A reversed-phase (RP) high performance liquid chromatography (HPLC) method for analysing melamine residue in cow's milk using 100% water in the sample preparation stage and as the eluent is described. HPLC was performed on an ODS column with an isocratic 100% water mobile phase and a photo-diode array (PDA) detector. The proposed method was validated by the analyses of spiked milk samples resulting in recoveries greater than 96.8% with a relative standard deviation (RSD) of less than 2.0%. The total analysis time was less than 15 min/sample and the quantitation limit was 0.05 µg/g. 

Melamine [1,3,5-triazine -2,4,6-triamine (C

)] is an inexpensive nitrogen-containing industrial chemical that is widely used in the manufacture of plastics and adhesives, but can also be used to alter tests to check the protein content of milk products (Figure 1). Adding melamine increases the nitrogen content of the milk and its apparent protein content measurement, which falsely increases the final price of the milk. 

 Figure 1: Chemical structure of melamine.			

Recent reports from several countries on food products containing milk contaminated with melamine have led to an increased interest in the analysis of this compund. Melamine contamination has been detected in various food products containing milk (1–3). Significant consumption of melamine can result in renal failure and, in rare cases, death. In fact, melamine contamination in milk products has resulted in illness in nearly 300,?000 babies and the deaths of at least six infants in China in 2008 (3–5). 

The European Union set a standard for acceptable human consumption (the tolerable daily intake — TDI) of melamine at 0.5 milligrams per kg of body mass (reduced to 0.2 mg per kg in April 2010) (6,7). Canada declared a limit of 0.35 mg and the US Food and Drug Administration's (FDA's) limit was put at 0.63 mg, but was later reduced to 0.063 mg daily. The World Health Organization's (WHO's) food safety director estimated that the amount of melamine a person could stand per day without incurring a bigger health risk was a TDI of 0.2 mg per kg of body mass (6,8). The Codex Alimentarius Commission has set the maximum amount of melamine allowed in food products containing milk to 2.5 µg/g and the amount of the chemical allowed in powdered infant formula to 1 µg/g (9), equivalent to the FDA's safety limit (3). Milk is an indispensable food and a base raw material of various foods because it is highly nutritious, inexpensive and readily available. Strict monitoring of the presence of melamine in milk is, therefore, an important activity as a means of guaranteeing food safety.  

The development of international standardized methods to determine chemical residues in foods is essential to guarantee equitable international trade in these foods and ensure food safety for consumers. Whether in industrial nations or developing countries, an internal harmonized method for residue monitoring in foods is needed. The optimal method for the routine monitoring of chemicals in foods should be simple, quick, economical in time and cost and, importantly, cause no harm to the environment and analyst.  

Eliminating the use of organic solvents and reagents is an important goal in terms of environmental conservation, human health and the economy. 

The reported methods for quantification of melamine in milk include enzyme-linked immunosorbent assay (ELISA) (10), GC–MS (10,11), HPLC–UV (12) or diode array detector (DAD) (13) and LC–MS–MS (14,15). The FDA issued new methods for the analysis of melamine in liquid formula, based on LC–MS–MS detection (16). The direct analysis of liquid milk by low-temperature plasma MS (LTP–MS) (17) and ultrasoundassisted extractive electrospray ionization (EESI–MS) (18) have recently been proposed. The facilities that GC–MS, LC–MS–MS, LTP–MS or EESI–MS system are available in are limited because these methodologies use complex, specific and expensive instrumentation. These are unavailable in a lot of laboratories for routine analysis, particularly in developing countries. All of the methods mentioned above consume organic solvents and/or reagents. No optimal method that satisfies the requirements for melamine analysis previously described has yet been identified.  

This article describes a safe and simple method to monitor melamine residue in milk using 100% water in sample preparation and HPLC separation without using organic solvent and reagents.  

 Distilled water (HPLC grade) and melamine standard were purchased from Wako Pure Chem. Ltd. (Osaka, Japan). A stock melamine standard solution of 100 µg/mL in water was taken into a cryo-vial, sealed and refrigerated until further use. Working standard solutions were prepared by diluting the stock solution with water.  

The following apparatus were used in the sample preparation stage: handheld ultrasonic-homogenizer (model HOM100, 2 mm i.d. probe, Iwaki Glass Co., Ltd., Funabashi, Japan); micro-centrifuge (Biofuge fresco, Kendo Lab. Products, Hanau, Germany); MonoSpin C18 as a centrifugal monolithic silica spin mini-column (GL Sciences Inc., Tokyo, Japan) (highly pure sol-gel silica gel monolith, sample throughput volume ≤ 300 µL). The spin column (GL Sciences) was pre-conditioned as follows: 100 µL of water was added and then centrifuged at 3,500 g for 1 min at room temperature.  

The following types of non-polar sorbent columns (5 µm 

 250 or 150 × 4.6 mm) for HPLC were used: Wakosil 5TMS (C1) (Wako); Mightysil RP-4GP (C4) (Kanto Chemical Co., Inc., Tokyo, Japan); Lichrospher 60 RP-selectB (C8) (Merck, Darmstadt, Germany); Mightysil RP-18 GP Aqua (C18) (Kanto); Mightysil RP-18 (C18) (Kanto); Inertsil HILIC (alkyl diol) (GL Sciences, Tokyo, Japan); Inertsil ODS-4 (C18) (GL Sciences) (Table 1).

Table 1: Physical/chemical specifications of the reversed-phase columnsa used and chromatographic melamine separation obtained under the HPLC conditions examinedb.			

The HPLC system included a model PU-980 pump and DG-980-50 degasser (both from Jasco Corp., Tokyo, Japan), as well as a model SPD-M10A 

 diode array detector (DAD) (Shimadzu Scientific Instruments, Kyoto, Japan). The analytical column was an Inertsil ODS-4 (octadecyl chemically bonded silica, C18) (250 × 4.6 mm, 5 µm 

  surface area, 100 Å pore diameter, 1.05 mL/g pore volume, 11% carbon load) column (GL Sciences) using a water mobile phase at a flow rate of 1.0 mL min

  at room temperature. The 25 °C DAD was operated at 190–300 nm: monitoring the wavelength for melamine was adjusted to 202 nm which is an absorption maximum for the analyte. The injection volume was 20 µL. 

 An accurate 0.1 mL sample was taken into a micro-centrifuge tube and homogenized with 0.6 mL of water with a handheld ultrasonic-homogenizer for 30 s. The capped tube was then centrifuged at 10000 g for 5 min. 100 µL of supernatant liquid was poured into a MonoSpin C18 mini-column and centrifuged at 3,500 g for 1 min. The eluate was injected into the HPLC system.  

 The ultrasonic-homogenization allowed the satisfactory extraction of melamine from samples with 100% water. The extract obtained was purified by subsequent centrifugal monolithic silica spin minicolumn: MonoSpin C18. The mini-column is a monolithic SPE column which is said to be excellent for small volume samples with easy and quick operation by centrifuge (19).  

The monospin method is not only used for deproteinization but can also be used for defatting. Since the recovery of the sample from the column varies with centrifugal acceleration (rotary speed), we investigated the optimum acceleration (> 1,000 g) to recover melamine from the spin mini-column. 

 In this test, blank milk sample extract processed with water was centrifuged as described earlier. The supernatant was then fortified with two-level melamine standard solutions and mixed. A 100 µL portion of the sample extract containing 0.25 µg or 0.5 µg of melamine was applied to the spin minicolumn. The centrifugal time was standardized at 1 min. The optimal acceleration that recovered melamine from MonoSpin C18 was at 3,500 g.  

There were no significant differences in data between two spiking levels (2.5 µg/mL and 5 µg/mL, n = 5 for each). The purification treatment by MonoSpin at 3,500 g for 1 min gave a satisfactory recovery (> 98 %) and reproducibility (RSD = 1.3 %, n = 10) for melamine. The time required for the sample preparation of a single sample was < 8 min. For sequential analyses, a batch of 24 samples could be prepared in 40 min. The eco-friendly procedure provided an easy-to-use and efficient purification and had a short operation time, resulting in high repeatability (Table 2) and great saving analysis costs. 

Table 2: Method validation data for melamine-fortified milk samples.			

In order to achieve separation with a 100% water mobile phase, this study tested eight types of non-polar sorbent columns. The physical and chemical specifications are listed in Table 1. We used 100% water as an isocratic mobile phase and examined column temperatures (≥20 °C) and flow rates (≥ 0.5 mL/min). The eight columns were compared and judged on how they separated between melamine and its interfering peaks and the sharpness of melamine peak obtained upon injection of equal amounts. The chromatographic separations obtained under these conditions are also presented in Table 1. 

Columns A–F had difficulty separating melamine and the interfering milk extract throughout the examined conditions. They separated melamine as a broadening or rounded peak. The complete separation of melamine and the interference peaks and its symmetrical peak was achieved with Column H using an isocratic mobile phase of water with a column temperature of 25 °C and a flow-rate of 1.0 mL/min.  

The melamine peak is clearly distinguished at 6.3 min (Figure 2). The present HPLC analysis accomplished optimum separation in a short time without the need for a gradient system to improve the separation and pre-column washing after an analysis. 

Figure 2: Chromatograms obtained from the present HPLC system. A milk sample spiked with melamine (1 ï¿½g/mL) (upper profile) and a blank milk sample (lower profile). Peak: 1, melamine (retention time = 6.3 min).			

Figure 2 shows that the resulting chromatograms were free from interfering compounds for quantitation and identification of melamine by HPLC, with the DAD set at 202 nm (giving the maximum absorbance for melamine).			

 The main validation parameters are summarized in Table 2. The linearity, accurary, precision and quantitation limit (QL) are well within acceptable criteria for the residue analysis that the Codex set up (20). The decision limit (CCα, α = 5%) and detection capability (CCβ, β = 5%) calculated in accordance with the EU regulation decision (2002/657/EU) are also displayed in this table. In the CCα and CCβ calculations, the Codex's maximum amount of melamine, 2.5 µg/mL in milk, equivalent to the FDA's safety limit (3,9), was taken as an established permitted limit (21). The other parameters are as follows: 

 The present HPLC system easily confirmed the peak identity of the target compound. Melamine was identified in the sample by its retention time and absorption spectrum. The separation, identification and quantification were achieved in a mere 6–7 min run without requiring a gradient system to improve the separation or pre-column washing after an analysis. The HPLC system did not need to use MS–MS–MS to identity the target compound.  

The robustness of the HPLC system was examined by using a spiked (2.5 µg/mL of melamine) milk sample obtained under the established procedure.  

Changes of ±5% units of the two chromatographic parameters, flow rate (1.0 mL/min) and column temperature (25 °C) were determined. The effects on the peak area and the validations in the retention time were evaluated. Changes of ±5% of the flow rate and the column temperature had no effect on the peak area, whereas the slight variations in the retention time were obtained with both parameters. Normal retention time for melamine was 6.3 min. At +5% flow rate, the retention time was decreased, and at –5% the time was increased. The decreasing and increasing retention times were 1.5 and 5.1%, respectively. By changing the column temperature by +5%, a decreasing retention time of 5.4% was obtained, however, no significant variations were observed with –5%. During these studies the target compound was separated.  

 The time and budget required for the analysis of one sample were < 15 min and roughly 4.72 Euro (¥500, as on 2 May 2012), respectively. These findings became a criteria in routine residue analysis. 

The present procedure was easy-to-use, rapid, spacesaving, and did not involve the use of organic solvents/reagents. It also resulted in high recovery and reproducibility with considerable saving of analysis time/cost. The procedure may be proposed as an international standardized method for routine residue monitoring of melamine in milk. 

The authors thank Mr Yoshimura (GL Sciences) for supplying MonoSpin C18 spin mini-column and Inertsil ODS-4 column. 

 (Doctor of Agriculture) is an associate professor with the Graduate School of Human Life Science, Osaka City University, Osaka, Japan. 

 is a food sanitation supervisor with the Okamura Oil Mill Co., Ltd., Kawahara, Osaka, Japan. 

(1) World Health Organization, Melamine contamination event, China, 2008, retrieved 2 December 2008, from 

http://www.who.int/foodsafety/fs management/infosan events/en/index.html

(2) World Health Organization, Melamine and Cyanuric acid: Toxicity, Preliminary Risk Assessment and Guidance on Levels in Food, 25 September 2008; updated 30 October 2008, from 

http://www.who.int/foodsafety/fs_management/Melamine.pdf

(3) HUFFPOST TECHNOLOGY, UN food safety meeting sets melamine limit, 6 July 2010, from 

http://www.huffingtonpost.com/huff-wires/20100706/un-un-melamine-limit/

(4) E.Y. Chan, S.M. Griffiths and C.W. Chan, 

(5) M.S. Filigenzi, B. Puschner, L.S. Aston and R.H. Poppenga, 

http://en.wikipedia.org/wiki/Melamine#cite_note-67

(7) Rory Harrington (15 April 2010), "EFSA cuts melamine TDI by 60 per cent". 

http://www.foodqualitynews.com/Legislation/EFSA-cuts-melamine-TDI-by-60-per-cent?utm_source=RSS_text_news

Articles

A novel "green chromatography" method for analysing melamine in milk is described. The method uses 100% water as the eluent and in the sample preparation stage and is reported to be a reliable and cost-effective method for laboratories in developing countries.

"Green" HPLC with 100% Water Eluent for Analysing Melamine in Milk

An excerpt from LCGC's e-learning tutorial on analytical method validation (AMV) at

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.  

 is the closeness of test results to the true value. For drug substances, accuracy measurements are obtained by comparing test results to the analysis of a standard reference material or to a second, well-characterized method. For drug products, accuracy is evaluated by analysing synthetic mixtures (containing all excipient materials in the correct proportions) spiked with known quantities of analyte. Guidelines recommend that data be collected from a minimum of nine determinations over at least three concentration levels covering the specified range. The data should be reported as the percent recovery of the known, added amount, or as the difference between the mean and true value with confidence intervals (such as ±1 SD). Acceptability criteria are defined by end users but rarely fall outside 97–103% of the nominal value. Statistical analysis can be applied using a one sample 

 measures the degree of agreement among test results when the method is applied repeatedly to multiple samplings of a homogeneous sample. Precision is commonly described in terms of repeatability, intermediate precision and reproducibility. Repeatability is investigated by analysing a minimum of nine determinations using the same equipment and sample, covering the specified range of the procedure, or a minimum of six determinations at 100% of the test concentration and reported as percent relative standard deviation (RSD). 

refers to the agreement among the results from a single laboratory, despite potential variations in sample preparation, analysts or equipment. 

 refers to the agreement among the results from different laboratories. Results are reported as % RSD, and the percentage difference in the mean values between the analysts must be within specifications. Less than 2% RSD is often recommended, but less than 5% RSD can be acceptable for minor components. 

(LOD) is the lowest concentration of an analyte in a sample that can be detected. The 

(LOQ) is the lowest concentration of an analyte in a sample that can be quantified with acceptable precision and accuracy under the stated operational conditions of the method. In a chromatography laboratory, the most common way to determine both the LOD and the LOQ is using signal-to-noise ratios (S/Ns), commonly 3:1 for LOD and 10:1 for LOQ. An appropriate number of samples must be analysed to fully validate the method performance at the limit. 

is the ability of a method to provide results that are directly proportional to analyte concentration within a given range. 

 is the interval between the upper and lower concentrations of an analyte that have been demonstrated to be determined with acceptable precision, accuracy and linearity using the method. The range is normally expressed in the same units as the test results obtained by the method (for example, nanograms per millilitre). Guidelines specify that a minimum of five concentration levels be used to determine the range and linearity, along with certain minimum specified ranges depending on the type of method. Data to be reported generally include the equation for the calibration curve line, the coefficient of determination (

 is a measure of a method's capacity to obtain comparable and acceptable results when perturbed by small but deliberate variations in procedural parameters; it provides an indication of the method's suitability and reliability during normal use. During a robustness study, method parameters (such as eluent composition, gradient and detector settings) are intentionally varied to study the effects on analytical results. Common chromatography parameters used to measure and document robustness include critical peak pair resolution (

) or peak width in gradient elution, retention time (

), peak area (and height) and concentration.  

(1) US Food and Drug Administration. Guidance for Industry, "Analytical Procedures and Method Validation." 

(2) International Conference on Harmonization, Harmonized Tripartite Guideline, Validation of Analytical Procedures, Text and Methodology, Q2(R1), November 2005. 

The Essentials

An excerpt from LCGC's e-learning tutorial on analytical method validation (AMV) at Chromacademy.com.

Analytical Method Validation

Will an internal standard always improve data quality? 

A reader recently contacted me after receiving reviewer comments for a paper that they had submitted for publication. The reviewer suggested that the author would get much better data quality if they used an internal standard with their liquid chromatography (LC) method. The reader wasn't sure how an internal standard would improve the results, because they had never used this method of calibration before, so they asked me for some help. I think this is a good opportunity for a general discussion on this topic, because even though many workers use internal standards on a daily basis, many others may never have the need or, as is the present reader's case, may not know enough about internal standardization to make an informed decision about its use. 

The concept of an internal standard (IS) is quite simple — you just add a known amount of the IS to every sample, both calibrators and unknowns, and instead of basing the calibration on the absolute response of the analyte, the calibration uses the ratio of response between the analyte and the IS.  

Most of us are familiar with how external standardization is used, but a review won't hurt. To construct a calibration plot, a series of calibration solutions are made containing known concentrations of reference standard. For the present discussion, let's refer to these as C

 for concentration of analyte, and make solutions at 0, 2 ng/mL, 4 ng/mL, 6 ng/mL, 8 ng/mL and 10 ng/mL. A series of chromatograms are run where the same volume of each solution (for example, 10 µL) is injected, and the response of the analyte (

) is recorded (usually peak height or peak area). If the responses for the respective solutions are 0, 2, 4, 6, 8 and 10 area units, we can construct a calibration plot, such as that shown in Figure 1. Each point on the calibration line corresponds to a calibrator concentration and its response, as illustrated for the 4-ng/mL calibrator and its 4 area-unit response (dashed line in Figure 1). Then samples of unknown concentration are prepared using the same sample preparation procedure, injected and the area of the resulting peak is measured, an area of 8, for example. This value is located on the vertical axis and a horizontal line is drawn to the right (dashed line) until it intersects with the calibration line. From this point, a vertical line is dropped to determine the concentration of analyte in the unknown sample (8 ng/mL in the present example). Of course, with a modern data system all this is done mathematically in the background, so manual plots such as that of Figure 1 are rarely constructed. 

 Figure 1: Calibration plots of concentration of analyte (CA) versus response (RA) for external standardization (plain text) and concentration ratio (CA/C IS) versus response ratio (RA/R IS) for internal standardization (labels in parentheses).			

For IS calibration, an IS is chosen (according to the criteria discussed below), and is added at the same concentration to every sample. For example, to generate an internally standardized calibration plot for the data discussed above, we might make a concentrated IS solution of 100 ng/mL of the IS. Then an aliquot of this IS solution would be added to a specific volume of a reference standard solution to obtain a known ratio of concentrations, 

. For example, 900 µL of a 4-ng/mL solution of reference standard plus 100 µL of the 100-ng/mL IS solution would yield 1 mL of a solution with a ratio of 0.36 

, which is equal to ([4 ng/mL × 900 µL]/1000 µL)/([100 ng/mL × 100 µL]/1000 µL). When this solution is injected, it will give a chromatogram with two peaks, one each for the reference standard and IS. The area ratio for these two peaks is paired with the concentration ratio to construct the calibration plot as in Figure 1, where the arrows highlight the 0.36 

 concentration ratio and its corresponding 0.36 

 response ratio (I've assumed the analyte and internal standard have the same response characteristics). Now, if the same sample preparation procedure is used to add IS to unknown samples, the 

 ratio can be used to determine the concentration ratio (arrows for 0.8 

 in Figure 1). Because the concentration of IS (

) added to the sample is known, the concentration of analyte in the sample (

Next, let's see when an internal standard is and is not effective at improving method performance. In particular, we're interested in the accuracy and the amount of uncertainty (imprecision) of the measured result for a sample. We'll consider three different cases as illustrations. 

If the sample preparation is quite simple and the LC equipment, especially the autosampler, is working very well, an IS may not add any benefit. For example, if a liquid sample is taken and diluted 1:10 for injection using volumetric pipettes and glassware, there isn't much that can go wrong. In such cases, chromatograms may appear like those in Figure 2. In these partial chromatograms, the analyte peak (A) is eluted first, followed by the IS peak, with the areas and peak-area ratios noted in the caption. You can see that the IS peak size is almost identical for all three samples, as might be expected with such a simple sample preparation procedure and a high-precision autosampler. For this kind of sample, external standardization would be preferred for several reasons. First, it is more convenient and less expensive because the extra step of adding the IS during sample preparation is eliminated. Second, the chromatogram is simpler, so there will be less concern about interfering peaks that might compromise the results. Third, the data are easier to process because there is only one peak to measure in each chromatogram. And fourth, the uncertainty of the results may be smaller because the variability in IS addition and IS peak measurement is eliminated. 

Figure 2: Partial simulated chromatograms for an analyte (A) and internal standard (IS). Area and response ratios (RA/R IS) of (a) 6012/10056 = 0.6, (b) 7995/9996 = 0.8 and (c) 9997/10040 = 1.0.			

An internal standard will benefit the method most commonly when there are multiple sample preparation steps, especially when volumetric recovery at each step may vary. (In the past, internal standards sometimes were used to correct for variability in injection volume, but today's autosamplers generally have a volumetric imprecision of <0.5%, so such corrections are not necessary.) Consider the case of a method used for determination of a drug in plasma using a liquid–liquid extraction technique for sample preparation. For such cases, smallvolume samples are used to minimize sample collection issues, reduce reagent expenses and simplify the handling of perhaps hundreds or thousands of samples in a drug development study. Let's say that 10 µL of IS solution is added to a 200-µL plasma sample, vortexed, and mixed with 100 µL of high-pH buffer. Next, 100 µL of this solution is extracted with 500 µL of methyl-

-butyl ether (MTBE). The MTBE phase is removed, evaporated to dryness and the residue is reconstituted in 50 µL of injection solvent, followed by injection of 10 µL.  

There are several steps where small errors may increase the variability of results. Any error in addition of the buffer will be passed along to the exaction step when an aliquot of the sample-buffer mixture is taken. When the MTBE layer is removed, typically a little MTBE is left behind so that none of the aqueous phase is included; the amount of MTBE recovered therefore will vary slightly. Evaporation and reconstitution offer additional opportunities for variability in the process. The net result is that the absolute fraction of the original sample that ends up in the injection vial will vary much more with this multistep procedure than with the simple dilution used in the previous example. If an IS is used, it should be added as early in the process as possible, then the subsequent volumetric losses will be compensated.  

Consider the example of Figure 3, where three replicate preparations were made of a homogenous sample. Because of sample loss during sample preparation, the absolute peak areas vary by 40% between the samples, but the 

 ratios are constant, so imprecision is low even with widely varying peak sizes. In other words, because we're interested only in the ratio of peak sizes, the IS technique corrects for variability that would be difficult to control otherwise. 

Figure 3: Partial simulated chromatograms, as in Figure 2 for three replicate extractions of a homogenous sample. (a), (b) and (c) each have RA/R IS = 1.0.			

There are cases, however, where the use of an internal standard may not only be ineffective, but may also be misleading. An example of this is shown for another set of three replicate samples in Figure 4, similar to those of Figure 3. In Figure 4, you can see that the peak size of the IS peak and the 

 ratio varies significantly. If the IS were working properly, the IS peak size wouldn't matter, because the response ratio would be constant. If the IS were not needed, the analyte peak response should be constant. 

Figure 4: Partial simulated chromatograms, as in Figure 3 for three replicate extractions of a homogeneous sample.RA/R IS for (a) = 0.6, (b) = 1.0 and (c) = 0.8.			

When we see a problem like this, the IS procedure is not working properly, usually because of the way the IS is used or because an inappropriate IS was chosen (see discussion on how to choose an IS further on). If we assume that the IS was selected correctly, what could cause the problems of Figure 4? The basic assumption with internal standardization is that once the IS is in the sample, any losses should affect the IS and analyte proportionally, so the ratio of analyte to IS stays constant. This means that most likely the problem is related to something that happened to the sample before or during the addition of the IS. For example, if the sample was inappropriately assumed to be homogeneous before aliquotting and adding the IS, the IS would not correct for lack of sample homogeneity. 

On the other hand, if the IS were not added properly, problems could occur. I saw this once in a client laboratory when the mechanical pipette used to add the internal standard was out of calibration and not reproducible. Tracking down such problems can be tedious, but by breaking the process into steps and using matrix-free samples, you can analyse the solution at each step in the process and isolate the problem step. 

If it is determined that an IS is likely to improve the data quality, a suitable compound needs to be chosen for the IS. The sidebar contains a list of characteristics that are often cited as requirements for a good internal standard. Let's take a moment and consider each of these characteristics. 

 This seems logical — after all, if the internal standard is always or occasionally present in untreated samples, how can you tell the difference between the response of the native IS and the added IS? Although absence of the IS is an ideal, if the amount of IS present in normal samples is much less than the amount of added IS, it is unlikely that there will be a problem. For example, if the native IS peak is never more than 2% of the added IS peak, the results will probably show no adverse effects. 

 The chromatographic system needs to be able to independently measure the size of the analyte and IS peaks. For most detectors, this means that there should be baseline resolution between the peaks. For liquid chromatography–mass spectrometry (LC–MS), however, the added selectivity of the MS detector will often allow independent measurement of the analyte and IS if they do not have the same molecular weight. This is one reason why stablelabel, or isotopically labeled, versions of the analyte are used with LC–MS methods — the peaks can be coeluted, yet they can be separately measured by the detector. 

 This is a desirable characteristic rather than a necessary one. If the IS elutes after the analyte, it can offer additional information on the quality of the separation. For example, if the retention time of the IS is correct, you know that the chromatographic conditions were correct up to that point in the chromatogram and you can be confident in the retention times of all earlier-eluted peaks. On the other hand, if the IS comes out before the analyte, you may know that everything up to that point in the chromatogram was ok, but something could have happened immediately afterward to affect the remaining peaks. 

At a minimum, the IS needs to be sufficiently stable so that it does not degrade during the sample preparation and chromatographic analysis processes. Although not essential, more stability will mean more convenience, so that stock solutions will last longer, refrigeration or freezing may not be needed and so forth. 

 This requirement is a bit misleading. It would be better to state that any impurities present will not be coeluted with the analyte or otherwise interfere with the process. For example, an IS with 80% purity could be used if the 20% impurity did not interfere with the method. Higher-purity internal standards are usually preferred, but 100% purity is not required, and in some cases is very difficult to obtain. For example, in the case of stable-label or isotopically labeled internal standards that are common with LC–MS methods, 100% purity is almost never the case. The reason is that it is nearly impossible to synthesize a stable-label IS without at least some unlabeled analyte present. As long as the amount of unlabeled material is so small that it does not affect the results, it will not matter. The rule of thumb for the analysis of drugs in biological matrices, such as plasma, is that the unlabeled analyte from the IS solution present in the sample needs to generate <5% of the response of the analyte at the lower limit of quantification (LLOQ). With this small amount of unlabeled analyte present, the performance of the method will not suffer. 

Adequate detector response is certainly necessary. However, the response does not have to be the same as the analyte in terms such as response/gram or detection wavelength, but it does need to be easily detectable. 

It is often stated that the IS needs to have a close chemical relationship to the analyte of interest. This really is not necessary — if benzene has all the other characteristics needed, it might be an adequate IS for a drug analysis method. However, because the IS needs to have similar extraction characteristics, retention times, stability and detector response as the analyte, it is highly likely that it will be a compound of similar structure. Most internal standards are existing compounds with close structural relationships to the analyte, specially synthesized modifications of the analyte or stable-label compounds for LC–MS, where several carbon-13 or deuterium atoms are substituted into the unlabeled analyte. 

We have seen that the addition of internal standards can be a useful tool to help improve the precision and accuracy of some LC methods, but not in every case. In the example of Figure 2, we saw that the IS added no benefit to the method, so external standardization is likely to be more convenient and less expensive. In the second case (Figure 3), the IS did exactly what it was supposed to, correcting for uncontrolled sample losses during sample preparation; this method benefits by the use of an IS. In the final example (Figure 4), the IS did not correct for method variability. This may be caused by a poor choice of IS, faulty equipment, addition of the IS after sample variability is influenced or some other problem that will need to be isolated. Whether or not an IS is to be used for this method, work needs to be done to isolate and correct the excess variability or the analytical results will not be very useful. 

Even if you choose to use an IS, I suggest that you run at least one batch of validation (or prevalidation) samples where the data are worked up using both external and internal standardization. Let the data help you decide which technique gives better method performance. 

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 

LC Troubleshooting

Will an internal standard always improve data quality?

When Should an Internal Standard Be Used?

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 (

U.S. Pharmacopeia and American Herbal Pharmacopeia

); 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 post-chromatographic 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 is often 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 sevenmobilephase 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. 

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

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 

After drying the developed plate, compound zones are detected on the layer by their natural colour, natural fluorescence under 366nm UV light, quenching of fluorescence under 254nm UV light on an "F" layer containing a phosphor (termed an UV indicator), or as coloured, 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 vapours (for example, iodine). Chemical detection reagents can be nondestructive universal (iodine), destructive universal (for example, charring with sulphuric 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 analyse 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 colour, or scan the TLC plate to document it. However, a commercial system for photography is needed if documentation of a combination of coloured, fluorescent and quenched zones is required. Camag, Desaga and Analtech all offer photodocumentation systems.  

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 headbased 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 vs. 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 highperformance 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 sulphate) 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 colour 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 NanoSILHPTLC plates (Macherey-Nagel) have enhanced brightness because of increased levels of the F

The differences between TLC and HPTLC can be seen in Figure 3 and Table 1. 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' websites (EMD/Millipore, Macherey-Nagel and Analtech). 

Table 1: A comparison of the physical and operational parameters of TLC and HPTLC.			

Plates with Channels and Preadsorbent Zone

Kieselguhr (diatomaceous earth) and silica 50000 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. 

Figure 4: Cutting tools for scoring and breaking glass TLC plates: (a) pistol-grip glass cutters and (b) grozier pliers. These items are available from hobby or stained glass stores. Images from author’s collection (F.R.).			

Silica gel (silica) is by far the most frequently used layer material in TLC. The structure is held together by bonded silicon and oxygen (siloxane groups), and separations take place primarily because of differential migration of sample molecules caused by selective hydrogen bonding, dipole–dipole interaction and electrostatic interactions with surface silanol groups (Si-OH). The intensity of these adsorption forces depends on the number of effective silanol groups, the chemical nature of the sample molecules to be separated and the elution strength of the mobile phase.  

Column Watch

The stationary phases currently in use for modern thin-layer chromatography (TLC) are reviewed.

Stationary Phases for Modern Thin-Layer Chromatography

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

In part 1 of this series, we introduced the concept of lipidomics as a new research strategy for analysing lipids using state-ofthe-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 analyse 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 analysers — that are more appropriate for answering the biological questions posed. In the second and final instalment of this series, we present and discuss the use of those technologies for lipid analysis.  

The high sensitivity of MS offers less labour-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 can also 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 phenylsulphonic 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 postionization 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 muchused 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" (MDMSSL).  

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

), determined on high-resolution mass spectrometers such as time-of-flight (TOF), Fouriertransform 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:  					

: 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. 

 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. 

: 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 analyser (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.

 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 

  = 97.0 (sulphate) can identify the [M-H]

  ions of sulphatides whereas neutral loss scans for 

Innovative technologies are rapidly helping to measure the numerous and diverse lipids present in biological samples. In this second and final instalment of the series, the authors present and discuss the use of these technologies for lipid analysis.

LCGC Europe-06-01-2012

"Green" HPLC with 100% Water Eluent for Analysing Melamine in Milk

Analytical Method Validation

When Should an Internal Standard Be Used?

Stationary Phases for Modern Thin-Layer Chromatography

New Frontiers for MS in Lipidomics (Part 2)

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