Asymmetrical flow field flow fractionation (AF4) has great potential for characterizing intermolecular interactions, such as complexation and aggregation, in multifunctional polymers. High molar mass aggregates or polymer–drug complexes are separated with low or no destruction by shear forces. AF4 combined with light scattering detection leads to information on the scaling properties of these species. A particular advantage of this method is the filtering effect of the membrane, which enables quantitative characterization of the complexation. Complexation and encapsulation properties of dendritic materials are important for medical and pharmaceutical development of drug delivery systems.  

Aggregation, complexation and encapsulation are important processes in the development of drug delivery systems for biomedical and pharmaceutical applications. Dendritic polymers are promising materials for such applications because of their extremely broad number of functionalities, which can be modified arbitrarily (1). They also have threedimensional architectures with a high number of peripheral groups that contain different cavities within the dendritic scaffold (Figure 1). 

Figure 1: Schematic representation of two types of dendritic polymers. Dendrimers are synthesized stepwise and have a globular, perfectly branched structure formed by dendritic and terminal units. Hyperbranched polymers are the product of a one-pot reaction leading to statistical type of branching with a number of linear together with dendritic and terminal units.  

Dendritic polymers have been intensively investigated for their potential to transport bioactive molecules and metal ions (2–4), and also as artificial enzymes (5), biomimetic structures (6) and templates for nanoparticles (7). Higher generation dendrimers that are modified to obtain a soft core and a hard shell can be used as dendritic boxes for guest molecules (8). Furthermore, polymers with a hydrophobic core and hydrophilic shell can carry hydrophobic substances under physiological conditions (9). The formation of dendritic complexes with guest molecules promises long-term drug delivery. However, the encapsulation and release are dependent on the type of modification of the endfunctionalities. In all these applications biocompatibility and non-toxicity should be seriously considered and this is often achieved by modifying with sugar, polyetheleneglycol or oligo-peptide units. For example, polyglycerols with a core-shell structure can carry paclitaxel and doxorubicin molecules, which are used in the cancer therapy (4). However, the encapsulation properties are often investigated using model guest molecules, such as Rose-Bengal, because they cost less and are easier to detect (10,11). By varying the concentration, temperature or pH value it is possible to control the release of the bioactive substances. 

Reliable investigation of these complex structures is essential but not easy for various reasons. Light scattering or microscopy techniques deliver only average values on sizes. Big aggregates could be easily destroyed by shear forces when using size exclusion chromatography (SEC) or are too big for separation and size determination using this technique. Qualitative determination of complexes by ultraviolet-visible (UV-vis) measurements does not lead to precise results because of their chemically complex coreshell architectures. The quantitative characterization of encapsulation and complexation of model substances can be made by liquid chromatography (12,13), spectroscopy (14) or electrophoresis (15). In most of the cases, UV-vis of the absorption shift of the guest molecules after complexation is used in batch (16) or coupled to SEC (12). Both methods are limited: UV-vis does not lead to correct results because of the very complex chemical structure of the polymer carrier. The separation of the complex by SEC could lead prejudicially to strong interactions with the column material.  

The separation of polymer complexes or aggregates leading to information on their distribution by size and complexation capacity is essential for their later medical use. Therefore, a gentle separation technique with reduced interaction is required. Asymmetrical flow field flow fractionation (AF4) has the potential to meet all these requirements. AF4 separates the sample along a thin and long channel, in which a laminar flow drives the molecules or particles. The interplay of both diffusion and cross flow field leads to the distribution of the molecules within the laminar flow according to size, density or surface properties, leading to their separation (17).  

AF4 has been used to successfully separate biopolymers (18), nanoparticles (19), proteins (20) and viruses (21). Recently, in the field of biocomplexes, characterization of the distribution of triglycerides and cholesterol in plasma proteins was reported (20, 22). The potential of AF4 for complex characterization is huge.  

In this work our recent developments in the characterization of dendritic biocomplexes by AF4 will be demonstrated to investigate the aggregation of dendronized biomolecules and of the quantitative determination of guest molecules in a dendritic complex. Both topics are based on dendritic structures with a significant number of end functionalities which are modified with oligosaccharides.  

The dendronization of a linear polymer chain leads to high concentration of functional groups along the linear backbone. In contrast to dendrimers, where dendrons are coupled to a core and the size and functionality of the molecules are limited, dendronized linear structures enable the engineering of a portfolio of nano-objects varying in length, branch point frequency and dendron generation. This huge functionality is suitable to control complexation and to tune aggregation by varying the conditions within the physiologically relevant values (23). The effective control over functional activity needs clarification of the conditions influencing the aggregation, that is, the concentration, generation number and the pH value. The determination of the aggregation mass and size distribution, as well as shape analysis, were possible by AF4. 

The same parameters are responsible for the encapsulation and release properties of the dendritic host. However, the amount of the complexed guest molecules in these systems should be determined considering their size, which could be in strong contrast to the host. Additionally, the stability of the complexes could be limited. These facts call for hyphenation of techniques allowing simultaneous, fast and smooth complex quantification. In this work a simple method to determine the non-bounded guest molecules by filtration through the AF4 membrane and calculation of the size of the complex by fractionation in the AF4 channel is demonstrated. 

All AF4 measurements were performed on an Eclipse 3 system (Wyatt Technology Europe, Dernbach, Germany). A poly(tetrafluoroethylene) (PTFE) channel spacer was used. The thickness of the channel was 350 µm and the channel dimensions were 26.5 cm length and 2.1–0.6 cm width. The membranes used in the accumulation wall and filtration device had different molecular weight cut-offs (MWCOs) of 5 kDa and 10 kDa and were made of regenerated cellulose. Flows were controlled with an Agilent Technologies 1200 series isocratic pump (Agilent Technologies, Wilmington, DE,s USA) equipped with a vacuum degasser. The detection system consists of a variable wavelength UV-Vis detector (UV) (Knauer, Berlin, Germany) tuned at 550 nm, an RI-detector Dn 2010 (WGE Dr. Bures, Dallgow-Doeberitz, Germany, λ = 620 nm, 25 °C) and a three-angle laser light scattering detector (MiniDAWN from Wyatt Technology, Santa Barbara, CA, USA). The UV detector was connected to the cross-flow outlet to determine the amount of free dye. The channel flow rate was maintained at 1.0 mL/min for all AF4 measurements. Samples were injected during the focusing/relaxation step with 0.2 mL/min during 2 min. 

For dye filtration the focus flow was set to 3 mL/min for 20 min. The fractionation of the complex within this measurement followed in elution mode with linear cross flow gradient from 2 mL/min to 0 mL/min in a period of 20 min. For the determination of the molar mass of the complex focusing at 1.5 mL/min focus flow over a time period of 2 min and elution under a linear cross flow gradient from 2 mL/min to 0 mL/min for 20 min was performed. In the same way the separation and detection of the aggregates of dendronized polymers was performed. However, exponential cross flow gradient was applied.  

For dye@polymer complex characterization deionized, UV-treated and ultra-filtrated pure water containing 0.02% sodium azide (v/v) was used. For the characterization of the aggregation of dendronized polymers, an additional pH adjustment in NaNO

The dendronized glycopolymers are composed of a linear maleimid (MI) backbone at which lysine-dendrons are coupled with a nearly complete modification of every MI unit. The dendrons are in three generations, whereas the number of the end functionalities increases with the generation number. The end-functionalities of the dendronized polymers are then completely modified with maltose (Figure 2) (24). The Maltose-modified end-functions lead to pronounced aggregation. The aggregation ability is amplified at higher concentrations in water. The generation number strongly influences the aggregation too because when the generation number increases the amount of end groups for each of the molecules increases substantially. 

Figure 2: Schematic representation of dendronized, maltose-modified polymer of third generations.  

This property is also dependent on their concentration in a water solution. Additionally, the aggregates are very sensitive towards pH changes and this property should be easy to adjust for their future application conditions. The investigation of the aggregates and their distribution can be easily performed by AF4. Other common techniques, such as SEC, are not suitable because the molar mass of the dendronized polymers is extremely high. Another difficulty is their strong affinity to the stationary phase because of their strong polar functionality. Additionally, shear forces formed during elution through the column could destroy the aggregates and cannot lead to reliable data. In contrast to this the shear forces in AF4 are strongly reduced. The lack of stationary phase reduces undesirable interactions and allows for easy change of the pH of the eluent. This enables systematical aggregation study at variable conditions (pH, concentration and generation number of the dendrons).  

Systematic investigation of concentration, pH value and generation number showed that the first generation dendronized copolymer does not show significant aggregation, whereas the third generation starts to aggregate at very low solution concentrations (Figure 3). 

Figure 3: Light scattering signal at 90Â° of AF4 separation and Rg/elution volume plot of three generations of the dendronized polymers at pH 3.5 and c = 1 mg/mL.  

Further analysis of the size of the molecules using static and dynamic light scattering indicated a strong dependence of the size of the species on the generation number. The pH dependence of the radius of gyration and the hydrodynamic radius of the aggregates is clearly visible in Figure 4, where the behaviour of the third generation dendronized MI is shown. Whereas big aggregates are formed at acidic conditions, at pH value of 7 the aggregation is strongly reduced and is increasing again at higher pH.. Further interpretation of the molar mass and radius of gyration combined with AFM measurements showed that stiff, laterally ordered aggregates are formed under these conditions. 

Figure 4: pH dependent aggregation of the third generation dendronized MI at c = 0.5 mg/mL. Rg (z-average) determined by AF4-MALLS and was Rh determined by dynamic light scattering.  

Complexation Characterization of Dendritic Glycopolymers

Oligosaccharide-modified hyperbranched poly(ethylene imine) (PEI) possesses strongly reduced toxicity compared to the non-modified PEI and high ability to carry drug molecules (25). Dendritic polymers with an oligosaccharide shell can therefore form host-guest systems with dye molecules as models system to study drug delivery ability (Figure 5). 

Figure 5: Schematical representation of encapsulation and release of RB by PEI-Mal.  

Core-shell molecules of hyperbranched PEI as a core and oligosaccharide shell were mixed with Rose Bengal (RB) resulting in dye@PEI-Mal complexes (25, 26). Their complexation properties are strongly dependent on the pH value which was found by the shift of the UV-vis absorbtion maximum of RB: a typical indication for interaction of the dye. Commonly, the separation and determination of the free dye from the complex is performed by means of dialysis, ultra-filtration or ultracentrifugation followed by a UV-vis analysis in batch. However, for this specific system these methods provide nonquantitative information about the complexation capacity as a result of different possible chemical moieties binding the dye such as PEI or maltose, which does not allow for reliable calibration of the UV-vis signal shift of the RB. Therefore, we considered the ultra-filtration effect of AF4 as a suitable alternative for this quantification. During the focusing phase ultra-filtration is taking place, filtering the species smaller than the cut-off of the membrane (Figure 6) (26). During this step the non-bound Rose Bengal is flushed out from the channel while the complex of Rose Bengal and PEI-Mal remains inside. 

Figure 6: Filtration of free RB and separation from the complex RB@PEI-Mal.   

The quantitative analysis of the free dye filtrated through the membrane requires few optimization steps. The fact that RB interacts with the membrane of regenerated cellulose could appear as a significant problem during the experiments. Our membrane tests showed that the modification of the membrane with 250 µg RB is sufficient for the formation of stable, layer allowing for quantitative analysis over several days. Under those conditions the calculation of the filtrated dye was made by calibration of a UV-vis signal from a detector connected to the waste line of the cross flow. Further parameters, for example, membrane material, molecular weight cut-off and stability of pure RB and RB@PEI-Mal complexes, were investigated and verified, showing high reproducibility of the performed measurements as shown in Table 1. 

Table 1:  Recovery test for the quantitative determination of RB calibration (UV-vis detection).  

The development of the quantitative determination of free RB allowed us to determine the complex capacity of the PEIMal molecules. Figure 7 represents the data obtained for different mixtures of RB and PEI-Mal. The amount of the RB in the complex was determined in two ways: 

Figure 7: RB complexation capacity of PEI-Mal determined by calculating the free RB (red symbols) and by the molar mass increase of the complexes (blue crosses). RB withMn = 1018 g/mol and PEI-Mal with Mn = 200,500 g/mol.  

Subtracting the free RB detected after filtration during the focusing from the RB in the injected mixture.

Increasing the molar mass of the complex compared to the pure PEI-Mal, which was determined by AF4 separation in the channel. 

The development of the molar mass distributions depending on the PEI-Mal:RB ration in the mixture is shown in Figure 8. The complexation studies based on both those methods are in very good agreement. The capacity of the complexes was found to reach 271 RB guest molecules for each host PEI-Mal. This value was supported by additional experiments using other ultra-filtration techniques.  

Figure 8: Differential molar mass distribution of free PEI-Mal (black) compared to complexes RB@PEI-Mal with initial ratio of 1:41 (red), 1:96 (green), 1:246 (blue), 1:657 (cyan) PEI-Mal:RB.  

The AF4 technique is suitable for comprehensive analysis of intermolecular interactions in multifunctional polymers as aggregation and complexation with guest molecules. Effective molar mass determination of these polymers is possible as a result of reduced interactions with channel material leading to full size separation. The investigation of the aggregation behaviour of big polymers is advantageous, mainly because of weak shear forces during the separation and the possibility to couple this technique to size sensitive detectors. Furthermore, the ultra-filtration effect of AF4 can be effectively used for quantitative determination of the amount of complexed small guest dye molecules in core-shell polymers. This feature of AF4 has a big potential for upgrading to separation and quantification of drugs encapsulated in polymers and makes the AF4 technique very promising for the analysis of drug delivery systems. 

 received her PhD in Physical Chemistry of Polymers in 1999 from Max-Planck-Institute of Polymer Research and University Mainz. She has been leader of the Polymer Separation Group at IPF Dresden since 2007. The main focus of her research is molecular properties of dendritic and multifunctional polymers and the development of new separation methods for branched polymers.  

 joined the Polymer Separation Group at IPF Dresden in 2006 after studying Chemical Engineering at the University of Applied Sciences in Dresden. She is finishing her PhD thesis devoted to the separation of branched and functional polymers using FFF and alternative chromatographic approaches. 

 A. W Bosman, .H. M Janssen and E. W. Meijer, 

Articles

Describes how asymmetrical flow field flow fractionation (AF4) can be used to characterize intermolecular interactions in multifunctional polymers used in drug delivery.

Asymmetrical Flow Field Flow Fractionation for Investigating Intermolecular Interactions of Multifunctional Polymers

This is the third instalment in a series focusing on some of the basic principles of troubleshooting liquid chromatography (LC) methods. First, we looked at some general practices for troubleshooting any LC problem (1). Then we looked at problems whose symptoms are related to pressure changes (2). This month, we'll concentrate on problems exhibited as abnormal retention times. As a means to organize the discussion, let's look at situations where retention times are too long, too short or inconsistent. 

Before we look at specific problems, let's take a moment to consider the things that influence retention. We can categorize these as the mobile phase, the stationary phase (column), the hardware, the environment and the sample. Let's simplify this discussion and assume that nothing has happened to the sample, such as degradation or other chemical changes.   

The mobile phase can change because of an error in formulating it, such as a mistake in volumetric measurement or adjustment of the pH. If an error in formulating the mobile phase is suspected, it is best to make a new batch to see if it fixes the problem. Some mobile phases can change over time because of chemical degradation, selective evaporation of one component or microbial growth in highly aqueous mobile phases. Again, reformulation is the best way to verify this problem source. Most of us use on-line mixing to prepare isocratic mobile phases. An error in instrument settings or hardware failure can be the cause of errors in on-line mixing. Check for proper degassing and pump operation, as well as the correct control-programme settings. Sometimes hand-mixed mobile phases can be used as a check for on-line mixing, or alternate mixing channels can be used for both isocratic and gradient methods (for example, use the C and D solvent reservoirs instead of A and B in a four-solvent LC system). 

The stationary phase in the column has a finite lifetime, generally in the 500–2000 sample range (or more), depending on the nature of the sample. Every column will die eventually, and some methods are harder on columns than others. For example, mobile phases outside the pH 2–8 region accelerate the degradation of silica-based columns. Some column types have shorter lifetimes than others. For example, cyano and amino columns are unlikely to last as long as C8 or C18 columns, which tend to be quite robust. In addition to changes in retention, column failure usually is accompanied by a rise in system pressure and an increase in peak tailing. Replacement of a suspect column with a new one is the easiest way to check for column-related problems. 

System hardware problems that generate symptoms of changed retention most commonly are associated with pump malfunctions or leaks. Pump problems can be checked with a simple flow-rate measurement with a stop watch and volumetric flask. A secondary symptom of pump problems may be high, low or fluctuating pressure. Low flow may be associated with faulty check valves, worn pump seals, air bubbles in the pump or errors in pump settings. Cleaning, component replacement or degassing should correct such problems. High flow rates usually are a result of improper settings. 

The most common environmental cause of retention changes is a change in column temperature. This effect is common if the column oven is not used or is not working properly. Methods that operate under ambient conditions are highly susceptible to failure, especially if the laboratory temperature is not well controlled. In my travels I have encountered laboratory temperatures ranging from 10 °C (central China in January) to 35 °C (Tel Aviv in June). If we use the rule of thumb that retention can change by 2% with each 1 °C change in temperature, you can imagine the result if the same method were run in both of those laboratories under ambient conditions! Use the column oven and make sure that it is operating properly. 

One tool that can be very useful in diagnosing the source of retention problems is the retention factor (also called the capacity factor, 

) is the column dead time, usually measured by the first disturbance in the baseline (the "solvent front"). Another useful calculation is the selectivity, or relative retention, α 

-values for the first and second peaks of an adjacent peak pair, respectively. 

Notice that changes in flow rate, whether intentional or due to a leak, will change both 

 will remain constant for such changes. On the other hand, chemical changes will change only 

 value is changed, too. Generally, when the 

 value is changed it does not change exactly the same for all peaks in the chromatogram. One way to confirm chemical changes in the system is to check the α value; if α changes, a chemical source of the problem is most likely. Because 

 and α are so useful in distinguishing between flow-related and chemical changes, it is a good idea to make these measurements a part of the process for troubleshooting retention-time problems. 

-values stay constant, a flow-rate problem is indicated. Double-check the flow-rate setting to be sure you didn't make a mistake. Leaks and pump problems are the two most common remaining causes. Check for leaks throughout the system; these may or may not be accompanied by low system pressure, depending on the magnitude of the leak. Several possible problems related to the pump could be sources of increased retention. Air bubbles in the pump will also cause pressure fluctuations; thorough degassing of the mobile phase and purging the pump should correct such problems. If problems persist, check to be sure the frit in the mobile phase reservoir is not restricting flow to the pump. Faulty check valves or pump seals also can result in low flow and long retention times. Sonication of check valves will usually restore their function. Pump seal leakage often is accompanied by liquid dripping from the drain hole just behind the inlet check valve on most pumps. Check the maintenance records — if the pump seal has been in use for a year or more, replace it. 

 values (and often α) is observed, you have evidence that a change in system chemistry is responsible for an increase in retention. The easiest way to check this is to make a new batch of mobile phase. If this does not correct the problem, replace the column. 

A final possible source of increased retention is a drop in the column temperature. As mentioned above, a 2% increase in retention for a 1 °C drop in temperature is common. Based on the magnitude of the retention change, you can estimate the temperature change and see if it is a reasonable cause of retention. Has the oven failed, did you forget to turn it on or are you relying on ambient operating conditions? Any of these sources can account for increased retention. 

When retention times are smaller than they normally appear, a flow-rate change is highly unlikely, unless you made an error in one of the settings. This is because decreases in flow due to leaks or other malfunctions are not uncommon, but there are no corresponding causes that result in higher-than-normal flow rates that are necessary for reduced retention. 

As with retention times that are too long, do a stepwise elimination of problem sources by first making up a new batch of mobile phase. If this approach doesn't fix the problem, replace the column. Temperatures that are higher than normal will cause a drop in retention; the sources of temperature problems are the same as for excess retention. 

Usually, an increase or decrease in retention will not be an abrupt change. If it is, the cause is likely related to operator intervention, such as improper formulation of a new batch of mobile phase, installing the wrong column or changing a column-oven setting. More commonly, retention will gradually increase or decrease over tens, hundreds or thousands of samples, or it cycles over a 24-h period. Cycling retention is commonly correlated with ambient methods and a laboratory temperature that changes throughout the day and night. 

Retention drift that occurs over hundreds or perhaps thousands of injections is most likely because of normal column aging. Drift from column aging usually will be accompanied by a gradual increase in pressure and an increase in peak tailing. Often, a shift in relative retention also will be observed when α-values are calculated. If the column is suspected, replace it to see if the problem is corrected. 

Shorter-term retention drift may be caused by the mobile phase. Although fairly rare, it is possible to selectively evaporate a volatile component of the mobile phase, especially if helium sparging is used for degassing. Retention drift resulting from a pH shift can occur if the buffer is outside its optimal buffering region, generally ±1 pH unit from its p

a. The use of volatile buffers, as is common with LC–mass spectrometry (MS) mobile phases, may shorten the stable lifetime of the mobile phase, so daily reformulation may be a wise practice. Make up a new batch of mobile phase if the mobile phase is suspected, and be sure to adjust the pH before any organic solvent is added. 

To avoid retention problems, make sure to keep the instrument in good operating condition by servicing it regularly. A routine preventive maintenance session should be done on an annual basis at a minimum, and more often for heavily used LC systems. 

Because column temperature changes can have such a profound influence on retention time, it is wise to always use a column oven. Many ovens do not control the temperature well near room temperature; a good practice is to use a minimum operating temperature of 30–35 °C so that good temperature control is ensured. It may take 30 min or longer for the column oven to reach a stable temperature. Be sure to use the solvent preheater that is included with most column ovens. The most common preheater implementation is a piece of capillary tubing that is embedded in the aluminum block of the oven. 

Columns will usually last for more than 1000 injections. When this number of samples has been analysed, the cost-per-sample for the column is less than 5% of the overall per-sample cost of analysis. My feeling is that at this point it isn't worth my time to do anything more than flush the column with strong solvent (for example, acetonitrile or methanol); if this doesn't restore the column, replace it. Guard columns or sample preparation will both extend the column life, but they have their associated costs, which may make the economics of their use questionable. A 0.5-µm in-line filter between the autosampler and column will help keep particulate matter from blocking the column inlet frit, but it will not influence retention-related problems. Another good practice for extending column life is to use a single column for each method. When the same column is used for multiple methods, sometimes unimportant sample components from one method will interfere with another method. 

Mobile phases have finite lifetimes too. My recommendation is to replace any buffer at least once a week and organic-based mobile-phase components at least monthly. It is a good idea to replace the reservoir with a clean one whenever the mobile phase is replaced so that prior contamination doesn't get transferred to the new mobile phase.  

If you pay close attention to patterns in retention changes, correlations with mobile-phase use and column history, you can establish expected replacement cycles for each component of each method. After such patterns are defined, you can put in place preventive-maintenance and component-replacement practices that will help you avoid most retention-related problems. Armed with an understanding of which variable most strongly influences retention in your particular method, you'll be able to more quickly identify and correct problems when they occur. 

, is vice president of LC Resources in Walnut Creek, California, USA. He is also a member of LCGC's editorial advisory board. Direct correspondence about this column should go to "LC Troubleshooting", LCGC Europe, 4A Bridgegate Pavilion, Chester Business Park, Wrexham Road, Chester, CH4 9QH, UK or e-mail the editor, Alasdair Matheson, at 

What causes peaks to appear where they do not belong?

Troubleshooting Basics, Part 3: Retention Problems

In this instalment of "Sample Preparation Perspectives," Norwegian authors from the University of Oslo describe the practical aspects of hollow fibre liquid-phase microextraction in the three-phase mode (HF

 LPME). The guest authors highlight important practical issues related to the supported liquid membrane, the hollow fibre and the extraction itself. They also discuss practical work with electromembrane extraction (EME), which is related to HF

 LPME but uses an electrical potential as the driving force for the extraction.

This month's instalment describes practical aspects of hollow-fibre liquidphase microextraction in the three-phase mode (HF

 LPME is a microscale sample preparation technique (1) where target analytes are extracted from an aqueous sample through a supported liquid membrane (SLM) that is immobilized in the pores of a porous polymeric material and into a volume of acceptor solution (typically, 10–30 μL). In this context, the porous polymeric material is a hollow fibre. Here, we highlight important practical issues related to the SLM, the hollow fibre and the extraction itself because these issues are important for successful HF

 LPME. We also discuss practical work with electromembrane extraction (EME), which is related to the HF

 LPME device but uses an electrical potential as the driving force for the extraction (2). 

 LPME can be used for extraction of basic or acidic analytes from aqueous samples. Figure 1 illustrates a setup for HF

 LPME. The sample is contained in a sample vial and the pH is adjusted in the sample before extraction to keep the analytes in their uncharged state. For basic analytes, the sample is made alkaline, and for acidic analytes, the sample is acidified. A small piece of a porous hollow fibre, typically made of polypropylene, is closed in one end and dipped in an organic solvent immiscible with water. In a few seconds, this organic solvent is immobilized in the pores in the wall of the hollow fibre by capillary forces, forming an SLM. 

Figure 1: Illustration of a typical hollow-fibre  liquid-phase microextraction in the three-phase mode (HF3LPME) setup. 

A 10–30 μL volume of aqueous acceptor solution is then injected into the lumen of the hollow fibre. For basic analytes, the acceptor solution is acidic, whereas it is alkaline for acidic analytes. The hollow fibre is finally placed into the sample and the whole assembly (sample vial and hollow fibre) is agitated for typically 15–45 min. During this time, analyte molecules are extracted in their uncharged state from the sample into the SLM, and further into the acceptor solution. In the acceptor solution, the analyte molecules become ionized, which prevents them from re-entering the SLM. After extraction, the acceptor solution is collected and analysed directly by high performance liquid chromatography (HPLC), capillary electrophoresis (CE), mass spectrometry (MS) or other related analytical techniques.  

Direct compatibility with HPLC, CE and MS

Low solvent consumption (10–30 μL of solvent for each extraction).

Advantages, as well as limitations, of HF

 LPME have been discussed substantially in the literature and several reviews discussed a broad range of applications (4–11). 

 LPME are typically made of polypropylene (4). Most work published in the literature has been accomplished with a polypropylene hollow fibre from Membrana (Wuppertal, Germany) termed "Q3/2" that has an internal diameter of 600 μm, a wall thickness of 200 μm and a pore diameter of 0.2 μm (4). The hollow fibre is connected to a pre-cut pipette tip or a medical syringe needle. The pipette tip or syringe needle serves as a guide tube to facilitate the injection and withdrawal of the acceptor solution, as illustrated in Figure 2. The fibre can be arranged either in the loop configuration with connections in both ends (Figure 2b) or in the rod configuration with a connection in one end and the other end closed (Figure 2a). To close the hollow fibre, mechanical pressure with a pair of pincers can be used — no heat or glue is required. In our laboratory, we use pipette tips for the connections. In this case, we carefully heat the connection between the tip and the fibre with a soldering iron to prevent disruption. 

Figure 2: Different configurations for hollow-fibre liquid-phase microextraction in the three-phase mode (HF3LPME): (a) rod configuration, (b) loop configuration and (c) hollow fiber attached directly to the needle of a microsyringe. 

Alternatively, the hollow fibre can also be attached directly to the needle of a microsyringe for easy injection and withdrawal of the acceptor solution, as illustrated in Figure 2c (12). Alternative fibre dimensions can also be used, but generally the thickness of the wall of the fibre should not exceed 200–300 μm because the extraction speed and the recovery are dependent on the thickness of the SLM. Fibres with internal diameters less than 600 μm have been reported to speed up HF

 LPME with small volumes of acceptor solution for high enrichment (3), and hollow fibres with internal diameters larger than 600 μm have been used for easier injection and withdrawal of the acceptor solution (13,14). 

 LPME, the SLM is an intermediate extraction medium. Analytes should be extracted rapidly and efficiently into the SLM, but transport out of the SLM and into the acceptor phase should also be efficient to avoid substantial trapping of analytes in the SLM itself. Substantial trapping in the SLM is undesirable because it reduces the extraction recovery. In other words, finding the optimum SLM solvent for the application is an important step. 

 LPME, dihexyl ether and 1-octanol have been the most popular SLM solvents (4). As seen in Table 1, these two solvents have a high boiling point (> 195 °C), and when the hollow fibre is dipped in the solvents, little or almost no evaporation of the SLM is observed during 2 min of air exposure (Table 1). This observation is important because the hollow fibre with the immobilized SLM is normally exposed for a short time (< 2 min) to open air before it is placed in the sample. After the hollow fibre with the SLM is inserted in the sample in a capped vial, evaporation is no longer an issue because the SLM is protected by the aqueous sample and the system is closed. Volatile solvents such as toluene and 1-chloropentane may be difficult to use in HF

 LPME because they evaporate quickly and give an unstable SLM (see Table 1). In general, it is recommended not to use solvents with a boiling point below 190–200 °C for HF

Table 1: Boiling point, evaporation, water solubility, and dissolution data for various solvents for hollow-fibre liquid-phase microextraction in the three-phase mode (HF3LPME). 

In addition to the volatility of the solvent, the water solubility is also important. For dihexyl ether, the water solubility is low (< 110 μg/mL), therefore, SLMs made from this solvent are very stable during extraction. With 1-octanol, the water solubility is higher (1200 μg/mL) and this solvent may leak into the sample (and the acceptor solution) in significant amounts. As seen in Table 1, about 11% of a 1-octanol SLM may theoretically leak into 1 mL of sample based on the water solubility, resulting in a significant reduction of the SLM. This level of SLM leakage has been verified experimentally in our laboratory by analysing the sample solution after HF

 LPME with gas chromatography–mass spectrometry (GC–MS) to determine traces of 1-octanol. This loss may be even greater if the sample volume is increased or if surfactants and other emulsifying agents are present in the sample. In general, it is preferred not to use solvents exceeding 200–400 μg/mL in terms of water solubility. In addition to dihexyl ether, solvents such as 1-decanol, dodecyl acetate, 2-nitrophenyl octyl ether and 1-nonanol fulfill the criteria discussed above and have been used in HF

 LPME studies (Table 1) (4). Some new solvents that have also been tested recently in our laboratory for HF

 LPME are included in Table 1 and may be interesting SLM candidates for the future.  

How Much Does the SLM Affect the Extraction?

Table 2 (basic drugs as model analytes) and Table 3 (acidic drugs as model analytes) illustrate recent examples from our laboratory in which the extraction recovery in HF

 LPME was strongly affected by the type of solvent used in the SLM. For the basic model analytes, the highest recoveries were obtained with dodecyl acetate, 2-octanone and isopentyl benzene as the SLM solvent. For the acidic drugs, dodecyl acetate, isopentyl benzene and 1-decanol were the top three SLM solvent candidates. A closer look at the results in Tables 2 and 3 indicate several important aspects. First, recoveries were generally higher for the acidic model analytes than for the basic model analytes. The reason for this result was probably that the selected acidic model analytes were slightly more polar than the basic ones. From earlier experience (15), it is well known that extraction recoveries in HF

 LPME are highest for analytes with log P values in the 2–4 range, whereas the extractability decreases somewhat for analytes with log P values exceeding 4. 

Table 2: Hollow-fibre liquid-phase microextraction in the three-phase mode (HF3LPME) performance ranking of solvents for basic model analytes. 

The basic model analytes in Table 2 where highly hydrophobic (log P in the 3.1–5.3 range), and extraction from the organic SLM and into the aqueous acceptor phase was somewhat limited by partition. Because of this, selection of the solvent was very important for the basic model analytes. Second, the extraction performance of each of the solvents was checked against the Snyder solvent selectivity classification system (16). The two top solvents, namely dodecyl acetate and 2-octanone, were both class VI solvents (aliphatic ketones and esters) and the next two solvents were both class VII solvents (aromatic hydrocarbons). Although relatively different in terms of chemical structure, class VI and VII solvents are close to each other in terms of solvent selectivity properties with relatively strong proton acceptor and dipole characteristics. 

Table 3: Hollow-fibre liquid-phase microextraction in the three-phase mode (HF3LPME) performance ranking of solvents for acidic model analytes. 

A somewhat different picture was observed for the acidic model analytes in Table 3. Because these analytes were less hydrophobic, with log P values in the 2.9–4.3 range, they were more easily extracted from the organic SLM and into the aqueous acceptor phase. Therefore, the selectivity of the solvent was less critical for these analytes. Thus, the five top solvents, all of which provided average recoveries of 70% or more, belonged to Snyder classes I, II, VI and VII. These solvents have substantial differences in terms of solvent selectivity, proton acceptor, proton donor and dipole characteristics.  

 LPME has been performed mainly by trial and error, testing a limited number of candidates including dihexyl ether and 1-octanol. Most likely, more systematic approaches will be developed in the future that will take further solvent properties into consideration. It also should be mentioned that HF

 LPME is best suited for analytes with log P > 2. For analytes with log P < 2, extraction is more challenging and requires the addition of a carrier such as an ion-pair reagent either to the sample or the SLM (15).  

How Should the Dry Hollow Fibre Be Handled?

 LPME typically are purchased as bundles from the manufacturer. No hollow fibres are currently manufactured specifically for HF

 LPME; instead they are industrial products for totally different applications. We recommend storing the hollow fibres in a closed bag protected from light, because air and light exposure over long time periods might degrade the mechanical stability of the hollow fibre and make it more fragile. Before use, the hollow fibre needs to be cut to form pieces of appropriate length. We recommend using gloves when cutting the fibre to avoid contaminating it. It is important that each piece be cut to exactly the same length. If fibres are of different lengths, the volume of the SLM will vary from extraction to extraction, and this may give some variation in the results. In two-phase hollowfibre liquid-phase microextraction it is common to wash the hollow fibre with acetone before extraction to remove additives in the polymeric material (17). This is less important in HF

 LPME because the acceptor solution is aqueous and most polymer additives are not soluble in aqueous solution. However, for extraction of very hydrophobic analytes, recoveries may be slightly improved if the hollow fibre is prewashed with acetone (18). Each piece of hollow fibre is for single use and should always be discarded after extraction to avoid carryover from one extraction to another. 

The SLM normally is prepared by dipping the hollow fibre into the organic solvent for 5–10 s. The organic solvent is immediately immobilized in the pores in the wall of the hollow fibre by capillary forces. This procedure is very simple, but the exact amount of organic solvent is unknown, and excess organic solvent may be located on the surface of the hollow fibre. In such cases, it is recommended to remove excess solvent from the hollow fibre. This can be accomplished either by wiping the fibre with a medical wipe or exposing the fibre and the SLM to ultrasonification in a water bath for 5–10 s. The former method (medical wipe) is recommended, as this procedure has been reported to yield the most reproducible SLM (18). 

Alternatively, the organic solvent can be injected into the lumen of the hollow fibre using a microsyringe. In this procedure, the SLM is coated from the inside of the hollow fibre. The advantage here is that the volume of the SLM solvent is controlled more exactly; this approach may be interesting for future automation of HF

 LPME. The typical volume of the SLM solvent in one piece of hollow fibre is 10–30 μL. 

It is recommended to immobilize the SLM solvent in the hollow fibre in the shortest amount of time possible before the extraction. This is done to avoid partial evaporation of the SLM solvent and so that the SLM solvent is not gradually swelled into the polymer itself. With solvents like dihexyl ether and 1-octanol, swelling may totally interrupt the SLM after a few days of storage. Other solvents have been found to be as highly stable as the SLM solvent, and they may be immobilized for up to 60 days before use. These solvents include silicone oil AR 20 (polyphenylmethylsiloxane), 2-nitrophenyl octyl ether and dodecyl acetate (18). 

How Should the Acceptor Solution Be Loaded?

When the SLM is prepared, the acceptor solution has to be injected into the lumen of the hollow fibre. This is accomplished with a microsyringe. Loading exact and constant volumes of acceptor solution from extraction to extraction is important to obtain the highest repeatability. Make sure that injection of the acceptor solution is performed slowly. Rapid injection of the acceptor solution into the narrow hollow fibre may cause air bubbles, which results in small segments of the hollow fibre containing no active acceptor solution. Air bubbles in the hollow fibre from rapid injection can affect the results significantly and sacrifice repeatability (18).  

How Should the Actual Extraction Be Performed?

Extraction is initiated at the time when the hollow fibre, containing both the SLM and the acceptor solution, is placed in the sample. Exact timing of the extraction is important, and the time for each extraction should be measured from the point when the fibre is placed in the sample. Immediately, the whole assembly (sample plus the hollow fibre) should be transferred to an agitator. Agitation (or stirring) is important to facilitate extraction and constantly replenish the sample in close contact with the SLM. Normally, we recommend agitating the entire extraction unit (sample vial and hollow fibre) at 800–1200 rpm, but stirring with small magnetic stir bars may also be used. Normally, HF

 LPME is accomplished at room temperature with no external temperature control. 

When the extraction has been completed at an exact stop time, the hollow fibre should be removed immediately from the sample to stop the extraction. This is especially important if extractions are not performed to equilibrium as shown in Figure 3. Additionally, the acceptor solution should be removed immediately from the hollow fibre to avoid partial backextraction into the SLM and loss of analyte. 

Figure 3: Recovery versus extraction time for haloperidol. Supported liquid membrane: 2-nitrophenyl octylether; sample: 1 mL of 25 mM ammonia buffer pH 10 containing 1 Î¼g/mL haloperidol; acceptor solution: 25 Î¼L of 10 mM HCl. 

The acceptor solution is normally removed with a microsyringe and transferred to a sample vial for the final analysis by HPLC or CE. Because the acceptor solution volumes typically used are low, the acceptor solution should be protected from evaporation. Therefore, storage at low temperature in a closed vial is highly recommended. When the acceptor solution is removed from the hollow fibre, it is also important to check the volume of the acceptor phase. Occasionally, the volume collected after extraction is different compared to what was injected into the hollow fibre before extraction; this volume difference is a clear indication of leakage in the system. In such cases, the acceptor solution should be discarded. 

 LPME of basic and acidic compounds, the pH of the sample and the acceptor solution is highly important. For basic analytes, the pH of the sample should be high to suppress ionization of the basic substances and promote their extraction into the SLM, whereas the pH of the acceptor solution should be low to ionize the basic substances upon arrival in the acceptor solution. The latter effect also prevents the analytes from re-entering the SLM. The strong pH-gradients across the SLM serve as the driving force for the extraction. For basic analytes, we normally recommend adjusting the pH 1–3 units above the pKa values of the analytes in the sample and 1–3 units below their pKa values in the acceptor solution. Typically, the sample is made alkaline with sodium hydroxide, whereas hydrochloric acid (10 mM) or formic acid (LC–MS friendly) is used as the acceptor solution (4). For acidic substances, the pH gradient is reversed, with acidic conditions in the sample and alkaline conditions in the acceptor solution. Usually, hydrochloric acid is used to acidify the sample, whereas 10 mM sodium hydroxide or ammonia solution (LC–MS friendly) is used as the acceptor solution (4). 

Sample Preparation Perspectives

Describes practical aspects of hollow-fibre liquid-phase microextraction in the three phase mode (HF3LPME) and also electromembrane extraction (EME).

Hollow-Fibre Liquid-Phase Microextraction in the Three-Phase Mode – Practical Considerations

Since the 1990s, chiral chromatography has been growing by leaps and bounds. In the pharmaceutical industry, chiral chromatography is now clearly established as an effective way to access gram and kilogram quantities of enantiopure substances. I was in my fourth year of pharmacology studies at the University of Barcelona when the syllabus covered medicinal chemistry. This remarkable area sparked an interest in the interaction of molecules with living systems, and it was at this point that I learned the importance of chirality in life science. 

At the time, analytical use of chiral chromatography was on the rise, and researchers had just begun investigating chiral chromatography for preparative separations. There were still many questions surrounding the science, but the technological advances were exciting. 

While finishing my pharmacy studies, I began working in a laboratory at the University of Barcelona with Cristina Minguillón — one of my former professors — on the preparation of chiral stationary phases (CSPs). After graduating in 1992, I started my doctorate with a focus on the preparation and use of polysaccharide derivatives as chiral stationary phases for high performance liquid chromatography (HPLC) under the supervision of Professor Minguillón and Laureano Oliveros.  

During this time, I studied the preparation, optimization, and subsequent testing of Pirkle CSPs, which were commercially introduced in 1980. These were named for William Pirkle, emeritus professor of chemistry at the University of Illinois. Pirkle CSPs can separate a variety of drug compound groups, including nonsteroidal anti-inflammatory drugs and beta blockers. Pirkle CSPs were only a portion of my thesis, however. I also researched the synthesis of immobilized polysaccharide-derived CSPs, which became the main topic of my doctorate work. 

My first chiral separation was with analytical resolutions of enantiomers with the first CSPs I had prepared in the laboratory. At that time, the equipment was not very sophisticated — manual injectors, integrators, and no computer control — but it still helped me understand the mechanisms. Computer control came soon enough, following which research advanced more quickly.  

My doctoral work led me to be published in several international journals. My research demonstrated the possibility of producing immobilized CSPs and using them systematically for separating enantiomers. Although the method still was not optimal, it provided good insight into the influence of various parameters that could be considered in producing these new phases, as well as their influence in chiral recognition abilities. 

I then completed two years of postdoctoral work with Wolfgang Lindner at the University of Vienna, as a Marie Curie Fellow from the European Commission. During this time, I researched the synthesis of chiral selectors that were mainly derived from cinchona alkaloids. In contrast to the work in Barcelona, which I dedicated mainly to liquid chromatography (HPLC and centrifugal partition chromatography), in Vienna I explored other techniques such as capillary electrophoresis and membrane separations. 

Dr. Lindner has deep knowledge of chiral recognition mechanisms and over the years has been developing successful systems for separating enantiomers using various techniques. Some of Dr. Lindner's CSPs have become commercially available, and he has made important findings, but I believe his biggest contribution is all the educational work he has done with chirality throughout his career. 

The study of chiral molecules is actually a branch of stereochemistry, which involves the study of the relative spatial arrangement of atoms within molecules. The term 

describes an object that can not be superimposed on its mirror image. In chemistry, chirality refers to two molecules. 

The mirror images of chiral molecules are called 

(from the Greek enantio, meaning "opposite"), and enantiomers can exhibit distinct chemical behaviour when subjected to a chiral environment. Chemists often designate these pairs as right- and left-handed. In fact, the term "chirality" is derived from the Greek word for h

Lord Kelvin coined the term chirality in 1873, almost 60 years after the phenomenon was observed by Jean-Baptiste Biot. The property gained considerable importance in the sugar and pharmaceutical industries as well as in the field of analytical chemistry. The term racemic, meaning a mix of both enantiomers, comes from the Latin racemus for "a bunch of grapes." The word originated with Louis Pasteur, who isolated racemic tartaric acid from wine. 

One of the most significant examples of chirality is the thalidomide disaster of the 1960s. Although thalidomide originally was used as a tranquilizer, physicians also administered the drug to pregnant women to prevent morning sickness. First developed in Germany in 1957, thalidomide was manufactured and sold as a racemic mixture of N-phthalylglutamic acid imide. The drug's desired physiological activity stems from the (

)-(+)-isomer. Once ingested, thalidomide undergoes racemization inside the human body. So, even if the safe isomer is taken, the other enantiomer is produced. Researchers discovered too late that the corresponding (

)-(–)-enantiomer could cause severe birth defects. It has been estimated that more than 10,000 babies were born with deformations because of this drug. 

The thalidomide disaster captured the interest of worldwide drug manufacturers and regulatory agencies. As a result, chiral research took off. The United States Food and Drug Administration later mandated that the isomers of all new drugs be studied individually for their physiological effects. This requirement created a demand for analytical and preparative techniques that were lacking, despite earlier work on CSPs and reagents. 

It was not until the 1990s, however, that pharmaceutical manufacturers realized they could use chiral separations to safely analyze pure enantiomers and bring them to market. Significant progress with chiral separations at that time resulted from four essential advances: the introduction of efficient CSPs; the advanced development of simulated moving bed (SMB) chromatography systems; the introduction of computer systems used to model SMB processes; and the integration of efficient evaporation technologies to recycle mobile phases. 

In the pharmaceutical industry today, chiral chromatography iis recognised as an efficient way to access gram and kilogram quantities of enantiopure substances. Its cost-effectiveness has been demonstrated at large scale with various drugs that have been approved for marketing. Today, it is well known that the drugs Lexapro (escitalopram), Zoloft (sertraline), Keppra (levetiracetam), and Xyzal (levocetirizine) use chiral chromatography in their industrial-scale production processes. 

My research and the studies of others have shown that polysaccharide-derived CSPs are versatile and durable when used for HPLC and supercritical fluid chromatography (SFC) separations. These qualities have made them the first choice for analytical and preparative separations of enantiomers. 

CSPs were first introduced to the market as coated materials on a silica support. A great deal of their development was carried out by Yoshio Okamoto at Nagoya University. Okamoto  found that he could achieve most enantiomeric separations using four different derivatives. Two are based on cellulose, and two are based on amylose. These polysaccharide phases were developed by deriving amylose or cellulose and coating these chiral polymers on a silica gel base. Among the various polysaccharides available, amylose and cellulose derivatives are the most useful as CSPs. Columns containing these CSPs are equally effective with normal-phase and reversed-phase solvents. 

In that format, however, the potential of CSPs was limited by restrictions in the solvent choice, which was dictated by polymer solubility. At that point, immobilized CSPs were created, and the first commercially available immobilized CSP was introduced in 2004. 

Many research groups investigated the best way to prepare immobilized CSPs. Researchers with Daicel and Chiral Technologies Worldwide, as well as E. Francotte at Novartis, were at the forefront of immobilization technology. Work by our colleagues at Daicel was essential to the development and Tong Zhang of Chiral Technologies Europe was deeply involved in studying immobilization technologies. 

Several methods have been developed to chemically bond, or immobilize, the polysaccharide-based polymer to the silica support, but only a few technologies have worked for a wide range of applications with high efficiencies and an unlimited choice of solvents, including ethyl acetate, dichoromethane, tetrahydrofuran, and acetone. 

Nevertheless, immobilizing a chiral polymer on a silica support offers several benefits. First, immobilized CSPs are not damaged when they come into contact with a solvent that swells or dissolves the polymer. Another benefit is that immobilized CSPs allow difficult separations to be performed with solvents that could not be used previously with CSPs. Additionally, new selectivity can be developed by using previously unusable chiral selectors. 

Most importantly, perhaps, immobilizing chiral polymers brings a new generation of polysaccharide-based CSPs in line with existing HPLC phases. Immobilized CSPs help extend column life and eliminate unexpected column failures. 

Since the 1990s, the use of chiral chromatography has grown significantly. The technology itself has improved vastly, especially in the area of preparative chiral chromatography. Now, chiral chromatography systems are becoming mandatory tools in chemical processing and research and development departments, particularly in the area of preparative separations. 

In the past, pharmaceutical manufacturers viewed preparative chromatography as a time-consuming and expensive process. Today, it has evolved to allow manufacturers to perform fast and cost-effective separations using automated equipment for techniques such as continuous chromatography and SFC. In fact, pharmaceutical companies have developed chiral chromatography methods — from feasibility study to production — in less than one year. 

Continuous chromatography, also referred to as SMB, has been around since the 1950s, but initially it was only applicable to manufacturing foods and sugars. It wasn't until it became possible to operate SMB systems at high pressures that pharmaceutical companies could start harnessing it for drug development. 

SMB is now a workhorse in Big Pharma companies like GlaxoSmithKline, Merck, and Pfizer. The largest SMB units process up to 200 metric tons of material annually. This is extraordinary, given that when I started in preparative solutions, we could only create 1 g at a time. 

Companies using SMB can now expect to complete method development in a few months as opposed to two to five years for asymmetric methods that also have a much higher cost. It has been projected that chiral chromatographies will continue to evolve in tandem with the growth of single-enantiomer, small-molecule drugs. 

A related technology that has been advancing is SFC. Although SFC systems were invented before the 1990s, initially the technology was not very well established. Today, SFC systems are more robust and it is becoming a well established technique. SFC is important for enantioselective separations, especially for preparative purification and isolation of individual enantiomers. 

SFC offers significant benefits, including faster separation and method development and improved column efficiency. But in larger separations, the benefits of SFC go beyond speed. Its use of carbon dioxide makes SFC a "green" technology because it doesn't add anything to the atmosphere, and the consumption of organic solvents is greatly reduced. Also, chemists can recover separated products more quickly because isolated fractions are collected in smaller volumes of liquid. 

Moving forward, the continued development of advanced CSPs and improved commercial-scale processes will ensure that preparative chiral chromatography is viewed as an equal to other technologies in the pharmaceutical industry. Pharmaceutical researchers are embracing new chiral technologies, and methods such as SFC are becoming more and more accepted. 

 is the manager of technical operations for Chiral Technologies Europe, illkirch Cedex France. Direct correspondence about this column should go to LCGC Europe, 4A Bridgegate Pavilion, Chester Business Park, Wrexham Road, Chester, CH4 9QH, UK or e-mail the editor, 

History of Chromatography

The development and application of chiral chromatography is described.

The Evolution of Chiral Chromatography

The 27th International Symposium on MicroScale Bioseparations and Analyses (MSB 2012) 

Starling Hotel and Conference Centre, Geneva, Switzerland 

The symposium traditionally focuses on fundamental aspects of separations on the 'micro'scale. This year, the organizers have added "Analyses" to the symposium title to emphasize that the symposium focuses not only on separation science, but also on the application of microscale separations for bioanalytical measurements in life science research, biopharmaceuticals development, forensics, toxicology, doping analyses, food safety and authenticity and environmental sustainability. 

The symposium will offer plenary and keynote lectures and submitted oral presentations by leading academic and industrial scientists, poster sessions, workshops, tutorials and seminars by equipment manufacturers.  

Leaders in the field delivering plenary lectures include Professor Ruedi Aebersold of the ETH Institute of Molecular Systems Biology in Zürich, Switzerland, who will present the opening lecture on 

Mapping and Measuring Proteomes in Systems Biology

 followed by an always entertaining lecture by Professor Mario Thevis from The Center for Preventive Doping Research in Cologne University, Germany, who will discuss 

Recent Challenges in Sports Drug Testing.

 Other plenary speakers will be Professor Barry Karger, director of the Barnett Institute at Northeastern University in Boston, who will speak on 

Translational Regulatory Science: Recent Advances in Microscale Bioseparations for Biotechnology Development,

 Dr Markus Bluemel of Novartis Pharma AG in Basel will speak on 

The Use of Micro-Scale Separation Methods in Development of New Biopharmaceuticals 

and Professor Thomas Hankemeier, director of the Netherlands Metabolomics Institute, will discuss 

The MSB symposia hosts the prestigious Arnold O. Beckman Medal and Award for 

Outstanding Scientific Achievements in the Field of Electrodriven Separation Techniques

. The first recipient of the award will be Professor Pier Giorgio Rhigetti for his outstanding contributions to the development of new electro-driven separation techniques, which have been key to the development of many microscale separation and analysis methods. The award is sponsored by Beckman-Coulter and comprises a medal, a book honouring the 100year anniversary of Dr A.O. Beckman and a cash prize. A plenary talk by the awardee will be the first lecture on the first full symposium day. 

The oral program layout will be arranged in two parallel sessions: one focusing on fundamentals and the other on applications of microscale (bio)separations. A strong workshop programme precedes the oral programme on Sunday. Further details can be obtained from the organisers. 

Planned tutorials include Dr Joerg-Ingo Baumbach on ion mobility spectrometry, Professor Alexandra Vos on dielectrophoresis and Dr Fritz Erni on Quality by Design for Method Development. 

A post-symposium course on capillary electrophoresis by Dr Cari Saenger-van de Grient will follow on 

. Important deadlines for the event include early bird registration: 

; Submission of posters for inclusion in the conference book: 

; and late contributions (not included in the conference book): 

CE Method Development and Validation Course

 The Convent Hotel, Amsterdam, The Netherlands 

 Oud-Sint-Jan' Conference Centre, Bruges, Belgium 

: Royal Flemish Chemical Society and Royal Society of Chemstry 

 Ornage County Convention Centre, Orlando, Florida, USA 

 New Munich Fair Trade Centre, Munich, Germany 

 Silesian School of Management, Katowice, Poland 

: Institute of Environmental Engineering of Polish Academy of Science 

Departments

Event News

In multivariate calibration and modelling a model is built between a large data matrix containing many variables for each sample, such as a spectrum or a chromatogram, and a property of the samples, such as a given activity. Variable selection or reduction in the large matrix is required, or can be recommended, to obtain better and simpler models.

In pharmaceutical data analysis multivariate calibration and modelling are often used to build predictive models. Many different modelling techniques can be used. However, these techniques usually require either a variable selection step to limit the number of variables to build the model or create so called latent variables to build simple models.  

Many variable selection methods have been applied in the literature. An overview of some of the most frequently applied will be discussed in this article In cases where latent variables are used, the models with latent variables based on the entire data set are usually not as good as when uninformative variables are eliminated first. Different possibilities exist to reduce the number of variables used in the calculation of the latent variables. The goal of feature selection or reduction is to select the most important but also to build simpler methods with a better predictive performance than when applying the entire data set. 

In multivariate calibration and modelling a model is built between a large data matrix containing many variables for each sample and the property of the sample.

LCGC Europe-12-01-2011

Asymmetrical Flow Field Flow Fractionation for Investigating Intermolecular Interactions of Multifunctional Polymers

Troubleshooting Basics, Part 3: Retention Problems

Hollow-Fibre Liquid-Phase Microextraction in the Three-Phase Mode – Practical Considerations

The Evolution of Chiral Chromatography

Event News

Variable Selection and Reduction in Multivariate Calibration and Modelling