On-line sample pre-concentration in microfluidic devices: A review

Analytica Chimica Acta 718 (2012) 11–24

Contents lists available at SciVerse ScienceDirect

Analytica Chimica Acta journal homepage: www.elsevier.com/locate/aca

Review

On-line sample pre-concentration in microfluidic devices: A review Braden C. Giordano a,∗ , Dean S. Burgi b , Sean J. Hart a , Alex Terray a a b

Naval Research Laboratory, Chemistry Division – Code 6112, 4555 Overlook Ave, SW, Washington, DC 20375-5342, United States dbqp, 767 Mahogany Ln, Sunnyvale, CA 94086-8638, United States

a r t i c l e

i n f o

Article history: Received 3 October 2011 Received in revised form 1 December 2011 Accepted 21 December 2011 Available online 2 January 2012 Keywords: Preconcentration Microfluidics Stacking Extraction

a b s t r a c t On-line sample preconcentration is an essential tool in the development of microfluidic-based separation platforms. In order to become more competitive with traditional separation techniques, the community must continue to develop newer and more novel methods to improve detection limits, remove unwanted sample matrix components that disrupt separation performance, and enrich/purify analytes for other chip-based actions. Our goal in this review is to familiarize the reader with many of the options available for on-chip concentration enhancement with a focus on those manuscripts that, in our assessment, best describe the fundamental principles that govern those enhancements. Sections discussing both electrophoretic and nonelectrophoretic modes of preconcentration are included with a focus on device design and mechanisms of preconcentration. This review is not meant to be a comprehensive collection of every available example, but our hope is that by learning how on-line sample concentration techniques are being applied today, the reader will be inspired to apply these techniques to further enhance their own programs. Published by Elsevier B.V.

Contents 1.

2.

3.

4. 5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. The microfluidic platform . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2. Preconcentration and enrichment options . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sample stacking in electrophoretic systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. First principles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.1. The Kohlrausch regulation function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2. Four fundamental modes of electrophoretic stacking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.3. Electroosmotic flow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.4. Other considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Electrophoretic preconcentration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.1. Field amplified stacking methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2. Moving boundary stacking methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.3. Isotachophoresis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.4. Isoelectric focusing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.5. Stacking in micellar-based separations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Extraction methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Solid-phase extraction (SPE) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.1. Biomacromolecule applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.2. Small molecule applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Non-SPE methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cell enrichment and preconcentration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

∗ Corresponding author. E-mail address: [email protected] (B.C. Giordano). 0003-2670/$ – see front matter. Published by Elsevier B.V. doi:10.1016/j.aca.2011.12.050

12 12 12 12 12 12 13 13 13 14 14 16 17 17 19 19 19 19 20 21 22 23 24

12

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

Braden C. Giordano is an analytical chemist with extensive experience in integrating functionality on microfluidic devices. He received his PhD from The University of Virginia where he worked on integrating PCR-based DNA amplification with sizedependent separations of diagnostic significance. His current efforts focus on understanding on-line sample stacking phenomena that occur in complex, high conductivity sample matrices in both capillary and microchip-based electrophoretic systems.

Sean J. Hart has extensive experience in a diverse range of research areas including spectroscopy, laser trapping and separation, optical engineering, physical chemistry, chemical and biochemical separation, statistical data analysis/calibration and artificial intelligence. He received his BS degree in Chemistry from Suffolk University in 1994 and PhD in Chemistry from Tufts University in 1998. Currently Dr. Hart has been pursuing novel particle separations using optical pressure. Applications include bulk sample bio-enrichment and purification, sample separation, detection and advanced characterization.

Dean S. Burgi is an analytical chemist working in the Bay Area bringing biotech products to market. He has been involved with capillary electrophoresis and on-line sample preconcentration techniques since the late 80s and was active in the Human Genome Project. Now he works on gene expression and protein analysis.

Alex Terray is a chemical engineer, obtained his M.S. in chemical engineering from the Colorado School of Mines in 2002. His research interests include the optical manipulation of microscopic biotic and abiotic particles inside microfluidic systems for applications including sample detection and separation.

1. Introduction 1.1. The microfluidic platform The microfluidic platform or microchip can be used to describe a large number of devices with various forms and functions. Typically, one envisions a planar substrate with a channel that mimics the function of the bare fused-silica capillaries used in capillary electrophoresis (CE) and small volume wells replacing sample and separation vials used in commercial instruments. The desire to integrate functionality on a single device has resulted in micro-total analysis system (␮TAS) devices that allow for multiple processes on a single platform to be performed in series. With this concept in mind, one may classify microfluidic devices as either a single purpose device or a multifunctional Lab-on-a-Chip device. A single purpose device would include standard cross-t configurations for separations, i.e. Ref. [1], or devices used exclusively for solid-phase extraction (SPE), i.e. Ref. [2]. A Lab-on-a-Chip device would include devices developed by several groups, for example, that incorporate solid-phase extraction, polymerase chain reaction (PCR)-based DNA amplification, and separation with detection and analysis on a single platform, i.e. Ref. [3]. In both cases, examples of on-line sample preconcentration and enrichment are desirable, and in some cases several instances of concentration enhancement are observed on a single platform. A more comprehensive description of microfluidic devices and there function, including the implementation of various integrated functionality options can be found in the Handbook of Capillary and Microchip Electrophoresis and Associated Microtechniques (3rd edition) edited by James P. Landers.

1.2. Preconcentration and enrichment options Sample preconcentration on a microfluidic device can be broadly classified into two categories – (1) electrophoretic methods and (2) nonelectrophoretic extraction methods. Electrophoretic methods describe techniques including field-amplified stacking (FAS) [4], isotachophoresis (ITP) [5], or isoelectric focusing (IEF)

[6]. Such techniques require careful consideration of electrical behavior in the sample matrix, and background electrolyte composition (BGE) and ion concentration. Other manipulations of channel dimensions, buffer composition, and timing allows analyte to preconcentrate at various points throughout the device. Preconcentration is also possible via non-electrical extraction techniques. These methods can take the form of on-chip vapor impinging [7], chips designed for liquid/liquid extraction [8], or the inclusion of packed beds for solid-phase extraction [2]. The design of the platform is only limited to the physical and electrical boundaries of the separation desired. Biological particles such as cells and spores have also been analyzed on-chip, with a focus on using microfluidics for preparative scale cell sorting [9] and sample clean-up [10]. Enriching samples and/or removing sample matrix components that could impede off-chip analyses are key features that serve to expand the utility of the microfluidic platform. Furthermore, the ability to select the cell type of interest based upon a variety of characteristics and concentrate it selectively will yield lower detection limits. These techniques are not classified as either electrophoretic or extraction-based preconcentration methods, but yet are no less important. 2. Sample stacking in electrophoretic systems 2.1. First principles Whether in a capillary-based system or on a microfluidic device, the bulk of on-line sample preconcentration techniques rely on manipulating sample and/or background electrolyte composition. This manipulation increases the concentration of an analyte of interest during the initial moments of a voltage being applied across the separation channel as the ions reach their steady state. 2.1.1. The Kohlrausch regulation function When the ionic solution is first placed in a shaped vessel, e.g. the capillary or microchannel, a numerical value, termed the

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

Kohlrausch value [11], related to the conductivity of the solution at every given point along the pathway, is established. The Kohlrausch value (ω) can be calculated as: ω=

cz  i i i  i

where ci is concentration of a given ion, zi is the relative charge of strong electrolytes, and i is the mobility. This value exists before the electric field is even applied and is maintained at each cross-section, ultimately determining how the ions will adjust their concentrations in effort to maintain that value during the period of time that a voltage is applied. Once the electric field is broken, a new Kohlrausch value can be calculated which will lead to a different movement of the ionic species once the electric field is reapplied. This movement of ionic species is the foundation of electrophoresis and it governs all the sample stacking phenomena observed in every named electrophoretic method including zone electrophoresis, moving boundary electrophoresis (MBE), isotachophoresis and isoelectric focusing. A very good review article by Hruˇska and Gaˇs on the Kohlrausch regulation function can be found in Ref. [12]. 2.1.2. Four fundamental modes of electrophoretic stacking Zone or stationary boundary electrophoresis is arguably the simplest electrophoretic system to understand. In this case, the sample matrix and the background electrolyte share a constant composition. Stationary boundaries form at the interface between the sample plug and the BGE, with the sample moving simply as a function of the electric field inside the sample plug; the flux across the sample matrix/BGE boundaries are zero. FAS occurs when the sample matrix is simply diluted background electrolyte; the sample moves faster in the high electric field associated with the sample zone and slows down upon encountering the sample plug/BGE interface. This mode of sample stacking is one of the most commonly used in microfluidic applications since it does not require a vast knowledge of the sample matrix composition, or complex microchip design in order to realize on-line sample preconcentration. Early examples of capillary-based FAS are summarized in a review by Chien and Burgi [13]. Moving boundary electrophoresis always occurs whenever there is a discontinuity in the constituents of the sample matrix and the BGE. The ions in the BGE sample matrix will move as a function of their mobility and stack according to the Kohlrausch values defined by the channel content prior to the application of a voltage. When a sample plug has a Kohlrausch value lower than that of the BGE, the constituents of the sample plug will stack at the BGE/sample plug interface, i.e. the front end of the sample plug. These stable moving boundaries are identified as system peaks in most separations and will move through the channel at the speed dictated by the ion mobility [14–17]. Isotachophoresis, as the name states, is a separation technique where all the ions are moving at the same speed. The interfaces are moving boundaries but have a finite time before they rearrange themselves to reach steady state. Inside the channel is a leading electrolyte (the fastest moving ion), followed by the sample and then the trailing electrolyte (slowest moving ion). The sample components separate from one another such that the fastest moving ion in the sample falls in line behind the leading electrolyte, then the next fastest ion, and so on and so forth until the slowest ion is nestled against the trailing electrolyte. Sample ion concentration will increase or decrease in order to maintain a constant speed for all ions flanked by the leading and trailing electrolytes. Transient ITP (tITP) is often used where ITP-based stacking is coupled to a zone electrophoresis-based separation of the stacked components. Various schematics of capillary-based tITP can be found in a review article by Bocek and co-workers [18].

13

Isoelectric focusing uses discrete pH zones set-up by the inclusion of ampholytes in the sample to produce a series of stable stationary boundaries that are defined by a particular pH. One end of the channel is filled with high pH solution and the other end is prepared with a low pH solution, such as 0.1 M NaOH and 100 mM phosphoric acid, respectively. Analytes of interest migrate through the pH zones until they reach a location of net neutral charge and stop moving. If diffusion moves the molecule out of its zone, the molecule will pick up a charge (either positive or negative) to move it back to its net zero point (pI) or isoelectric point. Incredibly narrow, high concentration bands are formed by this method. A very good review, by Righetti et al., discusses the genesis of IEF in capillary systems, including examples of various applications at the time the review was written [19]. All on-line electrophoretic-based techniques come from these four fundamental modes described above, and each is ultimately governed by the Kohlrausch value calculated prior to the application of an electric field. A discrete plug of analyte can be preconcentrated in an effort to satisfy this value as the compound of interest migrates through the separation channel. We encourage readers of any level to look to review articles indicated above and the original literature in order to develop a deeper understanding of the underlying electrophoretic principles that govern stacking in both capillary and microchip-based systems. 2.1.3. Electroosmotic flow The phenomenon called electroosmotic flow (EOF) is generated when a fluid comes in contact with a wetable surface [20]. The charge on the solid wall is counter balanced by a corresponding charged species in the fluid. For example, in the presence of an aqueous solution at a pH of 7.0, the wall is negativity charged and positive ions from the fluid strongly associate with that surface charge, coating the surface to form a double layer. This is also termed the Debye layer, defining a zeta potential that extends out into the fluid. When an electrical field is applied to the solution, the negative ions will move to the anode and the positive ions will move to the cathode, but since the wall cannot move in the electric field, a sheath of positive ions will move towards the cathode, carrying the rest of the fluid with it due to hydrogen bonding. Thus, when considering ions in a discrete sample plug, the sample’s positive ions reach the cathode first, neutral ions are carried by the flow, and the negative ions resist the flow due to their opposing mobility, albeit reaching the cathode provided their mobility is insufficient to overcome the EOF mobility. Unlike pressure driven flow systems where the flow profile is laminar, EOF can be best described as a plug flow; that is to say EOF has a flat flow profile. EOF mobility is defined as:

o =

ε 

where  is the zeta potential at the capillary wall, ε is the permittivity of the fluid in the capillary, and  is the viscosity of the solution. The ability to manipulate EOF in conjunction with the four modes of sample stacking described above are the key factors in the development of on-line electrophoretic stacking techniques. 2.1.4. Other considerations Another issue to consider when discussing electrophoretic preconcentration is the concept of adulteration of sample and separation wells used to hold liquid on the microchip substrate. Since electrokinetic injection is biased by ion mobility, the potential for depleting or adding ions to a well as a function of injection and separation time is a distinct possibility. This issue, often ignored when using commercial CE instruments, due to large vials to contain sample and BGE, may be exacerbated due to the much smaller volumes used in microfluidic devices (10–100 ␮L). To our knowledge, there

14

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

Fig. 1. (a) Cartoon illustrating the injection scheme and stacking associated with the FASS microchip. Colors indicate the solution present in the channel; black – separation buffer, gray – sample matrix, white – analyte. (b) Images of the development of the stacked analyte with a 10 fold difference in sample matrix and separation buffer conductivity. (c) Images of the development of the stacked analyte with a 100 fold difference in sample matrix and separation buffer conductivity. Separation buffer is 32 mM carbonate at pH 9.6, sample is to FITC labeled amino acids. (d) Cartoon illustrating the injection scheme for the revised microfluidic device for FASS. Sample is injected and stacked in a channel other than the one used for separation. Any disruptions due to EOF mismatch in the sample matrix and separation buffer is isolated to this other channel. (Reprinted from Lichtenberg et al., Sample preconcentration by field amplified stacking for microchip-based capillary electrophoresis, Electrophoresis, 22 (2001) 258–271, with permission from John Wiley and Sons).

has been no exhaustive study discussing this issue and should, at the very least, be understood as a phenomenon that may impact preconcentration performance and reproducibility. 2.2. Electrophoretic preconcentration 2.2.1. Field amplified stacking methods Field amplified sample stacking (FASS) is arguably the simplest form of on-line preconcentration in microfluidic devices. As described previously, the key to implementing FASS is that the conductivity of the sample matrix must be lower than the conductivity of the BGE. The primary issue faced when implementing FASS on a microfluidic device is how to manage the field gradient that exists in the injection region of the device. As a low conductivity matrix migrates into a microfluidic network during injection, it may be difficult to control the direction the ions move as a function of the disparate fields. Also of concern is a localized increase of temperature in the injection region due to this increase in electric field [21,22]. Lichtenberg et al. demonstrated one of the earliest examples of FASS on a microfluidic device [4]. They looked at a number of different microchip designs with an eye towards improving detection limits by injecting large volumes of low conductivity sample matrix into a standard separation channel. They noted that mismatched EOF velocities in the sample matrix and BGE resulted in the generation of Poiseuille flow effects. Some of these effects are illustrated in Fig. 1. The chip design is shown in Fig. 1a; position A is separation buffer, A is buffer waste (position A to A defines the separation channel), B is the sample reservoir, and C is the sample waste. During injection a potential is applied between B and C causing the sample matrix and FITC-labeled analytes to migrate

into the separation channel. Upon the application of the separation voltage between A and A , the sample plug begins to migrate into the separation channel. Fig. 1b shows the effect of mismatched EOF on a system where the BGE has conductivity 10 times greater than the sample matrix, and Fig. 1c shows a 100 times more concentrated BGE. As the difference between sample matrix and BGE conductivity increases, the effects of Poiseuille flow become more pronounced. In order to mitigate the effects of the conductivity mismatch on the overall separation performance, the authors designed the chip shown in Fig. 1d. This chip design allows for the sample stacking to develop in a channel other than the separation channel, thus limiting the disruption associated with the mismatched EOF velocities in the sample matrix and BGE brought on by the conductivity difference. The authors realized 65-fold improvements in signal when using this device for a pair of FITC labeled amino acids [4]. The relationship between chip design and the efficacy of fieldbased stacking techniques is very important. Chien and Yang designed a microfluidic chip better suited for field-amplified sample stacking [23]. Fig. 2 shows the injection scheme used in this device. The most important aspect of this chip design’s performance is the presence of the concentration boundary as indicated in Fig. 2a. This boundary is established by carefully controlling channel dimensions and the pressure above each reservoir of the device. Initially the entire fluidic network is filled with low conductivity sample matrix, then the high conductivity BGE is loaded in and the pressure above the individual reservoirs adjusted such that the boundary of SM/BGE is established as indicated in Fig. 2a. The remaining injection/separation scheme is illustrated in Fig. 2b and c. By controlling the location of the boundary prior to application of an injection voltage, a highly reproducible and effective separation,

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

15

Fig. 2. Cartoon illustrating the injection scheme of a microfluidic device for field amplified stacking. (a) The location of separation buffer and sample matrix after solution is loaded into device. (b) The application of voltage for the purposes of sample loading. (c) The application of voltage for the purposes of separation (separation channel is indicated by the – V applied). (d) Separation of two fluorescent dyes in the absence of sample stacking. (e) Separation of two fluorescent dyes with field amplified stacking implemented. (Reprinted from Yang et al., Sample stacking in laboratory-on-a-chip devices, Journal of Chromatography A, 924 (2001) 155–163, with permission from Elsevier).

as depicted in Fig. 2d and e, is possible. A non-stacked separation of two fluorescent dyes is presented in Fig. 2d, and the separation afforded after stacking is presented in Fig. 2e. The net effect of stacking was a near 100-fold improvement in sensitivity for some fluorescent dyes [23]. Santiago and co-workers focused on improving injection schemes in a device they developed that included a photoinitiated porous polymer [24]. The chip design, injection and separation scheme is illustrated in Fig. 3. The porous plug allows the authors to preload the injection cross-t with low conductivity sample matrix without compromising the effective length of the separation – the plug prevents hydrodynamic flow down the separation channel, but does not inhibit electromigration during separation. Sample is injected electrokinetically, and the anions of interest are detected after they have migrated through the plug. The unique design of this device allows for sample matrix/BGE conductivity mismatches in excess of 3 orders-of-magnitude and corresponding signal enhancements of 1000 fold for the dyes fluorescein and Bodipy [24]. An alternative to FASS is field amplified sample injection (FASI). In capillary systems FASI injections are implemented by injecting

a plug of water into the capillary prior to electrokinetic injection of sample. A reversed polarity is applied across the capillary during injection; the resultant high field generated in the water plug as compared to the field in the BGE causes anions in the sample to migrate into the capillary, while the water plug exits into the sample vial. These anions are effectively stacked and can be separated under normal polarity. Heineman and co-workers implemented the microfluidic equivalent of FASI using a standard cross-t the chip design. Low conductivity sample matrix is injected across the t, then down into the separation channel; the FASI injection was implemented by applying reversed polarity across the separation channel, allowing for both anion preconcentration and sample matrix migration out of the separation channel. After completion of the stacking step, normal polarity is applied and the separation performed. Sensitivity enhancements of as much as 160-fold were observed when compared to the traditional pinched injection (i.e. injection volume is the volume of sample at the cross-t intersection for several fluorescent dyes) [25]. The above examples illustrate that the implementation of fieldbased sample injections is not trivial and certainly not as easy to implement in a microfluidic device as they are on a capillary

16

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

Fig. 3. Cartoon illustrating separation buffer loading and sample injection scheme on a device with a porous plug in place to prevent sample matrix flow into the separation channel during pressure injections. Top cartoon illustrates the separation buffer loading step. (a) Illustrates pressure injection of the low conductivity sample. (b) The application of voltage allows for field amplified stacking. (c) Migration of analyte through the porous plug and separation of analytes. (Reprinted from Jung et al., Thousandfold signal increase using field-amplified sample stacking for on-chip electrophoresis, Electrophoresis, 24 (2003) 3476–3483, with permission from John Wiley and Sons).

system. Flow disruption at concentration gradients can adversely affect separation performance, with many of the injection schemes described above designed to mitigate those problems. Shiddiky et al. designed a microfluidic device that allowed for both FASS and FASI injections prior to the separation of phenolic samples [26]. Their chip design and injection scheme are shown in Fig. 4. Sample is introduced into the fluidic network by applying a potential between R1 and R2 (Image 1). FASS occurs in the first channel and during this time period a water plug was introduced into the second separation channel via syringe in order to prepare it for the FASI injection (Image 2). Anions are concentrated in the FASI channel and introduced into the separation channel (Images 3 and 4); and finally separated using MEKC (Image 5). Detection limits as low as 100 pM were achieved electrochemically [26]. As FASS and FASI methods have become more common, the type of samples analyzed has expanded. For example, Shim and co-workers used FASS and FASI in the analysis of endocrine disruptors [27], food dyes [28], DNA [29], and antibiotics [30]. Guan and Henry focused on improving detection limits of dopamine and various inorganic anions on microchips realizing detection limits in the single nM range [31,32]. The detection limit of 8 nM dopamine was made possible by coupling on-chip FAS with an end-column

Fig. 4. Cartoon illustrating the injection scheme for a device which has two stacking domains one for field amplified sample stacking and one for field amplified stacking injection. (1) Sample is loaded into the field sampled sample stacking domain. (2) Field amplified sample stacking occurs while water is loaded into the field amplified sample injection domain of the device. (3) Field amplified sample injection occurs and the stacked sample plug is allowed to migrate to the sample injection cross t under reverse polarity. (4) Sample is injected via cross t into the separation channel. (5) Separation occurs. (Reprinted from Shiddiky et al., Direct analysis of trace phenolics with a microchip: In-channel sample preconcentration, separation, and electrochemical detection, Analytical Chemistry, 78 (2006) 6809–6817, with permission from American Chemical Society).

electrochemical bubble cell [31]. Using the bubble cell increased the exposed area of the working electrode, thereby improving detector sensitivity. Zhang et al. used FASS in the analysis of monosulfated disaccharide isomers [33]. 2.2.2. Moving boundary stacking methods Moving boundary stacking was used first to separate proteins from blood [34,35]. The key to analyte stacking via the moving boundary method is that the molecule of interest is next to an interface of two differing ion compositions. The boundary forms due to the discontinuous nature of the ions at this interface. The formation of the boundary is associated with more than simply a change in concentration of BGE ions, as seen in FAS, but also a difference in the ion species; thus two components of the Kohlrausch value come into play (concentration and mobility). This discontinuous boundary migrates through the electric field and travels through all zones in its path. Stacking and destacking occurs before and after the sample interface. Vrouwe et al. show a heart cut method for lithium in blood using the device and injection scheme illustrated in Fig. 5 [36]. Complete understanding of the sample matrix composition is critical as other components of the sample matrix will be present in the sample if the cut is not tight enough. The authors manipulate moving boundary electrophoresis to

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

17

Fig. 5. (A) Cartoon illustrating microchip design for heart cutting injection of lithium from blood. (B) MBE injection of sample. (C) Injection of sample containing sodium and lithium along with the acetate counterion. (D) Electropherograms showing the separation of sodium and lithium as a function of injection time. Trace A – 30 s sample loading; Trace B – 240 s sample loading; Trace C – 300 s sample loading. (Reprinted from Vrouwe et al., Microchip analysis of lithium in blood using moving boundary electrophoresis and zone electrophoresis, Electrophoresis, 26 (2005) 3032–3042, with permission from John Wiley and Sons).

facilitate the injection of a diffuse zone containing both sodium and lithium as depicted in the electropherograms in Fig. 5B. Briefly, MBE is implemented and stable zones established. To the right of the boundary depicted by the double line in Fig. 5B, a diffuse zone of sodium and lithium is established and that zone of analyte is injected into the separation channel, where the sodium and lithium are detected. While it appears counter intuitive, it is necessary to inject a dilute sodium/lithium mixture or the lithium is not resolved from the sodium, as indicated in the 300 s injection (trace C) in Fig. 5D [36]. In that instance, the injection is so long, that a higher concentration plug of sodium and lithium was allowed to enter the separation channel. 2.2.3. Isotachophoresis ITP can take two forms in capillary and microchip-based electrophoresis, one where the separation mechanism is entirely ITP-based and the second where ITP occurs within a sample matrix during injection and that event is followed by separation. In both cases, stacking is made possible by the presence of a leading and trailing electrolyte (LE and TE, respectively) in the system. Park et al. used a secondary channel to enable the ITP event, first, and then pushed the sample into the separation channel, as depicted in Fig. 6 [37]. In Fig. 6A the leading and trailing electrolyte are noted with reservoirs labeled A and B, indicating the reactants used in the immunoassay. The reactants are mixed on the microchip by pulling vacuum at wells 4 and 5. In Fig. 6B, an electric field is applied, causing ITP-based stacking of the sample components flanked by the TE and LE. Monitoring the current allowed the researchers to switch the electric field at an appropriate time when ITP stacking had completed (Fig. 6C), enabling the separation of the components of interest (Fig. 6D) [37]. The dual channel design permits

clean concentration zones to be formed and subsequently pushed into a secondary channel for final separation or other types of chip functions like PCR or labeling. Alternatively, Santiago and co-workers used the ability to switch fluids in and out of the separation chamber to create a single channel device [5,38]. In one instance, the authors use both cationic and anionic LE and TE pairs for simultaneously stacking sample cations and anions [38]. While the authors are interested in the stacking and separation of anionic DNA fragments, the simultaneous stacking of cations is necessary to facilitate the separation of the stacked components. During ITP the two stacked zones migrate towards one another, causing a phenomenon dubbed “ion concentration shockwaves” by the authors [38]. The consequence of these shockwaves is the disruption of ITP-based stacking and subsequent electrophoretic separation of the stacked components. Fig. 7 shows the ITP-stacked DNA fragments and the resultant separation with a comparison to the traditional slab-gel separation. 2.2.4. Isoelectric focusing IEF is a powerful electrophoretic technique because it has a selfgoverning feedback loop in the separation. The separation space is filled with a pH-generating material, usually ampholytes, that enable a pH gradient. Each molecule in the sample has a dual ionic state in which it bears a point of neutral charge (pI). In traditional IEF separations, an immobile pH gradient is used and is typically placed onto a second gel-based separation media for size-based separations (2-D gel electrophoresis). Guillo et al. coupled on-chip pumping with IEF in order to facilitate detection [39]. Briefly, sample mixed with an ampholyte solution is loaded into a separation channel on a microfluidic device to which a voltage is applied. The pH gradient is established and analytes migrate to the region

18

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

Fig. 6. Schematic diagram of sample loading, stacking and separation. (A) Initial state of the microfludici device. Mixing of components A and B is afforded by pulling vacuum at positions 4 and 5. (B) Potential is applied to afford ITP-based stacking of sample matrix components. (C) Stacked analyte transitions from the ITP domain to the separation domain. (B) Separation of stacked analytes. (Reprinted from Park et al., Controlling data quality and reproducibility of a high-sensitivity immunoassay using isotachophoresis in a microchip, Analytical Chemistry, 80 (2008) 808–814, with permission from American Chemical Society).

corresponding to their pI. After separation is complete, the entire separation channel contents are mobilized to the detection point via pressure. Hirokawa and co-workers studied an important component of all microchip-based IEF separations, the establishment of the pH gradient during the application of the electric field [6]. They demonstrate that performance is improved when EOF is

suppressed in the separation channel. Fig. 8 shows the evolution of the separation of several proteins as a function of separation time using whole channel UV absorbance detection. This figure illustrates the time necessary for analyte to reach its appropriate position in the pH gradient. Detection limits of 0.8 ␮g mL−1 were observed for a mixture of carbonic anhydrase-I and carbonic anhydrase-II [6]. Liang et al. utilize a monolithic immobilized pH gradient with an eye towards coupling to other microchip-based separation techniques in the future [40]. Fig. 9 shows the resultant separation of a mixture of three FITC-labeled proteins. The quality of the overall sample stacking and separation performance is attributed

Fig. 7. Electropherograms showing the transition from (A) stacked DNA fragments to (B) the transition state due to ion concentration shockwaves, to (C) the separation of ten DNA fragments using bidirectional ITP.

Fig. 8. Time course of the IEF-based separation of a mixture of 11 proteins with whole-channel UV absorbance detection. Peaks are 1 – Trypsinogen, 2 – Lentil lectin (basic), 3 – Lentil lectin (middle), 4 – Lentil lectn (acidic), 5 – Myoglobin (basic), 6 – Myoglobin (acidic), 7 – Carbonic anhydrase B (human), 8 – Carbonic anhydrase B (bovine), b-Lactoglobulin A, 10–Trypsin inhibitor, 11 – Amylglycosidase.

(Reprinted from Bagha et al., Coupled isotachophoretic preconcentration and electrophorestic separation using bidirectional isotachophoresis, Analytical Chemistry, 83 (2011) 6154–6162, with permission from American Chemical Society).

(Reprinted from Xu et al., Investigation of the pH gradient formation and cathodic drift in microchip isoelectric focusing with imaged UV detection, Electrophoresis, 31 (2010) 3558–3565, with permission from John Wiley and Sons).

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

19

Research in this area has been focused in two primary areas: developing the methodology to perform SPE on microfluidic devices and developing the solid-phase material for inclusion within the device.

Fig. 9. Separation of three FITC-labeled proteins. Sample concentrations are 7.5 ␮g mL−1 ribonuclease B, 22 ␮g mL−1 myoglobin, and 26 ␮g mL−1 ␣-casein. Sample was focused for 2 min. (Reprinted from Laing et al., Microchip isoelectric focusing with monolithic immobilized pH gradient materials for proteins separation, Electrophoresis, 30 (2009) 4034–4039, with permission from John Wiley and Sons).

to a combination of the immobilized pH gradient, pressure mobilization, and the choice of detection point [40]. Visual inspection of the electropherogram indicates limits of detection in the ng mL−1 range. 2.2.5. Stacking in micellar-based separations While the above sections outline field-based and mobility-based stacking techniques, it is important not to ignore the preconcentration of neutral molecules on microfluidic devices. Neutral molecule preconcentration is made possible by inducing a mobility change in the molecule when it interacts with a surfactant micelle in the BGE, this separation mode is called micellar electrokinetic chromatography (MEKC). Briefly, neutral molecules move with electroosmotic flow in the sample matrix, until they interact with a micelle in the BGE and an equilibrium is established between the analyte and the micelle. The analyte exhibits an apparent mobility somewhere between that of EOF and the micelle – this mode of on-line preconcentration has been referred to alternatively as either sweeping or stacking. For example Newman et al. performed MEKC-based separations on a mixture of neutral alkaloids, establishing sample matrix conditions that afforded improved sensitivity due to a combination of micelle stacking at the sample matrix/BGE boundary and the traditional sweeping mechanism described above [41]. Several examples of on-chip preconcentration due to interaction with surfactant micelles are present throughout this review as stacking via the mechanism of micelle affinity is not limited to neutral molecules. The work presented in the previous section by Shiddiky et al., for example, includes a MEKC-based separation [26]. 3. Extraction methods 3.1. Solid-phase extraction (SPE) Microfluidic devices that incorporate on-chip extraction methodology are an important step in realizing Lab-on-a-Chip devices. Sample enrichment via an extraction step couples sample clean-up to the obvious benefits of preconcentration, where the elution volume is much lower than the sampled volume. SPE is the most common extraction method applied on microfluidic devices, with the most prominent role being the generation of polymerase chain reaction (PCR)-ready DNA from complex biological samples.

3.1.1. Biomacromolecule applications The Landers group has published a series of papers focusing on using microchip-based SPE for the purification of DNA. The goal is to provide DNA for on-chip PCR that requires not only the extraction of DNA from complex biological matrices, but also the removal of potential interferents from the DNA such as proteins that inhibit PCR. One of their initial efforts focused on developing a silica-bead based packed bed for SPE using sol–gels to immobilize the beads in a short microfluidic channel [42]. This device is very simple, requiring only a single syringe pump to transport liquid to and through the solid-phase material. The use of silica bead-beds for SPE gave way to sol–gel monoliths [43]. As with their initial efforts, the goal was to generate PCR-ready material. Fig. 10 shows the results of protein and DNA quantitation as a function of volume using the sample loading, washing, and elution step of the SPE process. The important feature of this solid-phase is that it has the capacity to capture not only the nanogram levels of DNA necessary for successful PCR (Fig. 10A and C), but also the microgram levels of protein that would otherwise inhibit PCR (Fig. 10A and B). The technique allows impurities to be removed from the sample in the wash step [43]. Additional efforts from the Landers group include large volume extraction and elution wherein as much as 1 mL of sample is processed [44,45], the use of PMMA devices functionalized with the polysaccharide chitosan for DNA and RNA purification [46,47], the use of SPE in conjunction with microfluidic devices fabricated from polyester [48], and a device for the extraction of two positional isomers of dihydroxybenzoic acid from saliva [49]. In addition to standalone SPE devices, efforts also included coupling SPE to PCR devices [50] that integrate all components for true sample-in-answer-out functionality [3]. The Harrison group published a series of manuscripts on SPE [2] and the integration of protein digestion and SPE on microfluidic devices coupled to mass spectra (MS) detection [51,52]. A protein digestion/SPE device is depicted in Fig. 11 [52]. Briefly, protein is digested at position A in the device, then the resultant fragments are preconcentrated at the SPE bed at position B; finally, the extract is injected and separated via microchip electrophoresis. Fig. 11B and C shows the results obtained with digestion followed by extraction in a 4 mm long SPE bed (B) and a 2 mm long SPE bed with subsequent injection of trypsin for on bed digestion of cytochrome C. The onchip SPE afforded better than ten-fold concentration enhancement and the total on-chip process took only 3 min compared to a time of 2 h required for bench top solution-based digests [52]. More recent efforts are focused on functionalizing the SPE bed with polycationic surface coatings in order to prevent protein adsorption and improve injection into the MS detector [51]. SPE is coupled to microchip-based HPLC by DeVoe and coworkers [53]. In their efforts, two devices are presented, as depicted in Fig. 12; device 1 (pictured in Fig. 12A and B) is for dynamic sample injection while device 2 (Fig. 12C and D) couples a front-end SPE step prior to separation. A comparison between the performances of the two devices is shown in Fig. 12E and F. The authors present the separation of FITC-labeled ribonuclease A (peaks 1a and 1b – multiple peaks due to variable labeling efficiency with FITC) and cytochrome C (peak 2). Clearly, the coupling of SPE to the front end of the separation is beneficial, especially considering that the analyte concentration in Fig. 12F is an order of magnitude lower than that presented in Fig. 12E. The increase in peak height for the cytochrome C relative to the ribonuclease peaks is due to a greater affinity of the cytochrome C for the stationary phase using the

20

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

Fig. 10. (A) Protein and DNA concentration profile as a function of volume. Protein elution occurs in the wash step; DNA elution occurs in the elution step. (B) Electropherogram of negative control (PCR amplification from a whole blood sample) (C) Electropherogram from DNA extracted using sol–gel SPE device. (Reprinted from Wu et al., Microchip-based macroporous silica sol–gel monolith for efficient isolation of DNA from clinical samples, Analytical Chemistry, 78 (2006) 5704–4710, with permission from American Chemical Society).

SPE domain, specifically a polymethacrylate monolith composed of butylmethacrylate and trimethylolpropane trimethacrylate [53]. The authors indicate that extraction efficiency is a function of analyte hydrophobicity.

3.1.2. Small molecule applications SPE extraction is not limited to biological applications; there are a number of examples where microchip-based SPE is performed for the analysis of small non-biological molecules. Ramsey and Collins

Fig. 11. (A) Fluidic device design including tryptic digest and SPE beds along with injection cross and separation channel coupled to MS detection. (B) Mass spectra of cytochrome C digestion followed by extraction onto a 4 mm long SPE bed. (C) Mass spectra of cytochrome C digestion followed by extraction onto a 2 mm long SPE bed – during elution for the SPE bed additional trypsin was added to induction on-bed digestion of any protein adhering to the walls of the device. Black closed-circles indicate peptide assigned to cytochrome C digestion, T indicates autodigestion of trypsin, and I represents interferent peaks. (Reprinted from Wang et al., Multifunctional protein processing chip with integrated digestion, solid-phase extraction, separation and electrospray, Electrophoresis, 31 (2010) 3703–3710, with permission from John Wiley and Sons).

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

21

Fig. 12. (A and B) Microchip design and instrument schematics for microchip-based LC (C and D) Microchip design and instrument schematics for microchip-based SPE coupled to LC. (E) Separation of FITC-labeled ribonuclease A (peak 1a and 1b) and cytochrome C (peak 2) on the HPLC microfluidic device. (F) Separation of FITC-labeled ribonuclease A (peak 1a and 1b) and cytochrome C (peak 2) on the HPLC microfluidic device with integrated SPE. (Reprinted from Liu et al., Polymer microchips integrating solid-phase extraction and high-performance liquid chromatography using reversed-phase polymethacrylate monoliths, Analytical Chemistry, 81 (2009) 2545–2554, with permission from American Chemical Society).

Fig. 13. (A) Image of the microfluidic device including a SPE domain and a separation domain. (B) Close-in view of the SPE domain illustrating sample and elution inlet and waste reservoirs and the separation channel. (C) Electropherogram showing the resultant separation after the extraction and elution of coumarin 334 and 314. The dotted lines indicate when extraction, elution, and injection into the MEKC-based separation domain occur. (Reprinted from Ramsey et al., Integrated microfluidic device for solid-phase extraction coupled to micellar electrokinetic chromatography separation, Analytical Chemistry, 77 (2005) 6664–6670, with permission from American Chemical Society).

coupled SPE to the MEKC-based separation of fluorescent dyes [54]. The authors incorporated a photopolymerizable frit in the column chamber region of the device depicted in Fig. 13A and B; C-18 beads are packed against the frit and solvent-locked in place resulting in an on-chip SPE bed. Careful control of the applied potential on the device allowed for injection onto the SPE column, elution from the column, and subsequent injection and MEKC-based separation. Fig. 13C shows the timing associated with all steps of the extraction and separation process along with the successful separation of two fluorescent dyes. This technology allows for the detection of as little as 100 pM rhodamine B with signal-to-noise ratios sufficient to calculate a LOD as low as 60 fM, with sample load times of 300 s [54]. The use of SPE for on-line preconcentration can also be implemented seamlessly with microchip-based electrochromatography separations. Collins and co-workers show multiple column volume injections of nitroaromatic and nitroamine explosives and their

degradation products using a sol–gel based stationary phase [55]. Samples are prepared in an aqueous matrix; the analytes affinity for the stationary phase is such that the analytes are retained on the head of the column while the sample matrix continues to migrate down the column. A similar injection scheme is implement by this group on a microfluidic device that employs an EOF pump as the motive force for on-chip LC [56]. Hennion and co-workers used head column extraction on a microfluidic device for the analysis of a number of compounds [57]. 3.2. Non-SPE methods Extraction methods demonstrated on the microfluidic platform are not limited to SPE. There are a growing number of examples of liquid–liquid extraction on microfluidic devices. A device designed by Chen et al. is shown in Fig. 14A [58]. A small recess within the

22

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

Fig. 14. (A) Cartoon depicting the liquid–liquid extraction microchip. Initially organic solvent is allowed to flow through the device (hexanol) followed by aqueous solution; only the hexanol trapped in the side-reservoir remains in the channel. (B) Time lapse images of the extraction of butyl rhodamine B. Initially the orange color intensifies, then the color transitions to purple due to photobleaching; the loss of hexanol due to dissolving into the aqueous media is noted at approximately 6 min. (Reprinted from Chen et al., Microfluidic chip-based liquid–liquid extraction and preconcentration using a subnanoliter droplet trapping technique, Lab on a chip, 5 (2005) 719–725, with permission from Royal Society of Chemistry).

channel is formed to hold a small volume of organic solvent. Sample in an aqueous matrix flows through the channel, and analyte is extracted into the small volume of organic solvent. Enrichment factors in excess of 3000 are possible with this chip configuration, as shown in Fig. 14B [58]. The transition from orange to purplish color is due to photobleaching at high fluorescent dye concentrations. While the above example demonstrated extraction at a single point within a device, efforts by Kitamori and co-workers have used a flow based system, where organic solvent flows through a channel in parallel and in contact with urine samples for the extraction of amphetamines [8]. These samples are prepared for eventual off-chip analysis via GC. Tipple et al. demonstrated on-chip vapor impinging for the analysis of cyanide [7]. Fig. 15A shows a schematic diagram for this simple device. Vapor is introduced into the device and is allowed to bubble through a liquid suitable for vapor extraction. A snapshot of gas bubbles moving through the extraction liquid is shown in Fig. 15B. The presence of cyanide is detected off-chip fluorescently, based upon a derivatization reaction with naphthalene-2,3-dicarboxaldehyde. Detection of vapors

with cyanide concentrations as low as 1.9 mg m−3 are demonstrated with a calculated LOD of 0.486 mg m−3 with a one minute sample time at a sample flow rate of 2 mL min−1 [7]. 4. Cell enrichment and preconcentration On-line preconcentration and enrichment is not limited to small molecules or biological macromolecules such as protein and DNA. Cells and cell-based separations are becoming more common on microfluidic devices with an eye towards removing contaminants that may impede other chip-based assays and/or increasing the concentration of a particular cell-type of interest. Interestingly, the transition from small molecules, DNA, and proteins to particles has resulted in the development of new technologies and bulk separation techniques tailored to working with cells. Landers and co-workers demonstrate that a simple straight channel device is sufficient to separate sperm and epithelial cells from one another in order to address the large backlog of samples associated with sexual assault [9]. Due to a physical property difference between the two cell types, the epithelial cells settle to the

Fig. 15. (A) Cartoon of vapor impinge microfluidic device. Vapor enters the microchip at (f) and transitions through position (a) and travels through the array of channels (b). Bubbles pass through the reservoir (c). A Teflon membrane (d) prevents liquid from escaping the device and the vapor stream continues out of the chip (e). (B) Image of vapor bubbles passing through the extraction liquid at position (c) in the device. (Reprinted from Tipple et al., Development of a microfabricated impinge for on-chip gas phase sampling, Analytica Chimica Acta, 551 (2005) 9–14, with permission from Elsevier).

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

23

Fig. 16. Left side (top) – Cartoon of optical chromatography chip for sample clean up. Arrows indicate fluid flow direction and laser path. Left side (bottom) – close up of capillary connector between two sides of the microfluidic device. Spores are pinned on the bottom region in this picture. Right side – demonstration of RT-PCR from spores prepared in a mixture of humic substances including appropriate positive and negative controls. (Reprinted from Hart et al., Preparative optical chromatography with external collection and analysis, Optics Express, 551 (2008) 18782–18789, with permission from The Optical Society).

bottom of the reservoir in which a sample is added, while the sperm cells flow through the microchannel to a second reservoir where no epithelial cells are detected. Toner and co-workers have used antibody coated microposts in silicon devices to isolate circulating tumor cells (CTCs) from whole blood samples [59]. Capture on the microposts was strongly dependent upon sample flow rate through the device, with efficiencies in excess of 65% when rates were maintained between 1 and 2 mL h−1 . Subsequent efforts replaced the microposts with a herringbone-pattern on the surface of one side of the microchannel coated with appropriate antibodies [60]. This structure promoted mixing and in large-scale experiments, capture efficiencies as high as 91% were observed. Hoshino et al. developed a microchip-based immunomagnetic assay for the detection of CTCs [61], where CTCs are labeled with magnetic nanoparticles and selectively captured in a microfluidic device via magnetic field. Flow rates of 10 mL h−1 were possible, with capture efficiencies at or near 90% (dependent on cell type). Hart et al. focused on using optical chromatography as a facilitator in sample cleanup of Bacillus anthracis Stern strain spores from known PCR inhibitors [10] using the microfluidic device shown on the left hand side of Fig. 16. Optical chromatography uses an optical force generated by a mildly focused laser beam to propel particles along its axis of propagation [10]. The beam is aimed directly against a fluid flow; the balance of fluid drag force and optical pressure results in the stable trapping of particles within the beam. Larger size or greater refractive index particles experience greater optical pressure and are, thus, propelled further than smaller or lower refractive index particles. This results in unique positions (separation) along the beam axis for particles of different size and composition (refractive index). This device consists of two

layers of microfluidic channels connected via a laser etched transverse microchannel. The laser beam is introduced into the device through this microchannel against the fluid flow, wherein the laser power is sufficient to pin the spores against the wall of the microfluidic device while the sample matrix continues to flow out of the microfluidic network. The matrix can be replaced with one suitable for RT-PCR. One such example is shown on the right hand side of Fig. 16, where the spores are prepared in a matrix containing humic substances (the color observed in the sample vial depicted in panel A is due to the presence of these humic materials). In the presence of humics, RT-PCR is not possible; however, by using preparative scale optical chromatography, the sample spores are isolated from the humic matrix and successful RT-PCR occurs [10]. This technology has been extended to allow for bulk fractionation of mixtures of biological agents including Escherichia coli cells, B. anthracis spores and pollen with 80–90% efficiencies [62]. 5. Conclusions Our hope in writing this review article is to present the community with some of the more common modes of on-line preconcentration in microfluidic devices, along with a few of the not so common. Several orders-of-magnitude improvements in detection limits are available with appropriate implementation of a number of these techniques. Our desire was to primarily focus on microchip design and sample preconcentration implementation so that a person unfamiliar with many of the preconcentration options could quickly discern what available methods might best suit their particular research needs. Clearly, the work presented herein illustrates that there is no single technique by which sensitive detection of all

24

B.C. Giordano et al. / Analytica Chimica Acta 718 (2012) 11–24

analytes of interest is possible. Ultimately, effective on-line sample pre-concentration is dependent upon the analyte of interest and the matrix in which it is contained. Coupling multiple stacking modes on a single device seems to be the direction in which many research groups are moving, which is very encouraging. Ideally, future works will continue on this track, with an added focus on understanding how to deal with complex sample matrices and how those matrices effect stacking performance. By no means is this review a complete picture of all techniques available, however, it does serve as a useful primer for the microfluidic novice. As detection limits continue to match and exceed existing technologies, the field of microfluidics will continue to grow and, ideally, establish itself as a robust tool for researchers.

[29] [30] [31] [32] [33] [34] [35] [36] [37] [38] [39] [40] [41]

References [1] K. Seiler, D.J. Harrison, A. Manz, Anal. Chem. 65 (1993) 1481. [2] A.B. Jemere, R.D. Oleschuk, F. Ouchen, F. Fajuyigbe, D.J. Harrison, Electrophoresis 23 (2002) 3537. [3] J.P. Landers, C.J. Easley, J.M. Karlinsey, J.M. Bienvenue, L.A. Legendre, M.G. Roper, S.H. Feldman, M.A. Hughes, E.L. Hewlett, T.J. Merkel, J.P. Ferrance, Proc. Natl. Acad. Sci. U.S.A. 103 (2006) 19272. [4] J. Lichtenberg, E. Verpoorte, N.F. de Rooij, Electrophoresis 22 (2001) 258. [5] S.S. Bahga, G.V. Kaigala, M. Bercovici, J.G. Santiago, Electrophoresis 32 (2011) 563. [6] Z.Q. Xu, N. Okabe, A. Arai, T. Hirokawa, Electrophoresis 31 (2010) 3558. [7] C.A. Tipple, M. Smith, G.E. Collins, Anal. Chim. Acta 551 (2005) 9. [8] H. Miyaguchi, M. Tokeshi, Y. Kikutani, A. Hibara, H. Inoue, T. Kitamori, J. Chromatogr. A 1129 (2006) 105. [9] K.M. Horsman, S.L.R. Barker, J.P. Ferrance, K.A. Forrest, K.A. Koen, J.P. Landers, Anal. Chem. 77 (2005) 742. [10] S.J. Hart, A. Terray, J. Arnold, T.A. Leski, Opt. Express 16 (2008) 18782. [11] F. Kohlrausch, Ann. Phys. (Leipzig) 62 (1897) 209. [12] V. Hruska, B. Gas, Electrophoresis 28 (2007) 3. [13] R.L. Chien, D.S. Burgi, Anal. Chem. 64 (1992) A489. [14] P. Gebauer, P. Borecka, P. Bocek, Anal. Chem. 70 (1998) 3397. [15] M. Macka, P.R. Haddad, P. Gebauer, P. Bocek, Electrophoresis 18 (1997) 1998. [16] C. Desiderio, S. Fanali, P. Gebauer, P. Bocek, J. Chromatogr. A 772 (1997) 81. [17] P. Gebauer, P. Bocek, J. Chromatogr. A 772 (1997) 73. [18] M. Urbanek, L. Krivankova, P. Bocek, Electrophoresis 24 (2003) 466. [19] P.G. Righetti, C. Gelfi, M. Conti, J. Chromatogr. B 699 (1997) 91. [20] H.Z. Helmholtz, Ann. Phys. Chem. 7 (1879) 337. [21] B.C. Giordano, K.M. Horsman, D.S. Burgi, J.P. Ferrance, J.P. Landers, Electrophoresis 27 (2006) 1355. [22] A. Vinther, H. Soeberg, J. Chromatogr. 559 (1991) 27. [23] H. Yang, R.L. Chien, J. Chromatogr. A 924 (2001) 155. [24] B. Jung, R. Bharadwaj, J.G. Santiago, Electrophoresis 24 (2003) 3476. [25] M.J. Gong, K.R. Wehmeyer, P.A. Limbach, F. Arias, W.R. Heineman, Anal. Chem. 78 (2006) 3730. [26] M.J.A. Shiddiky, H. Park, Y.B. Shim, Anal. Chem. 78 (2006) 6809. [27] H.B. Noh, K.S. Lee, B.S. Lim, S.J. Kim, Y.B. Shim, Electrophoresis 31 (2010) 3053. [28] K.S. Lee, M.J.A. Shiddiky, S.H. Park, D.S. Park, Y.B. Shim, Electrophoresis 29 (2008) 1910.

[42]

[43] [44] [45] [46] [47] [48] [49] [50] [51] [52] [53] [54] [55] [56] [57] [58] [59]

[60]

[61] [62]

M.J.A. Shiddiky, Y.B. Shim, Anal. Chem. 79 (2007) 3724. K.S. Lee, S.H. Park, S.Y. Won, Y.B. Shim, Electrophoresis 30 (2009) 3219. Q. Guan, C.S. Henry, Electrophoresis 30 (2009) 3339. S.D. Noblitt, F.M. Schwandner, S.V. Hering, J.L. Collett, C.S. Henry, J. Chromatogr. A 1216 (2009) 1503. Y. Zhang, G.C. Ping, B.M. Zhu, N. Kaji, M. Tokeshi, Y. Baba, Electrophoresis 28 (2007) 414. A. Tiselius, Biochem. J. 31 (1937) 1464. A. Tiselius, Biochem. J. 31 (1937) 313. E.X. Vrouwe, R. Luttge, W. Olthuis, A. van den Berg, Electrophoresis 26 (2005) 3032. C.C. Park, I. Kazakova, T. Kawabata, M. Spaid, R.L. Chien, H.G. Wada, S. Satomura, Anal. Chem. 80 (2008) 808. S.S. Bahga, R.D. Chambers, J.G. Santiago, Anal. Chem. 83 (2011) 6154. C. Guillo, J.M. Karlinsey, J.P. Landers, Lab Chip 7 (2007) 112. Y. Liang, Y.Z. Cong, Z. Liang, Y.K. Zhang, L.H. Zhang, Electrophoresis 30 (2009) 4034. C.I.D. Newman, B.C. Giordano, C.L. Copper, G.E. Collins, Electrophoresis 29 (2008) 803. M.C. Breadmore, K.A. Wolfe, I.G. Arcibal, W.K. Leung, D. Dickson, B.C. Giordano, M.E. Power, J.P. Ferrance, S.H. Feldman, P.M. Norris, J.P. Landers, Anal. Chem. 75 (2003) 1880. Q.R. Wu, J.M. Bienvenue, B.J. Hassan, Y.C. Kwok, B.C. Giordano, P.M. Norris, J.P. Landers, J.P. Ferrance, Anal. Chem. 78 (2006) 5704. C.R. Reedy, J.M. Bienvenue, L. Coletta, B.C. Strachan, N. Bhatri, S. Greenspoon, J.P. Landers, Forensic Sci. Int.: Gen. 4 (2010) 206. C.R. Reedy, K.A. Hagan, B.C. Strachan, J.J. Higginson, J.M. Bienvenue, S.A. Greenspoon, J.P. Ferrance, J.P. Landers, Anal. Chem. 82 (2010) 5669. W.D. Cao, C.J. Easley, J.P. Ferrance, J.P. Landers, Anal. Chem. 78 (2006) 7222. K.A. Hagan, W.L. Meier, J.P. Ferrance, J.P. Landers, Anal. Chem. 81 (2009) 5249. G.R.M. Duarte, C.W. Price, B.H. Augustine, E. Carrilho, J.P. Landers, Anal. Chem. 83 (2011) 5182. D.J. Marchiarullo, J.Y. Lim, Z. Vaksman, J.P. Ferrance, L. Putcha, J.P. Landers, J. Chromatogr. A 1200 (2008) 198. K.A. Hagan, C.R. Reedy, J.M. Bienvenue, A.H. Dewald, J.P. Landers, Analyst 136 (2011) 1928. Y.J. Hua, A.B. Jemere, D.J. Harrison, J. Chromatogr. A 1218 (2011) 4039. C. Wang, A.B. Jemere, D.J. Harrison, Electrophoresis 31 (2010) 3703. J.K. Liu, C.F. Chen, C.W. Tsao, C.C. Chang, C.C. Chu, D.L. DeVoe, Anal. Chem. 81 (2009) 2545. J.D. Ramsey, G.E. Collins, Anal. Chem. 77 (2005) 6664. B.C. Giordano, A. Terray, G.E. Collins, Electrophoresis 27 (2006) 4295. J.F. Borowsky, B.C. Giordano, Q. Lu, A. Terray, G.E. Collins, Anal. Chem. 80 (2008) 8287. G. Proczek, V. Augustin, S. Descroix, M.C. Hennion, Electrophoresis 30 (2009) 515. H. Chen, Q. Fang, X.F. Yin, Z.L. Fang, Lab Chip 5 (2005) 719. S. Nagrath, L.V. Sequist, S. Maheswaran, D.W. Bell, D. Irimia, L. Ulkus, M.R. Smith, E.L. Kwak, S. Digumarthy, A. Muzikansky, P. Ryan, U.J. Balis, R.G. Tompkins, D.A. Haber, M. Toner, Nature 450 (2007) 1235. S.L. Stott, C.H. Hsu, D.I. Tsukrov, M. Yu, D.T. Miyamoto, B.A. Waltman, S.M. Rothenberg, A.M. Shah, M.E. Smas, G.K. Korir, F.P. Floyd, A.J. Gilman, J.B. Lord, D. Winokur, S. Springer, D. Irimia, S. Nagrath, L.V. Sequist, R.J. Lee, K.J. Isselbacher, S. Maheswaran, D.A. Haber, M. Toner, Proc. Natl. Acad. Sci. U.S.A. 107 (2010) 18392. K. Hoshino, Y.Y. Huang, N. Lane, M. Huebschman, J.W. Uhr, E.P. Frenkel, X.J. Zhang, Lab Chip 11 (2011) 3449. A. Terray, J.D. Taylor, S.J. Hart, Biomicrofluidics 3 (2009) 44106.