Review pubs.acs.org/ac
Fundamentals for LC Miniaturization Gert Desmet* and Sebastiaan Eeltink
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Department of Chemical Engineering, Vrije Universiteit Brussel, Pleinlaan 2, B-1050 Brussels, Belgium achievable in GC) and low flow rates, the latter being of special relevance to mass spectrometry (MS) detection. At low flow rates, smaller droplets are formed with higher surface-to-volume ratios, which lead to an improved ion desorption into a gas phase and thus to higher ESI-MS sensitivity. Additional advantages of miniaturized LC systems are their economy of use, as smaller amounts of stationary-phase material are needed, while also smaller volumes of mobile phase are being consumed. Given the global interest in greener operations, the latter advantage has become more and more prominent over the past decade. If (and only if !) the sample amount is limited (e.g., as is the case in some applications in proteomics, forensics, single-cell analysis, pediatric clinical analysis), an enhanced detection sensitivity can be expected, as the fixed (and limited) sample amount simply gets less diluted on the column.9,10 On the other hand, miniaturizing the column volume is only counterproductive when sufficient sample volumes are available, as column miniaturization also inevitably involves a reduction of the detector volume. The latter follows from the generally accepted design criterion that both the injector and detector volumes should not be larger than one peak standard deviation (σV). The latter is fully determined by the column volume Vcol (via σV = Vcol (1 + k′)/√N).11 Hence, the very small column volumes of miniaturized systems require the injection of small sample volumes and the use of small detectors. This severely compromises the detection sensitivity of miniaturized LC systems, because ultimately the signal-to-noise ratio (S/N) of any detection scheme (be it concentration or mass sensitivity) will always depend on the absolute number of molecules that can contribute to the signal.10,11 A notable exception to the loss in detection sensitivity inevitably accompanying the miniaturization of the column is electrospray ionization-mass spectrometry (ESI-MS), which performs optimally when flow rates in the 300 nL/min range are applied. A landmark paper demonstrating the superior ESI properties of low flow rates for sample-limited applications is that of Shen et al.12 Despite the observed “concentration-sensitive” behavior of ESI-MS detection, it should, however, be clear that also for this type of detector, a further miniaturization will eventually hit a limit where the flow rates become so small that they can only bring minute amounts of molecules to the detector. The other original major driver for the miniaturization of LC systems (i.e., the possibility to achieve higher efficiencies than with normal base columns) necessitates the distinction between packed and open-tubular columns. For packed-bed columns, the potential advantage of a diameter miniaturization (in turn leading to reduction of the column-to-particle diameter aspect ratio, see
CONTENTS
Recent Drivers for the Miniaturization of LC Systems Pursuit of Ultrahigh Pressures Improvements in Detectors (ESI-MS) New Machining Methods for Chips Multidimensional Separations Temperature-Assisted Separations Packed Capillaries Monolithic Capillary Columns Open-Tubular Columns Chip-Based Systems Chip Systems Packed with Particles or Monolithic Beds Chip Systems with Micromachined Supports Perspectives Author Information Corresponding Author Notes Biographies References Note Added after ASAP Publication
544 544 545 545 545 546 546 547 549 551 551 551 553 553 553 553 553 554 556
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hereas a huge surge in the development of miniaturized LC systems was marking the field in the 1970s and 1980s of the past century,1−3 the academic interest in the topic declined considerably in the years after. This was mainly because of the gloomy prospects for the development of novel detector schemes that would be able to get the same strong signal out of a 100− 10 000 times smaller amounts of molecules. Not many analysts want to make compromises on the detection sensitivity they can get out of their normal bore columns and the corresponding detector volumes. Fortunately for the field, a number of new areas emerged in the late 1990s and early 2000s (cf. the proteomics and metabolomics research), where usually only a few microliters of sample can be collected, even after massive efforts. In this case, the use of a miniaturized system is not even an option, it is simply mandatory. To meet the needs of these booming fields, quite a large number of miniaturized LC systems (nano-LC instruments, chip formats, packed- and monolithic-bed capillaries) have been commercialized in the past 10 years.4−7 Since about 2005, the academic research has shown a renewed general interest in the topic due to the emergence of a number of new drivers, as discussed in Recent Drivers for the Miniaturization of LC Systems. Apart from also providing a short historical background, the present review focuses on the developments that have occurred in this last period. This period is relatively large but is due to the fact that the last review on the fundamentals of miniaturized LC technology appearing in this series already dates back to 2002.8 The initial interest in miniaturized LC systems was spurred by the desire to combine high efficiencies (pursuing the high values © 2012 American Chemical Society
Special Issue: Fundamental and Applied Reviews in Analytical Chemistry 2013 Published: November 16, 2012 543
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Figure 1. Kinetic-performance limit curves calculated for packed-bed capillaries (based on the reduced Van Deemter parameters A = 0.8, B = 3, and C = 0.075, where h = A + B/ν + Cν, and ϕ = 800) and for open-tubular capillaries coated with a thin film. For both cases, a 1, 3, and 5 μm particle or capillary diameter is considered. The black and red straight lines represent the Knox and Saleem-limit of the packed-bed and OT-LC capillaries, respectively.
(1)
determines the plate height and the column permeability, whereas for packed or monolithic bed columns, both parameters are solely determined by the particle diameter or the domain size (except for potential differences in packing homogeneity and/or thermal effects). This has the direct consequence that the OT-LC system should have a diameter of a comparable size to that of the particle diameters used in their corresponding packed beds to produce the same efficiency. A more exact rule has been proposed by Guiochon,15 showing that the diameter of a cylindrical OT-LC system should not be larger than about 2 times the particle diameter used in the packed bed. A more comprehensive view on the achievable kinetic performance of packed-bed and open-tubular capillaries with different sizes is given in Figure 1, representing the kinetic performance limits16 of packed-bed and an open-tubular capillaries with different diameters (respectively, 1, 3, and 5 μm). When compared for the same analysis time, the OT-LC format theoretically clearly enables much larger efficiencies (factor of 10 or more) than the packed capillary format. However, the 1 and 3 μm i.d. columns needed for the OT-LC are so small that the peaks eluting from it would already be too large for a very small 1 nL detector cell.
The lower the Emin, the faster a given column's format can achieve a given number of theoretical plates. With the above cited values, the advantage of OT-LC versus packed-bed columns becomes readily apparent: Emin ≅ 20 to 30 for the OT-LC, whereas Emin ≅ 2000 to 3000 for packed-bed columns. Monolithic bed columns theoretically have the potential to produce Emin values that approach that of OT-LC13 because the Eminvalue is strongly dominated by the external porosity of the column, and monolithic columns can achieve very high external porosity values (on the order of 0.8−0.9). For highly dense monolithic columns (i.e., with a porosity of ε = 0.5 or smaller), the best possible Emin value can be expected to be closer to that of the packed bed of spheres, for which ε = 0.35−0.4 is typical. Whereas for packed-bed columns, the miniaturization option is rather the consequence of an economic issue or to overcome a sample limitation problem. The miniaturization of OT-LC systems is mandatory because the column diameter directly
RECENT DRIVERS FOR THE MINIATURIZATION OF LC SYSTEMS Pursuit of Ultrahigh Pressures. With its groundbreaking work on the implementation of ultrahigh pressures (up to 5000 bar and more) around the year 2000, the Jorgenson group has lead the chromatography field into the UHPLC area, enabling the use of 60 min. Focusing occurs when the mobile phase from the first dimension (at the time of elution) is a weak effluent in the second dimension. This allowed Wu et al. to develop an off-line LC/ × /LC workflow that involves a strong cation-exchange separation on a 300 μm i.d. column packed with 2.5 μm SXC particles by applying a 60 min 1D salt gradient followed by a 2D reversed-phase pressurized capillary electrochromatographic (pCEC) with a 100 μm i.d. capillary column.36 The same group combined capillary IEF (cIEF) with pCEC using a microinjection valve as the interface.37 After the cIEF separation, the focused bands were then electrically mobilized into the loop attached to the microinjector and sequentially analyzed. Cesla et al. developed an automated off-line 2D separation system by coupling a HPLC gradient separation on a bonded PEG column in the first dimension with a micellar electrokinetic chromatographic (MEKC) separation.38 Hence, the ease of integration between a pressure-driven and an electrically driven separation also seems to be another driver for the continued interest in miniaturized LC systems.
columns (corresponding to the other extreme in thermaloperating conditions, opposite of the adiabatic-operating conditions), it is well-known that the deleterious effect of the radial temperature gradients increases to the sixth power of the column radius.21 Hence, here also, column miniaturization constitutes a huge advantage. Therefore, in order to exploit the benefits of pressures in excess of the presently available 1000 to 1300 bar UHPLC pressures, a miniaturization of the diameter of packed-bed columns will be mandatory, except when other possible solutions such as the use of intermediate cooling strategies are invoked.22 Improvements in Detectors (ESI-MS). Another major trend marking the past decade is that the developments in LC technology have become increasingly more mass-spectrometry driven. This detector technology has made major strides in terms of speed and mass resolution and has become much more widely available. In most miniaturized LC applications, the most compatible format with the nano- to microliter flow rate prevailing is ESI-MS. The utility of this technique is based on the very low flow rates from nanoelectrospray needles (20− 50 nL/min) and the concentration-dependent response of the electrospray.20 Another interesting approach in this area is the development of parallel systems, such as the fully automated four-column capillary LC-MS system proposed by Livesay et al.23 The duty cycle of the LC system is defined as the ratio of the useful chromatographic separation time to the total time between the beginnings of subsequent chromatographic separations. To increase the duty cycle, the system was designed such to perform staggered parallel separations. An encoding translation stage was used for positioning the columns (all equipped with their own ESI emitters) sequentially at the inlet of the mass spectrometer. New Machining Methods for Chips. One particular technological improvement leading toward a renewed interest in miniaturized systems is the use of the photolithographic patterning and anisotropic dry etching techniques24,25 that have revolutionized the microelectronics field and have led to the advent of the lab-on-a-chip field. These techniques, together with other novel techniques such as the laser ablation of polymer films,26 allow for chromatographic microchannels with a high degree of accuracy and spatial resolution to be defined on the surface of a variety of planar substrates (see Chip-Based Systems). Also, the use of polymer imprinting or hot embossing technologies as a strategy toward the mass production of chipbased columns has been explored by the groups of Kutter and 27−29 ́ Romano-Rodriguez, among others. Here, the use of cyclic olefin copolymers (COC or topaz) deserve special attention as they show very promising properties, such as high chemical resistance, low water absorption, good optical transparency near the UV range, and ease of fabrication.30 Multidimensional Separations. In the past few years, the field of LC is also clearly witnessing an unprecedented high interest in multidimensional separations.31 In this area, miniaturized LC systems can play an important role in providing a better match between the peak volumes eluting from one dimension and the peak volumes which can subsequently be loaded onto the second dimension column, without overloading the latter. For this purpose, a miniaturized first dimension column feeding into a normal bore second dimension column seems to be a perfect match. However, this may have a dramatic effect on dilution and therefore detection sensitivity. VivoTruyols et al. described a Pareto-optimality approach providing the optimal parameters (particle sizes, column diameters, and 545
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Another area where capillary columns can contribute to the development of better performing multi-LC systems is by using them as trapping units. Due to their rapid temperature response, rapid heating and/or cooling cycles can be used as a means to enhance the trapping and/or remobilization process. Verstraeten et al. reported a proof-of-principle thermal-modulation experiment, which has the potential to enrich and remobilize smallmolecule analytes between subsequent separations in online LC × LC.39 Enrichment of neutral analytes eluting from the firstdimension column was performed using a capillary “trap” column packed with porous-graphitic carbon particles placed in a heating sleeve. By applying a heat pulse (temperature ramps up to +1200 °C/min), the molecules were remobilized. Compared to the nonmodulated signal, the presented thermal modulator yielded narrow peaks, and a concentration enhancement factor up to 18 was achieved. Eghbali et al. described the possibilities of cryogenic cooling in LC to trap components in LC.40 Trap experiments were conducted using an in-house built setup that allowed for a segment of the column to be cooled down to temperatures below T = −20 °C. It was illustrated that thermal peak focusing can be applied to achieve a better peak shape and an increased S/N ratio of the different components in a separation. The same concept was used to trap different components in one concentrated band, which is similar to the “cryofocusing” technique in GC. This opens the door to the use of this principle in LC for equivalent purposes as in GC, including heart cutting, 2D-LC modulation, and dual-stage cooling. Temperature-Assisted Separations. Other drivers, such as a further exploration of the possibilities of high-temperature pulsing to enhance the selectivity and resolution of certain critical pair separations, that have already been explored in the past41 are showing up again as well.42 Short temperature pulses improved resolution in discrete sections of chromatograms, as was demonstrated by the ion-exchange chromatography (IC) and hydrophilic interaction chromatography (HILIC) modes. Temperature programming for HILIC separations is particularly attractive as the narrow range of organic modifier concentrations that can be employed makes method development difficult. Since the time needed for temperature responses grows with the second power of the tube diameter,43 the use of a miniaturized capillary format is indispensable here as well. Another recent innovation taking advantage of the rapid heating and cooling cycles of miniaturized LC systems is the work of Collins et al., demonstrating the possibility of making a direct-contact thermoelectric array for capillary columns (chip-based format) that enables precise temperature control, using individually controlled sequentially aligned thermoelectric Peltier elements.44 The operating temperature range of the platform was between 15 and 200 °C for each of 10 aligned Peltier units, with a ramp rate of approximately 400 °C/min. The system allows for the imposition of temperature gradients with both linear and nonlinear profiles, including both static column temperature gradients and temporal temperature gradients.44 Low-thermal mass liquid chromatography (LTMLC) was introduced in 2009 by Gu et al.45 The LTM assembly utilizes the principle of resistive wire heating around a capillary column and a temperature sensor to accurately deliver heating (up to 1800 °C/min) or cooling (100−200 °C/min) rates. Both solvent gradients and temperature programs were demonstrated for selectivity tuning of model mixtures that contained basic compounds. This study also demonstrated the feasibility of applying oscillating temperature gradients for selectivity tuning in LC.
Review
PACKED CAPILLARIES
In the past, packed-bed capillaries have been claimed to lead to higher efficiencies than their normal-bore counterparts. As early as 1969, Knox and Parcher suggested that an additional improvement in efficiency can be obtained by optimizing the column diameter (dc) in relation to the particle diameter.46 With the use of columns packed with spherical glass beads that had an aspect ratio (dc/dp) as small as between 6 and 8, an increase in efficiency was reported for the existence of a dominant wall effect on the packing structure. Kennedy and Jorgenson packed capillary columns with inner diameters between 20 and 50 μm with 5 μm silica particles.47 The minimum reduced plate height (hmin) decreased from 1.4 to 1.0 for an unretained compound and from 2.4 to 1.5 for a retained compound with a retention factor of 2.7 over the range of decreasing column diameters. A decrease in the eddy-diffusion (A term) and mass-transfer contribution (Cm term) was observed as the column diameter decreased. When the aspect ratio is decreased, the distances across which the wall effect and other packing heterogeneities need to be leveled out become smaller and smaller. Knowing that the plate-height contribution associated with the transcolumn velocity gradients varies with the second power of the column diameter,48,49 the advantage of column miniaturization appears to be obvious. In addition, a slight downward trend in flow resistance was observed when the column diameter was decreased for HPLC columns. When the aspect ratio is decreased, the wall effect will influence the packing structure over the entire cross section and a better flow homogeneity can be obtained. In addition, a slight downward trend in flow resistance was observed when the column diameter was decreased for HPLC columns. Both the effect of the aspect ratio on hmin and the flow resistance (ϕ) was found to lead to a significant decrease in separation impedance (Emin).47 For a retained compound (resorcinol), Emin decreased from 4 000 to 1 300 for aspect ratios decreasing from 10 to 4. The same trend was observed for the unretained nitrite, but here the absolute values (Emin = 1 700 to 530) were even much lower (due to lower hmin values). In 2004, this was very convincingly confirmed by Patel et al., who showed a consistent decrease in the minimal plate height, going from about hmin = 2.5 to hmin = 1.3 when the diameter of a set of capillary columns packed with 1.0 μm nonporous C18-bonded particles was decreased from 150 to 10 μm i.d.18 Using fully porous particles, Eeltink et al., on the other hand, observed no significant improvements in efficiency or flow resistance for 150, 100, and 75 μm i.d. capillary columns packed with 10 μm particles (yielding hmin ∼ 2).50 When the aspect ratio was decreased by increasing the particle size, a decrease in reduced plate height was observed. However, the results of flow resistance measurements showed that the latter effect should be attributed to differences in packing and/or the particle-size distribution quality rather than to an aspect-ratio effect. In 2008, the packing density of 30−250 μm i.d. capillary columns packed with 5 μm silica particles was investigated using inverse size exclusion by Ehlert et al.51 For dc/dp > 35, the intraparticle pore space (εi) was determined at 0.36, while for the decreasing aspect ratios an exponential increase in εi was observed, reaching εi ∼ 0.47 at dc/dp = 5. They concluded that the wall effect covers a distance of 4−5 particles. The effect the aspect ratio on the packing density, permeability, and separation efficiency of packed microchips was studied theoretically by Khirevich et al.52 (see Figure 2) and experimentally by Jung et al.53 εi was determined at 0.47, 0.41, and 0.38 for microchips 546
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Figure 2. (A) SEM image of extruded section of a packed bed of 1.0 μm nonporous particles packed in a 10 μm i.d. capillary. Reprinted from ref 18. Copyright 2004 American Chemical Society. (B) Reconstructed sphere packing in a microchannel with low (ε = 0.42) and high (ε = 0.48) bed porosity. The front view is of the packing cross section (left), projection of particle centers (of the 100 000 particles in the front) onto the front plane (middle), and lateral porosity distribution averaged over the whole packing length (right). Reprinted from ref 52. Copyright 2009 American Chemical Society.
from a trapezoidal cross section (ca. 50 × 75 μm with a base angle close to 80°) packed with 10, 5, and 3 μm particles, respectively. Bruns et al. applied confocal laser scanning microscopy to visualize the interparticle void space of 10 to 75 μm i.d. capillary columns packed with 15. The larger-diameter columns yielded looser-packed wall regions than did the bulk regions in low-aspect-ratio packings. Apparently, size segregation of particles along the column radius also affected the packing structure. Both effects led to an additional variation in transchannel dispersion. This study also showed that the packing protocol and particle properties had an effect on the packing density, which may explain why a general consensus of the effect of the aspect ratio from previous studies was not reached. Very recently, the Jorgenson group55,56 presented some exciting results putting the packed capillary back at the forefront of the promising miniaturized systems. Using 1 m long capillaries with a diameter of 50 μm and packed with 1.9 μm fully porous particles, they demonstrated efficiencies in excess of 350 000 plates for retained components (hydroquinone, resorcinol, catechol, and 4-methyl catechol). Around the minimum of the van Deemter curve, reduced plate heights around h = 1.5 were obtained. Given the general experience that it is very difficult to pack homogeneous columns with 600 m2/g). On the precursor monolithic columns, the test mixture of alkyl benzenes eluted close together at the t0 marker as a single broad peak, whereas after hypercrosslinking the alkyl benzenes were baseline separated. Huo et al. suggested that surface-diffusion effects may explain the poor performance of (precursor) polymer monoliths for small-molecule separations.104 Whereas the surface diffusion in typical reversedphase bonded phases is remarkably fast,105 for polymer monoliths the strong increase of the plate heights with retention indicates unfavorable surface-diffusion mass-transfer effects.104 This hypothesis is further supported by the observation that adding THF to the mobile phase resulted in better separations. Due to the swelling of the methacrylate monolith (softening of the stationary-phase surface), the surface-diffusion mass transfer was enhanced. A possible explanation for the improved performance after hypercrosslinking is that the interaction with the stationary phase is no longer dependent on absorption but on
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OPEN-TUBULAR COLUMNS After a decade with a virtually complete inactivity, the Karger group revived the work on open-tubular LC columns around 2007, motivated by the possibility to enhance the detection limits when coupling to ESI-MS and applying ultralow flow rates.109 Yue et al. described the preparation and application of poly(styrene-co-divinylbezene) PLOT columns of 10 μm i.d. for the ultratrace LC-MS analysis of proteomics samples (Figure 5).109 A 4.2 m long PLOT column with a layer thickness of 0.5−1 μm yielded a peak capacity of approximately 400 for the separation of a complex tryptic digest mixture applying a 3.5 h gradient. The detection sensitivity was determined by coupling the PLOT column to a linear ion-trap MS, analyzing a 10 amol injection of BSA tryptic peptides. On the basis of the peptide yielding the highest MS response, the detection limit was determined in the low attomol range. Thakur et al. described the comparative proteomic LC-MS analysis of disease tissues which included the collection of 10 000 primary and metastatic breast cancer cells via laser capture microdissection, sample cleanup, and protein level fractionation using short-range SDS−PAGE, followed by LC-MS/MS analysis using a PLOT column.110 Informatics analysis of the resulting data indicated that vesicular transport and extracellular remodeling processes were significantly altered between the two cell types. A 3 m long PS-DVB PLOT column 549
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A generic way to partly alleviate the detection problem of OTLC systems mentioned in the introduction is to apply thick coating layers that allow for the injection of larger volumes. Some renewed interest in capillary columns coated with thicker films was shown in 2006 with practical investigations of the OTLC columns for capillary electrochromatography.116 Very recently, Collins et al. described the preparation of monolithic PLOT columns in 100 μm i.d. Teflon-coated capillaries using a controlled ultraviolet photoinitiated fabrication method.117 The approach allowed for the preparation of columns of varying length because of an automated capillary delivery approach, with precisely controlled and uniform layer thickness and monolith morphology, from controlled UV power and exposure time (Figure 6).
Figure 5. Gradient nanoLC/ESI-MS separation of a mixture of tryptic peptides of bovine serum albumin: β-casein, a Lys-C using a 4.2 m long × 10 μm i.d. poly(styrene-co-divinylbenzene) PLOT column applying a flow rate of 20 nL/min, and a 45 min aqueous acetonitrile gradient with 0.1% formic acid as an ion-pairing agent. Reprinted from ref 109. Copyright 2007 American Chemical Society.
was applied by Rogeberg et al. for the separation of intact proteins.111 A maximum peak capacity of 185 was reported with the application of a 90 min gradient. A proteomics strategy using a 2.5 m long × 10 μm i.d. coated (PS-DVB) capillary column coupled to a linear ion trap (LTQ) collision-induced dissociation/electron transfer dissociation mass spectrometer was developed by Wang et al. for the sensitive site-specific characterization of N-linked protein glycosylation.112 Three LC-MS data-dependent runs were initially employed for glycopeptide identification (ETD) and glycan structure evaluation and quantitation (CID), followed by four additional targeted LC-MS/MS analyses (CID-MS/MS) on additional glycoforms for further in-depth glycan structure determination. Finally, three additional runs were conducted for glycosylation occupancy determination. The PLOT column technology led to good glycopeptide recovery, and the long column provided a high resolving power. Furthermore, the ultralow flow rate of 20 nL/min significantly enhanced the glycopeptide ionization efficiency. Luo et al. described the preparation of a 10 μm i.d. amine-bonded poly(vinylbenzyl chloride-divinylbenzene) hydrophilic interaction (HILIC) chromatographic analysis of glycan mixtures.113 As an example of the high sensitivity of the column, MS6 characterization of glycan structures was possible from the injection of 10 fmol of a neutral and sialylated glycan. In addition, the high LC-MS sensitivity was demonstrated with the analysis of 3 ng of a PNGase F digest of ovalbumin, which allowed 28 N-linked glycans to be confidently identified from a single analysis. The Karger group also designed a 2D-LC setup using a long 2D PLOT column.114,115 The sample was first loaded onto a 100 μm i.d. 1D column packed with 5 μm SCX particles and eluted using a salt step gradient onto a precontration column (50 μm i.d. polymer monolithic column). Five ion-exchange fractions were then sequentially analyzed on a 3.2 m long high-efficiency 2D PLOT column applying a 45 min RP gradient at 20 nL/min. The applicability of the 2D-LC approach was demonstrated with the analysis of an in-gel tryptic digest sample of a gel fraction of SiHa cells.
Figure 6. Scanning electron micrograph of a porous polymer layer formed in a 100 μm i.d. capillary via UV irradiation. Reprinted from ref 117. Copyright 2012 American Chemical Society.
The fact that liquid chromatography can afford stationaryphase coatings that are relatively thick compared to gas chromatography is due to the fact that liquid-phase diffusion coefficients are much more comparable to the typical stationaryphase diffusion coefficients than the gas-phase diffusion coefficient. Poppe and co-workers already pointed this out long ago.118 In fact, it can be shown that an optimized OT-LC system is one where the mass transfer resistance in the mobile phase is comparable to that in the stationary-phase layer.119 Given that in liquid chromatography the diffusion in the mesoporous support layer is generally only a factor of 2 to 4 slower than that in the mobile phase, and given that the layer thickness appears to the second power in the mass transfer resistance term, this implies that the ratio between film thickness and the diameter of the channel lumen can be as large as 0.5 to 1 without running into significant additional band-broadening problems. In a recent study, the optimal layer thickness of wall-coated OT-LC systems has been investigated in great detail, taking into account the peculiar shape effect originating from the annular geometry of thick coating layers.120 The use of thick films also shifts the optimum column diameter to larger and more practical values (from 1 to 3 μm for thin film columns to 4−6 μm for thick-film columns). In this way, part of the potential kinetic advantage of OT-LC-columns compared to packed-bed columns is sacrificed, without losing it completely (in Figure 1, the performance of thick-film columns would be shifted more to the left than the represented OT-LC curves, which were plotted for the thin-film cases). Still, a factor of 2−4 in analysis time reduction is to be 550
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been conducted by the group of Tallarek.52,129 Ehlert et al.130 found that high bed densities are critical to separation performance in noncylindrical packed beds because the hydrodynamic dispersion in noncylindrical packings only comes close to that of cylindrical packings at low bed porosities. At higher bed porosities, the presence of preferential fluid channels in the corners of the noncylindrical packings adversely affects the hydrodynamic dispersion. Jung et al.128 found that the desired high packing densities can be much more easily achieved using 3 μm packings than with 5 μm particles, because of the poor accessibility to the corners of the conduit shape for these particles. Packed with the 3 μm particles and using a prototype UV detection cell of 50 μm i.d. installed behind the outlet frit, efficiencies corresponding to a reduced plate height of about hmin = 3 could be realized. Whereas packing the microchannels of chip-based systems proves to be tedious, the obvious solution to circumvent this problem is to resort to monolithic packings. Levkin et al. demonstrated the preparation of methacrylate-based and styrene-based monolithic phases with the confines of polyimide chips, featuring channels having cross sections of 200 × 200 μm, by a thermally initiated free-radical polymerization reaction.126 The SEM image of the cross section of the separation channel filled with styrene monolithic material shows that monoliths appear to adhere well to the wall surface of the channel. The chips were used for the separation of a test mixture of four intact proteins and a cytochrome c digest (Figure 7). Today, the flat
expected, while significantly increasing the mass loadability and also alleviating some more practical manufacturing problems (cf. the tendency to easily clog with the high pressure drop occurring when applying the often viscous coating solutions). Causon et al. also observed that the operation at elevated pressures shifted the optimal column diameters to even larger values (on the order of 5−9 μm), indicating that the first breakthrough of OT-LC applications can be expected in the hightemperature separation area.120 Assuming that a capillary diameter of 5 μm is at the edge of what is practically feasible from a manufacturing perspective, even these high-temperature applications will still be limited to the area of ultrahigh efficiency separations (requiring plate numbers in the 100 000 to 1 000 000 range), inevitably also requiring very long analysis times (on the order of one to several hours). Easier (and thus faster) separations still require capillaries with a diameter of only a few micrometer in order to compete with the newest generation of 1−2 μm fully porous or core-shell particles.14,55
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CHIP-BASED SYSTEMS Probably the most important new trend in the field of LC miniaturization introduced in the past decade is the increasing interest in so-called flat format columns, here, further referred to as chip-based systems. This new technology can be considered an offspring from the very successful lab-on-a-chip and microfluidics research areas.121−123 Its advantages are currently being exploited at two different levels. At the first level (discussed in Chip Systems Packed with Particles or Monolithic Beds), the flat format is preferred for its easy (in terms of manufacturing) and seamless integration with pre- and postcolumn separation and reaction steps, as well as for the minimal extra-column connection volumes that can be realized. Other advantages of the flat column format, as already mentioned in Recent Drivers for the Miniaturization of LC Systems, are the easy integration with localized heating/cooling elements.44 At the second level (discussed in Chip Systems with Micromachined Supports), the chip-based systems also involve a change in the support morphology, using the high resolution micromachining technology that is nowadays available to leave the perfectly ordered arrays of silicon or glass pillars. Chip Systems Packed with Particles or Monolithic Beds. An important breakthrough in the area of chip-based columns has been the elimination of the connection volumes using planar connections and the clamping force of commercial rotor-stator valves to directly clamp the chip between the rotor and the stator of the injection valve. The first such concept has been introduced by Yin et al.26,124−128 Using lasers to ablate small channels on polyimide chips, microchannels were fabricated that form a directly connected system of a column and an ESI-spray emitter. As there are no external connections (just a planar clamp-on interface) and as the connections within the chip require no fitting, this generic approach allows for the maximal elimination of dead volumes. The chips consist of a 75 μm i.d. column in a length of 43 or 150 mm and a 40 or 160 nL enrichment column, and they were initially packed with conventional 5 μm particles, which ensure easy method transfers from wide-bore columns. Packing the channels with particles to produce beds that are sufficiently homogeneous to achieve the same reduced plate-height performance as possible in cylindrical channels, however, proved less straightforward. This was mainly because of the trapezoidal form of the channels, the latter being the consequence of the employed laser ablation process. A complete study of the impact of this noncircular geometry has
Figure 7. SEM image of the cross section of the separation channel in a polyimide chip willed with a poly(styrene-co-divinylbenzene) monolith. Reprinted with permission from ref 126. Copyright 2008 Elsevier.
chip format and the clamp-on connection mode has also been adopted by other manufacturers.131 Chip Systems with Micromachined Supports. The concept of miniaturized LC systems using micromachined supports has been introduced by Regnier and co-workers in a series of landmark papers.132−134 Apart from potentially offering a column-to-column reproducibility that is much higher than possible with conventional packed-bed columns, the use of photolithography and reactive ion-etching technologies to create an array of supports (further referred to as pillars) on the surface of a silicon or glass wafer has the additional advantage that the 551
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A major problem with the use of etched pillars is their lack of a retention surface. A proposed first attempt to increase the pillar surface area has been the in situ depositon of thick silicon oxide layers using plasma-enhanced chemical vapor deposition (PECVD) or thermal growth processes.27,147 Both strategies at the same time also allow for the reduction of the width of the through pores formed between the pillars. Another approach has been the in situ growth of carbon nanotubes.148,149 Fonverne et al. could show preliminary chromatographic experiments with a mixture of two amines.150 Mogensen et al. demonstrated an electrochromatographic separation of two Coumarin dyes on a CNT column with an acetonitrile content of 90%.151 In yet another approach, De Malsche et al. used an electrochemical anodization process to produce pillars with a relatively thick layer of a mesoporous silicon layer, thus forming the column152 (layer depths could be varied from 0 to 0.6, 1.0, and 1.4 μm for a total pillar diameter of 5 μm). The obtained hmin values were significantly lower than in the packed beds with coreshell particles (h = 0.4−0.5 under nonretained conditions and hmin = 0.9 under retained conditions, even at retention factors up to k′ ≅ 12), as a reflection of the perfect homogeneity of the pillar beds (Figure 9). Detobel et al. proposed a hybrid approach combining the silica monolith and the pillar array technology, by performing the synthesis of siloxane-based monoliths in the presence of a twodimensional, perfectly ordered array of micropillars.153,154 They found that the formed structures were to a large extent nearly exclusively determined by the ratio between the bulk domain size of the monolith on the one hand and the distance between the micropillars on the other hand. When this ratio is small, the presence of the pillars has nearly no effect on the morphology of the produced monoliths (Figure 10a). However, when the ratio approaches unity and ascends above it, some new types of monolith morphologies are induced, two of which appear to have interesting properties for use as novel chromatographic supports. One of these structures (obtained when the domain size/ interpillar distance ratio is around unity) is a 3D network of linear interconnections between the pillars, organized such that all skeleton branches are oriented perpendicular to the micropillar surface (Figure 10b). A second interesting structure is obtained at even higher values of the domain size/interpillar distance ratio. In this case, each individual micropillar is uniformly coated with a mesoporous shell (Figure 10c).153,154 With the use of the latter
pillars in the bed can be arranged in a perfectly ordered conformation (Figure 8).135−142 The latter offers a large
Figure 8. SEM image showing the transverse section of a pillar-array column near the side-wall region of a bed packed with pillars of 4 μm in diameter. Reprinted with permission from ref 142. Copyright 2012 Wiley-VCH Verlag GmbH & Co. KGaA.
potential gain (about a factor of 2−3) as compared with a randomly packed-bed column.143 Apart from an improved efficiency, pillar array columns can also be designed so that the flow resistance can be drastically reduced compared to that in a packed bed of spheres.144 This is due to the fact that the distance between the pillars can be independently controlled outside of the pillar size, such that the external porosity, ε, is not restricted to values on the order of ε = 0.36 to ε = 0.4, as is inevitably the case for random packed-bed columns. When designing pillar array columns with an external porosity on the order of ε = 0.5− 0.6, much more permeable beds can be made. These higher porosities lower the separation impedance of the produced channels considerably. Using nonporous pillar array columns carrying only a C18 coating on the outer surface of the pillars, separation impedances on the order of 50 (nonretained analytes) to 150 (retained analytes with k′ = 0.23) have been realized.146 In terms of reduced plate heights, values as low as h = 0.2 to 0.9 were obtained.
Figure 9. Reduced van Deemter curves (h = H/pillar diameter) for a porous-shell pillar array (pillar diameter = 5 μm, shell thickness = 1 μm) recorded for a coumarin dye (C480) in different water−methanol compositions (65, 60, 55, and 45 v/v% water/methanol). Reprinted from ref 152. Copyright 2008 American Chemical Society. 552
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operate close to the Knox and Saleem limit of micropillar array columns in the region of the one million theoretical plate mark under the prevailing pressure restriction (350 bar). The obtained efficiency was slightly affected (some 15 to 20% around the optimal flow rate) by the turns that were inevitably needed to arrange a 3 m long column on a 4 in. silicon wafer.
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Figure 10. SEM images of three different morphologies obtained by the in situ synthesis of three different silica sol−gel mixtures yielding (a) a large, (b) an intermediate, and (c) a small domain size inside micropillararray columns with pillar diameters of 2.4 μm and an interpillar distance of 3.6 μm. Reprinted with permission from ref 153. Copyright 2009 Elsevier.
PERSPECTIVES On the academic front, today the field is clearly still pushing the limits of miniaturization. In pursuit of the ultimate degree of miniaturization, Singhal et al. very recently succeeded in bringing OT-LC into the nanometer realm.160 They demonstrated that even a single template-grown carbon nanotube can be used as a separation column to separate attoliter volumes of binary mixtures of fluorescent dyes.160 The employed columns were 40 μm long and were only 70 to 200 nm in outside diameter (o.d.) and 60 to 190 nm in inside diameter (i.d.). The detection mode was fluorescence microscopy. Other work on separations in nanochannels is from the Kitamori group.161 Using 1 μm wide and 400 nm deep channels, they realized a normal phase separation of a 10 fL sample with a significant number of theoretical plates (1 000 plates) and in a relatively short separation time (4 s). When shifting to other driving forces, such as shear-driven forces, even faster nanochannel separations come within reach.162 Closer to daily practice, it seems that without a radical breakthrough in detection technology sensitivity, miniaturized LC systems will for many years to come still only be used for special applications (limited sample volume, need for extreme peak capacity that can only be achieved by applying several 1 000 bar of pressures, use of rapid temperature pulses for trapping or selectivity enhancement, etc.). However, noticing a clear shift toward smaller diameters in the use of normal-bore columns (from 4.6 to 2.1 mm and a clear interest in 1 mm columns in pursuit of a better ESI-MS compatibility and a lower mobile phase consumption), it seems likely that both fields in the future will converge into one standard 0.5 to 1 mm i.d. column format, possibly packed with 1 μm core-shell particles and operated at several thousand bars. Given our poor understanding of the packing process, especially in this range of column diameters, a lot of work still needs to be done, however, before such columns can be expected on the market. For OT-LC, where intrinsic orders are faster than packed-bed columns but much more limited in detector sensitivity, the first applications are certainly expected at high temperatures, as these tolerate somewhat wider column diameters than low temperatures. For monolithic packings, the field is still waiting for synthesis procedures yielding structures with much smaller domain sizes in order to cash-in on the intrinsic advantage of their lower separation impedance (mainly originating from the lower flow resistance and the higher external porosity).
option to produce a rectangular pillar-array column (1 mm in width, 29 μm in height, and 33.75 mm in length filled with micropillars clad with a 0.5 μm thick porous-shell layer of mesoporous silica), minimal plate heights ranging between 3.9 (nonretaining conditions) and 6 μm (for a retention factor of 6.5) were obtained, corresponding to the domain size-reduced plate heights ranging between 0.7 and 1.2.154 Whereas the efficiency of pillar array columns is unprecedented in terms of reduced plate heights, the absolute plate heights are mainly still on the order of an ∼5 μm tolerance. Recently, Lavrik et al. pushed the minimal plate height even lower to 0.76 μm by fabricating pillars with diameters of only 1.9 μm and a spacing of about 1 μm, using PECVD deposition of SiO2 after the etching step to reduce the interpillar distance.140 To reach even smaller plate heights, smaller pillars and especially smaller interpillar distances are needed. However, it seems that when attempting to pursue these smaller interpillar distances, one runs into a very limiting case wherein the distance between the pillars becomes so small (order of 500 nm) that they can no longer be individually resolved, even when using the most refined deep-UV lithography equipment.145 Yet another hurdle that needs to be overcome before pillararray columns will be able to compete with the more conventional packed-bed capillaries is the fact that, currently, a large part of the efficiency measured on-chip is lost to the connection volumes when connecting a pillar array column to the injector and detector of a commercial capillary LC instrument.155,156 To alleviate the connection problem, the obvious strategy is to focus on long columns. Given the finite dimensions of a silicon wafer (usually 4 or 6 in. diameter), long column lengths can only be achieved by making several turns.157 Aoyama et al. published a study on a serpentine PAC with a total length of 110 mm, containing turns filled with micropillars and designed according to the work of Griffiths et al.139,158 The turns gradually decreased from a channel width of 400 μm to a circular turn with a width of 110 μm. With these turns, the plate height (corrected for injection volume) only increased from 2.7 μm (value measured in a straight part of the column) to 3.5 μm (value incorporating the effect of the turns) for the case of a nonretained component eluting at a mobile phase velocity of about 1.3 mm/s. In their latest work, the Tsunoda group have now also incorporated an online gradient elution system using a cross-Tesla mixer.157 Fluorescent derivatives of aliphatic amines were successfully separated within 110 s. The results show that the proposed system is promising for the analyses of complex biological samples. Very recently, De Malsche et al. succeeded in realizing over one million theoretical plates in a dead time of 20 min.159 This was achieved using a micropillar array column with an optimized pillar diameter (5 μm) and interpillar distance (2.5 μm) to
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Tel: +32-2-6293251. Fax: +32-26293248. Notes
The authors declare no competing financial interest. Biographies Gert Desmet has a Masters degree in chemical engineering (1990) and a Ph.D. in chemical engineering (1996), both from the Vrije Universiteit Brussel, in Brussels, Belgium. He is currently a full professor at the same 553
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(28) Illa, X.; De Malsche, W.; Gardeniers, H.; Desmet, G.; RomanoRodríguez, A. J. Chromatogr., A 2010, 1217, 5817−5821. (29) Illa, X.; De Malsche, W.; Gardeniers, H.; Desmet, G.; RomanoRodríguez, A. J. Sep. Sci. 2010, 33, 3313−3318. (30) Nunes, P.; Ohlsson, P.; Ordeig, O.; Kutter, J. Microfluid. Nanofluid. 2010, 9, 145−161. (31) Groskreutz, S. R.; Swenson, M. M.; Secor, L. B.; Stoll, D. R. J. Chromatogr., A 2012, 1228, 31−40. (32) Vivó-Truyols, G.; van der Wal, S.; Schoenmakers, P. J. Anal. Chem. 2010, 82, 8525−8536. (33) Sommella, E.; Cacciola, F.; Donato, P.; Dugo, P.; Campiglia, P.; Mondello, L. J. Sep. Sci. 2012, 35, 530−533. (34) Bedani, F.; Kok, W. Th.; Janssen, H.-G. J. Chromatogr., A 2006, 1133, 126−134. (35) Eeltink, S.; Dolman, S.; Vivo-Truyols, G.; Schoenmakers, P.; Swart, R.; Ursem, M.; Desmet, G. Anal. Chem. 2010, 82, 7015−7020. (36) Wu, Y.; Wang, Y.; Gu, X.; Zhang, L.; Yan., C. J. Sep. Sci. 2011, 34, 1027−1034. (37) Wei, J.; Gu, X.; Wang, Y.; Wu, Y.; Yan, C. Electrophoresis 2011, 32, 230−237. (38) Cesla, P.; Fischer, J.; Jandera, P. Electrophoresis 2010, 31, 2200− 2210. (39) Verstraeten, M.; Pursch, M.; Eckerle, P.; Luong, J.; Desmet, G. Anal. Chem. 2011, 83, 7053−7060. (40) Eghbali, H.; Sandra, K.; Tienpont, B.; Eeltink, S.; Sandra, P.; Desmet, G. Anal. Chem. 2012, 84, 2031−2037. (41) Chen, M. H.; Horváth, C. J. Chromatogr., A 1997, 788, 51−59. (42) Causon, T. J.; Cortes, H.; Shellie, R. A.; Hilder, E. F. Anal. Chem. 2012, 84, 3362−3368. (43) Verstraeten, M.; Desmet, G. J. Sep. Sci. 2011, 34, 2822−2832. (44) Collins, D.; Nesterenko, E.; Connolly, D.; Vasquez, M.; Macka, M.; Brabazon, D.; Paull, B. Anal. Chem. 2011, 83, 4307−4313. (45) Gu, B.; Cortes, H.; Luong, J.; Pursch, M.; Eckerle, P.; Mustacich, R. Anal. Chem. 2009, 81, 1488−1595. (46) Knox, J. H.; Parcher, J. F. Anal. Chem. 1969, 41, 1599−1606. (47) Kennedy, R. T.; Jorgenson, L. W. Anal. Chem. 1989, 61, 1128− 1135. (48) Knox, J. H.; Grant, I. Chromatographia 1987, 24, 135−145. (49) Broeckhoven, K.; Desmet, G. J. Chromatogr., A 2007, 1172, 25− 39. (50) Eeltink, S.; Rozing, G. P.; Schoenmakers, P. J.; Kok, W. Th. J. Chromatogr., A 2004, 1044, 311−316. (51) Ehlert, S.; Rosler, T.; Tallarek, U. J. Sep. Sci. 2008, 31, 1719−1728. (52) Khirevich, S.; Höltzel, A.; Ehlert, S.; Seidel-Morgenstern, A.; Tallarek, U. Anal. Chem. 2009, 81, 4937−4945. (53) Jung, S.; Ehlert, S.; Mora, J. A.; Kraiczek, K.; Dittmann, M.; Rozing., G. P.; Tallarek, U. J. Chromatogr., A 2009, 1216, 264−273. (54) Bruns, S.; Grinias, J. P.; Blue, L. E.; Jorgenson, J. W.; Tallarek, U. Anal. Chem. 2012, 84, 4496−4503. (55) Franklin, E.; Jorgenson, J. Long Capillary Columns Packed with Sub-2 micron Particles for Proteomic Applications. Presented at the 36th International Symposium on Capillary Chromatography (ISCC), Riva del Garda, Italy, May 27−June 1, 2012. (56) Franklin, E. Utilization of Long Columns Packed with sub-2 μm Particles Operated at High Pressures and Elevated Temperatures for High-Efficiency One-Dimensional Liquid Chromatographic Separations. Ph.D. Thesis, University of North Carolina at Chapel Hill, 2012 (57) Minakuchi, H.; Nakanishi, K.; Soga, N.; Ishizuka, N.; Tanaka, N. Anal. Chem. 1996, 68, 3498−3501. (58) Tanaka, N.; Ishizuka, N.; Hosoya, K.; Kimata, K.; Minakuchi, H.; Nakanishi, K.; Soga, N. Kuromatogurafi 1993, 14, 50−51. (59) Tennikova, T. B.; Svec, F.; Belenkii, B. G. J. Liquid Chromatogr. 1990, 13, 63−70. (60) Svec, F.; Fréchet, J. M. J. Anal. Chem. 1992, 64, 820−822. (61) Eeltink, S.; Geiser, L.; Svec, F.; Fréchet, J. M. J. J. Sep. Sci. 2007, 30, 2814−2820. (62) Hara, T.; Makino, S.; Watanabe, Y.; Ikegami, T.; Cabrera, K.; Smarsly, B.; Tanaka, N. J. Chromatogr., A 2010, 1217, 89−98.
University, where he also heads the department of chemical engineering and teaches courses on bioreactor design, reactor engineering, nano- and microbiotechnology, and chromatography in both the faculty of sciences and bioengineering and the faculty of engineering. His research focuses on the miniaturization of separation methods, the development of novel separation schemes, and the investigation and modeling of flow effects in chromatographic systems. Sebastiaan Eeltink received his Ph.D. in Chemistry (specialization Analytical Chemistry) in 2005 from the University of Amsterdam, The Netherlands. From 2005−2007, he conducted postdoctoral research at the University of California, Berkeley, California, and was a guest scientist at the Lawrence Berkeley National Laboratory. In 2007, he joined Dionex and conducted research on packed and monolith column technology for ultrahigh-pressure LC, two-dimensional LC, and nanoLC. In 2008, the National Fund for Scientific Research (FWO, Belgium) awarded him an Odysseus grant, and he currently holds a position as an assistant professor at the Department of Chemical Engineering at the Free University of Brussels.
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REFERENCES
(1) Ishii, D.; Takeuchi, T. Rev. Anal. Chem. 1982, 6, 87−112. (2) Novotny, M. Anal. Chem. 1981, 53, 1294A−1296A. (3) Knox, J. H. J. Chromatogr. Sci. 1980, 18, 453−461. (4) Chervet, J. P.; Ursem, M.; Salzmann, J. B. Anal. Chem. 1996, 68, 1507−1512. (5) Gritti, F.; Guiochon, G. J. Chromatogr., A 2012, 1236, 105−114. (6) Eeltink, S.; Dolman, S.; Swart, R.; Ursem, M.; Schoenmakers, P. J. J. Chromatogr., A 2009, 1216, 7368−7374. (7) Zhou, F.; Lu, Y.; Ficarro, S. B.; Webber, J. T.; Marto, J. A. Anal. Chem. 2012, 84, 5133−5139. (8) LaCourse, W. R. Anal. Chem. 2002, 74, 2813−2832. (9) Vissers, J. P. C. J. Chromatogr., A 1999, 856, 117−143. (10) Poppe, H. J. Chromatogr., A 1997, 778, 3−21. (11) Poppe, H. Chromatography, Fundamentals and Applicatons of Chromatograph and Related Differential Migration Methods. In Journal of Chromatography Library; Heftmann, E., Ed.; Elsevier: Amsterdam, 1992; 151−225. (12) Shen, Y.; Zhao, R.; Berger, S. J.; Anderson, G. A.; Rodriguez, N.; Smith, R. D. Anal. Chem. 2002, 74, 4235−4249. (13) Billen, J.; Desmet, G. J. Chromatogr., A 2007, 1168, 73−99. (14) Gritti, F.; Guiochon, G. J. Chromatogr., A 2011, 1218, 1915−1938. (15) Guiochon, G. Anal. Chem. 1981, 53, 1318−1325. (16) Broeckhoven, K.; Cabooter, D.; Lynen, F.; Sandra, P.; Desmet, G. J. Chromatogr., A 2010, 1217, 2787−2795. (17) MacNair, J. E.; Patel, K. D.; Jorgenson, J. W. Anal. Chem. 1999, 71, 700−708. (18) Patel, K. D.; Jerkovich, A. D.; Link, J. C.; Jorgenson, J. W. Anal. Chem. 2004, 76, 5777−5786. (19) Jerkovich, A. D.; Mellors, J. S.; Jorgenson, J. W. LC·GC Eur. 2003, 16, 20−23. (20) Tolley, L.; Jorgenson, J. W.; Moseley, M. A. Anal. Chem. 2001, 73, 2985−2991. (21) Desmet, G. J. Chromatogr., A 2006, 1116, 89−96. (22) Broeckhoven, K.; Billen, J.; Verstraeten, M.; Choikhet, K.; Dittmann, M.; Rozing, G.; Desmet, G. J. Chromatogr., A 2010, 1217, 2022−2031. (23) Livesay, E. A.; Tang, K.; Taylor, B. K. Anal. Chem. 2008, 80, 294− 302. (24) Madou, M. J. Fundamentals of Microfabrication and Nanotechnology; 3rd Ed.; CRC Press: Boca Raton, FL, 2011; Vol. 2. (25) Franssila, S. Introduction to Microfabrication; 2nd Ed.; John Wiley & Sons, Ltd: Chichester, West Sussex, England, 2010. (26) Yin, H. F.; Killeen, K. J. Sep. Sci. 2007, 30, 1427−1434. (27) Gustafsson, O.; Mogensen, K. B.; Kutter, J. P. Electrophoresis 2008, 29, 3145−52. 554
dx.doi.org/10.1021/ac303317c | Anal. Chem. 2013, 85, 543−556
Analytical Chemistry
Review
(63) Nunez, O.; Nakanishi, K.; Tanaka, N. J. Chromatogr., A 2008, 1191, 231−252. (64) Hara, T.; Kobayashi, H.; Ikegami, T.; Nakanishi, K.; Tanaka, N. Anal. Chem. 2006, 78, 7632−7642. (65) Tanaka, N.; Kobayashi, H.; Ishizuka, N.; Minakuchi, H.; Nakanishi, K.; Hosoya, K.; Ikega, I, T. J. Chromatogr., A 2002, 965, 35−49. (66) Morisato, K.; Miyazaki, S.; Ohira, M.; Furuno, M.; Nyudo, M.; Terashima, H.; Nakanishi, K. J. Chromatogr., A 2009, 1216, 7384−7387. (67) Miyamoto, K.; Hara, T.; Kobayashi, H.; Morisaka, H.; Tokuda, D.; Horie, K.; Koduki, K.; Makino, S.; Nunez, O.; Yang, C.; Kawabe, T.; Ikegami, T.; Takubo, H.; Ishihama, Y.; Tanaka, N. Anal. Chem. 2008, 80, 8741−8750. (68) Eghbali, H.; Sandra, K.; Detobel, F.; Lynen, F.; Nakanishi, K.; Sandra, P.; Desmet, G. J. Chromatogr., A 2011, 1218, 3360−3366. (69) Horie, K.; Sato, Y.; Kimura, T.; Nakamura, T.; Ishihama, Y.; Oda, Y.; Ikegami, T.; Tanaka, N. J. Chromatogr., A 2012, 1228, 283−291. (70) Iwasaki, M.; Miwa, S.; Ikegami, T.; Tomita, M.; Tanaka, N.; Ishihama, Y. Anal. Chem. 2010, 82, 2616−2620. (71) Dioszegi, T. A.; Raynie, D. E. J. Chromatogr., A 2012, 1261, 107− 112. (72) Vaast, A.; Broeckhoven, K.; Dolman, S.; Desmet, G.; Eeltink, S. J. Chromatogr., A 2012, 1228, 270−285. (73) Viklund, C.; Svec, F.; Fréchet, J. M. J.; Irgum, K. Chem. Mater. 1996, 8, 744−750. (74) Peters, E. C.; Svec, F.; Fréchet, J. M. J.; Viklund, C.; Irgum, K. Macromolecules 1999, 32, 6377−6379. (75) Svec, F. J. Sep. Sci. 2004, 27, 1419−1430. (76) Arrua, R. D.; Talebi, M.; Causon, T. J.; Hilder, E. F. Anal. Chim. Acta 2012, 738, 1−12. (77) Svec, F. J. Chromatogr., A 2010, 1217, 902−924. (78) Zhang, Z. B.; Wang, F. J.; Xu, B.; Qin, H. Q.; Ye, M. L.; Zou, H. F. J. Chromatogr., A 2012, 1256, 136−143. (79) Xie, S.; Allington, R. W.; Svec, F.; Fréchet, J. M. J. J. Chromatogr., A 1999, 865, 169−174. (80) Huber, C. G.; Krajete, A. Anal. Chem. 1999, 71, 3730−3739. (81) Wouters, B.; Vaast, A.; Treumann, A.; Ursem, M.; Ho, J.; Hornshaw, M.; Raes, M.; Terryn, H.; Eeltink, S. LC·GC Eur. 2012, 25, 10−17. (82) Eeltink, S.; Wouters, B.; Desmet, G.; Ursem, M.; Blinco, D.; Kemp, G. D.; Treumann, A. J. Chromatogr., A 2011, 32, 5504−5511. (83) Detobel, F.; Broeckhoven, K.; Wellens, J.; Wouters, B.; Swart, R.; Ursem, M.; Desmet, G.; Eeltink, S. J. Chromatogr., A 2010, 1217, 3085− 3090. (84) Xie, S.; Svec, F.; Fréchet, J. M. J. J. Chromatogr., A 1997, 775, 65− 72. (85) Chen, M.-L.; Li, L.-M.; Y., B.-F.; Ma, Q.; Feng, Y. J. Chromatogr., A 2012, 1230, 54−60. (86) Lubbad, S. H.; Buchmeiser, M. R. J. Chromatogr., A 2010, 1717, 3223−3230. (87) Buchmeiser, M. R. J. Sep. Sci 2008, 31, 1907−1922. (88) Urban, J.; Eeltink, S.; Jandera, P.; Schoenmakers, P. J. J. Chromatogr., A 2008, 1182, 161−168. (89) Detobel, F.; Broeckhoven, K.; Wellens, J.; Wouters, B.; Swart, R.; Ursem, M.; Desmet, G.; Eeltink, S. J. Chromatogr., A 2010, 1217, 3085− 3090. (90) Eeltink, S.; Dolman, S.; Detobel, F.; Swart, R.; Ursem, M.; Schoenmakers, P. J. J. Chromatogr., A 2010, 1217, 6610−6615. (91) Hutchinson, J. P.; Hilder., E. F.; Macka, M.; Avadolivic, N.; Haddad, P. R. J. Chromatogr., A 2006, 1109, 10−18. (92) Arrua, R. D.; Nordborg, A.; Haddad, P. R.; Hilder, E. F. J. Chromatogr., A 2012, DOI: 10.1016/j.chroma.2012.10.068. (93) Walsh, Z.; Levkin, P.; Jain, V.; Paull, B.; Svec, F.; Macka, M. J. Sep. Sci. 2010, 33, 61−66. (94) Badaloni, E.; Barbarino, M.; Cabri, W.; D’Acquarica, I.; Forte, M.; Gasparinni, F.; Giorgio, F.; Pierinine, M.; Simone, P.; Ursini, O.; Villani, C. J. Chromatogr., A 2011, 1218, 3862−3875. (95) Krajnc, P.; Leber, N.; Stefanec, D.; Kontrec, S.; Podgornik, A. J. Chromatogr., A 2005, 1065, 69−73.
(96) Aoki, H.; Kubo, T.; Ikegami, T.; Tanaka, N.; Hosoya, K.; Tokuda, D.; Ishizuka, N. J. Chromatogr., A 2006, 1119, 66−79. (97) Nischang, I.; Svec, F.; Fréchet, J. M. J. J. Chromatogr., A 2009, 1216, 2355−2361. (98) Nischang, I.; Svec, F.; Fréchet, J. M. J. Anal. Chem. 2009, 81, 7390−7396. (99) Trojer, L.; Bisjak, C. P.; Wieder, W.; Bonn, G. K. J. Chromatogr., A 2009, 1216, 6303−6309. (100) Greiderer, A.; Trojer, L.; Huck, C. W.; Bonn, G. K. J. Chromatogr., A 2009, 1216, 7747−7754. (101) Nischang, I.; Bruggemann, O. J. Chromatogr., A 2010, 1217, 5389−5397. (102) Nischang, I.; Taesdale, I.; Bruggemann, O. J. Chromatogr., A 2010, 1217, 7514−7522. (103) Urban, J.; Svec, F.; Fréchet, J. M. J. J. Chromatogr., A 2010, 1217, 8212−8221. (104) Hue, Y.; Schoenmakers, P. J.; Kok, W. Th. J. Chromatogr., A 2007, 1175, 81−88. (105) Miyabe, K.; Guiochon, G. J. Sep. Sci. 2003, 26, 155−173. (106) Li, Y.; Tolley, H. D.; Lee, M. L. J. Chromatogr., A 2011, 1218, 1399−1408. (107) Liu, K.; Tolley, H. D.; Lee, M. L. J. Chromatogr., A 2012, 1227, 96−104. (108) Svobodova, A.; Krizek, T.; Sirc, J.; Salek, P.; Tesarova, E.; Coufal, P.; Stulik, K. J. Chromatogr., A 2011, 1218, 1544−1547. (109) Yue, G.; Quanzhou, L.; Zhang, J.; Wu, S.-L.; Karger, B. L. Anal. Chem. 2007, 79, 938−946. (110) Thakur, D.; Rejtar, T.; Wang, D.; Bones, J.; Cha, S.; ClodfelderMiller, B.; Richardson, E.; Binns, S.; Dahiya, S.; Sgroi, D.; Karger, B. L. J. Chromatogr., A 2011, 1218, 8168−8174. (111) Rogeberg, M.; Wilson, S. R.; Greibrokk, T.; Lundanes, E. J. Chromatogr., A 2012, 1217, 2782−2786. (112) Wang, D.; Hincapie, M.; Rejtar, T.; Karger, B. L. Anal. Chem. 2011, 83, 2029−2037. (113) Luo, A.; Rejtar, T.; Wu, S.-L.; Karger, B. L. J. Chromatogr., A 2009, 1216, 1223−1231. (114) Luo, Q.; Yue, G.; Valaskovic, G. A.; Gu, Y.; Wu, S.-L.; Karger, B. L. Anal. Chem. 2007, 79, 6174−6181. (115) Liu, Q.; Gu, Y.; Wu, S.-L.; Rejtar, T.; Karger, B. L. Electrophoresis 2008, 29, 1604−1611. (116) Eeltink, S.; Svec, F.; Fréchet, J. M. J. Electrophoresis 2006, 27, 4249−4256. (117) Collins, D. A.; Nesterenko, E. P.; Brabazon, D.; Paull, B. Anal. Chem. 2012, 84, 3465−3472. (118) Tock, P. P. H.; Duijsters, P. P. E.; Kraak, J. C.; Poppe, H. J. Chromatogr., A 1990, 506, 185−199. (119) Desmet, G.; Baron, G. V. J. Chromatogr., A 2000, 867, 23−43. (120) Causon, T. J.; Shellie, R. A.; Hilder, E. F.; Desmet, G.; Eeltink, S. J. Chromatogr., A 2011, 1218, 8388−8393. (121) Verpoorte, E. Lab Chip 2003, 3, 60N−68N. (122) Arora, A.; Simone, G.; Salieb-Beugelaar, G. B.; Kim, J. T.; Manz, A. Anal. Chem. 2010, 82, 4830−4847. (123) West, J.; Becker, M.; Tombrink, S.; Manz, A. Anal. Chem. 2008, 80, 4403−4419. (124) Yin, H. F.; Killeen, K.; Brennen, R.; Sobek, D.; Werlich, M.; van de Goor, T. Anal. Chem. 2005, 77, 527−533. (125) Brennen, R. A.; Yin, H. F.; Killeen, K. Anal. Chem. 2007, 79, 9302−9309. (126) Levkin, P. A.; Eeltink, S.; Stratton, T. R.; Brennen, R.; Robotti, K.; Yin, H.; Killeen, K.; Svec, F.; Fréchet, J. M. J. J. Chromatogr., A 2008, 1200, 55−61. (127) Bynum, M. A.; Yin, H. F.; Felts, K.; Lee, Y. M.; Monell, C. R.; Killeen, K. Anal. Chem. 2009, 81, 8818−8825. (128) Jung, S.; Höltzel, A.; Ehlert, S.; Mora, J. A.; Kraiczek, K.; Dittmann, M.; Rozing, G. P.; Tallarek, U. Anal. Chem. 2009, 81, 10193− 10200. (129) Khirevich, S.; Höltzel, A.; Hlushkou, D.; Tallarek, U. Anal. Chem. 2007, 79, 9340−9349. 555
dx.doi.org/10.1021/ac303317c | Anal. Chem. 2013, 85, 543−556
Analytical Chemistry
Review
(130) Ehlert, S.; Kraiczek, K.; Mora, J. A.; Dittmann, M.; Rozing, G. P.; Tallarek, U. Anal. Chem. 2008, 80, 5945−5950. (131) Gerhardt, G. Chromatography Today 2011, 13, 6−8. (132) He, B.; Tait, N.; Regnier, F. Anal. Chem. 1998, 70, 3790. (133) Regnier, F. E. J. High Resolut. Chromatogr. 2000, 23, 19. (134) Slentz, B. E.; Penner, N. A.; Regnier, F. J. Sep. Sci. 2002, 25, 1011. (135) De Pra, M.; Kok, W. Th.; Gardeniers, J. G. E.; Desmet, G.; Eeltink, S.; van Nieuwkasteele, J. W. Anal. Chem. 2006, 78, 6519. (136) Eijkel, J. Lab Chip 2007, 7, 815−817. (137) Gustafsson, O.; Mogensen, K. B.; Ohlsson, P. D.; Liu, Y.; Jacobson, S. C.; Kutter, J. P. J. Micromech. Microeng. 2008, 18, 5502. (138) Mogensen, K. B.; Eriksson, F.; Gústafsson, O.; Nikolajsen, R. P. H.; Kutter, J. P. Electrophoresis 2004, 25, 3788. (139) Aoyama, C.; Saeki, A.; Noguchi, M.; Shirasaki, Y.; Shoji, S.; Funatsu, T.; Mizuno, J.; Tsunoda, M. Anal. Chem. 2010, 82, 1420−1426. (140) Taylor, L. C.; Lavrik, N. V.; Sepaniak, M. J. Anal. Chem. 2010, 82, 9549−9556. (141) Lavrik, N. V.; Taylor, L. C.; Sepaniak, M. J. Anal. Chim. Acta 2011, 694, 6−20. (142) Op de Beeck, J.; De Malsche, W.; De Moor, P.; Desmet, G. J. Sep. Sci. 2012, 35, 1877−1883. (143) Gzil, P.; De Smet, J.; Vervoort, N.; Verelst, H.; Baron, G. V.; Desmet, G. J. Chromatogr., A 2004, 1030, 53. (144) De Smet, J.; Gzil, P.; Vervoort, N.; Verelst, H.; Baron, G. V.; Desmet, G. Anal. Chem. 2004, 76, 3716−3726. (145) Op de Beeck, J.; De Malsche, W.; Tezcan, D. S.; De Moor, P.; Desmet, G. J. Chromatogr., A 2012, 1239, 35−48. (146) De Malsche, W.; Eghbali, H.; Clicq, D.; Vangelooven, J.; Gardeniers, H.; Desmet, G. Anal. Chem. 2007, 79, 5915. (147) Lavrik, N. V.; Taylor, L. C.; Sepaniak, M. J. Lab Chip 2010, 10, 1086−1092. (148) Mogensen, K. B.; Kutter, J. P. Lab Chip 2012, 12, 1951−1958. (149) Fonverne, A.; Ricoul, F.; Demesmay, C.; Delattre, C.; Fournier, A.; Dijon, J.; Vinet, F. Sens. Actuators, B 2008, 129, 510−517. (150) Fonverne, A.; Demesmay, C.; Ricoul, F.; Rouvière, E.; Dijon, J.; Vinet, F. Sens. Actuators, A 2011, 167, 517−523. (151) Mogensen, K. B.; Chen, M.; Molhave, K.; Boggild, P.; Kutter, J. P. Lab Chip 2011, 11, 2116−2118. (152) De Malsche, W.; Gardeniers, H.; Desmet, G. Anal. Chem. 2008, 80, 5391−5400. (153) Detobel, F.; Eghbali, H.; De Bruyne, S.; Terryn, H.; Gardeniers, H.; Desmet, G. J. Chromatogr., A 2009, 1216, 7360−7367. (154) Detobel, F.; De Bruyne, S.; Vangelooven, J.; De Malsche, W.; Aerts, T.; Terryn, H.; Gardeniers, H.; Eeltink, S.; Desmet, G. Anal. Chem. 2010, 82, 7208−7217. (155) De Malsche, W.; De Bruyne, S.; Op De Beek, J.; Sandra, P.; Gardeniers, H.; Desmet, G.; Lynen, F. J. Chromatogr., A 2012, 1230, 41−47. (156) De Malsche, W.; De Bruyne, S.; Op De Beek, J.; Eeltink, S.; Detobel, F.; Gardeniers, H.; Desmet, G. J. Sep. Sci. 2012, 0, 1−8. (157) Song, Y.; Noguchi, M.; Takatsuki, K.; Sekiguchi, T.; Mizuno, J.; Funatsu, T.; Shoji, S.; Tsunoda, M. Anal. Chem. 2012, 84, 4739−4745. (158) Griffiths, S. K.; Nilson, R. H. Anal. Chem. 2001, 73, 272−278. (159) De Malsche, W.; Op De Beek, J.; De Bruyne, S.; Gardeniers, H.; Desmet, G. Anal. Chem. 2012, 84, 1214−1219. (160) Singhal, R.; Mochalin, V. N.; Lukatskaya, M. R.; Friedman, G.; Gogotsi, Y. Nature 2012, DOI: 10.1038/srep00510. (161) Ishibashi, R.; Mawatari, K.; Kitamori, T. J. Chromatogr., A 2012, 1238, 152−155. (162) Clicq, D.; Vervoort, N.; Vounckx, R.; Ottevaere, H.; Gooijer, C.; Ariese, F.; Baron, G. V.; Desmet, G. J. Chromatogr., A 2002, 979, 33−42.
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NOTE ADDED AFTER ASAP PUBLICATION After this paper was published on the Web on December 7, 2012, a correction was made to the paragraph underneath Chip-Based Systems. The corrected version was reposted on December 19, 2012.
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