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Stephen Ball is product marketing manager, nanoparticle and molecular characterization, Malvern Instruments Ltd, Enigma Business Park, Grovewood Road, Malvern, WR14 1XZ UK, tel 44 (0) 1684 892456; firstname.lastname@example.org
The author describes a method to avoid protein aggregation when using light scattering systems.
As the number of biopharmaceutical formulations that use proteins as drug molecules continues to increase, the spotlight is being directed on the analytical methods used in their development, formulation, and quality control. Protein mobility is one property that has been identified as a promising indicator of formulation stability, viscosity, and behavior. This article describes an approach to using electrophoretic light scattering to make protein mobility measurements.
Light scattering techniques are widely used in protein characterization. Dynamic light scattering (DLS) is established in the measurement of particle and molecular size, and in studying the interactions between proteins. Electrophoretic light scattering (ELS) is used to measure the electrophoretic mobility of particles in dispersion or molecules (such as proteins) in solution. This mobility is often converted into zeta potential to enable comparison of materials under different conditions. In the case of proteins, the measurement of protein mobility allows the calculation of protein charge, which in turn relates to factors such as activity and reaction kinetics. Recent advances in instrumentation and methodologies are addressing the technical challenges of using light scattering to make mobility measurements on proteins.
The fundamental physical principle in ELS is that of electrophoresis. A sample is introduced into a cell containing two electrodes. An electrical field is applied and particles or molecules that have a net charge, or more strictly a net zeta potential, will migrate towards the oppositely charged electrode with a velocity, known as the electrophoretic mobility, that is related to their zeta potential. This velocity is measured using the laser Doppler technique, where the frequency shift or phase shift of an incident laser beam, caused by the moving particles, is measured as the particle mobility.
Martin McCarthy/Getty Images; Dan Ward
Experimentally, protein mobility measurements present two practical challenges. First, working with protein solutions often means working with dilute concentrations, low DLS count rates, and low electrophoretic mobilities. Second, the act of applying an electric field to the sample can damage the protein by stimulating aggregation, with resultant mobility measurements reflecting the aggregate molecules rather than the native protein.
A new approach combines a high sensitivity light scattering system (Zetasizer Nano ZSP, Malvern Instruments) with a diffusion barrier technique that separates the molecules in the sample from the electrodes, to avoid the risk of aggregation. A measurement protocol regulates voltage and temperature; and automatic size measurements before and after the electrophoretic mobility measurement verify that no aggregation has occurred.
Work by Corbett et al. (1) has shown that much of the aggregation that occurs during electrophoretic measurements of proteins in solution takes place at the electrodes. Standard guide ASTM E2865 (2), published in 2012, deals with the measurement of mobility and zeta potential in systems containing biological material such as proteins, DNA, liposomes, and other similar organic materials that possess particle sizes in the nanometer scale, below 100 nm. It indicates that these molecules can be quite labile and may absorb and decompose under an electrical field, and goes on to say that it is virtually impossible to eliminate such decomposition for some molecules unless specific routes are taken, such as isolating the electrodes from the biological molecules.
The diffusion barrier technique described here separates the sample molecules from the electrodes, minimizing the effect of the measurement technique itself on the sample. As a result, the voltage can be applied for a longer period to generate more data from the measurement. A folded capillary cell allows the introduction of approximately 20 μL sample, that is then separated from the electrodes by the same buffer in which the protein is dissolved (Figure 1). By creating this physical separation, the sample is protected for as long as it takes to diffuse to the electrodes, which may be many hours.
Figure 1: Folded capillary cells. Left-filled with blue dextran in 10 mM NaCl, and right-filled with 10 mM NaCl and loaded with 50 Î¼L blue dextran for measurement using the diffusion barrier technique. (ALL FIGURES ARE COURTESY OF MALVERN INSTRUMENTS)
In experiments conducted by application scientists at Malvern Instruments' headquarters in the UK, human serum albumin (HSA) was prepared in different buffers to a final concentration of approximately 2 mg/mL. The buffers were the original formulation buffer at pH 7 and a citrate buffer at pH 4.3. Samples were filtered to ensure there were no aggregates present at the start of the measurement. The diffusion barrier technique was performed by filling a disposable capillary cell with the appropriate buffer and then loading 20 μL of the diluted HSA into the optical area at the bottom of the cell with a gel-loading pipette tip. Size and mobility measurements were made in the Zetasizer Nano ZSP. The hydrodynamic size and mobility of HSA were measured using DLS and ELS, respectively. Both were measured at the same scattering angle to use the size results to confirm that mobility results corresponded with the protein and not with any aggregated material.
Figures 2A and 2D show the size distributions at the beginning of the experiment for samples at pH 7 and pH 4.3, respectively, and exhibit the single peaks expected from the filtered samples. Figures 2B and 2E are frequency plots from mobility measurements.
Figure 2: (A) Overlaid size measurement showing human serum albumin (HAS) at pH 7 before the mobility measurement. (B) Overlaid frequency plots during mobility measurement at pH 7. (C) Overlaid size measurements showing HSA at pH 7 after the mobility measurements. (D) Overlaid size measurement showing HSA at pH 4.3 before the mobility measurement. (E) Overlaid frequency plots during mobility measurement at pH 4.3. (F) Overlaid size measurements showing HSA at pH 4.3 after the mobility measurements.
These are useful in assessing measurement quality as they show the distribution of frequencies in the scattered light (3). Proteins have a significant Brownian component in their overall mobility, so the peak in the frequency plot is expected to be broad. Aggregates in the sample will have lower Brownian motion, resulting in a sharp peak in the frequency plot. The presence of a broad peak in the frequency plot is, therefore, indicative of high quality data whereas a frequency plot containing a sharp peak is highly likely to contain information derived from aggregates. In the results shown here, the broad shallow peaks from both samples indicate that the measurement is primarily of the protein and not aggregates.
The overall charge from the hydrodynamic size and electrophoretic mobility was calculated by the instrument software. The calculated charge for HSA at pH 7 was approximately -18 and at pH 4.3 approximately +10.
Figures 2C and 2F show the size distributions of the samples after the mobility measurements had been made and indicate that only a very small amount of aggregation occurred. In both cases, the primary peak was the dominant peak, accounting for 99% of the scattered light at pH 7 and 90% at pH 4.3. The results from these mobility measurements, therefore, have high confidence. The pH 4.3 sample was shown to be aggregating over time in the absence of an electric field; this is most likely because of its low pH and also accounts for the small amount of aggregation occurring in this sample during the measurement.
In conclusion, the mobility of HSA was measured successfully in different buffers at a concentration of 2 mg/mL and using a 20 μL sample.
Stephen Ball is product marketing manager, nanoparticle and molecular characterization, Malvern Instruments Ltd, Enigma Business Park, Grovewood Road, Malvern, WR14 1XZ UK, tel 44 (0) 1684 892456; email@example.com.
1. J. Corbett, M. Connah, and K. Mattison, Electrophoresis, 32 (14), 1787-94 (2011).
2. ASTM, ASTM E2865–Standard Guide for Measurement of Electrophoretic Mobility and Zeta Potential of Nanosized Biological Materials (West Conshohocken, PA, 2012).
3. Malvern application note: The Diffusion Barrier Technique, Practical Aspects and Data interpretation, www.malvern.com/diffusion-barrier (accessed April 10, 2013).