The stationary phases currently in use for modern thin-layer chromatography (TLC) are reviewed here, together with traditional and newer detection techniques. Advantages of the planar, off-line format of TLC relative to on-line column high performance liquid chromatography (HPLC), the steps in TLC analyses, and future prospects of TLC are also discussed. A limited number of applications are presented throughout the article, and many others can be found in the books, book chapters, and review articles cited.

This review discusses the current status of thin-layer chromatography (TLC), focusing mainly on commercial precoated layers that are in popular use. Bulk sorbents, for those who prefer to make their own TLC plates, are available but will not be covered.  

TLC is an increasingly widely used analytical method for the determination of many types of analytes in a variety of sample matrices. Its theory, practice, and applications have been documented in biennial reviews written by Sherma since 1970, the latest of which will be published in the 

 (2) gave the three largest industrial applications for TLC as clinical, pharmaceutical, and food testing. The most active research areas today are in the analysis of pharmaceutical bulk materials and dosage forms (3,4) and in natural phytochemical medicines (for example, traditional Chinese medicines and nutritional supplements [5,6]). 

The techniques and instruments of modern, instrumental TLC allow it to meet the guidelines for validated analytical methods in current good laboratory practice (cGLP) and current good manufacturing practice (cGMP); pharmacopeias of China, India, and the United States (

); international food regulations; official methods of AOAC International; International Conference on Harmonization (ICH) pharmaceutical analyses (7); and international food regulations (8,9). Here, we cover some special attributes of TLC relative to column high performance liquid chromatography (HPLC), briefly describe the steps of TLC analyses, focus on the stationary phases (layers) in use today, and speculate on prospects for TLC in the future.  

The use of a planar stationary phase rather than a column leads to a number of significant contrasts between TLC and HPLC. HPLC is a closed, on-line method with dynamic detection of eluted solutes, usually by UV absorption or mass spectrometry (MS). The predominant mode of HPLC is reversed phase on a C18 bonded silica gel column, and the predominant mode of TLC is normal phase on a silica gel layer; these diverse mechanisms allow TLC and HPLC to serve as methods with complementary selectivity for separations and analyte identification. An HPLC column is used for many analyses, but each thin-layer plate is used only once. This often leads to the need for less sample preparation (cleanup) for TLC compared to HPLC, because impure samples can be applied without concern for carryover and cross-contamination of samples that leads to additional (ghost) peaks in column chromatograms. 

Although it is not yet fully automated like HPLC, many of the steps in the TLC procedures are automated for faster sample analyses. TLC has the advantage of very high sample throughput because multiple standards and samples can be applied to a single plate and separated at the same time under essentially identical conditions. Each track on a TLC plate contains a complete chromatogram of an entire sample, including irreversibly sorbed substances (at the origin). On the other hand, in HPLC each chromatogram requires a separate injection and the chromatogram contains only those sample components that happen to be eluted and detected. Modern computer controlled scanning densitometers and automated sample application and layer development instruments allow validated quantification that is in many cases equivalent to HPLC (10). As shown later, TLC has the widest choice of stationary phases and mobile phases, making it the most versatile and flexible LC method. Solvents that can interfere with HPLC UV detection can be used because the TLC mobile phase is removed before detection. Mobile-phase consumption is low, minimizing solvent purchase and disposal costs, thereby making TLC a "green" method. The wide choice of development methods and pre- and postchromatographic detection methods leads to unsurpassed selectivity in TLC. Because TLC is an off-line method, the various steps can be carried out independently without time constraints; as an example of an advantage of this approach, zones can be scanned repeatedly with a densitometer using different parameters that are optimum for individual sample components. In addition, a TLC plate can be stored and reinvestigated at a later date if additional questions arise. 

Excellent discussions of sample preparation; sample application; mobile-phase selection; chromatogram development and documentation; and zone detection, identification, and quantification in modern TLC are presented in two books (11,12), two book chapters (13,14), and a review of TLC in drug analysis (15). This section contains brief discussions of some of these steps of the TLC analytical process, followed by a more detailed description of stationary phases. 

The most widely used methods for application of standard and sample solutions to a plate are manual with a capillary (for example, 10- or 25-µL Drummond digital microdispenser) and automated instrumental application. Application of narrow, homogeneous bands of controlled length has been shown to give superior chromatographic results compared to spot application. These automated application devices are available from Camag and Desaga. 

Mobile phases are less polar than the silica gel layer (normal-phase TLC) and are usually composed of nonpolar and polar constituents with or without an acid or base modifier to improve resolution. In reversed-phase TLC, the stationary phase is less polar than the mobile phase, which often is a combination of methanol, acetonitrile, or tetrahydrofuran with water. 

The mobile phase is optimized in terms of its strength and selectivity in relation to the sample and stationary phase. Snyder classified solvents into eight groups within a selectivity triangle based on proton acceptor, proton donor, and dipole properties (13). He later reviewed and expanded his studies of the role of the mobile phase in controlling selectivity for adsorption TLC and suggested a seven-mobile-phase experimental design for optimizing relative retention and selectivity with binary solvent mobile phases (16). 

Plates are most often developed in a conventional glass large volume chamber (N-chamber) in the ascending direction. Also available are twin trough chambers (TTCs), which allow the development of plates with a low volume of mobile phase and experimentation with chamber saturation conditions, and an automatic developing chamber (ADC 2) that provides complete control of all aspects of plate development (both from Camag). 

Horizontal development chambers are also available from Chromdes and Camag (HDC 2). An advantage of these chambers is the ability to develop the plate from both opposing sides towards the middle, permitting the number of samples to be doubled to achieve even greater sample throughput. 

Methods used to increase the resolution of complex mixtures include two-dimensional (2D) TLC, in which development of a single sample applied to the corner of a single-layer plate is performed in perpendicular directions, with drying in between, using two different mobile phases of about the same strength but complementary selectivities. A dual-layer plate (Multi-K, Whatman/GE Healthcare) that allows both reversed-phase and normal-phase development is also available. The use of either of these can spread sample components over the entire plate with high zone capacity. Figure 1 illustrates these two possibilities for 2D TLC. 

Figure 1: 2D TLC on a single plate. Top TLC plate: a single layer (silica, C18, or cellulose) developed in two directions with different mobile phases, 90Â° to one another, with drying between developments. Bottom TLC plate: a dual layer (C18 and silica, adjacent to one another) developed in two different modes to improve separation of complex mixtures.  

Note in each case, the components never need be removed from the plate before the second development is begun, saving time and potential loss. After development, other procedures and documentation as described below can be continued.  

The ease of application of 2D TLC is one example of an advantage of TLC compared to HPLC. It is much more difficult and costly to accomplish a successful 2D HPLC method. 2D HPLC often requires the collection of sample fractions or complex column switching. The biggest problem is often a mismatch of the mobile phase used in one mode compared to the second mode. If the mobile phases are immiscible, then drying and reconstituting each sample fraction is usually required before introduction into the second column. This is never a problem with 2D TLC because any initial mobile phase is evaporated from the plate before the second development is started.  

Multiple development either manually or with Camag's Automatic Multiple Development (AMD 2) instrument can also greatly improve the resolution of complex mixtures. The typical mode of automatic multiple development uses a multiple-step mobile-phase gradient with successive runs that have decreasing strength (less polar for a silica gel layer) and increasing development distance, with vacuum drying of the plate between steps. 

The development methods described above in this section involve migration of the mobile phase by capillary forces. There are now devices available for techniques that are categorized as forced-flow planar liquid chromatography (FFPLC) with which the mobile-phase migration is driven by other means. These include overpressured-layer chromatography (17), rotational planar chromatography (18), and pressurized planar electrochromatography (19).  

Detection (Visualization) of Zones and Documentation of Chromatograms

After drying the developed plate, compound zones are detected on the layer by their natural color, natural fluorescence under 366-nm UV light, quenching of fluorescence under 254-nm UV light on an "F" layer containing a phosphor (termed a UV indicator), or as colored, UV-absorbing, or fluorescent zones after reaction with an appropriate chemical reagent (postchromatographic derivatization). Chromogenic and fluorogenic reagents are applied by spraying or dipping a layer or exposing it to reagent vapors (for example, iodine). Chemical detection reagents can be nondestructive universal (iodine), destructive universal (for example, charring with sulfuric acid), or selective (for example, ninhydrin for amino compounds). Most chemical derivatization reactions require heating of the plate for completion. A forced air oven (with a solid glass or metal plate onto which the TLC plate is placed for even heating) or a Camag plate heater can be used. 

Effect directed analysis (EDA) is becoming more important in food, pharmaceutical, and phytochemical analyses. In EDA, there is no attempt to analyze all compounds in a sample, but instead the focus is on just the relevant (for example, harmful or bioactive) compounds detected by a bioassay coupled to TLC. These compounds can be the target analyte as well as unknown metabolites, side products, process contaminants, adulteration products, and residues. Bioassays used with TLC include microbiological assays (bioautography), bacterial assays (the Camag BioLuminizer bioluminescence detector utilizing 

), and biochemical detections using enzymes, including immunoassays. TLC–EDA was reviewed by Morlock and Schwack (20). 

If the chromatogram and its visualization produce visible results, it is possible to simply photocopy in black and white or color, or scan the TLC plate to document it. However, a commercial system for photography is needed if documentation of a combination of colored, fluorescent, and quenched zones is required. Camag, Desaga, and Analtech all offer photodocumentation systems. 

Figure 2: TLCâMS interface: (a) front of instrument and (b) schematic of extraction section. (Courtesy of Camag USA.)  

Separated zones are initially identified based on the correspondence of 

 values and detection characteristics between sample and standard zones on the same plate, but must be independently confirmed by other evidence, such as off-line and on-line coupling of TLC with various spectrometric methods. Scanning densitometers have a spectral mode that allows measurement of UV–vis spectra of zones directly on the plate and comparison with standards. In addition, peak purity can be examined by carrying out spectral studies at the peak start, middle, and end. 

One of the most important and active areas of current research is hyphenation of TLC with MS for identification or structure elucidation, which was reviewed by Morlock and Schwack (20,21). Many TLC–MS coupling approaches were covered in these reviews, but the elution head-based technique is the most interesting because of the commercial availability since mid-2009 of the Camag TLC–MS interface. Details of its use are available from Camag. The reviews by Morlock and Schwack also cover hyphenation of TLC with UV–vis, fluorescence, Fourier transform infrared (FT-IR), and surface-enhanced Raman spectrometry in various combinations to characterize samples as completely as possible. 

Most reported quantitative analyses by TLC involve reflectance scanning in the absorbance or fluorescence mode using a Camag TLC Scanner 3, a Desaga CD-60 densitometer, or a J&M diode array detector scanner. Standard zones are applied to a plate to create a calibration curve of peak area or height versus weight through linear, nonlinear, polynomial, or Michaelis–Menten regression, and weights of bracketed sample zones on the plate are interpolated from the curve. Videodensitometers with a visible or UV light source, camera, and software for image capture and quantification are also available from Camag and Desaga, but these do not allow measurement at an optimal wavelength as the slit scanning densitometers with a monochromator do.  

The chronology of the introduction of different TLC sorbents, plate formats, and instruments was reviewed in an historical article (22). As in column chromatography, the chemistry and size of the particles as well as the thickness of the layer affects the speed and resolution of the separations that are possible. Silica gel is the predominant TLC layer because of its high selectivity for a large range of compounds with different functional groups. Supposedly, similar layers according to the label from different manufacturers (for example, the most widely used layer by far, silica gel 60 F

) with the same layer thickness can give quite different results, perhaps because of the unstated binder type and content, and should be comparatively evaluated before substitution of plates in a given method.  

In addition to the chosen sorbent, precoated plates usually contain a binder to help the particles adhere to the support that serves as the backing for preparation of the layer. The most widely used commercial TLC and high-performance TLC (HPTLC) plates contain an organic polymeric binder, such as polymethacrylate at a concentration of 1–2%; these are harder, smoother, more durable, and generally give better separations than G layers. The classical TLC plates designated as "G" for gypsum (calcium sulfate) binder are still available from EMD/Millipore, Macherey-Nagel, and Analtech. The nature of the binder in a plate can affect separations and detection based on color reactions. 

Layers containing an indicator that fluoresces when irradiated with 254- or 366-nm ultraviolet light are designated as "F" or "UV" layers. These layers are used to facilitate the detection of compounds that absorb at these wavelengths and give dark zones on a bright background (fluorescence quenching). F

 indicators can give green (zinc silicate) or blue (magnesium tungstate) fluorescence. F

 indicators can be an optical brightener, fluorescein, or a rhodamine dye (23). Some precoated plates have both indicators to detect compounds that absorb at both wavelengths (F

 plates). Plates designated with an "s" have a UV indicator that is acid stable (F

 plates). Lux TLC plates (EMD/Millipore) and Adamant TLC and Nano-SIL-HPTLC plates (Macherey-Nagel) have enhanced brightness because of increased levels of the F

Table I: A comparison of the physical and operational parameters of TLC and HPTLC  

The differences between TLC and HPTLC can be seen in Figure 3 and Table I. The effect of the smaller particle size and the shortened development distances combine to give higher efficiency, shortened development times, increased sensitivity, and a larger number of samples processed simultaneously. 

Figure 3: Comparison of the separation of dansyl amino acids on (a) a classical TLC silica gel 60 plate and (b) an HPTLC silica gel 60 plate under identical conditions. Samples (bottom to top): N-Î±-dansyl-L-asparagine (lowest Rf), Î±-dansyl-L-arginine, dansyl-L-cysteic acid, N-dansyl-L-serine, dansyl-glycine, and N,N-didansyl-L-tyrosine (highest Rf). Sample volume: TLC, 4 ÂµL; HPTLC, 0.3 ÂµL; mobile phase: ethyl acetateâmethanolâpropionic acid (22:10:3,v/v/v); migration distance: TLC, 10 cm; HPTLC, 5 cm; analysis time: TLC, 42 min; HPTLC, 13 min, 45 s; detection: UV at 366 nm. (Chromatograms courtesy of Merck KGaA, Darmstadt, Germany.)  

TLC and HPTLC silica gel 60 plates with etched identification codes, spherical particle plates (not irregular silica gels as used in most standard layers), and thinner layers for special applications are also available. These can be checked out on the manufacturers' web sites (EMD/Millipore, Macherey-Nagel, and Analtech). 

Plates with Channels and Preadsorbent Zone

Kieselguhr (diatomaceous earth) and silica 50,000 are small surface area, weak adsorbents that are used as the lower 2–4 cm inactive preadsorbent (also called concentration zone) in the manufacture of silica gel and C18 preadsorbent plates. Samples applied to the preadsorbent zone run with the mobile-phase front and reform into sharp, narrow, concentrated bands at the preadsorbent–analytical sorbent interface. This leads to efficient separations with minimum time and effort in manual application of samples, and possible sample cleanup by retention of interferences in the preadsorbent, which can be modified by dipping (then drying) into reagents that can capture unwanted impurities once spotted. 

Channeled (laned) TLC and HPTLC plates have tracks scribed in the layer that are 9 mm wide with 1 mm between channels (19 useable channels across a 20-cm wide plate). Advantages include no possible cross-contamination of zones during development; exact location of zones during application, which facilitates alignment of a densitometer light beam for automated scanning; and easier removal of separated zones by scraping, before recovery by elution, without contaminating or damaging adjacent tracks. 

To remove extraneous materials that may be present because of manufacture, shipping, or storage conditions, it is recommended to preclean plates before use by development to the top of the plate with methanol (24). The plate is then dried in a drying oven or plate heater for 30 min at 120 °C. If the plate is being used immediately, it will equilibrate with the laboratory relative humidity (which should be controlled to 40–60% and recorded regularly) during sample application. Plates that are prewashed do not need activation by heating unless they have been exposed to high humidity. More complete suggestions for initial treatment, prewashing, activation, and conditioning of different types of glass- and foil-backed layers have been published (24).  

Glass plates can be cut to a smaller size using a commercial glass cutter and broken with grozier pliers for better results (Figure 4). Scissors, a razor knife, razor blade, or circular cutter (as used to cut fabric) can all be used to cut plastic or aluminum sheets. A straightedge should be used, and care must be taken to avoid damaging the sorbent layer and your fingers. 

Articles

Column Watch

Advantages of the planar, off-line format of TLC relative to on-line column HPLC, the steps in TLC analyses, and future prospects of TLC are discussed.

Stationary Phases for Modern Thin-Layer Chromatography

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 like liquid chromatography (LC) and gel electrophoresis, capillary electrophoresis (CE) is a proven powerful separation technique for biotechnology-derived proteins (4). CE analyses can be carried out 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 sheath-liquid 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 sheath-liquid 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 polybrene-poly(vinylsulfonic acid) (PB-PVS) and polybrene-dextran sulfate-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 analyzed. 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 TOF-MS for the analysis of aqueous solutions of rhGH. First, a standard solution of rhGH was analyzed using a BGE of high pH. A low-pH sheath liquid was used to allow efficient positive ESI. A narrow and symmetrical peak (

 ~ 100,000) was obtained for rhGH (Figure 2a, peak 0). Deconvolution of the mass spectrum obtained in the apex of the peak (Figure 2b, 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 analyzed (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 2b). 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 sulfonic acid groups may have been formed from a protein disulfide 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 sulfonated 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 3a). Figure 3b shows the deconvoluted mass spectrum obtained in the apex of the most intense peak (9.1 min), indicating a molecular mass of 22,375 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 22,375 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.  

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.  

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 analyzed 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 analyzed 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 5b the deconvoluted mass spectrum obtained for the most intense peak (25.4 min) is shown. A main mass of 29,888 Da is obtained. Also, a variety of other molecular masses are observed corresponding to glycoforms differing in number of HexHexNAc and SiA units (Figure 5b). 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 29,888 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 Figure 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 5c 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 fused-silica 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 8a). The reaction products (total protein concentration per preparation, 100 nM) were analyzed 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 8b). 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 8c). 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 8c). 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 (pH 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 8c). 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 sulfonic 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-molecular-weight 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, 

Article

CE–MS using noncovalent coated capillaries provides advantages for purity and stability analysis of biopharmaceuticals.

Capillary Electrophoresis–Mass Spectrometry for the Analysis of Biopharmaceuticals

Innovative technologies are rapidly facilitating our ability to measure the numerous and diverse lipids present in biological samples. In the second and final installment of this series, we present and discuss the use of those technologies for lipid analysis. 

In part I of this series, we introduced the concept of lipidomics as a new research strategy for analyzing lipids using state-of-the-art mass spectrometry (MS) technology. Together with genomics, transcriptomics, and proteomics, lipidomics is now routinely used in various scientific areas including drug and biomarker discovery, drug development, nutrition, biotechnology, and metabolic engineering research. Moreover, the future of lipidomics holds significant promise to further the development of personalized medicine. 

Researchers use lipidomics as a tool to analyze a selected set of lipids (targeted lipidomics). They also use it, in a more comprehensive way, to screen all lipids present in a biological sample (untargeted lipidomics). A lipidomic research strategy presupposes the choice of analytical solutions — sample preparation, chromatography, ionization sources, and MS analyzers — that are more appropriate for answering the biological questions posed. In the second and final installment of this series, we present and discuss the use of those technologies for lipid analysis.  

The high sensitivity of MS offers less labor-intensive lipid analyses. Nevertheless, the quality of sample preparation remains an important factor for success. Sample preparation includes lipid extraction from the biological matrix and elimination of nonlipid contaminants. Usually, lipid concentrations in biological samples are normalized by volume, tissue weight, cell number, or protein or DNA concentration. Quantification is done by lipids labeled with stable isotopes or nonendogenous structural analogs (internal standards) that are added before the extraction procedure. The methods for lipid extraction vary according to the sample matrix, the particular lipids under investigation, and the subsequent analytical method. The most common methods for lipid extraction are liquid–liquid extraction and solid-phase extraction (SPE).  

Jordi Folch and colleagues (1) and E.G. Bligh and W.J. Dyer (2) developed the most widely accepted methods for lipid extraction from biological samples. Those methods are based on chloroform–methanol protocols that include phase partitioning of lipids into the organic layer. For a large variety of physiologically relevant lipids, the protocols work relatively well, and they have been adapted toward recovering more complex lipid chemistries as well as low-abundance lipid species. Multistep extractions and pH alterations also can be used for recovering acidic phospholipids such as gangliosides and phosphoinositides. Alternative extraction procedures that use less toxic organic solvents such as methyl-

-butyl ether, hexane–isopropanol, and ethyl acetate–ethanol mixtures have been proposed for a wide range of tissues.  

SPE has become a frequently used analytical procedure for rapid, preparative separation of total lipid extracts into different lipid classes. SPE involves using diverse solvents and commercial, prepacked columns with various stationary phases including silica and reversed-phase materials. (The latter use chemically bonded octadecylsilyl [ODS, C18] groups or ion-exchange media such as those with bonded aminopropyl or phenylsulfonic acid moieties.) Lipid extraction from a complex matrix might be achieved by choosing conditions (solvents and stationary phases) so that the required analyte is retained on the column while the impurities pass directly through. Conversely, extraction occurs by allowing the analyte to be eluted through the column while the impurities are retained. Targeted lipidomic studies often use SPE for fractionating the low-abundance lipids species from the highly abundant species like phospholipids, creating dynamic-range and ion-suppression issues. 

These three MS-based lipidomics approaches are currently in use: 									

Direct infusion MS (for example, shotgun lipidomics)

On-line chromatographic separation-MS (for example, liquid chromatography–mass spectrometry [LC–MS])

Surface-based desorption ionization MS (for example, MS imaging) 

Ion mobility spectrometry, a post-ionization separation technique, can be coupled to any of these approaches. 

Directly infusing lipid extracts into the ionization source (usually electrospray ionization [ESI] or nanoESI) of the mass spectrometer without prior chromatographic separation provides for high-throughput analysis of lipid species in biological samples. This technique is known as "shotgun lipidomics" (3), by analogy to the much-used shotgun genomic sequencing. Direct infusion experiments are performed by continuous flow injection or loop injection, using automated robotic systems. "Intrasource" separation preferentially ionizes certain lipid classes by varying ESI parameter values, like source temperature and cone voltage, and doping the spray solvent with counterions, like lithium hydroxide. Additional ramping of fragmentation conditions, such as collision energy or other instrument settings, can increase the dimensions of separation of lipid species in what has been named "multidimensional MS-based shotgun lipidomics" (MDMS-SL).  

Similar to proteomics experiments, shotgun lipidomics experiments lend themselves to dividing into categories of top-down and bottom-up. In a top-down analysis, lipid species are identified by their accurate mass-to-charge ratio (

), determined on high-resolution mass spectrometers such as time-of-flight (TOF), Fourier transform ion cyclotron resonance (FT-ICR), and orbital trap (4–6). Although individual molecular species are not identified by MS-MS, top-down lipidomics has demonstrated potential in high-throughput screens. 

Figure 1: MS-MS operation modes for triple-quadrupole instruments. In triple-quadrupole instruments, ions pass through three successive quadrupoles (Q1, Q2, and Q3). Q2 is the collision cell, where precursor ions fragment upon collision-induced dissociation (CID) with an inert gas (helium or argon).  			

In bottom-up lipidomics, all lipid precursor ions are subjected to MS-MS analysis, and identified by 

 of characteristic fragments or neutral losses on triple quadrupole instruments (Figures 1 and 2) (7).  

Figure 2: Representative precursor ion scanning of phospholipid classes from a total lipid extract using direct infusion-MS in positive ESI mode. [M+H]+ ions for phosphatidylcholines (PC), lyso-PC, and sphingomyelin were detected by monitoring the characteristic fragment at m/z 184.1 with a triple-quadrupole instrument. 			

Figure 1 shows MS-MS modes for triple-quadrupole instruments. Four main MS-MS scan experiments use a triple-quadrupole instrument:  					

Product ion scan: A precursor ion is selected in Q1, fragmented in the collision cell (Q2), and then all resultant masses are scanned (Figure 1, panel A). This experiment is commonly performed to identify transitions used for quantification by tandem MS. 

Precursor ion scan: The product ion is selected in Q3, and the precursor masses are scanned in Q1 and fragmented in the collision cell (Figure 1, panel B). This experiment is performed to identify complex lipids containing common building blocks and therefore belonging to the same lipid class or subclass. 

Neutral loss scan: Q1 scans all the masses; all ions are fragmented in the collision cell. Q2 also scans, but at a set offset from the first mass analyzer (Figure 1, panel C). This offset corresponds to a neutral loss that is commonly observed for lipids loosing the same fragment on collision and which are, therefore, closely related.

Multiple reaction monitoring (MRM): Both Q1 and Q2 are set to a selected mass (Figure 1, panel D). This experiment offers a very selective and highly sensitive analysis used mainly for quantification in targeted lipidomic approaches. 

Multiple (tens) precursor-ion and neutral-loss scans could be monitored simultaneously to detect various classes of lipids for a more untargeted analysis. Indeed, most lipid species represent linear combinations of relatively few building blocks. Those building blocks include glycerol, sphingoid bases, polar head groups, and fatty acyl substituents, which can be released from the precursor ions upon collision-induced dissociation (CID) with an inert gas (argon or helium). For example, using ESI in positive mode, precursor-ion scans for 

 184.1 (phosphocholine, Figure 3) or neutral loss scans for 

 59.1 (trimethylamine) can identify the [M+H]

  ions that correspond to sphingomyelins, phosphatidylcholines, and lysophosphatidylcholines. Using ESI in negative mode, precursor-ion scans for 

  ions of sulfatides whereas neutral loss scans for 

 87.0 (serine) can be used to identify phosphatidylserines. 

Figure 3:  Representative data-independent analysis of a selected phosphatidylcholine species (1-stearoyl-2-oleoyl-sn-glycero-3-phosphocholine) using ESI in positive mode allows the simultaneous acquisition of precursor (top) and fragment ion (bottom) information in a single UHPLC run. 			

Shotgun lipidomics on triple-quadrupole instruments remains a low-mass resolution approach (1–2 Da), and only a single precursor or neutral-loss scan can be performed at a time. Hybrid, tandem mass spectrometers built on TOF, FT-ICR, and orbital trap technology now make it possible to acquire high-resolution tandem mass spectra from hundreds of lipid precursors (6,8). For example, a quadrupole-TOF instrument makes it possible to sequentially isolate precursor ions in unit or higher mass resolution windows by using a quadrupole working stepwise from a low to a high 

 ratio, transmit ions into a collision cell, and acquire accurate measurements of full product ion spectra for that window at each step of the quadrupole. The automated post-acquisition interpretation of the full product ion spectra effectively emulates an unlimited number of precursor and neutral-loss scans in a single analysis. With these modes of acquisition, no prior knowledge of the lipid ions of interest is required, which is ideal for the rapid analysis of unknown lipid mixtures in biological samples.  

Although direct infusion can offer a high-throughput lipidomic technique, it is nonetheless associated with drawbacks. It cannot distinguish isobaric and isomeric species, whose masses are identical, and they often display similar fragmentation profiles. Furthermore, simultaneous introduction of multiple analytes into the ionization source can induce undesirable effects such as ion-suppression. 

To overcome the drawbacks of direct infusion, a more traditional approach engages the power of the LC separation to divide a complex mixture of lipids before MS analysis. LC can reduce ion suppression caused by coeluted lipids and isobaric interferences, and it can often separate isomers. In particular, chromatographic separation before an MS analysis is still necessary for analyzing very low-abundance (picomolar and attomolar levels) signaling lipids, such as eicosanoids, docosanoids endocannabinoids, and 

-acyl amino acids, to name a few. Important structural information can be found in the elution order from LC columns. The elution order can significantly aid in the characterization of lipids, generating an orthogonal measure (retention time, factor, or index) that gives information on the physical-chemical properties of the lipid species analyzed. Furthermore, because lipids can differ in mass by only two units, a partial chromatographic separation helps avoid the isotopic effects, which affect the actual mass abundance during direct-infusion experiments. 

The combinatorial nature of complex lipids, however, sometimes makes separating isomeric species by conventional high performance liquid chromatography (HPLC) difficult. One approach intended to increase chromatographic resolution uses reduced particle sizes (<2 µm). That approach led to the development of ultrahigh-pressure liquid chromatography (UHPLC) (10). Sub-2-µm particles provide narrow chromatographic peaks (often <3 s), which results not only in better resolution but also in lower detection limits and shorter times for the chromatographic run compared with conventional HPLC (9,10). The high pressures (10,000–15,000 psi) needed to operate these columns, which require special instruments, constitute the downside of this approach. UHPLC is often coupled to hybrid-tandem MS, which allows data-independent acquisition (see part I of this series) for the simultaneous collection of both precursor and fragment ion information (Figure 3) in a single chromatographic run (9,10). 

Reversed-phase LC using C18 columns is often used to separate lipids according to their fatty acyl chain length and degree of unsaturation. Lipids species containing longer acyl chains are eluted from the LC column later than shorter chain lipids (for example, C18:0>C16:0>C14:0). Saturated acyl structures tend to be eluted later than long polyunsaturated structures do (for example, C18:0>C18:1>C18:2). Generally, the first double bond on an acyl chain reduces the effective chain length by a little less than two carbon units, and additional double bonds have smaller effects on retention (9,10). 

Although reversed-phase separation methods provide good separation of lipids, the complete separation of lipid classes is not achievable. Normal-phase methods typically allow separation and characterization of different lipid classes, but their lengthy elution and equilibration times make them time-consuming processes. Moreover, the mobile phases typically used for normal-phase separations, such as chloroform or hexane, are difficult to handle because of their volatility and toxicity, and they prove challenging for ionization and introduction into a mass spectrometer. Although hydrophilic interaction liquid chromatography (HILIC) is considered a variant of normal-phase chromatography, using a reversed-phase solvent system (organic-aqueous) can avoid these limitations. Indeed, because of the high percentage of organic mobile phase used in HILIC (for example, >80% acetonitrile), ESI may be improved through more efficient mobile-phase desolvation and compound ionization (11). 

The ability to perform two-dimensional LC in one injection can improve the separation of complex lipid mixtures. For example, by coupling HILIC with reversed-phase chromatography, it is possible to profile different lipid classes in the first dimension (HILIC) and then separate lipid species in the second dimension (reversed phase), with significantly increased sensitivity (12). With the advent of commercialized two-dimensional UHPLC systems that couple with MS instruments, multidimensional chromatography will increasingly become a more practical tool for deconvoluting the chemical complexity of lipidomic profiles.

Recent advances in technology have revived the exploration of supercritical fluid chromatography (SFC) as a viable analytical technique for hydrophobic molecules (13). Supercritical fluids are highly compressed gases, which share the physical properties of both gas and liquid, and exhibit low viscosity and high solute diffusivity. These properties promote lower back pressure, shorter re-equilibration times, and higher throughput. Supercritical fluids (for example, carbon dioxide) in association with a cosolvent (typically, methanol) are compatible with MS. Moreover, they have demonstrated their usefulness as tools for separating complex mixtures of lipids. Although still in its infancy, SFC–MS can potentially find various applications for analyzing complex lipid mixtures while reducing the use of toxic solvents and the cost of analyses. 

Progress in microfluidic LC has led to the integration of most of the nanoflow LC components into a single LC-chip, which avoids problems associated with capillary connections and the need to keep the system free of leaks, blockages, and excessive dead volume. Lipidomics researchers are now benefiting from the higher sensitivity of the integrated LC-chip–MS platforms when analyzing low-abundance lipids and infinitesimally small amounts of biological materials such as dried blood, skin, hair, saliva, tears, and sweat — all of which may be used for diagnostic purposes. 

The past decade has witnessed an exponential growth in surface-based desorption ionization (DI) techniques and their applications in lipid analysis (14). In DI, the ionization process begins by irradiating a defined spot on the solid-state sample using a focused, excitatory beam such as a laser, ions, or charged solvent droplets. Upon impact, the sample's surface releases a vapor of ionized molecules that can be directed into a mass spectrometer. Alternatively, acoustic or thermal desorption could initiate the ionization process. 

Notably, DI-MS spectra of the biological tissues feature ions corresponding mainly to lipids. Lipids, which by molar quantities are the most abundant ionic molecular species in tissues, ionize well under physiological conditions. As occurred with the first MS experiments in the 1950s (please see part I of this series), lipids are again proving useful as a molecular tool for evaluating the potential of new MS-based technologies. 

Except for matrix-assisted laser desorption ionization (MALDI), most DI techniques are matrix-free. While some operate mostly under vacuum, like MALDI, cluster-secondary ion MS (SIMS), and nanostructure-initiator mass spectrometry (NIMS), others can operate at atmospheric pressure (ambient MS techniques). There are more than 25 DI techniques currently being used, and they display unique advantages according to sample type and application (14). Generally, DI-MS provides a new level of description beyond the pure measure of lipid concentration, which is the detailed spatial distribution of lipid species on a surface. Information like this is often missed using traditional sample preparation and lipid extraction protocols for lipidomic analysis.  

Nevertheless, surface-based, DI-MS techniques are not immune to drawbacks such as the incapacity to separate isobaric and isomeric species and the limitation of being (frequently) nonquantitative by nature. Apart from those drawbacks, DI-MS approaches main advantage is the ability to generate lipid fingerprints for discriminating among phenotypes, for example, healthy from diseased and bacterium A from bacterium B. Therefore, for their practicality, DI-MS techniques offer a convenient solution for the development of devices for lipidomics-based diagnostics. 

Three main biological applications for DI are of interest for lipids analysis:  									

DI-MS techniques are useful for real-time, rapid, in situ screening of cells, biological fluids, and tissues such as leaves, bacteria, skin, biopsies, and dried blood spots. Many new DI technologies have been applied to this kind of lipid analysis, including direct-analysis in real time (DART), extraction electrospray ionization (EESI), probe electrospray ionization (PESI), and rapid evaporative ionization mass spectrometry (REIMS). Using these techniques, the in situ generation of a particular profile of lipid ions has been proposed for the real-time molecular differentiation of diseased versus healthy tissues.

DI-MS can be used for direct analysis of processed tissue specimens or of planar chromatography spots, for example, paper chromatography, thin-layer chromatography (TLC), and high performance TLC (HPTLC). For example, we can now analyze by DI-MS processed tissues such as lipid extracts with or without using special surfaces that favor the ionization process. Robotic systems for in situ lipid extraction (by either solvent or SPE) and deposition have been shown to increase the dynamic range of detection (15). Using DI-MS in conjunction with planar chromatography is another very convenient method to analyze lipid species. In the past, unknown lipids were scraped from the paper or TLC–HPTLC plate, eluted into a tube, and transferred into the mass spectrometer. Today, very convenient planar chromatography can be interfaced with DI-MS to automatically analyze the lipids on the paper or plates.

MS-The Practical Art

Mass spectrometry analysis of lipids in complex matrices is revolutionizing the way lipid research is conducted.

New Frontiers for Mass Spectrometry in Lipidomics, Part II

This installment describes combining diode-array detection (DAD) and coulometric electrochemical-array detection to overcome sensitivity and specificity limitations often encountered when analyzing natural products.

Plants contain an amazingly diverse range of secondary metabolites that may offer important health benefits or risks (1,2). The challenge facing analytical chemists seeking to identify and measure these compounds is twofold: first, to accurately discriminate between compounds sharing similar physicochemical characteristics; and second, to tease this information out of complicated matrices including botanicals, supplements, foods, and beverages.  

Gradient high performance liquid chromatography (HPLC) with diode-array detection (DAD) is often used to measure the major components present in botanicals. HPLC–DAD is a robust, relatively simple technique that quantifies known analytes that can be resolved in simple matrices, but it suffers from a lack of specificity whenever compounds with similar structures that cannot be deconvoluted spectrally are coeluted (Figure 1). Furthermore, this technique is generally not sensitive enough for the study of natural product metabolism in animals and humans. Electrochemical detection (ECD), on the other hand, is highly sensitive and selective and can distinguish between subtle changes in chemical structure (Figure 1). 

Figure 1: Comparison of the spectral and electrochemical properties of selected flavonoids. Compounds with similar structures have similar absorbance spectra and changes in functional groups have limited effect on this response (inset). Electrochemical detection, however, is extremely sensitive to the type, degree, and position of substitution as shown by the different current voltage curves (hydrodynamic voltammograms) of catechin (oxidizing at approximately +100 mV), hesperidin (oxidizing at approximately +400 mV), and naringin (oxidizing at approximately +700 mV). 			

				Although ECD is selective, many chemicals are electrochemically active, including compounds containing an aromatic backbone with alcohol, amine, or thiol groups attached directly to an aromatic ring, quinones, and conjugated polyenes (with three or more conjugated double bonds) (3). Phytochemical examples include some alkaloids (such as lysergic acid and morphine), carotenoids (such as carotenes and lycopene), fat soluble antioxidants (such as tocopherol and phylloquinine), phenolic acids, and various polyphenols such as the flavonoid group (for example, catechins, daidzein, and genistein). 			

The structure–activity relationships of polyphenolics found in foods and beverages and their antioxidant properties have been extensively studied and are correlated to the ease of electrochemical oxidation. For example, the structural characteristics of polyphenolic phytochemicals that influence their oxidation potential include the type of substitution (such as hydroxy and methoxy), degree of substitution (such as phenol, catechol, and gallo), and the position of substitution relative to other groups (that is, ortho, para, and meta). Thus, the oxidation potential for different substitutions can be ranked from the most labile (lowest oxidation potential) to the most stable (highest oxidation potential): trihydroxy > dihydroxy > methoxy > monohydroxy. Such differences in electrochemical behavior are presented in Figure 1, using catechin, hesperidin, and naringin as examples. Because the compounds require different applied potentials to produce an electrochemical response, this attribute can provide voltammetric resolution when using an array of coulometric electrochemical sensors. Such electrochemical arrays are typically used with the first electrode set to a low potential and each electrode in the series set at increasing, but fixed potentials. Easily oxidized compounds can be selectively detected at upstream, low-potential sensors, and compounds that are more difficult to oxidize respond at downstream sensors set at higher potentials. Coulometric electrochemical-array detection uses a series of as many as 16 highly efficient sensors to voltammetrically resolve those compounds that are coeluted chromatographically (Figure 2) and can identify these analytes based on their voltammetric profile (in an analogous fashion to spectral information obtained from DAD) (3,4). This is a practical and robust form of ECD. 

Figure 2: In this application both chlorogenic acid and 4-hydroxybenzoic acid are coeluted and cannot be resolved spectrophotometrically (UV at 254 nm). However, the electrochemical profiles of these two compounds differ substantially, so they can be resolved voltammetrically, with chlorogenic acid responding exclusively at an upstream (low potential) electrode shown in red and 4-hydroxybenozoic acid responding exclusively at a downstream (high potential) electrode shown in green. 			

				This approach has been successfully applied to the targeted measurement of numerous phytochemicals in a number of matrices such as botanicals (5,6), juice and alcoholic beverages (7–13), food (14–16), and tobacco (11,17). The sensitivity and selectivity of this technique are crucial when studying mammalian phytochemical metabolism (18–28). 			

				Gradient HPLC with coulometric array detection is capable of resolving and measuring several hundred analytes  simultaneously (3,29–32). Changes in the pattern of metabolites, when evaluated using chemometric modeling software, can be used to study product adulteration, contamination, composition, and stability, and, in the case of wine and fruit juice, the effect of growing region and differences between varietals (33). 			

Although electrochemical detection offers analysts many advantages, it is limited to the measurement of electrochemically active compounds only. The combination of DAD and coulometric electrochemical-array detection extends the range of compounds measureable by either technique alone. This platform, called the spectro-electro array, places the diode-array detector before the coulometric array, thereby enabling spectral characterization of analytes before their electrochemical detection. As discussed below, this technique was evaluated for its ability to resolve and quantify targeted phytochemicals in crude extracts of a variety of botanical supplements (for example, ginseng, black cohosh, and gingko), beverages (black and green tea), culinary herbs (oregano, rosemary, sage, and thyme) and spices (clove and nutmeg), as well as its ability to differentiate between samples based on their metabolite patterns.  

A Thermo Scientific Dionex LPG-3400BM pump with an SR-3000 solvent rack was used. The autosampler used was a Thermo Scientific Dionex WPS-3000TBSL. The UV detector used was a Thermo Scientific Dionex DAD-3000RS diode-array detector, with the following settings: channel 1: 218 nm; channel 2: 240 nm; channel 3: 254 nm; and channel 4: 275 nm. A Thermo Scientific Dionex CoulArray detector with Thermal Organizer was used as the electrochemical detector, consisting of a 16-channel array at potentials ranging from 0 to +900 mV in 60-mV increments. Mobile-phase A consisted of 20 mM monobasic sodium phosphate, 3% acetonitrile, and 0.2% tetrahydrofuran at pH 3.35. Mobile-phase B consisted of 20 mM monobasic sodium phosphate, 50% acetonitrile, and 10% tetrahydrofuran at pH 3.45. Mobile-phase C consisted of 90% methanol. The gradient was as follows: 0–2 min at 2%B/3%C, then 2–30 min ramp to 97%B/3%C, hold until 45 min at 97%B/3%C. A 150 mm × 3 mm, 3-µm Thermo Scientific Acclaim 120 C18 HPLC column was used with a flow rate of 0.65 mL/min and an injection volume of 10 or 20 µL. 

Data were analyzed using Thermo Scientific Dionex Chromeleon chromatography data system 6.8 (SR9) and CoulArray software 3.1. Electrochemical-array data were transferred to Pirouette chemometrics software (Infometrix, Inc.) for chemometric analysis using a CoulArray version 2.0 software utility (Pattern Recognition Setup Wizard).  

Stock standards, depending on solubility, were prepared in ethanol, methanol, or methanol and water solutions at 1000 µg/mL or 100 µg/mL. Working standards were prepared at 0.2, 0.5, and 1.0 µg/mL in 10% methanol containing 0.2% ascorbic acid and 0.02% EDTA. 

Figure 3: Measurement of standard mixture by UV detection at 218 and 254 nm (upper figure) and 16-channel coulometric electrochemical-array detection (lower figure). 			

Supplements and culinary herbs were prepared for analysis by extracting 100 mg of the material with 20 mL of methanol. The samples were sonicated for 30 min and subsequently centrifuged to obtain a clear solution. The solution was diluted 5× with a preservative solution (10% methanol containing 0.2% ascorbic acid and 0.02% EDTA) for injection into the HPLC system. Wine samples were diluted 1:50 (v/v) with the preservative solution.  

Table I: Analyte identity (UV1âUV6 are analytes that have strong UV but weak EC response) 			

As shown in Figure 3, numerous phenol and polyphenol analytes can be resolved using this approach (see Table I for identification). It should be noted that there are six compounds designated UV1–UV6 that showed poor response by ECD, but they are readily detectable by UV. The limits of detection were typically 10–50 pg on-column by ECD and 100–500 pg by UV. The limits of quantification were 200–1000 pg (on-column) by ECD and 500–5000 pg by UV. The response ranges were over seven orders of magnitude by ECD and five by UV. Typical 

				 values were ~0.999 or better for all compounds. The average interday retention time precision for all analytes averaged 0.55% RSD over a 10-day period, with a range of 0.30–1.22%. 

Table II: Abundance (Âµg/g) of different analytes in supplements and herbs 			

The targeted approach was used to measure the levels of phenols and polyphenols in a variety of supplements and culinary herbs. Analyte levels (Table II) were in good agreement with levels reported in previous publications (9,34–36). St. John's wort (

) is reportedly effective in the treatment of moderate depression in a number of clinical trials. The two major polyphenols in St. John's wort, hypericin and pseudohypericin, were easily measured using both UV detection and ECD (Figure 4, ~18 min). Rosemary is particularly rich in carnosic acid, a potent antioxidant and anticarcinogen that is a major constituent of the dried leaf. Another major component of rosemary is rosemarinic acid, which is also known for its antioxidant properties. 

Figure 4: Measurement of phytochemicals in St. John’s wort by UV detection at 254 nm (upper figure) and 16-channel coulometric electrochemical-array detection (lower figure). 			

Examination of the pattern of both known and unknown metabolites can be used to authenticate a sample or measure contamination and adulteration. Two simple experiments were used to evaluate the application of the spectro-electro array to metabolomic studies. The first examined the metabolite profiles of a selection of red wines. The general polyphenol method was used to analyze five red wines (four cabernet sauvignon samples and one burgundy sample). Several hundred analytes, including both known (see Table III) and unknown compounds, were measured in each sample. Levels of known compounds were similar to those previously published (8,9). Principal component analysis (PCA) was able to identify wine samples as either burgundy or cabernet sauvignon (Figure 5). Although this study is preliminary it does show the capability of the system to differentiate samples by grape varietal or blend and growing region. This approach is important when trying to authenticate a sample or for identifying product adulteration. 

Table III: Some of the more abundant analytes measured in different wine samples; wine 1: cabernet sauvignon, Argentina; wine 2: cabernet sauvignon, South Africa; wine 3: cabernet sauvignon, United States; wine 4: cabernet sauvignon, Chile; and wine 5: hearty burgundy, United States  			

In the second application, the metabolomics approach was investigated as a means to authenticate or detect adulteration of orange juice samples (33). The study was intended to detect the lowest level of adulteration (by blending with other juices or through the addition of orange peel or pulp-wash) that could be detected in orange juice samples. Blending of as little as 10% grapefruit juice into orange juice could easily be measured. Similarly, blending only 10% orange peel or 10% pulp-wash into orange juice could be detected. 

Figure 5: Initial study showing principal component analysis of wines. 			

There is considerable interest in the antioxidant capacity of botanical supplements, foods, and beverages with high levels of antioxidants purported to offer health benefits. There are a number of different approaches that vary in their ability to accurately measure the total antioxidant capacity of a given sample and these have been reviewed elsewhere (37,38). One approach, the oxygen radical absorbance capacity (ORAC) test, is routinely applied to foods. An advantage of the ORAC test is that it is relatively quick and easy to perform, however it can only provide data for the total antioxidant capacity of the sample rather than measuring the individual contribution of different antioxidants (39). The coulometric electrochemical array not only has the ability to identify and simultaneously quantify multiple antioxidants (and thus the contribution of individual compounds), but also the total antioxidant capacity of a sample can be calculated through the summation of the response of all analytes in the pattern. There is good correlation between ORAC and the coulometric electrode array (39,40). 

Gradient HPLC with spectro-electro array detection can quantify a wide variety of important phytochemicals at trace levels in complex matrices such as botanical preparations, spices, wine, and fruit juice. To maintain data quality, sample preparation is kept as simple as possible and stabilizers are added to prevent auto-oxidation of analytes before injection. The sensitivity, wide dynamic range, and resolution of this combined approach is superior to traditional spectroscopic techniques. 

Gradient elution and voltammetric resolution of similar compounds significantly extends the number of analytes capable of being measured simultaneously. The great utility of this method lies in the ability to separate and quantitate approximately 50 different phenolic and polyphenolic compounds in a wide variety of sample matrices. In addition, even unidentified compounds provide useful information when used in pattern recognition studies. These metabolite patterns are useful for evaluating product identity, purity, authenticity, and for differentiating among varietals, strains, and growing regions. 

The spectro-electro array can be used to measure the total antioxidant capacity of a sample and to identify individual contributing antioxidants.  

 is a product applications manager with Thermo Fisher Scientific in Chelmsford, Massachusetts. 

 is a product and support scientist with Thermo Fisher Scientific. 

 is director of customer and application support with Thermo Fisher Scientific. 

 is with technical support at Thermo Fisher Scientific. 

 "Innovations in HPLC" Editor Michael E. Swartz, PhD, is Principal Scientist in Analytical Development at Ariad Pharmaceuticals, Cambridge, Massachusetts, and a member of 

's editorial advisory board. Direct correspondence about this column to "Innovations in HPLC," at 

(1) A. Abdel-Rahman, N. Anyangwe, L. Carlacci, S. Casper, R.P. Danam, E. Enongene, G. Erives, D. Fabricant, R. Gudi, C.J. Hilmas, F. Hines, P. Howard, D. Levy, Y. Lin, R.J. Moore, E. Pfeiler, T.S. Thurmond, S. Turujman, and N.J. Walker, 

(2) Z.P. Xiao, Z.Y. Peng, M.J. Peng, W.B. Yan, Y.Z. Ouyang, and H.L. Zhu, 

Innovations in HPLC

HPLC with diode-array and electrochemical-array detection overcomes sensitivity and specificity limitations in natural products analysis.

The Spectro-Electro Array: A Novel Platform for the Measurement of Secondary Metabolites in Botanicals, Supplements, and Beverages

Analytical method validation (AMV) is sometimes referred to as "the process of providing documented evidence that a method does what it is intended to do." Laboratories in the pharmaceutical and other regulated industries must perform AMV to comply with regulations. Published guidelines (1,2) help laboratories understand expectations. Conducting AMV is also good science. 

In AMV, several performance characteristics may be investigated, depending on the type of method and its intended use. These are summarized below. 

 is the ability to measure accurately and specifically the analyte of interest in the presence of other components. In drug assays, specificity takes into account the degree of interference from other active ingredients, excipients, impurities, degradation products, or matrices, and ensures that a chromatographic peak corresponds to a single component. Specificity can be demonstrated by the resolution between peaks of interest. Modern chromatographic methods typically include a peak-purity test based upon photodiode-array detection or mass spectrometry.  

Analytical method validation is the process of demonstrating that a method does what it is intended to do.

LCGC North America-06-01-2012

Stationary Phases for Modern Thin-Layer Chromatography

Capillary Electrophoresis–Mass Spectrometry for the Analysis of Biopharmaceuticals

New Frontiers for Mass Spectrometry in Lipidomics, Part II

The Spectro-Electro Array: A Novel Platform for the Measurement of Secondary Metabolites in Botanicals, Supplements, and Beverages

Performance Characteristics Measured in Method Validation

When Should an Internal Standard be Used?

Vol 30 No 6 LCGC North America June 2012 Regular Issue PDF