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Review

Toward greener analytical techniques for the absolute quantification of peptides in pharmaceutical and biological samples Ann Van Eeckhaut a,∗ , Debby Mangelings b a Department of Pharmaceutical Chemistry and Drug Analysis, Center for Neurosciences (C4N), Vrije Universiteit Brussel, Laarbeeklaan 103, 1090 Brussels, Belgium b Department of Analytical Chemistry and Pharmaceutical Technology, Vrije Universiteit Brussel, Laarbeeklaan 103, 1090 Brussels, Belgium

a r t i c l e

i n f o

Article history: Received 11 February 2015 Received in revised form 19 March 2015 Accepted 23 March 2015 Available online xxx Keywords: Peptide Green Liquid chromatography Capillary electrophoresis Supercritical fluid chromatography

a b s t r a c t Peptide-based biopharmaceuticals represent one of the fastest growing classes of new drug molecules. New reaction types included in the synthesis strategies to reduce the rapid metabolism of peptides, along with the availability of new formulation and delivery technologies, resulted in an increased marketing of peptide drug products. In this regard, the development of analytical methods for quantification of peptides in pharmaceutical and biological samples is of utmost importance. From the sample preparation step to their analysis by means of chromatographic or electrophoretic methods, many difficulties should be tackled to analyze them. Recent developments in analytical techniques emphasize more and more on the use of green analytical techniques. This review will discuss the progresses in and challenges observed during green analytical method development for the quantification of peptides in pharmaceutical and biological samples. © 2015 Elsevier B.V. All rights reserved.

Contents 1. 2.

3.

4.

5. 6.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Challenges related to physicochemical properties of peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Purity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Solubility–nonspecific binding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Toward greener liquid chromatographic methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Miniaturization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Decreasing analysis time . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Solvent replacement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. Selected applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alternatives to liquid chromatography for greener analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Capillary electrophoretic methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Supercritical fluid chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Challenges related to mass spectrometric quantification of peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

00 00 00 00 00 00 00 00 00 00 00 00 00 00 00 00 00

Abbreviations: CE, capillary electrophoresis; HRMS, high resolution mass spectrometry; id, internal diameter; LC, liquid chromatography; MS, mass spectrometry; MS/MS, tandem mass spectrometry; RPLC, reversed phase liquid chromatography; SFC, supercritical fluid chromatography; UHPLC, ultra high performance liquid chromatography. ∗ Corresponding author at: Department of Pharmaceutical Chemistry and Drug Analysis, Center for Neurosciences (C4N), Vrije Universiteit Brussel, Laarbeeklaan 103, B-1090 Brussels, Belgium. Tel.: +32 2 477 47 46; fax: +32 2 477 41 13. E-mail address: [email protected] (A. Van Eeckhaut). http://dx.doi.org/10.1016/j.jpba.2015.03.023 0731-7085/© 2015 Elsevier B.V. All rights reserved.

Please cite this article in press as: A. Van Eeckhaut, D. Mangelings, Toward greener analytical techniques for the absolute quantification of peptides in pharmaceutical and biological samples, J. Pharm. Biomed. Anal. (2015), http://dx.doi.org/10.1016/j.jpba.2015.03.023

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1. Introduction Peptides and peptide-based drug products are attracting increasing attention as diagnostics and therapeutics [1,2], due to the enormous diversity of functions of peptides in the human body and their involvement in almost every essential physiological process [3]. Today, approximately 100 therapeutic peptides are on the market in the three major regions, Europe, USA and Japan [4]. In 2012, six peptides have obtained marketing approval in the USA and five in the EU [5]. Moreover, the number of peptides entering clinical trials every year has increased till around 20 [4]. Peptides have higher potency, selectivity and specificity compared to small molecules, and they also show less off-target toxicity and drug–drug interactions [1,6,7]. The major drawback of peptides is their general poor oral bioavailability due their low permeability, in combination with a low metabolic stability. Recent advances to overcome these issues include for example modifications of the amino acid backbone and chemical conjugation with polymers [1,4,6,8]. Peptides are therefore one of the most promising areas for the development of new drugs with original mechanisms of action [4,9]. In this review, we follow the definition of peptides as proposed earlier [10,11], i.e. containing maximal 50 amino acid residues and having a molecular weight below 6 kDa. The technique used for peptide synthesis depends largely on its size and chemical features [4,12]. Peptide manufacturing can be achieved entirely through chemical synthesis, recombinant DNA technology, cell-free expression systems, transgenic animals and plants or enzymatic synthesis [1,4]. Chemical synthesis is the most universal approach as it permits to include unnatural amino acids and pseudo-peptide bonds [1,4]. It remains the gold standard for the manufacturing of peptides between 5 and 50 amino acid residues [1,13,14]. Three major approaches are distinguished, namely solution-phase, solid phase and hybrid approaches [4,13]. Bioanalysis of peptides faces problems derived from the low concentration of these analytes in biological matrices, the large number of potential interferences and the limited sample volume [15]. Sample treatments usually involve isolation of the target analytes from the matrix in combination with their preconcentration, in this way improving sensitivity and selectivity of the assay [15,16] and reducing matrix effects [17,18]. Sample treatment has evolved in the last few years following three main trends, i.e. automation, miniaturization and simplification. This has led to the development of a number of microextraction techniques [15,19]. The characteristics of these microextraction techniques match with the requirements of bioanalysis. Indeed, automation results in improved throughput, allowing larger number of samples to be processed. Moreover, less sample volume is required due to the miniaturization. Finally, the reduction of number of steps in the sample preparation process, i.e. simplification, improves the precision of the method [15]. Sample pretreatment steps will not be further discussed, as they are outside the scope of this review. Recently, the interest in green analytical chemistry has grown tremendously. An increased number of manuscripts on green analytical method development can be found in literature [20]. This term is used to describe analytical approaches that minimize the consumption of reagents and energy, as well as the reduction in the generation of hazardous waste. For analytical method development the focus is oriented toward reduction or elimination of toxic solvents and decreasing the analysis time [21]. The increasing importance of peptide therapeutics necessitates performance improvements in (bio)analytical techniques to support biopharmaceutical drug development [3]. Indeed, analytical methods are primordial to assess drug substance and drug product quality and to obtain reliable pharmacological and toxicological data [3,11,22]. The development of sensitive, selective, high-throughput methods

for peptide analysis using greener analytical techniques is highly demanded [23]. Ligand-binding assays are still the current gold standard for peptide analysis, because they offer high sensitivity. Moreover, immunoassays are considered as green techniques as they provide rapid analysis, are mostly performed in aqueous solution, require simple sample pretreatment and low sample volumes [24,25]. However, the dynamic range is often limited and there can be large selectivity issues [3,26,27]. In addition, development of new antibodies can be time-consuming and costly [26]. Liquid chromatography (LC) coupled to mass spectrometry offers improved selectivity and reduced method development time, and can be used as alternative or as orthogonal assay [28]. LC remains today the method of choice for peptide analysis. However, both capillary electrophoresis (CE) and supercritical fluid chromatography (SFC) are complementary techniques which have also shown their usefulness for drug analysis [29,30]. This review will therefore focus on the progresses in and challenges observed during green analytical method development for the quantification of peptides in pharmaceutical and biological samples. 2. Challenges related to physicochemical properties of peptides Independent of the used separation technique, there are several challenges that should be taken into account when quantifying peptides. These include the purity of the peptide standard, solubility of the peptide and related adsorption issues and also stability of these biopharmaceutical compounds. These issues will be discussed in more detail below. 2.1. Purity One important and evident aspect necessary to perform quantitative analyses is the purity of the used peptide standard. Although the peptide manufacturers assure the peptide to be >95% pure, cases are reported where more than 50% of the peptide content comprised of impurities [12] or even a totally different peptide was present [31]. The following impurity classes can be distinguished: amino acid deletion or insertion, incomplete removal of protecting groups after synthesis, oxidation or reduction, diastereoisomerization, side- and end-chain impurities, dimers, peptide counter ions (usually trifluoroacetic acid), structurally unrelated contamination and miscellaneous impurities (for review, see [12]). The presence of any of these impurities has significant consequences on the biological function of the peptide and consequently on the outcome of the experiments. Thus, it is highly recommended to verify the quality of peptides before experimental use and to repeat the biological experiments with a second peptide batch, obtained from another source. 2.2. Solubility–nonspecific binding Aspecific adsorption is a known property of peptides and is detrimental for sensitivity and carryover in all separation methods. Reduction of nonspecific binding should be investigated at the beginning of every method development phase. The extent of this nonspecific binding depends upon the characteristics of the peptide, such as charge of the peptide and hydrophobicity and should therefore be optimized for every peptide studied. Aspecific adsorption can be minimized by choosing the right dissolution solvent, dilution solvent for further standard preparation and injection solvent, but also the vial material, the temperature during standard preparation and during analytical separation should be carefully selected. A widely applicable strategy to improve method sensitivity by intervening on all aspects of standard preparation, i.e. from

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Fig. 1. Multiple reaction monitoring total ion chromatogram of neurotensin (NT), neuromedin B (NMB) and neuromedin N (NMN) corresponding to a 100 pM standard solution which is injected before (red) and after (green) reduction of aspecific adsorption during standard preparation and optimization of the injection solvent. The red chromatogram corresponds to the protocol where all dilutions are performed with water, while the green one corresponds to the most optimal standard preparation conditions as optimized in [32]. The least adsorption was observed when dissolving the lyophilized peptide in water/acetonitrile/formic acid (45/50/5, V/V/V), followed by performing a dilution series with water/formic acid (95/5, V/V) in a regular Eppendorf to obtain a 10 nM standard solution. The obtained standard solution was spiked afterwards into a polypropylene vial containing 13.1% (V/V) acetonitrile and 4.4% (V/V) formic acid in dialysate matrix. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.) Reprinted with permission from [32].

dissolution of the powder until the injection of the sample, was developed in our laboratory [32]. In first instance, it is important to choose the correct dissolution solvent for the lyophilized powder as proper solubilization is the key to a successful analysis. The solubility properties of a peptide depend on a combination of the following properties: the total number of amino acids; the ratio of apolar to the total number of amino acids and the overall net charge, including the total number of charges. Solubility guidelines were defined based on these properties to allow the reader to select the appropriate solvent [33]. However, the dissolution solvent should always be experimentally determined because also other parameters such as the conformation of the molecule and the distribution of the amino acids within the molecule can affect solubilization [32]. Furthermore, both the solvent for subsequent dilutions of the stock solution and the type of surface material of the vial used should be simultaneously optimized. Indeed, we have observed that the optimal solvent for dissolution was not suitable for peptide dilution. The latter also differed depending on the type of surface material [32]. Next to the type of surface, also the temperature and the pipetting protocol affect adsorption. Secondly, the sample composition in the injection vial was optimized by means of an experimental design. Also in this case, contradictory findings were observed between the injection and dilution solvent. In our study, the injection solvent composition had the largest impact on sensitivity. Indeed, optimization of the previous described steps was necessary to enable detection of a 100 pM standard solution of our peptides of interest, as shown in Fig. 1. The most important conclusion and something one should keep always in mind when working with peptides is that a tailored investigation of solvent compositions and used material is needed for every peptide under investigation due to their unique nature. 2.3. Stability Most peptides are formulated as lyophilized products to prolong stability during storage. In general, peptides are more stable in their lyophilized form than in solution. However, degradation can also occur in the solid state [34]. Moreover, excipients added to

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the finished drug product can also interact with the peptide causing degradation [14]. Proteases and other enzymes present in blood or other biological matrices can rapidly metabolize and inactivate peptides [10,35]. They are therefore significantly less stable than proteins in in vitro and in vivo experiments [35]. Moreover, peptides can undergo various types of chemical degradation including deamidation, hydrolysis, peptide bond cleavage, oxidation, Maillard reaction, ␤-elimination, enantiomerization, isomerization and dimerization [10,14]. Chemical degradation of peptides is residue specific and thus depending on the amino acid sequence. Asparagine and aspartic acid have been shown to be among the most unstable amino acid residues [36]. It is therefore important to ensure and verify peptide stability in dissolution solvents and in biological matrices at the beginning of method development. Moreover, freeze–thaw cycles and prolonged exposure to atmospheric oxygen should be avoided. 3. Toward greener liquid chromatographic methods 3.1. Introduction To assess quality attributes such as identification, related substances and assay in peptide drug substance, mostly LC–UV methods are described [11]. Due to its selectivity, robustness and ease of use, LC remains the technique of choice. Coupling to mass spectrometry (MS) further allows to identify possible impurities [14]. For quantification in biological samples, LC tandem mass spectrometry (MS/MS) is becoming the method of choice because it provides excellent selectivity for the analysis of compounds in complex biological matrices, good accuracy and precision and a wide dynamic range. However, peptide adsorption leading to carryover effects and the longer analysis times for complex samples, resulting in an overall low sample throughput, still remain to be ameliorated. Several excellent reviews describe the work flow for quantification of peptides and proteins in biological samples with LC–MS/MS [3,10,17,37–39]. Reversed-phase LC (RPLC) is an important tool for peptide separation. In contrast to the well-understood chromatographic behavior of small molecules, research is still needed to understand the behavior of peptides under different chromatographic conditions. Indeed, several retention prediction models have been developed to predict the retention time of a certain peptide based on its properties and/or the experimental conditions [40], mostly to improve peptide identification in proteomics research. The chromatographic behavior of peptides has most often been related to their amino acid composition [41]. However, also the sequence of amino acids in the peptide, peptide length, overall hydrophobicity, pI value, nearest neighbor effect of charged side chains and propensity to form helical structures should be included [41]. Next to the peptide characteristics, also the LC parameters (stationary phase, mobile phase – including ion-pairing reagents, gradient slope, pH and temperature) impact retention, separation selectivity and also the peptide peak shape [40–44]. Greener LC methods can be obtained by minimizing solvent consumption through (i) modification of relevant column-related parameters such as miniaturization of the column internal diameter (id), reduction of the particle size and/or reduction of the column length, (ii) operating at elevated temperature and (iii) switching to greener solvents and additives [24,25,45,46]. 3.2. Miniaturization Miniaturization of the LC system to micro (1 mm id columns) or nano (75–150 ␮m id columns) dimensions presents undeniable

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advantages such as lower peak dispersion and reduced flow rates, leading to an increased sensitivity and a decreased use of organic solvents [47–49]. Of course, the latter does not apply if high volume pumps are used in combination with a flow splitter. Downscaling of the column dimension of the chromatographic system from id1 to id2 enhances the mass sensitivity by a theoretical factor proportional to the ratio of the squares of the diameters (f = id1 2 /id2 2 ), due to a decreased dilution of the analyte zone. This gain in sensitivity is only valid if other column parameters remain constant and when equal amounts of analytes are injected on both systems [50–52]. For nano systems, the increased mass sensitivity, obtained by miniaturization, can be counterbalanced by a lack of increased concentration sensitivity due to limited injection volumes and loading capacity of the nano column [50]. The use of a precolumn in the columnswitching set-up allows the injection of larger volumes and it is accompanied by a sample clean-up phase to remove the contaminants. Otherwise, similar detection sensitivity compared to the conventional system is obtained using smaller injection volumes. This can be important in studies where limited sample volumes are available, for example in studies with small animals or children [53,54], or when multiple analyses of the same sample are desired. Additionally, at these low flow rates, smaller droplets are formed with higher surface-to-volume ratios, leading to a tremendous increase in efficiency of the electrospray ionization process and therefore increased mass spectrometric detection [54,55]. Next to the miniaturization of the column id, also the injector and the detector cell volume along with the volumes of all connections between injector and detector should be adjusted to minimize extra-column band broadening [56]. Due to the small volumes and flow rates, the effect of dead volumes are more pronounced in these miniaturized systems [53], putting a high constraint on the tubing connections. In recent years, reliable miniaturized systems are commercially available improving the robustness of the systems for routine analyses [53]. The major strategy to minimize void volumes and accompanied efficiency drop was to integrate all chromatographic components on a chip [47,56], including the electrospray emitter for MS detection [57]. Yin and colleagues were the first to describe an integrated nano LC-electrospray polymer microfluidic chip. An enrichment column, a RPLC column and an electrospray tip were laser-ablated into a polyimide film [57,58]. Integrating all components in a single device eliminates the need for fluidic connections, as found in conventional nano LC. As a result delay and dead volumes are reduced as well as the post-separation volume, leading to a reduced total analysis time and decreased band broadening [57,58]. This first LC microfluidic device was commercialized by Agilent and is routinely used in proteomics research, but also for pharmaceutical applications and in food analysis [54]. Waters also developed an integrated capillary-flow microfluidic LC system. Their ionKey/MSTM system integrates the UHPLC separation (150 ␮m id column) directly into the source of the mass spectrometer. Several application notes on the analysis of peptides in plasma samples are described on their website. 3.3. Decreasing analysis time Recent developments in chromatographic supports and instrumentation for LC, such as ultra high performance liquid chromatography (UHPLC) and superficially porous particles, are enabling rapid and highly efficient separations [59–61]. UHPLC is well established for small molecule analysis to increase throughput, while maintaining or increasing resolution [17]. As demonstrated by Van Deemter, the use of small particles is one of the best solutions to improve chromatographic performance. Sub-2 ␮m particles give higher efficiencies over a wider range of linear velocities due to the low mass transfer resistance of these supports [60].

Fig. 2. Influence of the temperature on the resolution between six angiotensin (Ang) peptides. As can be observed from the figure, the change in retention time is peptidedependent and therefore the temperature for optimal resolution differs depending on the peptide pair under investigation. This results in significant selectivity differences depending on the temperature used. Chromatographic conditions: Stationary ˚ mobile phase, phase, Alltima C18 stationary phase (1.0 mm × 150 mm, 3 ␮m, 100 A); A: water/FA 100:0.1 (V/V) and B: water/ACN/FA 20:80:0.1 (V/V/V); LC gradient was 17–25% B in 1 min, followed by 25–30% B in 9 min, 30–17% B in 0.1 min and 17% B for 5 min; flow, 50 ␮L/min; injection volume, 5 ␮L; UV detection, 220 nm.

The sample analysis time can therefore be decreased by reducing the particle size and column length. Indeed, the same plate numbers can be obtained with a shorter column when using sub-2 ␮m particles, while higher flow rates can be applied without loss in efficiency [56,60,62]. In order to allow the use of longer columns, the mobile phase temperature can be increased to 60–90 ◦ C. At these elevated temperatures, the mobile phase viscosity and consequently the column-pressure drop are reduced [63]. Eugster et al. have demonstrated that an increased temperature of the mobile phase (90 ◦ C) enhanced chromatographic performance, especially for peptides due to the higher improvement of their diffusion coefficient compared to small molecules [64]. However, thermal stability of the peptides should be carefully investigated, the stationary phase should be able to resist these elevated temperatures and one should always keep in mind that for peptides significant changes in selectivity can occur due to the increased temperature [62,64,65]. Fig. 2 illustrates the influence of the temperature on the resolution between the six angiotensin peptides investigated in our laboratory. The change in retention time was peptide-dependent and therefore the temperature for optimal resolution differed depending on the peptide pair under investigation. In addition to these sub-2 ␮m particle size columns, also columns packed with superficially porous particles can be used to obtain efficient and fast separations. In comparison with fully porous particles, analytes spend less time diffusing in and out of the thin shell that surrounds the solid core. This shorter diffusion path allows a faster mass transfer [66–68]. They therefore provide the advantage of sub-2 ␮m particles, but without the high backpressure [46]. Since a few years also sub-2 ␮m superficially porous columns are available on the market. The combination of a small particle size and the superficially porous technology results in a significant increase of column efficiency and chromatographic resolution [69,70]. A dedicated instrument is however mandatory to fully benefit from these innovative stationary phases. It should include a reasonably low dwell volume, reduced extra-column variance, and short injection cycle times, in combination with a sufficiently high upper pressure limit and data acquisition rate [59,63,71]. 3.4. Solvent replacement Acetonitrile is mostly used as organic modifier in the mobile phase in RPLC due to the fact that it has a low viscosity, is miscible with water and possesses a low UV absorbance [46]. From a green

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analytical chemistry point of view however it is less attractive as it is flammable and toxic [46]. Green environment friendly alternatives such as water, ethanol and carbon dioxide can replace acetonitrile in some applications [25], but to the best of our knowledge this has not been described for peptide analysis. The use of ethanol instead of acetonitrile as organic modifier can be an option when the poorer eluotropic strength of ethanol and its higher UV cut-off value is not an issue [46]. 3.5. Selected applications A mixed-mode solid phase extraction method followed by UHPLC–MS/MS is described by Lame et al. [72] for the quantification of multiple amyloid beta peptides in human cerebrospinal fluid. The authors conclude that their assay approaches the sensitivity of ELISA, but there is a higher sample throughput and it is more cost-effective [72]. The same authors described the quantification of bradykinin in human plasma after solid phase extraction using the microfluidic LC separation device from Waters. The limit of detection of the presented method was 2.5 pg/mL [73]. Kuze et al. [74] have developed a UHPLC–MS/MS method to quantify a new oral metastin receptor agonist, TAK-448, in human plasma. Separation was performed on a phenyl column in a total run time of 10 min. An UHPLC–MS/MS to determine arginine vasopressin in small volumes of plasma and urine in preterm infants is described by Zhang et al. [75]. The developed method was sensitive enough to allow measuring arginine vasopressin levels in the endogenous range [75]. Chambers et al. [76] developed a method for quantifying therapeutic insulin analogs using microelution solid phase extraction and UHPLC tandem mass spectrometry with a run time of 3.5 min. Improved peak shapes were obtained using a charged-surface (CSH) chromatographic column [76]. However, to allow quantification of human insulin and lispro and in order to achieve detection limits required for clinical samples, a multidimensional method was developed. The method included protein precipitation, solid phase extraction, trapping and back elution and separation on a superficially porous, charged surface sub-2 ␮m column [77]. Another paper of the same group presents a UHPLC–MS/MS method for the quantification of teriparatide, the 1-34 fragment of human parathyroid hormone, in human plasma [78]. The challenges of analyzing large peptides in biological fluids were addressed by optimizing sample preparation using a 96-well plate for solid phase extraction, by using the CSH chromatographic column and by choosing the appropriate MS transitions [78]. Thomas et al. [79] determined 11 prohibited, small peptides in urine samples using nano UHPLC and high resolution mass spectrometry (HRMS). In a following publication, they described the use of immunoaffinity purification in combination with nano UHPLC and HRMS for the analysis of prohibited peptides in plasma and urine [80]. Fillet and co-workers [48] have optimized a capillary LC–MS/MS method (300 ␮m id column) for the analysis of hepcidin. Later, the same group developed a nano LC-chip MS/MS system for the quantification of this peptide [47] with a limit of detection of 0.07 ng/mL. Very recently, we have published a validated nano UHPLC–MS/MS method for the quantification of three neuromedinlike peptides in microdialysis samples. Using a Xevo TQ-S mass spectrometer we achieved lower limits of quantification of 0.5 and 3 pM, corresponding to 2.5–15 amol on column [81]. 4. Alternatives to liquid chromatography for greener analysis Although LC analysis still remains the technique of choice for peptide bioanalysis, scientists are looking for methods that are

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faster, more sensitive and can handle low sample volumes. The section below therefore discusses the alternative possible techniques that may fulfill these requirements. However, it must be mentioned that at this moment not all alternative procedures and techniques can be used on a routine basis or replace the methods used today. 4.1. Capillary electrophoretic methods Capillary electrophoresis (CE) has long been established as an alternative separation technique, well suited for the analysis of complex mixtures [82]. Its major advantages include high separation efficiency, ease of method optimization and the possibility to analyze small sample volumes. The sensitivity is inferior compared to LC, but tremendous improvements can be obtained using stacking methods [82]. A practical challenge when analyzing biopharmaceuticals is how to reduce wall adsorption by adequate rinsing steps and coating of the capillary wall [82]. Furthermore, significant adsorption of matrix components is also observed when analyzing biological samples, in this way affecting the electroosmotic flow and analyte migration [23,82]. Examples of quality control and quality assurance of peptide and peptidomimetic drugs with capillary zone electrophoresis are described in literature [83–85]. Two recent publications will be discussed as examples. Tamizi and Jouyban [86] have described a stability indicating method for the analysis of octreotide acetate, a synthetic analog of somatostatin. The method was applied for the quantification of the peptide in pharmaceutical formulations and to study its degradation kinetics under various stress conditions [86]. The group of Fillet has developed a simple and efficient micellar electrokinetic chromatography method to simultaneously determine human insulin, its five analogs, the main degradation products and the excipients usually present in injection formulations. A fast method with an analysis time of 3 min was validated for the analysis of human insulin in commercial formulations [87]. Quantification of peptides in biological matrices is more complicated as CE suffers from relative low concentration sensitivity. However, this can be solved by using alternative detection systems such as laser-induced fluorescence, electrochemical or MS detection, together with online sample enrichment techniques [83,84]. Moreover, the small sample volume needed, the high efficiency and thus high resolving power make these techniques highly interesting [83–85]. A more detailed description of possible applications can be found elsewhere [83,84]. 4.2. Supercritical fluid chromatography Supercritical fluid chromatography (SFC) has undergone major instrumental improvements over the past years, resulting in a renewed interest in this technique. A supercritical fluid is obtained by elevating the temperature and pressure above the characteristic critical values of a substance. This reversible physical state possesses unique and interesting properties to apply in chromatography, such as the low viscosity and high diffusivity. This phase has properties that are comparable to that of gaseous mobile phases, which results in lower generated pressures over the system and column. The density and solvating power are comparable with that of liquid mobile phases, which creates a broad application range [88,89]. The technique provides high separation efficiency, selectivity and fast analysis. In SFC, carbon dioxide in the subcritical or supercritical state is used almost exclusively as main eluent. Organic modifiers, such as methanol or ethanol, are often added to the mobile phase to enhance the solvent strength and to allow elution of the compounds in a reasonable time frame. In addition, small amounts of acidic or basic additives are included to improve the peak shape of the analytes [46,90,91]. Because of the short analysis times, limited environmental impact and low organic solvent consumption, SFC fulfills the requirements of green analytical

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chemistry approaches [25]. Indeed, carbon dioxide is considered a green solvent compared to other solvents classically used in analytical chemistry [91]. Recently, the interest for peptide analysis with packed column SFC is growing, due in part to the rather extensive analysis time required by LC [29]. A limited number of applications for peptide analysis with SFC have so far been reported. They are summarized in a recent review by Lesellier and West [91]. For the moment, only rather hydrophobic peptides have been successfully analyzed. Indeed, the low solubility of very polar and ionic compounds in carbon dioxide based mobile phases limits their analysis with SFC [91]. An interesting study on the feasibility of analyzing polypeptides containing a variety of hydrophilic, basic and acidic amino acids was performed by Zheng et al. [92]. Moreover, Tognarelli et al. [29] obtained the separation of five peptides ranging in molecular weights from 238 to 1046 by SFC in a nearly fivefold decreased analysis time compared to LC. 5. Challenges related to mass spectrometric quantification of peptides The adoption of MS-based methodologies for peptide analysis has been relatively slow. This is in part due to the difficulty of achieving the same level of sensitivity of immunological methods [35]. The main cause of low sensitivity is that the peptide ion current is divided amongst the multiple charge states that are commonly observed in electrospray ionization [35]. Summation of different transitions can compensate to some extent the loss in signal intensity inherent to the appearance of multiple charge states [28]. The combination of an effective sample preparation method and the use of up-to-date instruments can enable peptides to be detected with sensitivities similar to immunologicalbased approaches [35]. For peptide compounds, the formation of specific multiple charged precursor ions and their relative abundance is influenced by both flow rate and solvent composition [76]. Tuning should therefore always be performed using a peptide solution in the mobile phase combination at which it elutes and using the flow rate of the LC method. Moreover, the improvement in MS instrumentation is an important step forward for sensitive analyses. We have recently compared two generations of triple quadrupole instruments for the quantification of three neuromedin-like peptides in rat brain microdialysis samples. A 20 to 30-fold improvement in sensitivity was obtained on the new generation instrument, allowing in vivo analysis of these peptides [81]. Furthermore, electrospray ionization is sensitive to matrix effects and these should therefore be studied early in method development. Indeed, co-eluting matrix components, which are not observed in the chromatogram, can have a detrimental effect on the analysis, since they can cause ion suppression or enhancement of the analyte [18]. The addition of a stable isotope labeled internal standard can correct for these matrix effects. However, even if matrix effects can be compensated by the use of an appropriate internal standard, efforts should be made to eliminate co-eluting compounds, since their presence will reduce method sensitivity. Possible solutions to reduce or eliminate matrix effects are described elsewhere [18]. 6. Concluding remarks The increasing importance of peptide therapeutics necessitates performance improvements in bioanalytical techniques to support biopharmaceutical drug development. The development of sensitive, selective, high-throughput methods for peptide analysis using green analytical techniques is highly demanded. Developing

bioanalytical methods for peptides remains complicated, since their behavior under various sample preparation and chromatographic conditions is still not well understood. Proper solubilization of the peptides is fundamental to obtain the required sensitivity and to diminish nonspecific binding. More environmentally friendly chromatographic developments include miniaturization, faster analysis and the use of less toxic solvents. Moreover, capillary electrophoresis and supercritical fluid chromatography are useful alternatives and can also be used as orthogonal separation assays compared to liquid chromatography. Although some of these techniques seem really promising, still a lot of research should be performed before they can be applied routinely in a (regulated) peptide bioanalysis laboratory. References [1] P. Vlieghe, V. Lisowski, J. Martinez, M. Khrestchatisky, Synthetic therapeutic peptides: science and market, Drug Discov. Today 15 (2010) 40–56. [2] J. Reichert, A. Tartar, M. Dunn, Development Trends for Peptide Therapeutics – A Comprehensive Quantitative Analysis of Peptide Therapeutics in Clinical Development, Summary Report, Peptide Therapeutics Foundation, 2010. [3] I. van den Broek, W.M. Niessen, W.D. van Dongen, Bioanalytical LC–MS/MS of protein-based biopharmaceuticals, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci. 929 (2013) 161–179. [4] T. Uhlig, T. Kyprianou, F.G. Martinelli, C.A. Oppici, D. Heiligers, D. Hills, X.R. Calvo, P. Verhaert, The emergence of peptides in the pharmaceutical business: from exploration to exploitation, EuPA Open Proteomics 4 (2014) 58–69. [5] A.A. Kaspar, J.M. Reichert, Future directions for peptide therapeutics development, Drug Discov. Today 18 (2013) 807–817. [6] L. Diao, B. Meibohm, Pharmacokinetics and pharmacokinetic–pharmacodynamic correlations of therapeutic peptides, Clin. Pharmacokinet. 52 (2013) 855–868. [7] R. Lax, The future of peptide development in the pharmaceutical industry, PharManufacturing: Int. Pept. Rev. (2010) 10. [8] P.W. Latham, Therapeutic peptides revisited, Nat. Biotechnol. 17 (1999) 755–757. [9] A.K. Sato, M. Viswanathan, R.B. Kent, C.R. Wood, Therapeutic peptides: technological advances driving peptides into development, Curr. Opin. Biotechnol. 17 (2006) 638–642. [10] M. Ewles, L. Goodwin, Bioanalytical approaches to analyzing peptides and proteins by LC–MS/MS, Bioanalysis 3 (2011) 1379–1397. [11] V. Vergote, C. Burvenich, C. Van de Wiele, B. De Spiegeleer, Quality specifications for peptide drugs: a regulatory-pharmaceutical approach, J. Pept. Sci. 15 (2009) 697–710. [12] M. D’Hondt, N. Bracke, L. Taevernier, B. Gevaert, F. Verbeke, E. Wynendaele, B. De Spiegeleer, Related impurities in peptide medicines, J. Pharm. Biomed. Anal. 101 (2014) 2–30. [13] L. Andersson, L. Blomberg, M. Flegel, L. Lepsa, B. Nilsson, M. Verlander, Largescale synthesis of peptides, Biopolymers 55 (2000) 227–250. [14] S. Van Dorpe, M. Verbeken, E. Wynendaele, B. De Spiegeleer, Purity profiling of peptide drugs, J Bioanal. Biomed. S6 (2011) 1–15. [15] R. Lucena, Commentary Microextraction Techniques in Bioanalysis http:// www.bioanalysis-zone.com/articles/commentary-microextractiontechniques-in-bioanalysis/ [16] G. Lasarte-Aragonés, R. Lucena, S. Cárdenas, M. Valcárcel, Nanoparticle-based microextraction techniques in bioanalysis, Bioanalysis 3 (2011) 2533–2548. [17] J.W. Howard, R.G. Kay, S. Pleasance, C.S. Creaser, UHPLC for the separation of proteins and peptides, Bioanalysis 4 (2012) 2971–2988. [18] A. Van Eeckhaut, K. Lanckmans, S. Sarre, I. Smolders, Y. Michotte, Validation of bioanalytical LC–MS/MS assays: evaluation of matrix effects, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci. 877 (2009) 2198–2207. [19] F. Pena-Pereira, I. Lavilla, C. Bendicho, Liquid-phase microextraction techniques within the framework of green chemistry, TrAC: Trends Anal. Chem. 29 (2010) 617–628. [20] C. Ghosh, Green bioanalysis: some innovative ideas towards green analytical techniques, Bioanalysis 4 (2012) 1377–1391. [21] J. Namiesnik, M. Tobiszewski, Developments in green chromatography, LC GC Europe 27 (2014) 405. [22] L. Tang, A.M. Persky, G. Hochhaus, B. Meibohm, Pharmacokinetic aspects of biotechnology products, J. Pharm. Sci. 93 (2004) 2184–2204. [23] R. Haselberg, G.J. de Jong, G.W. Somsen, CE–MS for the analysis of intact proteins 2010–2012, Electrophoresis 34 (2013) 99–112. [24] S. Armenta, S. Garrigues, M. de la Guardia, Green analytical chemistry, TrAC: Trends Anal. Chem. 27 (2008) 497–511. [25] H. Shaaban, T. Górecki, Current trends in green liquid chromatography for the analysis of pharmaceutically active compounds in the environmental water compartments, Talanta 132 (2015) 739–752. [26] G. Hopfgartner, A. Lesur, E. Varesio, Analysis of biopharmaceutical proteins in biological matrices by LC–MS/MS II. LC–MS/MS analysis, TrAC: Trends Anal. Chem. 48 (2013) 52–61.

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Please cite this article in press as: A. Van Eeckhaut, D. Mangelings, Toward greener analytical techniques for the absolute quantification of peptides in pharmaceutical and biological samples, J. Pharm. Biomed. Anal. (2015), http://dx.doi.org/10.1016/j.jpba.2015.03.023