized in the form of several EPDs, Fig.6. In this method, the data are normalized and presented in the form of changes in color, facial features, or distorted polygons. Detailed descriptions of these visualization methods are presented elsewhere.19 Using a clustering technique to define apparent phase boundaries, all methods of presentation resolved five distinct phases. Using the three-index EPD, the nature of each phase can be at least partially assigned. This can be supported by direct analysis of the data in Fig. 5 that was used to construct the EPDs, as shown in Fig. 6. In summary, the native state of the viral particle appeared to be present at pH 6–7 below 45◦ C. Alteration in the tertiary structure of the viral proteins was evident at pH 5 and 8 with further changes at lower pH (3–4) with reduced thermal stability (25◦ C–32◦ C). Major changes in protein secondary structure (CD spectra) were most evident at pH 5–7 above 50◦ C. Aggregation appeared above 60◦ C at both high (8) and low (3–5) pH and appeared at lower temperature (95%), there is almost no detectable signal increase even up to 90◦ C, indicating little or no formation of large aggregates. Trehalose (25%) was almost as effective as D2 O at inhibiting aggregation. Sucrose and sorbitol produced similar results to those for trehalose. IPV3 samples in mannitol and glycerol have similar onset temperatures of aggregation to that of the control. IPV3 samples in urea (5%) aggregated only at much higher temperatures than those noted for samples with other excipients, indicating that urea might solubilize the virions to some extent DOI 10.1002/jps.23801
and thereby reduce aggregation even at temperatures above the Tm . Samples in sodium citrate generated significant noise so that a clear transition in SLS signal during heating could not be discerned. Because D2 O has previously been reported to be highly effective at increasing polio virus thermal stability,17,18 IPV3 samples in D2 O were further studied by DLS and CD spectroscopies. With DLS, there was almost no size change observed even up to 90◦ C (Fig. 7a). A single peak with a diameter of approximately 30 nm dominated the DLS results without appearance of a significant amount of larger species. By comparison, at the same solution temperatures, aggregates of approximately 1 :m with at a volume percentage of approximately 50% appeared for IPV3 samples in PBS buffer alone. The thermal transition temperature observed with CD spectroscopy for IPV3 in D2 O (open circle) was approximately 5◦ C higher than samples in PBS buffer (closed square), which is quite close to that observed by fluorescence (Fig. 7b). Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
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The use of the extrinsic fluorescence of SYPRO Orange was unsuccessful in terms of distinguishing excipient effects either by isothermal incubation or thermal ramping stress (see Suppl. Material). For all samples examined for 1 h incubation at 50◦ C (Suppl. Fig. S6), the fluorescence intensity dropped to approximately 50% of the original signal at 20◦ C without peak shape change. During thermal ramping (Suppl. Fig. S3D), the SYPRO Orange intensity for IPV3 samples in the absence and presence of the tested excipients were fairly close to each other. After normalization, all intensity trend lines were overlapping and could not be distinguished from one another. The effect of pH on IPV3 ELISA antigen content was also examined immediately after dialysis against the phosphatecitrate buffer at pH values from 3.0 to 8.0. As shown in Fig. 9a, IPV3 only maintained approximately 20% and 60% of the ELISA antigen content at pH 3 and 4, respectively. In contrast, the ELISA antigen contents were not significantly different from each other at pH 5, 6, 7, and 8, with more than 80% of the content remaining. The losses of ELISA antigen content at low pH are readily explained by the results shown in Fig. 5. At acidic pH, IPV3 samples formed large virion aggregates and lost both tertiary and secondary structure compared with results at higher pH. The fluorescence Tm of IPV 3 was also obtained in an attempt to correlate it with the ELISA antigen content (Fig. 9b). At pH 3, no clear transition could be seen because of the extensive unfolding of the capsid proteins. IPV3 manifested lower Tm values at pH 4 (∼44◦ C) and pH 5 (∼50◦ C) than at pH 6–8 (∼56◦ C–57 ◦ C). Thus, the thermal stability of IPV3 under different pH condition also correlated well with the remaining ELISA antigen content. All of the biophysical characterizations of IPV3 in the above pH conditions were included in the EPD (Fig. 6). For clarity and direct comparison, fluorescence, CD, SLS, and DLS data of IPV3 at different pH values at 25◦ C are summarized in Suppl. Fig. S7 and could be referred from Fig. 5.
DISCUSSION
Figure 7. The effects of selected excipients on IPV3 thermal stability and potency. (a) Remaining titer of IPV3 samples incubated at 50◦ C for 1h (open bar). The remaining percentage of titer was calculated based on the fresh dialyzed and 1h incubated 50◦ C PBS samples as 100% and 0%, respectively. Tm was determined by subtracting the Tm of IPV3 samples in PBS from that of IPV3 in the presence of the excipients. All the Tm values were fitted with a two-state model of the intensity ratio of 350–330nm of Trp intrinsic fluorescence. (b) The correlation between the Tm and remaining relative titer. (c) SLS of IPV3 with selected excipients. Triplicates were performed to the above experiments and the mean values were plotted with the standard deviation as the error bar for panel a and b. Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
We have characterized IPV3 in terms of size, morphology, tertiary, and secondary structure and ELISA antigen content, and defined the most stable conditions as pH 7 at less than 40◦ C. Because of the complex structure of the virion and low resolution of the techniques used here, we are not able to provide detailed structural information. Rather, to facilitate formulation development, our goal is to identify physical methods that can measure IPV thermal stability and provide a good correlation between stability and ELISA antigen content. Intrinsic fluorescence was identified as the most sensitive technique among those examined, with fluorescence peak position the most useful property for the desired studies. To achieve high throughput, the fluorescence/light scattering-based Avacta system was selected for excipient formulation studies. This work is not intended to discern the protective mechanisms of effective excipients, but aspects of potential mechanisms can be considered. Saccharides and polyols probably exert their protective effects through the preferential exclusion mechanism. This mechanism has been thoroughly described in the literature.30,31 Briefly, because of the preferential exclusion of the stabilizer from the surface of protein molecules, the free energy of the protein is increased. The magnitude of the free energy increase varies directly with the protein surface area DOI 10.1002/jps.23801
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Figure 8. The effect of high-concentration deuterium on the size (a) and CD (b) of IPV3 upon thermal ramping. (a) DLS of IPV3 in D2 O (>95%) at different temperatures. (b) Wavelength of minimum intensity of IPV3 in D2 O (>95%) (open circle) upon thermal ramping, as compared with IPV3 in PBS (closed square).
Figure 9. (a) The relative titer remaining of IPV3 samples dialyzed at varied pH. (b) The fitted fluorescence peak position transition temperature of IPV3 samples at different pH. The star notified that no clear transition temperature could be fitted for IPV3 samples at pH 3. Triplicates were performed in the above experiments and the mean values were plotted with the standard deviation as the error bars.
exposed to solvent, and thus is much larger for the unfolded than for the native state of the protein. As a result, in the presence of a preferentially excluded excipient, the native state is stabilized and the free energy of unfolding is increased.31,32 Stabilization through the preferential exclusion mechanism requires relatively high concentrations of solutes. This requirement is consistent with our observation of protective effects of trehalose, sucrose, and sorbitol at higher concentration (25%, w/w) but almost no increase in IPV3 stability with relatively low concentration (5%). Glycine, mannitol, and glycerol might also exert their effects on IPV3 through preferential exclusion,30,33,34 but are typically less effective than disaccharides. Mannitol and glycerol did not inhibit aggregation, whereas glycine actually promoted self-association at earlier times (see Suppl. Fig. S5). Sodium citrate greatly enhanced IPV3 thermal stability as detected by fluorescence and ELISA antigen content. This may also be attributed to the preferential exclusion mechanism but binding mechanisms are also possible. As expected, urea decreased the Tm of IPV3 but it also delayed the onset of aggregation. Urea is well known to unfold proteins as well as increase protein solubility.35–37 The precise mechanism of this activity is still under investigation, with either preferential binding to the unfolded state or altering of its solvent environment as commonly suggested explanations.36
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The stabilizing effects of D2 O on proteins and vaccines have been widely reported.38–41 This phenomenon has generally been attributed to enhanced hydrophobic interactions within or between proteins and/or the presence of stronger hydrogen bonding in D2 O.39,40 In the case of polio vaccine, the protective effect of D2 O was discovered more than a decade ago.18,42,43 In a previous study, Trp fluorescence polarization indicated that poliovirus existed in a more rigid environment in D2 O than H2 O.14 In addition, the amount of water penetrating into the polio virus virion was substantially reduced by D2 O with such an effect attributed to a reduction in the swelling of the virus in response to temperature increases.14 In the same study, however, the observed CD signals were low and noisy because of low virus concentration, and the effects of the D2 O were not clearly demonstrated.14 In the current study of IPV3, we further provide clear evidence of stabilization by D2 O. The Tm of IPV3 from fluorescence and CD measurements increased by approximately 5◦ C, with no significant size changes seen even at very high temperature (∼90◦ C) based on both SLS and DLS studies. This suggests that IPV3 did not swell or aggregate appreciably during heating, perhaps because of an increase in virion rigidity in D2 O. We speculate that CD spectra were better resolved than in earlier studies because of the high purity and concentration of the IPV3 sample used in this study.
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The capsid proteins of IPV3 responded to pH and thermal stress somewhat differently. For instance, both an acidic environment and 50◦ C isothermal incubation significantly reduced the ELISA antigen content. The size distribution and biophysical properties under the two treatments were completely different. At pH 3, with DLS analysis, a peak over 1000 nm in diameter appeared, along with low Trp fluorescence intensify and diminished CD signal (Suppl. Fig. S7). These changes account for the loss of antigen content. In contrast, with isothermal incubation at 50◦ C, we did not see significant changes in IPV3 size and fluorescence peak position (Fig. 2 and Suppl. Fig. S2). This observation may be because of the fact that all of the physical data are of lower resolution and reflects averaged signal over many viral components. Thus, the antigen binding site may have gone through local environment changes that cannot be detected with our methods, such as fluorescent spectroscopy, because of the large number of Trp residues. Nevertheless, we were able to use fluorescence-based Tm values to compare the IPV3 thermal stability under various solution conditions. The relative IPV3 stabilities measured with fluorescence spectroscopy matched well with those obtained with ELISA antigen content. Thus, biophysical assessment of thermal stability with fluorescence spectroscopy potentially provides a valuable tool for formulation development of IPV3 and perhaps other virus-based vaccines as well.
ACKNOWLEDGMENTS The authors thank Dr. Sangeeta Joshi for her kind help on the pH and thermal stability screening study, Dr. Hardeep Samra in MedImmune for his advice on SYPRO Orange dye application, and Dorothy Dill for her patient help on TEM. We are grateful to the Pall Corporation (Port Washington, New York) for the loan of the Optim 1000 and accessories. We also thank Sanofi Pastuer for funding and the supplies of the IPV3 and antibodies for the ELISA.
REFERENCES 1. WHO. 2003. Global Polio eradication initiative strategic plan 2004–2008. 1–40 http://www.polioeradication.org/content/publications/ 2004stratplan.pdf last accessed Nov. 21, 2013. 2. WHO. 2011. Pliomyelities fact sheet http://www.poliofree.org/ polio-fact-sheet.htm last accessed Nov. 21, 2013 . 3. Dowdle WR, De Gourville E, Kew OM, Pallansch MA, Wood DJ. 2003. Polio eradication: The OPV paradox. Rev Med Virol 13(5):277– 291. 4. Friedrich F. 1997. Rare adverse events associated with oral poliovirus vaccine in Brazil. Braz J Med Biol Res 30(6):695–703. 5. Kew OM, Sutter RW, de Gourville EM, Dowdle WR, Pallansch MA. 2005. Vaccine-derived polioviruses and the endgame strategy for global polio eradication. Annu Rev Microbiol 59:587–635. 6. Minor P. 2009. Vaccine-derived poliovirus (VDPV): Impact on poliomyelitis eradication. Vaccine 27(20):2649–2652. 7. Salas-Peraza D, Avila-Aguero ML, Morice-Trejos A. 2010. Switching from OPV to IPV: Are we behind the schedule in Latin America? Expert Rev Vaccines 9(5):475–483. 8. Tajikis WHOCO, Europe WRO, Contro ECDP. 2010. Outbreak of poliomyelitis in Tajikistan in 2010: Risk for importation and impact on polio surveillance in Europe? Eurosurveillance 15(17):5–7. 9. Atkinson W, Hamborsky J, McIntyre L, Wolfe S. 2008. Epidemiology and prevention of vaccine-preventable diseases. 10th ed. Department of Health and Human Services, Washington D.C. Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
10. Chen DX, Kristensen D. 2009. Opportunities and challenges of developing thermostable vaccines. Expert Rev Vaccines 8(5):547–557. 11. Hogle JM, Chow M, Filman DJ. 1985. Three-dimensional structure of poliovirus at 2.9A resolution. Science 229(4720):1358–1365. 12. Bubeck D, Filman DJ, Cheng N, Steven AC, Hogle JM, Belnap DM. 2005. The structure of the poliovirus 135S cell entry intermediate at 10angstrom resolution reveals the location of an externalized polypeptide that binds to membranes. J Virol 79(12):7745–7755. 13. Brandenburg B, Lee LY, Lakadamyali M, Rust MJ, Zhuang X, Hogle JM. 2007. Imaging poliovirus entry in live cells. PLoS Biol 5(7):e183. 14. Chen CH, Wu R, Roth LG, Guillot S, Crainic R. 1997. Elucidating mechanisms of thermostabilization of poliovirus by D2 O and MgCl2 . Arch Biochem Biophys 342(1):108–116. 15. Grimmel M, Zibirre R, Koch G. 1983. Fluorescence spectrophotometric study of structural alterations in the capsid of poliovirus. Arch Virol 78(3–4):191–201. 16. Westdijk J, Brugmans D, Martin J, van’t Oever A, Bakker WAM, Levels L, Kersten G. 2011. Characterization and standardization of Sabin based inactivated polio vaccine: Proposal for a new antigen unit for inactivated polio vaccines. Vaccine 29(18):3390–3397. 17. Milstien JB, Lemon SM, Wright PF. 1997. Development of a more thermostable poliovirus vaccine. J Infect Dis 175:S247–S253. 18. Verheyden B, Andries K, Rombaut B. 2001. Capsid and RNA stabilisation of the oral polio vaccine. Vaccine 19(15–16):1899–1905. 19. Kim JH, Iyer V, Joshi SB, Volkin DB, Middaugh CR. 2012. Improved data visualization techniques for analyzing macromolecule structural changes. Protein Sci 21(10):1540–1553. 20. Kueltzo LA, Ersoy B, Ralston JP, Middaugh CR. 2003. Derivative absorbance spectroscopy and protein phase diagrams as tools for comprehensive protein characterization: A bGCSF case study. J Pharm Sci 92(9):1805–1820. 21. Maddux NR, Joshi SB, Volkin DB, Ralston JP, Middaugh CR. 2011. Multidimensional methods for the formulation of biopharmaceuticals and vaccines. J Pharm Sci 100(10):4171–4197. 22. Rezapkin G, Martin J, Chumakov K. 2005. Analysis of antigenic profiles of inactivated poliovirus vaccine and vaccine-derived polioviruses by block-ELISA method. Biologicals 33(1):29–39. 23. Ferguson M, Wood DJ, Minor PD. 1993. Antigenic structure of poliovirus in inactivated vaccines. J Gen Virol 74(4):685–690. 24. Boublik M, Drzeniek R. 1977. Structural subunits of poliovirus particles by electron microscopy. J Gen Virol 37(1):127– 134. 25. Eftink MR. 1991. Fluorescence techniques for studying protein structure. Methods Biochem Anal 35:127–205. 26. Volkin DB, Burke CJ, Marfia KE, Oswald CB, Wolanski B, Middaugh CR. 1997. Size and conformational stability of the hepatitis A virus used to prepare VAQTA, a highly purified inactivated vaccine. J Pharm Sci 86(6):666–673. 27. Filman DJ, Syed R, Chow M, Macadam AJ, Minor PD, Hogle JM. 1989. Structural factors that control conformational transitions and serotype specificity in type-3 poliovirus. Embo J 8(5):1567–1579. 28. Goldberg DS, Bishop SM, Shah AU, Sathish HA. 2011. Formulation development of therapeutic monoclonal antibodies using highthroughput fluorescence and static light scattering techniques: Role of conformational and colloidal stability. J Pharm Sci 100(4):1306– 1315. 29. Ausar SF, Chan JD, Hoque W, James O, Jayasundara K, Harper K. 2011. Application of extrinsic fluorescence spectroscopy for the high throughput formulation screening of aluminum-adjuvanted vaccines. J Pharm Sci 100(2):431–440. 30. Arakawa T, Timasheff SN. 1982. Mechanism of stabilization of proteins by glycerol and sucrose. Seikagaku 54(11):1255–1259. 31. Lee JC, Timasheff SN. 1981. The stabilization of proteins by sucrose. J Biol Chem 256(14):7193–7201. 32. Foster TM, Dormish JJ, Narahari U, Meyer JD, Vrkljan M, Henkin J, Porter WR, Staack H, Carpenter JF, Manning MC. 1996. Thermal stability of low molecular weight urokinase during heat treatment. 3. Effect of salts, sugars and Tween 80. Int J Pharm 134(1–2):193–201. DOI 10.1002/jps.23801
RESEARCH ARTICLE – Pharmaceutical Biotechnology
33. Haque I, Singh R, Moosavi-Movahedi AA, Ahmad F. 2005. Effect of polyol osmolytes on DeltaG(D), the Gibbs energy of stabilisation of proteins at different pH values. Biophys Chem 117(1):1– 12. 34. Santoro MM, Liu YF, Khan SMA, Hou LX, Bolen DW. 1992. Increased thermal-stability of proteins in the presence of naturallyoccurring osmolytes. Biochemistry 31(23):5278–5283. 35. Timasheff SN. 2002. Protein-solvent preferential interactions, protein hydration, and the modulation of biochemical reactions by solvent components. Proc Natl Acad Sci USA 99(15):9721–9726. 36. Bennion BJ, Daggett V. 2003. The molecular basis for the chemical denaturation of proteins by urea. Proc Natl Acad Sci USA 100(9):5142– 5147. 37. Chen J, Liu Y, Li X, Wang Y, Ding H, Ma G, Su Z. 2009. Cooperative effects of urea and L-arginine on protein refolding. Protein Expr Purif 66(1):82–90.
DOI 10.1002/jps.23801
151
38. Sen A, Balamurugan V, Rajak KK, Chakravarti S, Bhanuprakash V, Singh RK. 2009. Role of heavy water in biological sciences with an emphasis on thermostabilization of vaccines. Expert Rev Vaccines 8(11):1587–1602. 39. Cioni P, Strambini GB. 2002. Effect of heavy water on protein flexibility. Biophys J 82(6):3246–3253. 40. Efimova YM, Haemers S, Wierczinski B, Norde W, van Well AA. 2007. Stability of globular proteins in H2 O and D2 O. Biopolymers 85(3):264–273. 41. Verheul M, Roefs SP, de Kruif KG. 1998. Aggregation of betalactoglobulin and influence of D2 O. FEBS Lett 421(3):273–276. 42. Milstien JB, Lemon SM, Wright PF. 1997. Development of a more thermostable poliovirus vaccine. J Infect Dis 175 Suppl 1:S247–S253. 43. Wu R, Georgescu MM, Delpeyroux F, Guillot S, Balanant J, Simpson K, Crainic R. 1995. Thermostabilization of live virus vaccines by heavy water (D2 O). Vaccine 13(12):1058–1063.
Qi et al., JOURNAL OF PHARMACEUTICAL SCIENCES 103:140–151, 2014
