Developments in the fields of protein chemistry and pharmaceutical biotechnology have increased the demand for suitable analytical techniques for the characterization of intact proteins. Capillary electrophoresis (CE) coupled to mass spectrometry (MS) has proven to be a powerful tool for this purpose because it combines high separation efficiency with mass selectivity. This article provides an overview of CE–MS method development work in our laboratory, for biopharmaceutical analysis applying noncovalent capillary coatings to prevent protein adsorption. Technological aspects with respect to the use of CE–MS interfaces and types of noncovalent capillary coatings applied are treated. Furthermore, some typical examples are highlighted to demonstrate the versatility of CE–MS for the analysis of biopharmaceuticals. These include the analysis of recombinant human growth hormone, recombinant human interferon-

-1a, recombinant human erythropoietin and protein–drug conjugates. We concluded that there is strong potential for CE–MS systems using noncovalent coated capillaries for purity and stability analysis of biopharmaceuticals. 

Biopharmaceuticals have an increasingly important position in drug development and production (1). As for any drug designed for human use, biopharmaceuticals have to meet stringent quality requirements to ensure efficacy and safety. However, because of their macromolecular nature, the full characterization of biopharmaceuticals in terms of identity, content, purity, stability, conformation and function poses a great challenge (2,3). Moreover, multistep biotechnological production processes may show variability, introducing product diversity and different isoforms that can affect activity. With the appearance of biosimilars that are supposed to be copies of the innovator product (3), the demand for an effective quality assessment of therapeutic proteins has increased even further. 

Next to techniques such as liquid chromatography (LC) and gel electrophoresis, capillary electrophoresis (CE) is a proven powerful separation technique for biotechnology-derived proteins (4). CE analyses can be performed under mild conditions without the need for organic solvents or very high salt concentrations. This allows the study of proteins without causing conformational changes or protein degradation during analysis. In CE, separations are based on differences in the charge-to-size ratio of the analytes. Therefore, subtle differences among proteins, such as those caused by post-translational modifications like glycosylation, deamidation, or phosphorylation, can be differentiated with CE. Mass spectrometry (MS) has developed into one of the most popular and useful detection techniques in separation science. MS is well suited for protein analysis because of its sensitivity and selectivity. Furthermore, MS detection with high mass accuracy and resolution, such as provided by time-of-flight (TOF) instruments, can considerably enhance the utility of CE by providing information about the identity of the separated compounds. Therefore, coupling CE to MS creates a powerful analytical tool for the characterization of intact proteins (5,6). 

Despite its potential, the application of CE–MS for the analysis of biopharmaceuticals has been relatively limited. CE–MS methods that distinguish natural human growth hormone (hGH) from recombinant hGH (7) and reveal degradation products in hGH after heat exposure and prolonged storage (8,9) were introduced. Furthermore, the usefulness of CE–MS for the intact glycoform analysis of erythropoietin has been demonstrated (for example, references 10–12 and the references therein). During the last few years, our group has been working on the development of CE–MS methodologies for the characterization of intact proteins and biopharmaceuticals in particular. In this paper some representative data is provided showing the strength of the combination of CE with high-resolution TOF-MS for the characterization of protein drugs.  

: Electrospray ionization (ESI) is the main ionization technique used for the CE–MS analysis of proteins. ESI produces multiple charged protein ions in the gas phase. As a result, a so-called protein charge envelope comprising ions with mass-to-charge ratios of typically 500–3000 are obtained. Protein molecular masses can be determined by deconvolution of the ESI mass spectrum. Interfacing CE with MS via an ESI source is generally performed with a so-called sheathliquid interface (Figure 1). In the sheath-liquid interface, the CE capillary is surrounded by a second capillary in a coaxial arrangement. A sheath liquid is led through this second capillary, which merges with the CE effluent at the capillary outlet. A typical sheath liquid contains an organic solvent, water, and an acid, and has a flow rate between 2–5 μL/min. The terminating CE voltage is applied to the sheath liquid to close the electrical circuit. Often, a third coaxial tube delivers a gas flow that helps with nebulization into the ESI source. An important feature of this type of interface for protein analysis is the sheathliquid composition (13,14), because it might influence the signal intensity as well as the shape and position of the protein's charge envelop. In the studies described below, a Beckman Coulter MDQ CE instrument was coupled to a Bruker Daltonics micrOTOF mass spectrometer using a CE–MS interface from either Agilent Technologies or Beckman Coulter. 

Figure 1: Schematic representation of a sheath-liquid  CEâMS interface. 

: Because high-molecular-weight species like proteins have relatively low diffusion coefficients, theoretically, very high plate numbers can be obtained. However, an important limitation encountered in the CE analysis of proteins is their tendency to adsorb to the fused-silica capillary wall (15). The most common approach in CE to prevent protein adsorption is to coat the inner capillary wall with nonadsorptive agents. An elegant and simple way to create effective coatings is by applying charged polymers to the fused-silica surface (16). We have demonstrated the usefulness of noncovalent polyelectrolyte coatings of polybrenepoly(vinylsulphonic acid) (PB-PVS) and polybrene-dextran sulphate-polybrene (PB-DS-PB) for CE–MS analysis of intact proteins (8,9, 17–19). The negatively charged PB-PVS coating is used in combination with a background electrolyte (BGE) of medium or high pH for the analysis of acidic proteins. For basic proteins, the positively charged PB-DS-PB coating is applied in combination with a low-pH BGE. These coatings provide highly reproducible and efficient separations for both acidic and basic proteins. Typical migration time RSD values are below 1.6% (

 = 40), whereas plate numbers are generally around 100?000. The performance of these coatings in CE–MS was thoroughly studied (18) and their effectiveness against protein adsorption was also investigated in situ (20). Because of the permanent charge of the coatings, a relatively high and pH-independent electro-osmotic flow (EOF) is obtained. However, this high EOF also decreases the effective separation window, which can be limiting when more complex samples have to be analysed. Polyacrylamide coatings can be very suitable for reducing the EOF velocity to virtually zero and thereby improving the CE resolution (10). After applying such a coating, the EOF is reduced significantly, improved resolutions are obtained, and the analytical performance is not compromised.  

: Recombinant human growth hormone (rhGH) is a therapeutic protein used in the treatment of retarded growth and dwarfism. The monograph of rhGH in the

 includes a CE–UV method using bare fused-silica capillaries for detection of charge variants of this acidic protein. The use of PB-PVS coated capillaries for rhGH analysis by CE improves the method's repeatability, resolution, and efficiency. We developed an MS-compatible CE method using a volatile BGE of ammonium formate that provides a reliable determination of potential degradation products (8,9).  

The latter method was used in combination with TOFMS for the analysis of aqueous solutions of rhGH. First, a standard solution of rhGH was analysed using a BGE of high pH. A low-pH sheath liquid was used to allow efficient positive ESI. A narrow and symmetrical peak (

 ~ 100000) was obtained for rhGH [Figure 2(a), peak 0]. Deconvolution of the mass spectrum obtained in the apex of the peak [Figure 2(b), mass spectrum 0] confirmed the presence of rhGH (

 22,124). Next, a solution that was exposed to a temperature of 40 °C for 24 h and stored for one year at -18 °C was analysed (Figure 2). Remarkably, in this sample, no unmodified rhGH was detected although the main component (peak 1) had virtually the same migration time as rhGH. Based on the observed mass difference of 32 Da with respect to rhGH, this main degradation product was identified as doubly oxidized rhGH. The three additional degradation products could also be assigned [Figure 2(b)]. Peak 2 is clearly separated from the oxidized rhGH and differs by only 0.9 Da in mass. Most likely, this compound results from a combination of oxidation (double) as well as deamidation (single) of rhGH. Note that CE–TOF-MS can distinguish protein species that differ by one charge and 1 Da only. The molecular mass of the degradation product migrating at 12.2 min (peak 3) is 128 Da higher than rhGH, which in combination with its migration after compound 2, indicates that two sulphonic acid groups may have been formed from a protein disulphide bridge in addition to the double methionine oxidation. Considering its molecular mass and migration time, the component causing peak 4 most likely is the singly deamidated form of the sulphonated and oxidized product. The use of TOF-MS detection clearly adds important selectivity to the highly efficient CE separation, thereby enhancing the reliability of the method. Peaks 1 and 3 would probably erroneously have been assigned to unmodified rhGH and a bisdeamidated product, respectively, when using CE–UV. 

Figure 2: (a) Base-peak electropherograms obtained during CEâTOF-MS of rhGH (grey trace) and heat-exposed and stored rhGH (blue trace) using a PB-PVS coated capillary in combination with a background electrolyte of 75 mM ammonium formate (pH 8.5). The pound symbol indicates an unknown nonproteinaceous compound. (b) Mass spectra obtained in the apexes of the indicated peaks. After deconvolution, the degradation products (peak 1 to 4) were assigned based on their molecular mass and relative migration with respect to rhGH (peak 0). Adapted from reference 19. 

: The drug recombinant human interferon-β-1a (rhIFN-β) is a 23-kDa N-glycosylated protein (Asn

) that is used for the treatment of multiple sclerosis. It has been demonstrated that the glycosylation of rhIFN-β may significantly affect its biological activity (21). Therefore, establishing the glycoform heterogeneity among batches of rhIFN-β is important.  

rhIFN-β is a basic protein and, therefore, to avoid protein adsorption to the capillary wall, a PB-DS-PB coated capillary was selected in combination with an acidic BGE. The base-peak electropherogram (BPE) obtained after CE–MS analysis of rhIFN-β shows a distinct pattern of partially resolved bands [Figure 3(a)]. Figure 3(b) shows the deconvoluted mass spectrum obtained in the apex of the most intense peak (9.1 min), indicating a molecular mass of 22375 Da. It has been reported that the major glycoform rhIFN-β is a fucosylated disialylated biantennary structure with four hexose and five 

-acetylhexosamine units (21). This glycoform, including the amino acid chain of rhIFN-β, has a molecular weight of 22375 Da, which agrees well with the observed mass. To assign the glycoforms present in the rhIFN-β sample, deconvoluted mass spectra were constructed throughout the peak pattern. From these mass spectra a total of 10 distinctive masses could be determined (Figure 4). The masses all differ either 291 Da or 365 Da with respect to each other, indicating the loss or gain of either a sialic acid (SiA) unit or a hexose 

-acetylhexosamine (HexHexNAc) unit, respectively; unglycosylated rhIFN-β was not detected. With CE–MS, five out of the six glycoforms described in the 

 monograph of IFN-β, which uses direct ESI-MS analysis, were observed. Moreover, CE–MS revealed additional low-abundance glycoforms with more SiA and HexHexNAc units than described in the 

. This underlines the importance of an efficient separation before MS analysis to prevent signal suppression commonly observed in ESI-MS and allows for reliable and more detailed assignment. 

Figure 3: (a) Base-peak electropherogram obtained during CEâTOF-MS of rhIFN-Î² using a PB-DS-PB coated capillary in combination with a background electrolyte of 50 mM acetic acid (pH 3.0). (b) Deconvoluted mass spectrum obtained in the apex of the peak migrating at 9.1 min (denoted with an asterisk). The insert shows the assigned glycan structure with molecular weight 22,375 Da. Symbols: â/â, hexose; â², fucose; â , N-acetylhexosamine; â sialic acid. Adapted from reference 19. 

For each assigned glycoform an extracted ion electropherogram (EIE) was constructed using the most abundant 

 value of the glycoform's respective charge envelope. From the EIEs, relative peak areas for all 10 detected glycoforms were obtained leading to a quantitative glycoform distribution (Figure 4). The glycoform with the lowest abundance is almost 1% in area with respect to the most abundant isoform, but could still be detected reliably with a signal-to-noise ratio of 16 (

 = 3). The relative abundance of glycoforms may be useful for in-process and quality control of rhIFN-β batches or to compare biosimilar products. 

Figure 4: Migration time, molecular mass, relative peak area and glycan composition for rhIFN-Î² glycoforms as observed with CEâTOF-MS (see Figure 3). Symbols: â/â, hexose; â², fucose; â , N-acetylhexosamine; â, sialic acid. Adapted from reference 19. 

: Recombinant human erythropoietin (rhEPO) is a glycoprotein hormone that is frequently used for the treatment of anaemia. The protein has three N-glycosylation sites (Asn

). The total glycan content can make up about 40% of the protein's molecular weight and introduces high variation in both protein structure and mass. 

Figure 5: (a) Base-peak electropherogram obtained during CEâTOF-MS of rhEPO (epoetin-Î² product) using a polyacrylamide-coated capillary in combination with a background electrolyte of 2 M acetic acid (pH 2.0). (b) Deconvoluted mass spectrum obtained in the peak migrating at 25.4 min (denoted with an asterisk). (c) Composed extracted ion electropherogram (EIE) of different sialic acid isoforms with identical HexHexNAc content (Hex22HexNAc19Fuc3). The EIE was composed from the major ions of the respective charge distribution. 

Although rhEPO can be analysed on charged coatings in relatively short time, the separation of its various glycoforms is not optimal (10). To obtain a more efficient glycoform separation, epoetin-β was analysed with CE–TOF-MS using a neutrally coated capillary in combination with an acidic BGE (2 M acetic acid) (Figure 5). A cluster of overlapping peaks between 18 and 30 min was obtained in this EPO product. In Figure 5(b) the deconvoluted mass spectrum obtained for the most intense peak (25.4 min) is shown. A main mass of 29888 Da is obtained. Also, a variety of other molecular masses are observed corresponding to glycoforms differing in number of HexHexNAc and SiA units [Figure 5(b)]. Deconvoluted mass spectra were constructed throughout the peak profile resulting in a total of 70 distinctive masses (Figure 6). It is impossible to derive an exact glycoform composition based solely on the intact glycoform's mass. However, from literature it is known that the glycoform with a molecular weight of 29888 Da consists of the EPO protein backbone with 22 Hex, 19 HexNAc, 3 fucose (Fuc) and 13 SiA units (10). This provides a starting point allowing for the assignment of the other glycoforms (Figure 6). Next to the different glycoforms, signals that differ either 16 Da or 42 Da from the main mass were observed in the mass spectrum (data not shown) which could be assigned to oxidation and acetylation products. Overall, approximately 200 different isoforms could be discerned in this single sample. In a different batch of the same product, a cluster of glycoforms was observed that comigrated with the SiA

 glycoforms, but showed an increased mass of 146 Da. It was concluded that these glycoforms contain an additional Fuc residue (Figure 6). 

Figure 6: Main rhEPO glycoforms and their respective carbohydrate composition as derived from CEâTOF-MS (see FigureÂ 4). The molecular weights of the glycoforms with an additional fucose unit are indicated in italics. Their carbohydrate composition is indicated with the grey box. 

To gain a better insight into the obtained glycoform separation, EIEs were created for various rhEPO glycoforms. Figure 5(c) shows a composed EIE obtained for a species containing various numbers of SiA residues, but the same amount of HexHexNAc residues (Hex

). Clearly, these glycoforms are all well separated with increasing migration times for each additional negatively charged SiA to the glycan structure. The separation of HexHexNAc glycoforms bearing the same number of SiA residues is apparent, but less efficient (data not shown). Indeed, the addition of HexHexNAc residues merely results in a size difference of the protein because it does not bear any charged groups.  

Sensitivity Enhancement by Sheathless Interfacing: 

 In the studies described above, a sheath-liquid ESI interface to couple CE to MS detection was used. However, because of the dilution of the capillary effluent by the sheath liquid, the detection sensitivity is compromised. Still, in CE–MS of intact proteins there is a definite requirement for increased sensitivity. Multiple charging of proteins occurring during ESI distributes the overall signal intensity over many charge states, thereby increasing the achievable detection limits for intact proteins. To improve the sensitivity, recently a novel sheathless interface to couple CE with ESI-TOF-MS was tested in our group for the analysis of intact proteins (22,23). The interface was developed in the laboratories of Beckman Coulter, based on a design introduced by Moini (24). In this design (Figure 7), 3–4 cm of the terminating end of the fusedsilica separation capillary is etched with hydrofluoric acid, producing a ~5-μm thick porous capillary wall allowing electrical contact when immersed in BGE. Limits of detection (LODs) between 0.5 and 1.3 nM were obtained for intact proteins with the sheathless CE–MS system using a nanoESI source, which was 50- to 140-fold better than sheath-liquid interfacing using the same capillary (22). The gain in sensitivity was the result of both reduced noise levels and increased analyte responses as obtained with sheathless CE–MS. 

Figure 7: Schematic representation of the sheathless CEâMS capillary positioned in a stainless steel capillary. Adapted from reference 22. 

The sheathless CE–MS system was used for the analysis of drug–protein conjugates (23). Kinase inhibiting drugs, such as erlotinib, were coordinatively bound to lysozyme using a BOCmethionine and a platinum-based linker [Figure 8(a)]. The reaction products (total protein concentration per preparation, 100 nM) were analysed by CE–TOF-MS. The BPE of the erlotinib-lysozyme preparation showed a pattern of peaks with migration times between 8 and 9 min [Figure 8(b)]. For all detected components, EIEs were constructed using the signal for the most intense protein charge state of the respective protein species, showing distinct and well-resolved peaks [Figure 8(c)]. Deconvolution of the mass spectra obtained in the apices of the peak provided the molecular weights of the components found. Taking into account the relative migration of the different components and the molecular masses of the platinum linker, the kinase inhibitors, the BOCmethionine residue, and lysozyme, all detected components could be assigned [Figure 8(c)]. The efficient CE separation of the highly related conjugate products could be obtained because of protein charge changes induced by drug conjugation (net reduction of positive charge). On the other hand, conjugation of the protein to the drug-platinum complex containing Pt

  results in an increase in charge-to-mass ratio of the overall lysozyme conjugate. 

Figure 8: (a) Schematic representation of the production of drug-protein conjugates. (b) Base-peak electropherogram obtained during CEâTOF-MS of the erlotinib-ULS-BOCmet-lysozyme preparation using a positively-charged coating in combination with a background electrolyte of 100 mM HAc (pHÂ 3.1). (c) Extracted ion electropherograms for the main ion of peaks 1â6 constructed at the indicated m/z values, with interpretation and quantitation of the observed species on the right. After deconvolution, the reaction products (peaks 1â3, 5 and 6) were assigned based on their molecular mass and relative migration with respect to unreacted lysozyme (peak 4). Adapted from reference 23. 

It is important to note that no BOCmethionine–lysozyme variants were detected at the same migration times of the drug–lysozyme products. This means that the platinum-coordinated drug–lysozyme conjugates do not dissociate during ESI. Furthermore, for all components discrete symmetrical peaks are obtained, which indicates that the conjugates were stable during CE separation. In addition, no shift in protein mass spectra upon drug conjugation was observed. From these observations, it is concluded that the CE–TOF-MS results accurately reflect the actual qualitative and quantitative composition of the drug–lysozyme conjugate preparations. Therefore, EIEs were used to determine peak areas for all detected components allowing a quantitative estimation of the relative composition of the preparations [Figure 8(c)]. Based on the relative peak area of each component we can derive that the different components were present in concentrations of 5–29 nM, illustrating the ability of the sheathless CE–TOF-MS system to detect low-abundant conjugates, which would not have been possible with sheath-liquid CE–MS.  

During the past few years, we have demonstrated that CE–TOF-MS using noncovalently coated capillaries represents an efficient and reproducible tool for the characterization of biopharmaceuticals. From the presented results it can be concluded that CE is especially suitable to probe protein modifications that lead to charge differences. For example, degradation products as a result of deamidation or sulphonic acid formation could be efficiently resolved from the parent compound. Also, protein glycoforms that differ in number of sialic acid groups could be separated. In cases where modifications do not affect the overall protein charge, the high mass resolution and accuracy of TOF–MS still allowed for the assignment of components. Overall, we believe that CE–TOF–MS provides a highly promising means for the qualitative and quantitative assessment of the heterogeneity of (glycosylated) biopharmaceutical products. With state-of-the-art CE equipment routinely present in laboratories of the contemporary biopharmaceutical industry, and high-quality TOF mass spectrometers becoming very accessible and affordable, we expect CE–MS to find widespread application in the near future. The application of CE–MS for the comparison of profiles of biopharmaceutical biosimilars would be very interesting and highly significant with respect to regulatory, commercial, and therapeutic issues connected to the entrance of more and more "generic" biopharmaceuticals on the market. The excellent performance provided by the CE–TOF-MS methods might allow for reliable comparisons between different batches and products. Furthermore, CE–MS analysis of monoclonal antibodies, which represent the most significant part of today's biopharmaceuticals, has hardly been explored so far. As many monoclonal antibody modifications involve a change in net charge and size of the protein, CE–MS could also be a very suitable tool for the characterization of these proteins. However, creating gas-phase ions by ESI from such high-molecularweight compounds after a CE separation will require thorough optimization of both separation and ionization parameters.  

This research was supported by the Dutch Technology Foundation STW, Applied Science Division of NWO and the Technology Program of the Ministry of Economic Affairs. Vera Brinks, Robbert Jan Kok, Stefan Harmsen and Emmy Dolman of the Department of Pharmaceutics of Utrecht University and Chitra Ratnayake, Jeff Chapman and Hans Dewald of Beckman Coulter Inc. are kindly acknowledged for their input in parts of the presented work.  

Rob Haselberg, Gerhardus J. de Jong and Govert W. Somsen

 work at the Biomolecular Analysis group of Utrecht University in The Netherlands. Please direct correspondence to: 

(1) Pharmaceutical Research and Manufacturers of America, 

(2)  D.J.A. Crommelin, G. Storm, R. Verrijk, L. de Leede, W. Jiskoot and W.E. Hennink,

(5) R. Haselberg, G.J. de Jong and G.W. Somsen, 

(6) R. Haselberg, G.J. de Jong and G.W. Somsen, 

(7) A. Staub, S. Giraud, M. Saugy, S. Rudaz, J.L. Veuthey and J. Schappler, 

(8) J.R. Catai, J. Sastre Toraño, G.J. de Jong and G.W. Somsen, 

(9) J.R. Catai, J. Sastre Toraño, P.M.J.M. Jongen, G.J. de Jong and G.W. Somsen, 

(10) E. Balaguer, U. Demelbauer, M. Pelzing, V. Sanz-Nebot, J. Barbosa and C. Neusü , 

(11) A. Taichrib, M. Pelzing, C. Pellegrino, M. Rossi and C. Neusü , 

(12) E. Giménez, F. Benavente, J. Barbosa and V. Sanz-Nebot, 

(13) G. Brenner-Weiss, F. Kirschhofer, B. Kuhl, M. Nusser and U. Obst, 

Articles

An overview of the important and versatile role that capillary electrophoresis coupled to mass spectrometry plays in the biopharmaceutical analysis.

Capillary Electrophoresis-Mass Spectometry for the Analysis of Biopharmaceuticals

The solvent chosen for injection can affect the appearance of chromatographic peaks.

A recent observation in the laboratory of one of the authors (B.A.) of this column highlights a problem that has been mentioned in "

" discussions before, but its importance justifies additional discussion about the selection of the injection solvent to use with a liquid chromatography (LC) method. In this particular case, it was observed that the peak areas were constant with two different injection solvents, but the peak heights were not. Similar results were observed with methods for two different compounds (A and B). Let's see what the likely cause is for this problem. 

We can't share the complete details of the methods because they are proprietary, but the key elements follow. A 10 ppm concentration of sample is prepared in distilled water or in 100% methanol, and 20 μL of this solution is injected. A 250 mm × 4.6 mm C18 column packed with 5-μm diameter particles is used with UV absorbance detection at 210 nm. For compound A (method A), a mobile phase of 60:40 methanol–water is used at a flow rate of 1.1 mL/min; for compound B (method B), the mobile phase is 70:30 (methanol–water) with a flow rate of 1.4 mL/min. The data are summarized in Table 1. Both compounds (A and B) are well retained with 

-values of approximately 2.6 in each case. You can see that the peak area differences between injection in methanol and water are ≤0.1% for both methods. However, the peak heights are approximately 30% larger for the water injections. 

Table 1: Chromatographic results for two methods A and B. 

The chromatograms of Figure 1 illustrate the influence of the injection solvent on the appearance of peaks in the chromatogram for another sample (1). In each case, the reversed-phase column was operated with an 18% acetonitrile–buffer mobile phase, where the buffer was 81:1 water–acetic acid. Injection volumes of 30 μL were made in various injection solvent compositions. 

Figure 1: Chromatograms for caffeine (first peak of each chromatogram) and salicylamide (second peak). Injections of 30 Î¼L of ~1 mg/mL of each analyte dissolved in (a) acetonitrile, (b) mobile phase, (c) 81:1 waterâacetic acid, and (d) water. Column: 250 mm Ã 4 mm, 10-Î¼m dp; mobile phase: 18:81:1 acetonitrileâwaterâacetic acid; flow rate: 1 mL/min. Adapted from reference 1. 

First, consider the cases where the injection solvent is no stronger than the mobile phase [Figures 1(b)–1(d)]. In each case, the retention times of each peak are approximately the same. Although we don't have data for the peak areas in each case, we'll assume they are the same, because the same volume of the sample at the same concentration was used in each case. Note, however, that the peak heights differ. Water is the weak mobile-phase solvent in this example, and the less water that is present in the injection solvent, the shorter the peak heights are. These observations are consistent with the problem presented at the beginning, where peak areas and retention times were constant, but peak heights dropped when less water was used in the injection solvent. 

Next, notice what happens when the sample of Figure 1(a) was injected in 30 μL of 100% acetonitrile — the peaks are broader, distorted and at shorter retention times. This example continues the pattern of Figures 1(b)–1(d), where the peaks are broader and shorter as less water is used in the injection solvent.  

Let's think on a conceptual basis about what is going on with the various examples of Figure 1. If the sample is injected with mobile phase as the injection solvent, it does not have to undergo any chemical changes, so analyte molecules should immediately begin to interact with the stationary phase in the column and undergo normal retention behaviour. However, if the injection volume is large enough, you can imagine that some band broadening will take place during the injection. For example, if the injection is extremely large, such as 1 mL, the first sample molecules will travel a significant way through the column before the last of the sample molecules arrive at the inlet. The distance between the first and last sample molecules in a given band determine the peak width, so a broad peak would be observed. At the other extreme, an infinitely small injection would have none of this injection-related broadening, so the peak would be much narrower. As we'll see later, we generally can tolerate an injection volume of approximately 15% of the peak volume of the first peak of interest before we find the injection-related band broadening objectionable. 

When the injection solvent is changed, we have another variable in addition to the injection volume mentioned above. In this case, the injected solvent plug must be diluted to the same composition as the mobile phase before "normal" retention can occur. If we think of the injection plug as a ball of solvent at the inlet of the column, it will shrink as it travels through the column, with the outside edges being diluted to the mobile-phase concentration before the centre of the ball. While they are inside the ball of the injection solvent, molecules will travel down the column as if that solvent was the mobile phase, whereas those diluted into the mobile phase will now travel at the normal rate in the mobile phase. So if the injection solvent is stronger than the mobile phase, it will tend to sweep analyte molecules through the column more quickly than normal. Because some analyte molecules travel more quickly (in the middle of the injection plug) than others (on the outer edges), injection in a solvent stronger than the mobile phase tends to smear the sample along the column, causing peak distortion. On the other hand, if the injection solvent is weaker than the mobile phase, analyte molecules inside the injection plug will move more slowly than normal, which tends to concentrate them at the column inlet. This is often referred to as on-column concentration and is a technique that can be used to inject large volumes of dilute samples without excessive band broadening. And, as you might expect, the influence of the injection solvent depends on both its volume and how much its composition differs from the mobile phase. A small volume of a strong solvent will be less detrimental than a large volume of the same injection solvent, and at a small enough volume, even a very strong injection solvent will get diluted very quickly, so it will be of little consequence. 

It can be shown (see discussion of reference 2) that if you want to have <1% loss in resolution because of injectionrelated peak broadening, the injection volume should be <15% of the peak volume of the first peak of interest if mobile phase is used as the injection solvent. If you are willing to tolerate a 10% loss in resolution, you can increase this to <40% of the peak volume. You can determine the peak volume of a peak by measuring its baseline width (or half-height width × 1.7) and multiplying by the flow rate. Alternatively, you can get a good estimate of the peak volume with a few simple calculations that follow. We think it is a good idea to make this calculation and compare the results with the actual measurements as a doublecheck to make sure there isn't excessive peak broadening because of other causes, such as a bad column. 

We can use the data of Figure 1 as an example. The expected width can be estimated from the column plate number, 

 is the peak width at baseline between tangents drawn to the sides of the peak. Equation 1 can be rearranged to   

, which for real samples under practical separation conditions can be estimated as  

 is the column length (in millimetres) and 

 is the particle diameter (in micrometres). The column of Figure 1 is 250 mm long packed with 10-μm particles, so 

 ≈ 7500. Injected in the weakest solvent, water [Figure 1(d)], retention times are 4.72 and 9.05 min. With equation 2, this translates to expected peak widths of 0.0545 and 0.1045 min, respectively. (The usual disclaimer applies here: Values have been rounded for convenience, so if you try to reproduce these calculations, your results may vary somewhat.) Multiply by the flow rate (1 mL/min) to convert this to peak volumes of 55 and 105 μL. 

If we use the <15% rule mentioned at the beginning of this section, maximum recommended injection volumes in mobile phase are then 0.15 × 55 = 8 μL and 0.15 × 105 = 16 μL. If we are interested in the first peak, this will be the limiting case, so injection of 5–10 μL in mobile phase would be expected to cause less than 1% loss in resolution. If we are more liberal with the tolerance for loss of resolution, as might be justified in the case of Figure 1, where resolution >>2, the <40% rule should be adequate, or 0.4 × 55 = 22 μL. Similar calculations can be performed for the data of Table 1 and we would see that the <15% rule would allow approximately 10-μL injections and the <40% rule would allow approximately 30-μL injections. 

Now, the final test has to do with whether or not all these estimates line up with what is observed in reality. First, let's consider the data of Figure 1. Resolution is directly related to peak width, so a 1% or 10% loss in resolution would correspond to a 1% or 10% increase in peak width, respectively. Also, any increase in peak width should give a corresponding decrease in peak height, so we can indirectly measure any loss of resolution by a reduction in peak height. The 30 μL injections of Figure 1 are approximately 50% larger than the 22-μL injection volume estimated above for a 10% loss in resolution. You can see that the peak heights for injection of 30 μL of sample in mobile phase [Figure 1(b)] are 60–70% of those when 30 μL of water is injected [Figure 1(d)]. So some broadening is occurring, as expected –partly because of the injection in mobile phase rather than a weaker solvent and partly because of the larger than recommended injection volume. Note that in all cases where the mobile phase or weaker injection solvent was used [Figures 1(b)–1(d)], the retention times were unchanged (within normally expected sample-to-sample variability). 

Contrast the results of Figures 1(b)–1(d) with those of Figure 1(a). In the case of Figure 1(a), both too large a sample was injected and too strong an injection solvent was used. This resulted in two problems. First, the peaks broadened unacceptably, especially for the first peak, which is severely distorted. Second, some of the sample molecules are swept down the column during the injectionsolvent dilution process, giving shorter retention times. These problems could be solved by injecting a much smaller sample volume or using a more dilute injection solvent. 

The data of Table 1 show the same pattern, but less severely. This is because of at least two factors. First, the injected volumes (20 μL) are less than those of the <40% rule, and the difference between the injection solvent strength and the mobile phase is less. For Figure 1(a), 30 μL of acetonitrile is injected into an 18% acetonitrile mobile phase, whereas in Table 1, 20 μL of methanol is injected in a 60% or 70% methanol mobile phase. 

We have seen that the combination of injection solvent strength and injection solvent volume must be chosen carefully to avoid unwanted band broadening and the resulting loss of resolution. In the examples of Table 1 and Figures 1(b)–1(d), the injection conditions were responsible for some loss in resolution (measured indirectly from loss in peak height), but no change in retention was observed. The peaks of Figure 1(a), however, were distorted and lost retention because too much of too strong a solvent was injected. 

The <15% and <40% rules discussed above are guidelines that seem to work fairly well. But, as with all guidelines, it is a good idea to give the actual results a reality test. One way was discussed here, where estimates of peak volumes and injection conditions were compared with actual experimental conditions. Another technique is simply empirical – try increasing or decreasing the proposed injection volume by twofold. In the same manner, injection solvent concentrations that are closer to the mobile phase than the 100% strong or weak solvent conditions used in Table 1 and Figure 1(a) would be expected to have a less detrimental result. For example, injection in 70% methanol into a 60% methanol mobile phase would be expected to be less of a problem than an injection solvent of 100% methanol. If significant differences in the chromatograms are observed, you should adjust the conditions so that you are not operating too close to the reliability limits of the method. 

is a PhD student in the laboratory of Dr Hugh Flowers at Glasgow University in Scotland. 

 is vice president of LC Resources, Walnut Creek, California, USA. He is also a member of 

editorial advisory board. Direct correspondence about this column should go to "LC Troubleshooting", 

4A Bridgegate Pavilion, Chester Business Park, Wrexham Road, Chester, CH4 9QH, UK, or e-mail the editor, Alasdair Matheson, at 

(2) L.R. Snyder, J.J. Kirkland and J.W. Dolan, 

Introduction to Modern Liquid Chromatography, 3rd Ed. 

 (Wiley, Hoboken, New Jersey, 2010), section 2.6.1. 

LC Troubleshooting

The solvent chosen for injection can affect the apearance of chromatographic peaks.

The Role of the Injection Solvent

Using a molecular diversity model to choose test probes from a chemical library, guest authors Ray McClain and Matt Przybyciel characterized 12 supercritical fluid chromatography (SFC) achiral stationary phases with 60 compounds from four chemical classes — amines, amides, alcohols and carboxylic acids — to establish guidelines for the rational selection of the optimum stationary phase for separations. Efficiency, selectivity and peak shape were used as the evaluation criteria.

The use of supercritical fluid chromatography (SFC) as a separation technique continues to grow (1–6). SFC offers a number of advantages to chromatographers for both analytical and preparative separations. These advantages include the "green" aspect of using carbon dioxide as a mobile phase, the ease of isolation of purified components from preparative separations, and decreased analysis times because of the high diffusivity and low resistance to mass transfer encountered in SFC. For many years, SFC has proven extremely useful for chiral preparative separations (7–12). Achiral SFC separations utilize stationary phases from "normal-phase" high performance liquid chromatography (HPLC), such as unmodified silica, diol, amino and cyano (13). During the past 10 years, a number of commercially available stationary phases have been developed specifically for SFC applications (14). However, not all of these columns are well-suited for all structural classes of compounds, presenting a bottleneck when arrays of chromatographic data must be acquired and interpreted to find a suitable match. Recently, chemometric approaches have been developed to predict retention and selectivity for a variety of stationary phases used for SFC (1,15–19). These approaches can be a useful tool for column selection with a particular set of analytes. Despite this, many chemometric approaches don't provide any guidance regarding peak shapes of the separated components. In addition, the chemometric models generally require physical or chemical information for the compounds of interest, such as excess molar refraction, dipolarity and polarizability, hydrogen bond donor (acidity), hydrogen bond acceptor ability (basicity) and McGowan's volume (19); for many analytes of interest, information such as this is not readily available. 

Many pharmaceutical laboratories are involved in the characterization and purification of large chemical libraries that are employed in high-throughput screening for new drug development. These libraries are particularly challenging in the chemical diversity that they represent (20,21). Library molecules may contain a variety of different functional groups and may differ substantially in properties such as p

, hydrophobicity, molecular weights and molecular surface area, to name just a few. SFC–mass spectrometry (MS) has been shown to be a useful tool for the purification of chemical libraries (3,20,22–25); however, the selection of the appropriate stationary phase for the separation of chemicals in the library carrying out these separations can be difficult, especially with the variety of SFC stationary phases that have recently been commercialized (14). Furthermore, most chromatographers performing purifications prefer to avoid using mobile phase additives, such as amines, because it complicates compound recovery and can lead to product degradation. This poses a unique challenge for chromatographers performing purifications on a vast array of molecules. Chromatographers separating chemical libraries would like chemical compound selectivity, as well as good peak shapes, to optimize their purifications without the use of mobile phase additives. Ideally, one would like to have a set of screening stationary phases that the chromatographer could employ to separate a wide variety of molecules containing various functional groups. To this end, we would like to identify a limited set of stationary phases that can separate a wide variety of molecules and maintain good peak shape without mobile phase additives. 

Utilization of a Molecular Diversity Model

We used the large and structurally diverse chemical building block library available at Merck (Whitehouse Station, New Jersey, USA) to help identify a set of stationary phases. We queried this collection of chemicals to obtain representative compounds in four distinct functional group classes — carboxylic acids, amines, alcohols and amides. These four distinct functional group classes are viewed as important reactive groups for the synthesis of larger molecules. We identified 15 chemicals within each functional group class with maximal Tanimoto dissimilarity (26) using Merck's in-house developed software. These 15 compounds represent diverse chemical space for each chemical class based upon the combination of the large number of compounds in the building block collection and the computational calculated molecular diversity model. The structure of these proprietary compounds, serving as the test probes in this study, will not be disclosed but an example collection of commercially available amines with structures shown will be displayed later in the article. Figure 1 is an example of a screen shot from the custom written "Small Molecule Browser" software used to identify structurally diverse reagents within each functional group class. This projection is a two-dimensional nonlinear map of the ECFP4-derived Tanimoto similarity matrix (26). In rough terms, and allowing for distortions in the projection, molecularly similar compounds end up grouped together and molecularly different compounds tend to be farther apart. The red dot in the upper right quadrant was randomly selected as a starting point and the software was instructed to find the 14 most diverse carboxylic acids, relative to the starting point. The resulting selections are highlighted in blue.  

Figure 1: Example of a screen shot from the custom-written "Small Molecule Browser" used to identify structurally diverse reagents and analytes within each functional group class. This projection is a two-dimensional nonlinear map of the ECFP4-derived Tanimoto similarity matrix. 

Figure 2 highlights the many calculated parameters used in arranging the carboxylic acid collection into its multidimensional chemical space. We did not elect to use filters restricting the range of certain molecular parameters within a specific range to ensure the most random and diverse collection as possible. The figure displays the extensive range of each parameter used. The same molecular parameters were also used for the amine, alcohol and amide chemical classes and resulted in the selection of our test chemicals for each chemical class.  

Figure 2: A typical screen shot highlighting the large number of molecular parameters, specified as row headers on the left of the image, used to arrange the functional classes into their respective chemical spaces. The vast range of each of the parameters is highlighted by red bars in each of the lines representing an individual molecular parameter in the image. The lighter red bars signify a smaller number of building blocks or analytes contained in the collection possessing a specific magnitude of the molecular parameter specified, and an intense red bar signifies a greater population of analytes possessing that magnitude of the specified parameter. Such tools were used to provide a rational selection of test probes for the SFC columns. 

The Selection of the SFC Conditions and Stationary Phases Evaluated

The 15 compounds for each of the four chemical classes were analysed on seven different achiral stationary phases, many of which were in their endcapped and nonendcapped forms (listed in Table 1). A silica column was included as one of the seven, because this column has been extensively used for achiral SFC separations (13). An endcapped silica column (TMS) was also evaluated, but coverage of the polar surface of the silica virtually eliminated the retentive capabilities of the material, as expected, so it was eliminated early in the study. All of the phases evaluated were bonded to the same 5-μm, 350-m

/g, and 120-Å pore size base silica and the packed columns had dimensions of 250 mm × 4.6 mm. To maximize the interaction with the stationary phase, isocratic conditions were selected (27). As indicated by the column temperature of 25 °C, the study was performed in the subcritical state to minimize fluid compressibility and thus minimize density changes along the column that impact diffusion coefficients (19). All chromatographic data presented and discussed in this study were acquired using the following chromatographic conditions:  

Detection: UV absorbance at 214 nm coupled with positive atmospheric-pressure chemical ionization (APCI) MS for compound identification

Table 1: Columns evaluated (silica and column dimensions are described in the text).    

The 15 carboxylic acids presented by our diversity model were analysed on our column set. Of the columns tested, the 2-ethyl pyridine (EP) and GreenSep Basic columns provided the best results for the carboxylic acid test probes. Both stationary phases produced acceptable peak shape for all of the peaks on the endcapped EP, nonendcapped EP, endcapped basic and nonendcapped GreenSep Basic columns. The half-height peak efficiency values in plates per column for each peak in a specified chromatogram were averaged together to give one average efficiency per column. The values were 7770, 11998, 8489 and 8864 plates per column, respectively. A chromatogram of the carboxylic acid mixture on the endcapped and nonendcapped EP column is shown in Figure 3. The nonendcapped EP column exhibits additional selectivity as evident by the separation of critical pair peaks 10/11 and 3/6, as well as additional retention as evident by increased retention on all peaks (except peak 7). The nonendcapped version of the GreenSep Basic column did not demonstrate a change in selectivity or an increase in retention when compared to its endcapped version, as shown in Figure 4. The absence of increased retention for the nonendcapped GreenSep Basic column compared to its endcapped version suggests the free silanols are not able to compete with the imidazole for interacting and retaining the acidic test probes as experienced with the EP column. This is supported by the increase in p

 of imidazole compared to pyridine, 7.0 to 5.25, respectively. This behaviour was very interesting and requires further investigation. In addition, we discovered that diethyl amino propyl (DEAP) showed excessive retention for the acids (presumably because of the involvement of an ion-exchange retention mechanism).  

Figure 3: Carboxylic acid mix analysed under standard chromatographic conditions on the endcapped (top) and nonendcapped (bottom) 2-ethyl pyridine column. 

The study found that the nonendcapped EP is the column with optimal performance for the analysis of carboxylic acids. Previous studies have reported excessive retention of acidic compounds on the EP column (3). The carboxylic acid test probes used in the present study were largely unaffected by this phenomena, but presumably the use of an endcapped version of the column could be helpful in cases where retention is unacceptably long.  

Figure 4: Carboxylic acid mix analysed under standard chromatographic conditions on the endcapped (top) and nonendcapped (bottom) Basic column.  

The DEAP column provides excellent peak shape for the alcohols, as shown in Figure 5, even for compound 1 eluted at 25 min (see insert). The average half height peak efficiency for the alcohol test mix analysed on the DEAP column was calculated to be 9775 plates per column and the average USP tailing factor was calculated to be 1.19. This is a small but diverse data set for alcohols, and the DEAP column appears to have a vast degree of selectivity for this data set. The extremely efficient peaks may be a result of this column possessing only one retention mechanism, hydrogen bond accepting capabilities. The ideal selectivity, controlled retention and elution, and extremely efficient peaks make the DEAP column the preliminary column of choice for alcohol analysis.  

Figure 5: Alcohol mix analysed under standard chromatographic conditions on the nonendcapped DEAP column. 

All columns tested gave acceptable results for the chemical diverse amide compound test set. The best column for the amide compound set was found to be the nitro phenyl nonendcapped column, as shown in the chromatogram in Figure 6. This column provided excellent peak shape with an average USP tailing factor of 1.34 and good selectivity for the amides analysed. The peaks are very efficient with an average half height efficiency of 9279 plates per column with peak 9 providing good peak shape, with a half height peak efficiency of 9304 plates per column at 20 min.  

Figure 6: Amide mix analysed under standard chromatographic conditions on the nonendcapped nitro column. 

The SFC separation of amines is particularly challenging without the use of additives. The results for the amine test mixture varied greatly. Some of the columns tested produced very poor peak shape, had very poor selectivity and had excessive retention, and of these some exhibited both poor peak shape and poor selectivity. The GreenSep Basic nonendcapped column showed the best peak shape for most of the amines analysed, as shown in Figure 7, with the average USP tailing factor being 1.65 vs. the average tailing factor of 2.31 observed for the amines analysed on the endcapped version of the column. An additive such as diethyl amine could be added to the cosolvent in an attempt to sharpen peak shape and minimize further tailing. The use of additives is out of the scope for this article but Blackwell and Stringham (28) cover this topic more comprehensively. In our amine compound mixture, eight of the compounds are primary amines, suggesting that there would be little hydrogen bond accepting character displayed by these compounds on the GreenSep Basic column. The amine test mixture exhibited longer retention on the nonendcapped GreenSep Basic column, suggesting strong π-π interactions as all but one of the amine test mixture compounds possess aromatic systems. One of the poorest columns for amines was found to be silica with only one of the 15 amine compounds in our mixture eluted within a 60-min chromatographic analysis.  

Figure 7: Amine mix analysed under standard chromatographic conditions on the endcapped (bottom) and nonendcapped (top) Basic column. 

The structures of the Merck compounds used in this study could not be disclosed so a collection of commercially available amines (listed in Table 2) was selected to serve as an operational test for our approach of column assignment. Figure 8 shows a chromatogram of these commercially available amines analysed on the nonendcapped GreenSep Basic column according to the standard conditions described earlier in this article. The half height peak efficiency along with USP tailing factors for all seven of the commercial amine mixture components are shown in Table 2. Several of the components have a tailing factor greater than 2; however, all components were eluted in less than 12.5 min. Even with greater tailing factors, for some components this separation would be useful for preparative chromatography, especially considering that the conditions were isocratic and no additives were used. This mixture provides an operational test for a chemical diversity model. Based on our amine diversity mixture and our commercially available amine mixture, we believe that the nonendcapped GreenSep Basic column is the preliminary column of choice for the analysis of amines.  

Table 2: Commercially available amines used as test probes. 

One of the complexities routinely discussed about achiral SFC revolves around identification of a single stationary phase to resolve the majority of sample mixtures encountered in our laboratories. This desire could stem from the common application of C18 stationary phases in reversed-phase applications and the vast success encountered with its use. With the study presented in this column instalment, we hope to limit the experimental work necessary for column screening and data interpretation — there does not appear to be a current universal stationary phase for achiral SFC applications. Figure 9 displays the data generated for all four chemical classes on the nonendcapped nitro phenyl column in this study. The highly variable results for the four classes on this single column summarize the need for development of a universal phase for achiral SFC. The amide chromatogram displays controlled retention, controlled elution, efficient peaks and a vast degree of selectivity, making this phase the preferred suggestion for the analysis of amides. The carboxylic acid chromatogram reveals broad, inefficient peaks. This class of compounds also displays excessive retention as evident from the peaks eluted at 50–60 min and even several compounds that were not eluted from the column. The alcohol chromatogram displays the worst peak shape of all four classes of compounds on this column. Several broad peaks are observed in the UV data and alcohols 12 and 13 had to be extracted from the mass spectrometer signal to be visible. The amine chromatogram is self explanatory in the fact that this class of compounds is too strongly retained.  

Figure 8: Commercially available amines analysed under standard chromatographic conditions on the nonendcapped Basic column according to the standard chromatographic conditions used in this study. 

A library of test analytes with a broad range of molecular properties was used to characterize the suitability of several investigational stationary phases as general tools for SFC separation. Although no single column was identified with broad utility across the entire set of compounds, amides generally produced the most consistent peak shape across all stationary phases evaluated and amines tended to have the worst peak shape. The amines were also slightly less retentive, but frequently yielded inefficient peak shapes with severe tailing (for example, in the case of silica). Carboxylic acids were on average the most strongly retained class of compounds on most of the columns evaluated. An estimated, normalized retention order would be carboxylic acids < amines < alcohols < amides with the carboxylic acids being the most strongly retained. From the endcapping study, it appears the silanol groups on the nonendcapped stationary phases play a role in maintaining the quality of separations and demonstrates that silanols do play a significant role in the separation mechanisms in SFC.  

Figure 9: Example of a single phase not being universal. All four data files were generated using standard chromatographic conditions on the nonendcapped nitro column. Top: amide mix; second from top: carboxylic acid mix; third from top: alcohol mix; and bottom: amines. 

We developed the colour-coded chart shown in Figure 10 to summarize our findings. This chart is based upon the four chemical classes and the columns studied. In the chart, we identified the "best" column for each chemical class by taking into account selectivity, retention and peak shape. These "best" columns can be used as a first line for separating chemical in the four chemical classes:  

Ethyl pyridine (nonendcapped) for carboxylic acids

Figure 10: Summary table of recommended stationary phases based on analyte structural classes.  

In the future, more chemical classes of compounds will be subjected to these stationary phases to enable matching of additional chemical classes with its appropriate stationary phase. Ultimately, the approach described in this study will lead to the development of new stationary phases for SFC and may lead to the development of a universal SFC stationary phase.  

Column Watch

The authors characterized 12 SFC achiral stationary phases with 60 compounds from four chemical classes to establish guidelines for th rational selection of the optimum stationary phase for separations.

A Systematic Study of Achiral Stationary Phases Using Analytes with a Molecular Diversity Model

Some promising reversed-phase widepore stationary phases (organic monolithic, fully porous and coreshell type) were recently introduced for the fast and high resolution separations of large biomolecules. It was demonstrated that kinetic performance, retention capability, selectivity for closely related proteins and loading capacity, as well as recoveries, were satisfactory with recent core-shell and sub-2 μm fully porous materials. This article focuses on the achievable throughput and resolution for the characterization of biomolecules. As shown, fast separation of a therapeutic protein (~19 kDa) and some of its degradants (related proteins) was achieved in less than 1.5 min with a 50-mm column length while a peak capacity of more than 150 was obtained in about 15 min using two columns of 150 mm coupled in series. Similar results were also attained with intact therapeutic monoclonal antibodies (mAbs) of ~150 kDa and their fragments by varying the column length and gradient time. However, the recovery of intact mAbs was found to be unacceptable in reversed-phase liquid chromatography (RPLC) except when using elevated mobile phase temperature (80–90 °C).

The importance of therapeutic proteins and monoclonal antibodies is growing in the pharmaceutical field and expectations are high for this new class of compounds (1,2). Indeed, it has been established that such large biomolecules could present some significant benefits for the patient such as high efficacy, specificity, wide therapeutic range and limited side effects. 

All biopharmaceuticals are inherently variable as they are produced from living organisms. This variability exists within batches, from batch to batch, and when production processes are improved or changed between manufacturers. The variability of biopharmaceuticals is greater than that typically observed for conventional pharmaceuticals and applies to originator reference products as well as biosimilars. There is consequently a need to develop analytical tools that are able to characterize such complex molecules (3,4). The analytical step is quite difficult and is based on the combination of various analytical strategies to determine identity, impurity content, heterogeneity and activity before release. Even if capillary electrophoresis and biophysical techniques (for example, circular dichroism, light scattering and fluorescence spectrophotometry) are considered as reference methods, liquid chromatography (LC) is also widely applied in analytical laboratories (5,6). Today, the analyst has the choice between (i) historical techniques, such as size-exclusion chromatography (SEC) and ionexchange chromatography (IEC) that provide suitable selectivity for charge variants and aggregates, respectively, and (ii) reversed-phase liquid chromatography (RPLC) that produces lower selectivity between closely related proteins but compensated by the sharpness of the peak because of the fast kinetic interactions in RPLC. In addition, RPLC is more directly compatible with electrospray ionization mass spectrometry (ESI–MS) detection than SEC or IEC (7).  

Recently, some noteworthy improvements were brought to conventional RPLC columns to improve the performance achieved with large proteins. This includes the commercialization of various types of widepore phases such as organic monoliths with macropore sizes of 1–5 μm, core-shell particles of 2.7–5 μm and fully porous sub-2 μm UHPLC columns, which have been described in the literature. (8). In this article, two recent technologies of widepore RPLC columns were evaluated and compared for the fast and high resolution analysis of therapeutic proteins and mAbs (9,10).  

 Acetonitrile and methanol (gradient grade) were purchased from SigmaAldrich (Buchs, Switzerland). Water was obtained with a Milli-Q Purification system from Millipore (Bedford, Massachusetts, USA). Recombinant human granulocytecolony stimulating factor (G-CSF or filgrastim, MW~18.8 kDa) was obtained from Amgen (Zug, Switzerland). IgG monoclonal antibodies including rituximab (mabThera) and bevacizumab (avastin) were purchased from Roche (Roche Pharma, Basel, Switzerland). For fragmenting the monoclonal antibody, dithiothreitol (DTT) and papain (from Carica papaya) were obtained from Sigma-Aldrich (Buchs, Switzerland). Trifluoroacetic acid (TFA), 30% hydrogenperoxide and methionine were purchased from Sigma-Aldrich (Buchs, Switzerland). 

Acquity BEH300 C18 and C4 columns with a particle size of 1.7 μm (50 × 2.1 mm and 150 × 2.1 mm, 1.7 μm, 300 Å) were purchased from Waters (Milford, Massachusetts, USA). Aeris Widepore (WP) C18 columns packed with 3.6 μm core-shell particles (150 × 2.1 mm, 3.6 μm) were supplied by Phenomenex Inc. (Torrance, California, USA). The Acquity BEH300 columns are packed with 1.7 μm fully porous particles with an average pore size of ~300 A (11,12), while the Aeris WP is based on core-shell technology and its particles consist of a ~3.2 μm solid inner core surrounded by a ~0.2 μm porous layer (total particle size of 3.6 μm).  

In this study, two columns of 150 × 2.1 mm BEH300 were coupled in series to generate a 30-cm long column, while the 45-cm column length was prepared by coupling two columns of 150 × 2.1 mm BEH300 and one Aeris WP 150 × 2.1 mm columns in series (this was due to pressure considerations).  

All measurements were performed using an Acquity UPLC (Waters) system equipped with a binary solvent delivery pump, an autosampler and UV detector and/or fluorescence detector (FL). The Acquity system includes a 5 μL sample loop, a 0.5-μL UV flow-cell and a 2 μL FL flow-cell. The loop is directly connected to the injection switching valve (no needle seat capillary). The connection tube between the injector and column inlet was 0.13 mm i.d. and 250 mm long (passive preheating included), and the capillary located between the column and detector was 0.10 mm i.d. and 150 mm long. Data acquisition and instrument control was performed by Empower Pro 2 Software (Waters, Milford, Massachusetts, USA).  

Scanning electron microscopy (SEM) was performed using a JEOL JSM 5500LV instrument (Budapest, Hungary). 

 For the gradient separation of related proteins and mAbs fragments, the mobile phase "A" consisted of 0.1% TFA in water, while the mobile phase "B" was 0.1% TFA in acetonitrile. The chromatographic conditions were optimized by a systematic variation of the gradient span and steepness, and of the column temperature and flow rate. 

The intact mAbs were injected (0.5 μL) without dilution directly from the concentrated commercial solutions (10 mg/mL rituximab and 25 mg/mL bevacizumab). 

The intact filgrastim was directly injected as purchased (Neupogen concentrated solution 0.3 mg/mL). Reduced and oxidized protein samples were prepared. The intramolecular disulphide bonds of filgrastim were reduced by adding a small amount of dithiotreitol (DTT) to its solution and incubating at 30 ºC for 60 min. The oxidation was performed by adding 2% (v/v) hydrogen peroxide (30%) to the protein solution. After a 3 h incubation time (at 30 °C), the oxidation was stopped by adding 0.5 mg methionine to the solution. 

The IgG monoclonal antibodies possess intramolecular disulphide bonds which were also reduced with DTT. Experimentally, 0.05 mg of DTT was added to 100 μL mAb concentrated solution followed by an incubation at 30 ºC for 60 min. The proteins were completely converted to the light chain and heavy chain components of the antibodies.  

Papain is a thiol protease that cleaves IgG antibodies at the heavy chain hinge region into three fragments, one fraction crystallizable (Fc) and two identical fraction antigen-binding (Fab) fragments. The digestion of rituximab and bevacizumab was initiated by the addition of papain (diluted to 100 μg/mL with water) to give a final protein:enzyme ratio of 100:1 (m/m%). The final digestion volume was 200 μL and was directly injected with the chromatographic system using low volume insert vials. The digestion was carried out at 40 °C for 3 h.  

Scanning electron microscopic (SEM) images and Image-J (image-processing software) were used to show the roughness and sphericity of the particles and to determine the particle size distribution (PSD) of the different materials. 

 When dealing with therapeutic proteins, the RPLC approach is usually employed to determine the amount of impurities as well as degradants (for example, oxidized, deamidated or reduced forms) within the sample. In practice, it generally means that proteins that have very similar molecular weights and nearly identical structures (conformations) must be separated. The difference in molecular structure is relatively low; therefore, similar retention behaviours of the different forms are expected with RP phases. Selectivity is often very poor in RPLC, thus there is a need to work with experimental conditions that provide elevated kinetic performance. In this context, the last generation of RPLC columns packed with core-shell or sub-2 μm fully porous particles should be used preferentially, and elevated temperature could be a key tool because it improves the mass transfer process of large analytes as well as the secondary ionic interactions kinetics. 

Preliminary evaluation of recent widepore RPLC column technology:

 In a recent series of papers (9,10), the performance and possibilities of both widepore coreshell and fully porous sub-2 μm particles columns were investigated in terms of kinetic performance, selectivity, retention behaviour and loading capacity for intact proteins with molecular weight comprised between 5.8 and 66.4 kDa. 

In the first instance, these materials were characterized by their physical and physico-chemical properties (for example, PSD, pore volume, surface coverage, silanol activity, ligand densitiy, loadability and retention capability) (10). Figure 1 shows a SEM representation of the widepore core-shell and fully porous sub-2 μm particles and illustrates the porous layer of ~0.2 μm at the surface of the core-shell particle. The particle sizes were precisely determined as 3.6 μm (particle size distribution dp90/10 = 1.11) for the widepore core-shell material and 2.0 μm (particle size distribution dp90/10 = 1.44), instead of the 1.7 μm announced by the provider, for the fully porous sub-2 μm particles. Superficially porous particles generally displaying a very narrow PSD can lead to unprecedented low minimal reduced plate heights. It is, however, unclear whether these findings can be directly related as there are other factors that might influence the packing quality. Superficially porous particles have a higher density and some of them are rougher than fully porous particles. This might also have had an influence on the achieved packing quality and homogeneity. Two strategies were applied to evaluate the kinetic performance of these stationary phases in the gradient mode. The first one consists of calculating the peak capacities for various gradient times and for two constant flow rates. The second one maintains a similar gradient steepness for all flow rates and provides gradient kinetic plot representations based on a method proposed by Broeckhoven 

 (13). These experiments showed that both stationary phase technologies provide excellent performance with small and large proteins. In addition, these phases significantly outperform some medium pore-size materials or other reference HPLC phases of 300 A pore size. This is one of the main reasons why these two recent stationary phase technologies were selected for the characterization of therapeutic proteins and mAbs in this study. 

Figure 1: Scanning electron microscope (SEM) images of (a) BEH300 C18, (b) Aeris WP C18 and (c) a cut Aeris WP C18 particle. 

In addition, these two phases provide a very similar retention capability and a reasonable loading capacity for proteins because of the low porous volume on the silica-based widepore core-shell material (around 30% of the total particle volume). This provides a rather limited specific area and is probably compensated by a significant contribution of secondary (ion-exchange and H-bonding) mechanisms, compared to the hybrid type fully porous sub-2 μm material (10).  

Application to the fast and high resolution analysis of therapeutic proteins

: Figure 2 illustrates the analysis of native filgrastim (~18.8 kDa) and its related proteins (that is, oxidized and reduced forms). This type of experiment is useful in the biopharmaceutical field, to evaluate the stability and purity of therapeutic protein samples. As shown in Figure 2(a), the baseline resolution of these different compounds can be obtained in less than 1.4 min using a 50-mm column length, at a reasonable flow rate. The peak capacity was also acceptable and this application demonstrates that high throughput analysis can be carried out even with large biomolecules. On the other hand, because of the sample complexity and the need to also separate some protein isoforms or variants, it is also mandatory to achieve high resolution separations. The separation was therefore performed with a 300mm column length at a flow rate of only 0.25mL/min. Compared with the experiment illustrated in Figure 2(a), the column length was extended by 6-fold and flow rate reduced by 2×, to provide reasonable back pressure. Then, to maintain the same gradient steepness between Figure 2(a) and 2(b), the gradient time was 12× longer. The resolving power was drastically improved with this new configuration and peak capacity was increased above 150 for an analysis time of less than 15 min. It was also possible to observe a few additional peaks of weak intensity, eluted closely to the peaks of proteins that should correspond to protein isoforms. 

Figure 2: Fast and high-resolution separation of filgrastim related proteins. Column: Acquity BEH300 C18 2.1 mm, 1.7 Î¼m (above: 50 mm long and below: 150 mm + 150 mm = 300 mm). Mobile phase A: 0.1% TFA in water, mobile phase B: 0.1% TFA in acetonitrile. Gradient: 47â62% B, Temperature: 80 Â°C, Fluorescence detection: 280â350 nm. Injected volume: 1 Î¼L. Peaks: 1,2: oxidized filgrastim, 3: filgrastim, 4: reduced filgrastim. 

 There are different ways of characterizing therapeutic monoclonal antibodies using RPLC. The dominant method is to completely digest these large glycoproteins of ~150 kDa, using an enzyme such as trypsin (peptide mapping approach or bottom up). The analysis is then carried out on peptides of limited size (0.5–3 kDa), but the chromatogram remains complex (because of its size, the mAb is digested into a significant number of peptides) and the digestion protocol is long and tedious. An alternative approach has been developed in the last few years and involves performing limited proteolysis (LP approach). mAb is digested with enzymes such as papain to obtain large fragments of ~50 kDa (Fc and Fab). Alternatively, pepsin, lys-C or Fabricator [a modified cysteine protease (14)] can also be employed as enzymes to obtain different fragments. Finally, the mAb can be analysed under its intact form, to determine the impurity content of the produced mAb and under its reduced form to obtain two heavy chain fragments of ~50 kDa and two light chain fragments of ~25 kDa and determine the integrity of mAb under denaturing conditions. 

Adsorption of mAbs under RPLC conditions:

 One major issue when dealing with mAb analysis by RPLC is the adsorption phenomena. Indeed, we have recently demonstrated that adsorption of mAbs onto the stationary phase was significantly higher compared to proteins of equivalent size (15). In addition, the adsorption rate was particularly relevant with intact mAbs of 150 kDa. It decreases significantly with mAbs fragments of 50–100 kDa and becomes almost negligible with small fragments of only 25 kDa. Our previous findings also showed that the adsorption phenomenon was mostly due to secondary interactions and only to a limited extent to the pure hydrophobic interaction mechanism that occurs in RPLC. For this reason, there is no significant difference in recovery between a C4 and a C18 bonding, while there are some important differences depending on the silica matrix technology (hybrid or bare silica). To decrease the amount of secondary ionic interactions and improve recoveries of mAbs, it is necessary to increase mobile phase temperature up to 80–90 °C.  

This behaviour is illustrated in Figure 3 which shows some chromatograms of intact bevacizumab [Figure 3(a)] and reduced bevacizumab [Figure 3(b)] at various mobile phase temperatures between 40 and 80 °C (the maximum temperature that the column can withstand). As shown, the adsorption was obvious with the intact mAb and no peak was observed for temperatures below 50 °C, while the recovery was acceptable at 70–80 °C. In the case of heavy and light chains of 50 and 25 kDa, respectively, the adsorption was less pronounced and this was particularly true with the light chain [first eluted peak in Figure 3(b)]. Even if elevated mobile phase temperature is a good solution to overcome adsorption, this parameter should be used with caution as it could practically denaturate large biomolecules. Thus, a compromise has to be found between temperature and residence time. According to our previous study (15), an analysis time of less than 15 min and temperature in the range 70–90 °C was a good compromise between recovery and stability of columns and analytes. 

Figure 3: Representative chromatograms on the recovery of the intact, light and heavy chain of Bevacizumab. Column: BEH300 C4 (150 mm Ã 2.1 mm). Mobile phase A: 0.1% TFA in water, mobile phase B: 0.1% TFA in acetonitrile. Gradient: from 31% to 40% B in 6 min, flow rate: 0.3 mL/min. Temperature: from 40 ÂºC up to 80 ÂºC, injected volume: 0.5 Î¼L, Fluorescence detection: 280â360 nm. Peak 1: light chain, peak 2: heavy chain. 

Application to the fast and high resolution analysis of therapeutic mAbs:

 Despite the size of therapeutic mAbs and adsorption phenomena that could occur, it is possible to analyse intact mAbs and obtain sharp peaks using recent widepore stationary phases. Figure 4 demonstrates the possibility to perform some ultra-fast separations of mAbs and shows the analysis of intact and papain-digested commercial solution of bevacizumab. For this purpose, a column of 50-mm length was employed and gradient time was reduced down to only 1.6 min. Under these conditions, the peak shape remains acceptable for the intact mAb and it was also possible to separate the Fc and Fab fragments [Figure 4(b)]. However, the resolving power in these conditions was too low to adequately analyse a mAb sample. 

Figure 4: Fast separation of some variants of intact Bevacizumab and the Fc and Fab fragments (from papain digested sample). Column: Acquity BEH300 C4 50 mm Ã 2.1 mm, 1.7 Î¼m. Mobile phase A: 0.1% TFA in water, mobile phase B: 0.1% TFA in acetonitrile. Gradient: 31â38% B in 1.6 min, flow rate: 0.5 mL/min. Temperature: 80 Â°C, Fluorescence detection: 280â360 nm. Injected volume: 1 Î¼L. Peaks: 1: intact Bevacizumab, 2: Fc of Bevacizumab, 3: Fab of Bevacizumab. 

In this context, longer columns and gradient times should be used when dealing with therapeutic mAb samples. Figure 5 shows the analysis of intact rituximab using various column lengths and gradient times at a temperature of 80 °C. In Figure 5(a), a C18 column of 150 mm was selected. The gradient span was limited (31–39% ACN — this is always the case in mAbs analysis), the analysis time remained as short as ~5 min. In this case, the peak capacity was quite low (n

 of 55) but it is possible to significantly increase the performance by extending column length to 300 or 450 mm. With the 450-mm column length configuration, the analysis time was still acceptable, around 15 min, but peak capacity was 2-fold higher compared with the experiments on the 150mm column length. As shown in Figure 5(c), various charge variants can be separated around the peak of rituximab, but the resolution was still not sufficient to adequately characterize the commercial rituximab sample. 

Figure 5: RP-HPLC separation of intact Rituximab variants. Columns: Acquity BEH300 C18 150 mm Ã 2.1 mm, 1.7 Î¼m and Aeris WP C18 150 mm Ã 2.1 mm. Mobile phase A: 0.1% TFA in water, mobile phase B: 0.1% TFA in acetonitrile. 3.6 Î¼m. Gradient: 31â39% B, flow: 0.35 mL/min. Temperature: 80 Â°C, Fluorescence detection: 280â360 nm. Injected volume: 1 Î¼L. 

To better visualize the heterogeneity in the peptide chains of the rituximab sample and to determine the amount of isoforms, mAb was reduced with DTT and the produced fragments (light and heavy chains) were analysed using various column configurations. The results are presented in Figure 6. Compared with Figure 5, the peak capacity values were improved by about 20%, probably because the size of the heavy and light chains was more reasonable (~25–50 kDa) than that of intact rituximab (~150 kDa), leading to better kinetic performance. The chromatograms of Figure 6 show two major peaks which corresponds to the light chain fragment eluted first as a sharp peak, and the heavy chain fragment eluted later as a more heterogeneous peak with slightly resolved isoforms. Again, the resolving power of a 150-mm column length packed with 1.7 μm was limited (n

 of 68). On the other hand, when extending the column length to 450 mm and adjusting gradient time to maintain an equivalent gradient steepness [Figure 6(c)], the quality of the separation was significantly improved (n

Figure 6: RP-HPLC separation of the light chain and the heavy chain variants of Rituximab. Columns: Acquity BEH300 C18 150 mm Ã 2.1 mm, 1.7 Î¼m and Aeris WP C18 150 mm Ã 2.1 mm, 3.6 Î¼m. Mobile phase A: 0.1% TFA in water, mobile phase B: 0.1% TFA in acetonitrile. Gradient: 31-39 %B, flow: 0.35 mL/min. Temperature: 80Â°C, Fluorescence detection: 280â360 nm. Injected volume: 1 Î¼L. Peaks: 1: light chain, 2: heavy chain. 

As demonstrated in this article, despite the large molecular mass of recombinant biomolecules, it is possible to attain sharp peaks in RPLC, thanks to the recent advances in column technology, such as the introduction of widepore fully porous sub-2 μm and widepore core-shell particles.  

Depending on the goal of the method, ultra-fast (less than 2 min for a 50-mm column length) or high resolution analysis (peak capacity higher than 100 for a column of 300–450 mm) were demonstrated with these two columns for therapeutic proteins as well as recombinant mAbs. Except for the quality of the chromatographic columns, it is clear that mobile phase temperature is a key parameter for improving kinetic performance as well as limiting adsorption phenomena with mAbs. As a rule of thumb, a temperature of 70–90 °C is always useful but should be used only in combination with reasonable analysis times (up to 20 min) to avoid thermal degradation of proteins. 

The authors wish to thank Dr Tivadar Farkas (Phenomenex Inc.) for providing the Aeris Widepore columns and Róbert Berky and Prof Jenö Fekete (Budapest University of Technology and Economics, Department of Inorganic and Analytical Chemistry) for the SEM measurements. 

 holds a PhD degree in analytical chemistry from the University of Lyon, France. Since 2004, he is a lecturer in the laboratory of pharmaceutical analytical chemistry of the University of Geneva in Switzerland, under the direction of Prof Jean-Luc Veuthey. Guillarme is co-author of more than 60 scientific papers related to pharmaceutical analysis. His expertise includes liquid chromatography, UHPLC, HILIC, LC–MS, analysis of proteins and mAbs and more recently supercritical fluid chromatography. 

 gained his PhD in analytical chemistry from the group of Prof Jeno Fekete at the Budapest University of Technology, Hungary. He has worked for over 10 years in the pharmaceutical industry in the analytical R&D field. He currently works at the University of Geneva in Switzerland under the supervision of Prof Jean-Luc Veuthey and Dr Davy Guillarme. Fekete is co-author of 30 scientific papers on liquid chromatography and pharmaceutical analysis. His main interests lay in column technology, mass transfer processes, method development, pharmaceutical and protein analysis. 

 A. Staub, D. Guillarme, J. Schappler, J.L. Veuthey and S. Rudaz, 

 Guideline on bioanalytical method validation EMEA/CHMP/EWP/192217/2009, 

http://www.ema.europa.eu/docs/en_GB/document_library/Scientific_guideline/2011/08/WC500109686.pdf

 S. Eeltink, B. Wouters, G. Desmet, M. Ursem, D. Blinco, G.D. Kemp and A. Treumann, 

Two promising wide-pore stationary phases were recently introduced for the fast and high resolution separations of large biomoleculaes: the recently launched widepore 3.6?m superficially porous particle and the hybrid-type sub 2-?m fully porous particle. This article focuses on the achievable throughout and resolution for the characterization of biomolecules.

Reversed-Phase Liquid Chromatography for the Analysis of Therapeutic Proteins and Recombinant Monoclonal Antibodies.

In this month's instalment John Hinshaw discusses solid-phase microextraction.

Gas chromatography (GC) resolution and sensitivity are often limited by matrix effects in "real world" samples that originate outside of a laboratory. GC separation and detection may encounter interference from nonvolatile constituents, side-effects of large sample volumes and the chemical activity of matrix compounds and derivatization residues. A number of sample preparation techniques increase analyte concentrations and detector response while reducing matrix effects, including classical liquid–liquid extraction, chemical derivatization and sample preconcentration, as well as headspace, thermal desorption and large-volume sample injection.  

Solid-phase microextraction (SPME) is a relatively new sample extraction technique — first described in the 1990s by Pawliszyn (1,2) — that brings some unique capabilities to bear on the chromatographic analysis of dilute solutions in difficult matrices, both liquid and gaseous. Essentially, SPME consists of two discrete steps: solute absorption from the sample matrix into a thick layer of silicone or related adsorptive material, followed by transfer of the absorbed analytes into a chromatography inlet system by thermal or liquid desorption.  

SPME has been applied to both GC and liquid chromatography (LC) separations It eliminates the need for large-volume sample transfer into a GC column by concentrating analytes into the fibre coating while leaving the bulk of the solvent and nonvolatile residues behind. SPME uses orders of magnitude less solvent and has significant potential to greatly reduce or eliminate solvent consumption and the concomitant issues of used solvent disposal as part of sample preparation. 

Chromatographers should not confuse SPME with solid-phase extraction (SPE), a related predecessor with similar applications. The principal difference is that SPE is carried out with a relatively large sorptive wafer the size of a small filter paper and requires post-sorption liquid-phase extraction of analytes; SPME is accomplished with a small fibre or tube coated with sorptive material. SPME is applied both to gas-phase and liquid-phase extraction, whereas SPE is limited to extraction from liquid-phase samples. This 

 instalment primarily discusses SPME using absorptive polymeric coatings, although the principles also apply to adsorptive SPME onto active solid layers. 

A related technique, stir-bar sorptive extraction (SBSE), uses a magnetic stir bar coated with a thick layer of absorptive polymer. The stir bar is exposed to sample solution for a time, during which solutes are absorbed into the polymer coating. Subsequently the stir bar is removed, dried and then thermally desorbed for GC injection, or the absorbed analytes can be back-extracted with a different solvent. SBSE uses a larger volume of absorbent than SPME and, thus, is more efficient at extracting analytes with less absorbent solubility. Therefore, SBSE is generally more sensitive than SPME.  

SPME applications cover a broad range that includes flower scents (3), chemical warfare agents (4), pharmaceutical process impurities (5), the determination of organochlorine pesticides in Chinese teas (6), volatile compounds in acidic media (7) and cheese (8), volatile phenols in wine (9), environmental pollutants in water samples (10), chloroanisoles in cork stoppers (11), volatile aliphatic amines in air (12) and phenylurea herbicides in aqueous samples (13). 

SPME relies on the extraction of solutes from a sample into an absorptive layer on the SPME fibre. After a sampling period — during which extraction ideally reaches equilibrium — the SPME fibre and captured solutes within its absorptive layer are transferred into an inlet system that then desorbs the solutes into a gas (for GC) or liquid (for LC) mobile phase. Success relies on choosing conditions so that the desired solutes favour the SPME absorptive layer as much as possible in the presence of bulk sample, and then subsequently the absorbed solutes are released as quickly and completely as possible for chromatographic analysis. Secondary trapping and release of desorbed solutes after SPME is sometimes required when the initial desorption is too slow and the resulting broadened peaks do not yield full utilization of column resolving power. This may be accomplished with a discrete thermal trap or with column stationary phase trapping by injecting onto a cold column and subsequently temperature programming for solute elution. It is also possible to make multiple SPME extractions and desorptions into a secondary trap for enhanced sensitivity.  

Figure 1: Cross-sectional diagram of SPME extraction devices. Shown are (a) a fibre device with external sorptive coating and (b) a tube device with sorptive coating on the inside.  

For bulk samples contained in vials or otherwise easily accessed, SPME is performed conveniently with a short absorptive film–coated fibre, as shown in Figure 1(a). A short tube coated on the inside with an absorptive layer [Figure 1(b)] may be used as well for samples that are amenable to pumping. The choice of the absorptive layer chemistry and film thickness strongly influences the degree of absorption and the subsequent efficiency of desorption. SPME fibres are available in syringe-integrated assemblies that are conveniently handled either manually or by robotic autosampling systems. Used fibres often can be cleaned up for reuse by solvent rinsing or baking.  

Figure 2: SPME extraction from a sealed vial. Shown are diagrams representing sampling (a) from the liquid phase and (b) from the headspace (gas phase). 

For the extraction step with an externally coated SPME absorptive layer, the layer is exposed to sample in the liquid [Figure 2(a)] or gas [Figure 2(b)] phase. In the case of SPME within a coated tube, liquid or gaseous sample is circulated through the tube until equilibrium is reached. Stopped-flow flow-through sampling is also possible. Over time, the amounts of the solutes in the SPME layer reach an equilibrium level with their surroundings, which represents the maximum solute amounts that can be absorbed and withdrawn under a given set of sampling conditions. The amount of solute in the SPME layer at equilibrium (

) can be approximated by the following equation: 

 is an aggregate solute distribution constant between the SPME absorptive layer and the sample, 

is the solute concentration in the sample before performing SPME sampling. 

Equation 1 assumes that the sample volume is much greater than the volume of the SPME layer. SPME coatings typically have thicknesses of about 10–100 μm — around 10 times the film thickness range normally encountered in capillary GC columns. The volume, 

, of a 1-cm long by 100-μm thick annular coating on a 0.56-mm o.d. (24-gauge) fibre [as shown in Figure 1(a)] is approximately 2 μL: 

This is an update to a 2003 article with information about new developments in SPME.

LCGC Asia Pacific-11-02-2012

Capillary Electrophoresis-Mass Spectometry for the Analysis of Biopharmaceuticals

The Role of the Injection Solvent

A Systematic Study of Achiral Stationary Phases Using Analytes with a Molecular Diversity Model

Reversed-Phase Liquid Chromatography for the Analysis of Therapeutic Proteins and Recombinant Monoclonal Antibodies.

Solid-Phase Microextraction (SPME)

Application Note: Molecular Weight Determination of Low-Molecular-Weight Heparins: SEC/MALS vs. SEC/UV-RI

Vol 15 No 4 LCGC Asia Pacific November 2012 Regular Issue PDF