Current advances and challenges in microfluidic free-flow electrophoresis—A critical review

Accepted Manuscript Current advances and challenges in microfluidic free-flow electrophoresis—A critical review Pedro Novo, Dirk Janasek PII:

S0003-2670(17)30930-3

DOI:

10.1016/j.aca.2017.08.017

Reference:

ACA 235389

To appear in:

Analytica Chimica Acta

Received Date: 28 March 2017 Revised Date:

10 August 2017

Accepted Date: 11 August 2017

Please cite this article as: P. Novo, D. Janasek, Current advances and challenges in microfluidic freeflow Electrophoresis—A critical review, Analytica Chimica Acta (2017), doi: 10.1016/j.aca.2017.08.017. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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A review on microfluidic free-flow electrophoresis Microfluidics Fabrication methods Materials

Injection

Geometry

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Flow rates Residence times

Dimensions Design/shape Arrangement

Electrodes

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Materials Integration Separation

Separation of analytes

Output strategies

Separation modes Analytes separated Efficiency

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Chip-world interface Hyphenation

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Current Advances and Challenges in Microfluidic Free-Flow Electrophoresis—A critical review Pedro Novo and Dirk Janasek∗

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Leibniz-Institut f¨ ur Analytische Wissenschaften – ISAS – e.V, 44227 Dortmund, Germany, Otto-Hahn-Str. 6b

Abstract

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The research field on microfluidic free-flow electrophoresis has developed vast amounts of devices, methods, applications and raised new questions, often in analogy to conventional techniques from which it derives. Most efforts have been employed on device development and a myriad of architectures and fabrication techniques have been reported using simple proof-of-principle separations. As technological aspects reach a quite mature state, researchers’ new challenges include the development of protocols for the separation of complex mixtures, as required in the fields of application. The success of this effort is extremely dependent on the capability to transfer the device’s fabrication to an industrial setting as well as to ensure interfacing simplicity, namely at the solutions’ supply and collection, and actuation such as electric potential application and temperature control. Other advanced applications such as direct interfacing to downstream systems such as mass spectrometry, integration of sensing and feedback controls will require further development in the laboratory. In this review we provide an overview on the field, from basic concepts, through advanced developments both in the theoretical and experimental arenas, and addressing the above details. A comprehensive survey of designs, materials and applications is presented with particular highlights to most recent developments, namely the integration of electrodes, flow control and hyphenation of microfluidic free-flow electrophoresis with other techniques.

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Keywords: Microfluidic free-flow electrophoresis, materials, designs and fabrication techniques, miniaturization, device-world or device-device interface

∗

Corresponding author

Preprint submitted to Analytica Chimica Acta

August 19, 2017

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List of abbreviations ABS acrylonitrile butadiene styrene BGE background electrolyte

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CE capillary electrophoresis CGE capillary gel electrophoresis CIEF capillary isoelectric focusing CITP capillary isotachophoresis

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COP cyclic olefin polymer

EOF electroosmotic flow ESI electrospray ionization FFE free-flow electrophoresis

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CZE capillary zone electrophoresis

FF-FSE free-flow field-step electrophoresis FF-IEF free-flow isoelectric focusing

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FF-ITP free-flow isotachophoresis

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FF-ZE free-flow zone electrophoresis FITC fluorescein isothiocyanate

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FSE field-step electrophoresis GE gel electrophoresis

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HEPES 4-(2-hydroxyethyl)-1-piperazine ethane sulfonic acid IEF isoelectric focusing ITP isotachophoresis LE leading electrolyte LIF laser induced fluoresecence µFFE microfluidic free-flow electrophoresis

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µFF-IEF microfluidic free-flow isoelectric focusing µFF-ITP microfluidic free-flow isotachophoresis

µFF-ZE microfluidic free-flow zone electrophoresis MS mass spectrometry NOA 81 Norland Optical Adhesive 81 OEG-DA oligoethyleneglycol diacrylate

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PCB printed circuit board

PET polyethylene terephthalate pI isoelectric point

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PDMS polydimethylsiloxane PEEK polyether ether ketone

PMMA polymethylmethacrylate PS polystyrene

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PTFE polytetrafluoroethylene PVA polyvinyl alcohol

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µFF-FSE microfluidic free-flow field-step electrophoresis

SDS sodium dodecyl sulphate

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TE terminating electrolyte

TGF temperature gradient focusing

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ZE zone electrophoresis

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Contents 1 Introduction

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3 Theory

Free-Flow Electrophoresis 7 . . . . . . . . . . . . . . . . . 7 . . . . . . . . . . . . . . . . . 8 . . . . . . . . . . . . . . . . . 10 . . . . . . . . . . . . . . . . . 10 . . . . . . . . . . . . . . . . . 13

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2 Principle and Modes of Separation in 2.1 Zone Electrophoresis Mode . . . . . . 2.2 Field-Step Electrophoresis . . . . . . 2.3 Isotachophoresis Mode . . . . . . . . 2.4 Isoelectric Focusing Mode . . . . . . 2.5 Other Separation Modes . . . . . . .

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4 Approaches and Designs for Introduction of the Electric 4.1 Direct Electrode Integration . . . . . . . . . . . . . . . . . 4.2 Isolation of Electrodes by Geometrical Structures . . . . . 4.3 Gels as Salt Bridges . . . . . . . . . . . . . . . . . . . . . . 4.4 Integration of Commercial Membranes . . . . . . . . . . .

Field . . . . . . . . . . . . . . . .

. . . .

17 20 21 25 29

5 Materials 31 5.1 Materials for Chips . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 5.2 Materials for Electrodes . . . . . . . . . . . . . . . . . . . . . . . . 33

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6 Applications 34 6.1 Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34 6.2 Analytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35

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7 Interfaces 35 7.1 Preparative Outlets to the Macro-World . . . . . . . . . . . . . . . 36 7.2 Hyphenation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38 39

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8 Conclusion

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1. Introduction

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Electrophoresis is a popular technique in the laboratory for the separation of charged species upon application of a voltage generating an electric field across a separation compartment filled with gel matrix in case of gel electrophoresis (GE), or free liquid solution in case of capillary electrophoresis (CE) . For instance, it is routinely used for the separation of nucleic acids and proteins. Freeflow electrophoresis (FFE) is a variation of CE in which analytes separate in a hydrodynamically-driven liquid matrix flowing in a direction perpendicular to the electric field. Using liquid matrices allow for fast separation and direct sample collection, while compromising parallelization of the separation, meaning only one mixture/sample is separated at a time. Miniaturization through microfluidics leads to microfluidic free-flow electrophoresis (µFFE) which allows: (i) increase of separation speed which results in reduced sample residence times, (ii) high control of minute amounts of solutions in the 10–100 µL range (namely that of the sample), and (iii) portability, integration of several operational units and cost reduction. These advantages relate to: (i) the high area-to-volume aspect ratios which allow for fast heat dissipation and the introduction of high electric field strengths, (ii) the laminar flow regime in channels of characteristic dimensions in the micrometer range (reduced dead volume, e.g. in the pL to µL range) allow for precise flow control, such as demonstrated by the monolithic integration of a pH gradient generator with a µFFE device [1], and (iii) small device footprint, respectively. On the other hand, the use of microfluidics might be prohibitive when large sample preparation capacity is required due to the reduced flow rate possible (up to several hundreds of µL/min). However, a compromised size reduction might still be advantageous as compared to macro-scale FFE, namely the use of millifluidics which allows flow rates up to 2 mL/min [2]. A vast number of works on µFFE is already available in the literature (fig. 1). With it, a vast set of fabrication arrangements and methods, as well as materials used and separations performed have been demonstrated. Despite this large set of information, practical use of µFFE devices in the laboratory, for instance, has not yet been established. Most materials and fabrication methods that are not compatible with industrial manufacturing, and the lack of device-world and device-device interfaces are significant issues contributing to the problem. These practical aspects have not been in the scope of the reviews by Kohlheyer et al. [3] and Turgeon et al. [4]. However, the research community starts gaining momentum towards solving these issues. Several works have been published recently, setting new development trends which will be covered in this review. Here, we will provide an overview of the state-of-the art, highlighting most recent developments while critically addressing its characteristics, potentials and issues that hinder the transfer of the technology to an environment of routine use.

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Citations in Each Year

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Published Items in Each Year

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Figure 1: Web of Science citation report using the “microfluidic free-flow electrophoresis” topic. On the 6th June 2017.

2. Principle and Modes of Separation in Free-Flow Electrophoresis

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In FFE, the electric field is set perpendicular to a hydrodynamically pumped liquid system which consists of a sample fed into a carrier electrolyte (also named as background electrolyte (BGE), carrier buffer or separation buffer) flowing through a channel in the laminar regime. Each charged analyte molecule or particle i is deflected from the flow direction by the interplay of the drag velocity ~vhd and their specific electrophoretic velocity ~vi,ep . The magnitude of the individual ~vi,ep is a function of the type of analyte, the local buffer environment and electric field strength. In general terms and for charged species, the resulting analyte velocity vector, which is the sum of both velocities mentioned above, will result in diagonally flowing analytes and therefore in a two-dimensional separation with subsequent fractionation at the outlets. FFE can be applied to all separation modes known from CE [5]. Kaˇsiˇcka et al.[6] described the theoretical basis of the correlation between analytical capillary zone electrophoresis (CZE) separation and free-flow zone electrophoresis (FF-ZE) in preparative processes, which was also applied to free-flow isotachophoresis (FF-ITP) [7, 8] and free-flow isoelectric focusing (FF-IEF) [9]. The difference between the separation modes lies in the number and the composition of the BGE as well as gradients generated or externally applied (tab. 1). 2.1. Zone Electrophoresis Mode The simplest separation mode is zone electrophoresis (ZE) in which only one kind of BGE is used. The properties of the BGE, e.g. concentration, conductivity and pH, is comparable to those of the sample. Hence, no gradients are formed during operation resulting in constant velocity vectors ~vhd and ~vi,ep for each given analyte, at any time or local point in the separation bed.

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Because of its simplicity, this separation mode is often used for inaugural proofof-concept experiments demonstrating the capability and performance of conceptually new devices [1, 10–15], as also done by Raymond et al.[16] for the first-ever developed µFFE device. A variant of ZE can be established if the BGE is modified with a sieving matrix. Then, comparable to gel electrophoresis, the analyte size as an additional separation parameter comes into play. However, since the hydrodynamic flow is essential in FFE, the use of a cross-linked gel has to be omitted. As known from capillary gel electrophoresis (CGE) linear sieving matrices, e.g. polyethylene glycol or dextranes can be used. Novo et al.[17] utilized a BGE containing 0.5% 2-hydroxyethyl cellulose and 0.5% glycerol in order to separate a 5-fragment DNA ladder.

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2.2. Field-Step Electrophoresis In field-step electrophoresis (FSE) the deployed BGE varies from the sample in terms of conductivity causing a step in electric field [18]. The sample is dissolved in an electrolyte with low electrical conductivity (S and B1, respectively, in the FSE separation scheme in tab. 1) and introduced into the separation bed between two sheath buffers with higher conductivity (B2). This means that the electric field is high in the zone of sample buffer, causing fast migration of the charged analytes inside that zone towards the electrodes. Due to the step in the electric field at the boundaries between sample buffer and sheath buffer, separated analytes are strongly decelerated and concentrated there. The concentration factor strongly depends on the conductivity ratio between the buffers. Because of the flow scheme set-up only two concentrated streams at maximum can be obtained. Although many examples applying FSE are known for desktop FFE [18, 19, and references within] only few have been presented for µFFE [3, 17]. Kohlheyer et al.[3] showed the first implementation separating a mixture of rhodamine and fluorescein which concentrated four-fold at the boundary to the higher-conductive sheath buffer. However, only limited information is available in that publication, which makes it difficult to reproduce the results shown. To our knowledge only one additional demonstration of FSE in µFFE has been published. Recently, Novo et al.[20] conducted a proof-of-principle experiment for the concentration of fluorescein isothiocyanate (FITC) using a microfluidic free-flow field-step electrophoresis (µFF-FSE) device. The sample buffer was joined only at the anodic side by a buffer with 10-fold higher conductivity so that the negatively charged FITC was concentrated at that boundary. The concentration factor was only about 1.6 since the conductivity ratio between the neighboring buffers, which is strongly affecting the concentration efficiency, was only 10-fold which is not high enough according to [18].

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Table 1: Characteristics of the different separation modes employed with µFFE Gradient in

Separation Parameter

1-buffer system

Electrophoretic mobility

1-buffer system with sieving matrix

Electrophoretic mobility and size

Separation Scheme Sample

Buffer

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Zone Electrophoresis

BGE System

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Separation Mode

2-buffer system

Electric field

Electrophoretic mobility

Isotachophoresis

2-buffer system

Electric field

Electrophoretic mobility

Isoelectric Focussing

Ampholyte system

[1, 10–16] [17]

B2

B1 S

B1

Terminating Electrolyte

B2

Leading Electrolyte

[17, 98]

[15, 22–24]

pH

Isoelectric point (pI)

+ 0 - 33 2 5- 4- 1 2 54 4 3- 0 3- 6- 35 23 250 4- 4 4 20 -3 1 2 53 0 3 4 2 1 2- 3 0 -

6-

2-

1-

0

0 0 0 0 0 0 0 0 0 0 0 0

Temperature

1

-

2-

1-

21+

1- 2 + 2- 1 -

+ 10- 2

2+ 1

-

+ 1- 2 0

2+

1-

0

0 3+ 1+

4+ 1+

+

0

+

0 1+ 1 0 1 1- + 0 + 0 10 1 2 0 0 + 1 1 1 + 10 1 10 1 1+ 1 1+ 0 0 1-

0 10 0 0 0 0

0

1+ 2

[1, 11, 26–29, 30–35]

+

2+ 1

+

0 0 0

0 0

0 0 0 0 0 0 0 0 0 0

1+ 2 1+ 1+ +

3

3+

0 1 1+ 0

-

0

0

0 0 0 0 0 0 0 0 0

Electrophoretic mobility

[39] Hot

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1-buffer system with temperature depending conductivity

+

1+

0 0

Temperature Gradient Focussing

0

0 0 0

1 1- 2

Cold

TE

D

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Field-Step Electrophoresis

References

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2.3. Isotachophoresis Mode In isotachophoresis (ITP) a mixture of analytes in sample buffer is introduced between a 2-BGE system which differ in their electrophoretic mobility (see tab. 1). The leading electrolyte (LE) and terminating electrolyte (TE) are chosen to have higher and lower electrophoretic mobility than any analyte of interest in the mixture, respectively. As a result, analytes concentrate into adjacently stacked bands as a function of their electrophoretic mobility. According to Kohlrausch’s regulating function [21], the analyte concentration at each band adapts to that of the LE, causing a concentrating effect of originally low-concentrated samples and vice-versa. Additionally, after full separation, the single zones contain only individual analytes ions and the (preferable volatile) counter-ion, making it an optimal mode as sample preparation for direct coupling to mass spectrometry. The resolution of ITP in matrix-free systems, such as CE and FFE, is compromised by the electroosmotic flow (EOF); thus EOF has to be avoided or reduced by the choice of the device’s material (see section 5.1) or by dynamic surface coatings. For more information the reader is referred to [22]. Although extensively employed in desktop FFE [5, and references therein], only a few works have been reported in µFFE [15, 23–25]. Janasek et al.[23] used ITP in their microfluidic device to separate and concentrate fluorescein from eosin G using acetyl salicylic acid as a non-fluorescent spacer in a proof-of-concept experiment. In a follow-up experiment of the same work, the products of a labeling reaction of myoglobulin with FITC were separated in order to remove excess of unreacted FITC molecules, using serin as a non-fluorescent spacer. Furthermore, a ca. 5fold concentration increase was obtained at an electric field of 480 V/cm and for a residence time of 50 s. Park et al.[25] utilized microfluidic free-flow isotachophoresis (µFF-ITP) as a sample pretreatment prior to direct coupling to mass spectrometry (MS) (see also section 7.2). A system of formic acid as LE and propionic acid as TE was chosen in order to separate and concentrate two fluorescence dyes (Alexa Fluor 488 and fluorescein), glycolic acid and citric acid when applying a total flow rate of 10 µl/min at an electric field of 520 V/cm. Fu et al.[15] employed µFF-ITP, besides microfluidic free-flow zone electrophoresis (µFF-ZE), as a performance demonstration of their device. A sample mixture of Alexa Fluor 591 and fluorescein was introduced between a buffer system of hydrocloric acid as LE and HEPES as TE, at an electric field of 300 V/cm. 2.4. Isoelectric Focusing Mode As introduced previously, BGE gradients formed in the direction of the electric field result in analyte focusing effects. In isoelectric focusing (IEF), a gradient in the pH value is generated (see tab. 1), which is used to exploit the fact that multiple charged molecules exhibit different net charge as a function of the local

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pH. The molecule’s net charge is positive in acidic environments and negative in alkaline ones. The transition from one charge state to the other occurs at the molecule’s so-called isoelectric point (pI), corresponding to a zero net charge state. Thus, molecules migrate inside the pH gradient towards the electrode of opposite polarity until its surrounding pH equals its pI. At this location, the molecule no longer deviates due to electrophoresis. However, the molecule may migrate, due to diffusion, into adjacent regions with pH values different from their pI and re-acquire charge which in turn will cause them to re-concentrate back to the original pI region also known as the self-sharpening effect [26]. The pH gradient is formed from the BGE consisting of synthetic carrier ampholytes, which is a mixture of several hundred low-molecular aliphatic oligoaminooligocarboxylic acids, under the effect of an electric field. Similar to ITP, EOF compromises the performance of IEF and therefore has to be taken into consideration (compare section 2.3). IEF has been employed extensively in CE as well as in FFE due to their great importance in the biological and pharmaceutical field [5, and references therein]. IEF has been applied in µFFE to a great extent as well [1, 11, 27–36]. Xu et al.[27] showed with their device for the first time the validity of the scaling laws [37] for microfluidic free-flow isoelectric focucusing (µFF-IEF) when they separated and focused a mixture of two peptides within half a second at an electric field of 135 V/cm. The Jensen’s group reported on the use of µFF-IEF for the separation and enrichment of mitochondria and nuclei out of a mixture of whole cell lysate and intact cells [28] as well as for the focusing of proteins and protein complexes [29, 30]. Kohlheyer et al. employed µFF-IEF for the separation of fluorescent IEF markers using an ampholyte system [11] as well as a pre-separated ampholyte system [31] resulting in a more efficient pH gradient and hence highly improved separation resolution. Albrecht et al.[30] demonstrated the application of a cascaded µFF-IEF design (fig. 2) in order to obtain a separation resolution equivalent to conventional bench-top FF-ITP devices but with shorter residence times, and employed it for the focusing of fluorescent IEF markers, model protein mixtures and whole cell lysates. Wen et al. used their triangular shaped µFFE device (see fig. 12) for IEF separation of fluorescent pI markers [32] and protein mixtures [33] showing the comparability between results using the microfluidic system and standard 2D gel electrophoresis. The Nagl group employed a dedicated chip having an integrated fluorescence layer for real-time pH imaging. A near-infrared pH dependent fluorescence probe immobilized in a ∼9 µm thick polymeric sensor layer at the bottom of the µFFE separation bed was used to quantify the pH values across the separation bed during operation [34, 35]. At the same time label free [35] as well as on-chip fluorescently labeled analytes [36] could be detected.

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Figure 2: Cascaded µFF-IEF device design: (a) rendering of the cascaded device. (b) Detail of square channel posts and trapezoidal support array. (c) Photo of the device under operation. Reprinted with permission from [30]. Copyright 2007 American Chemical Society.

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2.5. Other Separation Modes In temperature gradient focusing (TGF) separation mode, as the name suggests, a temperature gradient is applied. The temperature gradient can be generated by an external source or by inherent Joule heating within the buffer [38]. The BGE used for TGF is characterized by a strongly temperature dependent ionic strength and consequently conductivity. Hence, a gradient in temperature will cause a gradient in electric field which in turn affects the electrophoretic velocity of charged analytes. The electrophoretic velocity gradient across a separation bed is counterbalanced by the bulk buffer velocity caused by EOF. Analytes will focus at the separation bed position at which both velocity forces are equal in magnitude, which is equal to an overall apparent velocity of null. Analytes differing in electrophoretic mobility will focus at different points along the gradient and thereby separate. In contrast to IEF which is applicable only to molecules with an accessible pI, or to ITP applicable to analytes with electrophoretic mobilities values in the LE-to-TE range, TGF can be applied to any charged analyte. For an overview of TGF in capillaries the reader is referred to [39]. The first implementation of TGF separation mode in µFFE was demonstrated by Becker et al.[40]. The separation bed was mounted on top of a pair of peltier elements which set temperatures of 10 ◦C and 80 ◦C. As thermometry investigations revealed, the external temperature gradient resulted in a buffer temperature gradient from ca. 25 ◦C to ca. 75 ◦C when the linear buffer velocity inside the separation bed was around 1.2 mm/min which corresponds to a 12.5 ◦C/mm temperature gradient. With this setup fluorescent dyes and proteins were separated and concentrated by a factor of 2. Compared to the higher concentration factors of up to 100 in capillary TGF, the only two-fold concentration was due to the short residence time of the sample inside the separation bed of only 10 seconds.

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In CE there is a mode called CGE which is a sub-mode of CZE and combines the size-depending separation in gels with the advantages having liquid buffers in CE. The BGE employed in that separation sub-mode contains a sieving matrix consisting of either cross-linked or linear polymers. CGE has been widely used for the separation of e.g. proteins, DNA, antibodies [41] and virus capsoid proteins [42]. The reader is cordially referred to some reviews in this area [43–45]. To our knowledge only one approach to implement a sieving matrix in FFE/µFFE separations has been reported [17]. Recently, Novo et al. used a BGE containing hydroxycellulose as a sieving matrix in order to successfully separate a 5-fragment DNA ladder as a proof-of-concept experiment. Advanced separation modes might be obtained by the combination of existing modes, namely for performing 2D separation as reviewed by Chen et al.[46].

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3. Theory

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As for all separation methods, the analytical performance depends on the resolution power which is related to the broadening of the analyte zones during the migration through the separation bed. For FF-ZE separations, the overall stream variance σTOTAL 2 is the sum of variances due to several factors, with the most prominent being injection, diffusion and hydrodynamic broadening [47]:

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σTOTAL 2 = σINJ 2 + σDIFF 2 + σHD 2 + . . . wINJ 2 σINJ 2 = 12 L σDIFF 2 = 2Dt = 2D v 2 2 h d v σHD 2 = 105DL

(1) (2)

(3) (4)

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where wINJ is the width of the sample inlet channel, D the diffusion coefficient, t the time, L the length of the separation chamber, v the linear velocity, h the height of the separation compartment and d the migration distance of the analyte species in direction of the electric field. The variance due to injection σINJ 2 depends in FFE from the width of the sample inlet channel and can be expressed analogously to the injection length in CE [48] (eq. 2). The variance σDIFF 2 is based on the molecular diffusion of the analyte with a compound-specific diffusion coefficient D while traveling with the linear velocity v through the separation chamber with the length L in a time t (eq. 3) [48]. The variance resulting from hydrodynamic broadening σHD 2 is caused by the parabolic flow profile of the hydrodynamic flow through the separation compartment. As the molecules in the middle between top and bottom plate of the FFE chamber move faster than the ones more toward the wall, the fastermoving molecules are affected by the electric field for a shorter time and thus do electrophoretically migrate a shorter distance compared to molecules closer to the walls (eq. 4 [47]. There are other factors contributing to band broadening which however in FFE and µFFE have smaller or no impact: 1) Electrodynamic broadening is based the electroosmotic flow which leads to a flow in the proximity of the wall towards the cathode whereas molecules in the cross-sectional center of the chamber are relatively unaffected. If the electroosmotic flow is opposite to the direction of the electrophoretic flow it can offset the band broadening caused by hydrodynamic distortion [49]. 2) Electrohydrodynamic distortion results from shear stresses caused by differences in the conductivity between sample and BGE. It can be exploited to

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advantage when the conductivity of the sample stream is lower than the surrounding BGE since it will lead to a concentration of the sample [49]. 3) Joule’s heating due to electrical current passing the FFE chamber results in band broadening because of thermal convection. The high surface-to-volume ratio in in microfluidic structures promotes a good heat dissipation minimizing that effect in µFFE. 4) Effects of surface adsorption and desorption of analyte molecules to the wall do not contribute to spatial band broadening in FFE separations as demonstrated by Geiger et al.[50].

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In order to design optimal devices, theoretical descriptions, mathematical models and simulations both for desktop devices and miniaturized approaches have been published [4, 28, 47, 49, 51–55]. Fonslow et al. identified the effect of hydrodynamic broadening as one of the major sources of band broadening in FF-ZE separations [47]. They deduced a general equation for band broadening in µFFE devices (eq. 5) analogous to the van Deemter equation: h2 · d2 · v winj 2 2 · D · L + + (5) 12 v 105 · D · L where winj is the width of the sample inlet channel into the separation bed, D the diffusion coefficient, L the lateral displacement between the sample inlet and the detection zone (or outlet from the separation bed), v the linear velocity of the buffer, and d the lateral migration distance of the analyte. They specified an optimal velocity vopt that minimizes the band width of an analyte at a particular position (eq. 6) as well as the resolution, RS , of a pair of analytes (identified with the indices 1 and 2 in the equation) for any combination of linear velocities and electric fields (eq. 7). √ 210 · D · L (6) vopt = h·d

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(µtotal1 − µtotal2 ) · E · t q 2 winj h2 ·d2 ·v + 2·Dv1 ·L + 105·D11 ·L + + 2·Dv2 ·L + 12

h2 ·d22 ·v 105·D2 ·L



(7)

where µtotali is the total mobility of the i th analyte. The results obtained by Fonslow et al. have been confirmed by Dutta’s methodof-moments formulation except for a factor 15 by which the contribution of hydrodynamic dispersion under non-ideal settings exceeds the theoretical predictions [53]. Therefore, Dutta augmented the description in order to quantitatively characterize the electrodynamic dispersion component based on transport equations [55]. In addition to the pressure-driven flow in the axial direction, an α fraction

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Figure 3: (a) Scheme of the FF-ZE fractionation process with a pressure-driven back-flow in the transverse direction. (b) Top view of a microfluidic FF-ZE device relevant to the mathematical analysis presented by Dutta. Reprinted from Ref. [55], Copyright 2015, with permission from Elsevier.

TE

D

of the net-transverse electrokinetic flow is assumed to be blocked by the channel sidewalls yielding in a pressure-driven cross-flow countering the EOF in the FF-ZE channel (fig. 3). The optimum flow velocity vopt at which the stream variance is minimized is then re-written as: s   2 · S2 · D D · L · (1 − α)2 · (8) vopt = 105 · 2 · D · L + L S 2 · d2

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where S = µE(1 − α)L/vhd , with vhd corresponding to the axial pressure-driven flow speed. α is a fraction of the net transverse electrokinetic flow, which is assumed to be blocked by the channel sidewalls [55]. Matsumoto et al. combined macroscopic models of electroosmotic flow, heat generation and viscosity’s change with the Navier-Stokes equation model and energy equation model in order to simulate temperature distribution inside the separation bed of a µFFE module. With this hybrid model simulation, heterogeneous temperature distributions seen in experimental results could be confirmed [56]. Lu et al. set up two pseudo 2D models and a 3D model to simulate molecular IEF and IEF of mitochondria, respectively [28]. The 2D simulations showed that the time scale for pH gradient establishment is on the order of 20 s when a potential of 20 V across an 1 mm wide microchannel is applied. Due to a smaller diffusion coefficient, the concentration of mitochondria is much slower, however

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Figure 4: Simulations of free-flow isoelectric focusing. (A) 3D simulation of µFF-IEF of mitochondria using a reaction-diffusion model. Adapted with permission from ref. [28]. Copyright 2004 American Chemical Society. (B) Numerical simulation of FF-IEF for fully open (Case A) and partially blocked (Case B) channel when focusing hemoglobin B (HBB) and G (HBG). Reprinted from ref. [54], with the permission of AIP Publishing.

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with a narrower peak width at steady state. Therefore, it can be assumed that the focusing of the mitochondria happens in an already established pH gradient. The simulation using the 3D model gave the same time scale for focusing of mitochondria, furthermore revealing that top and bottom of the velocity field are more focused than the middle (fig. 4A) owing to the longer residence time in these positions as a result of the parabolic velocity profile [28]. Yoo et al. developed a mathematical model implementing a finite volume based numerical scheme to simulate a two-dimensional FF-IEF in a microfluidic chip considering 48 ampholytes and two proteins for an applied electric field range of 133 to 533 V/cm. Furthermore, it was shown that a partial flow blockage caused by the insertion of a post creates significant cross-flow which improves the separation resolution of proteins with very close pI by increasing the separation distance between the focused peaks (fig. 4B:Case B). However, the higher separation resolution is achieved at the expense of the final concentration of the focused proteins, as it can be seen from the comparison between the fully open (fig. 4B:Case A) and the partially blocked channel (fig. 4B:Case B) [54]. 4. Approaches and Designs for Introduction of the Electric Field Besides the general need for electrodes to set a potential at a separation bed, literature in µFFE devices is quite diverse at different levels. For instance, different methods for introducing the electric field, values of applied potential and resulting electric field, separation bed dimensions, number of inlets/outlets, materials and

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fabrication procedures, sets of buffers, chemicals and analytes used, integrated sensing and actuation, among others, are available. We review each main aspect of µFFE devices in independent subsections below in order to address details of each configuration and for simplicity of comparison, which is often not possible or not compatible in the case of whole devices. A comprehensive overview of the several approaches is listed in tab. 2.

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Table 2: Different µFFE devices presented in literature, classified by design, operation and features.

Small connection channels

10 µm×(4×12) mm2

×

× Direct integration Small connection channels 146 µm glass membrane as dielectric barrier Direct integration or recessed electrode compartment

20 µm×(10×50) mm2

Two-depth channels

17 µm×(4×12) mm2

Acrylamide gel

15 µm×(3×10) mm2

30 µm×4.4×12.2 mm2

Direct integration Two-depth channels

25 µm×(20×30) mm2

Direct integration on side chimneys Direct integration Solid wall electrodes

Pt wire

Microfabrication Wet etch Anodic bonding

Low voltage breakdown

[16]

NA

20 s

2 V, 30 µA

20

100**

4

Glass SU8

Ti/Au thin-film

Microfabrication Electrodeposition

Organelle separation

[28]

1.7-3.4 nl/s

2s

1.75 kV, 140 µA

135

<12*

Pt wire

Microfabrication Soft-lithography

Aspiration at the outlet

10 µl/min

93.7 s

1.4 kV

350

10*

25-70 µl/min

50 s*

Up to 562 V, Up to 310 µA

Up to 283

50

1

Glass/ amorphous silicon

Cr/Au thin-film

20 µl/min

143 s*

150 V

180

50

64

Glass/glass

Woods alloy

3.13 ml/min

2 s*

>1000 V

Up to 370

100**

3

Polystyrene Carbon fiber loaded PS

Carbon fiber loaded PS

4.3 µl/min

370 s$

3.15 kV

119

10*

11

Glass/glass and PDMS

Pt wire

∼ 4 µl/min

∼ 3s

150 V, 50 µA

200-250

42-46*

3-5

Glass/glass

Protein pI markers Cytochrome C BSA, Hemoglobin

∼ 17 µl/min*

∼ 12 min

×

FITC-labeled aminoacids

∼ 0.17 µl/min

1.5 s*

200 µm×(100×150) mm2

×

Fluorescein Rhodamine B & 6G

∼ 5 ml/min

6 s*

25 µm×(633 µm×5 mm)

×

∼ 4.2 µl/min

∼ 1s

35 µm×(1.8×14) mm2

Microfabricated pillars External electrodes

70 µm×(10×20) mm2

Packed microbeads

13 µm×(1×5) mm2

×

70 µm×(10×20) mm2

50 µm×(14×28) mm2

80 µm×(10×25) mm2

Microfabricated pillars External electrodes

5 µm×(23×15) mm2

Acrylamide gel

20 µm×(11.5×20) mm2

Polycarbonate membranes

50 µm×(17×33) mm2

0.5×(50×100) mm3 Normal filter membranes * Calculated from presented data ** Assumed due to direct electrode integration $Estimated

×

× ×

50 µm×(8×12) mm2 (h NA)×(1×10) mm2

Fluorescein Rhodamine 6G Protin pI markers Ovalbumin Avidin pI markers Different albumins Blood serum pI markers AlexaFluor 488

× ×

×

×

×

outlets

Silicon/ oxide nitride/ glass

100-300 s$

×

Refs

NA

NA

×

Other

NA

Fluorescein Dunnite; TNP; 2,4-D

Polymer salt bridges

Direct electrode integration, two-depth channels

×

Fabrication

NA

∼ 56 µl/min*

∼ 1 s*

∼ 5 µl/min*

56 s*

2 µl/min

154-204 V

TE

Triangle: 160 µm×(46×56)/2 mm2

×

Amaranth, Acid green Bromophenol blue Glutamate Myoglobin Trypsin inhibitor Rhodamine B IEF markers

Electrodes

Up to 50 V

∼ 4 s*

Sulforhodamine Rhodamine 123 & B

∼ 9 µl/min

Fluorescein Rhodamine B AlexaFluor 488 & 591

∼ 10 µl/min

Proteins Peptides

∼ 14-30.6 µl/min ∼ 14-29 s*

∼ 14 s*

FITC, Sulforhodamin B Rhodamine 120, 6G & B Coumarin, Aspartic acid × NA Glycine, 2-butylamine B-phycoerythrin, StrII-eGFP Fluorescein Rhodamine 110 & 123 × 1001 µl/min Myoglobin, Cytochrome C Fluorescein AlexaFluor 488 × 10 µl/min Citric acid Glycolic acid Rhodamine 101, 123 & B Fluorescein Reagents and products × 19.5 µl/min in the synthesis of pyrrolobenzoxazoles FITC, Rhodamine B Pyrinin Y × × × × 15-102 µl/min Protein pI markers DNA × × × × × 10s-100s ml/h # Not a µFFE device. It is a capillary electrophoresis on chip and mentioned here to highlight the integration level NA Not available

1

67-88*,**

PDMS/glass

PDMS/PDMS

[10]

[23]

100**

1

Glass/glass

NA

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PDMS/glass

Pt wire

NOA 81/glass

Pt strips

1.8 kV, 2 mA

370±20

15-80

230*,**

115

100**

500 V

50

100

1

PMMA

Pt wire

Up to 10 V, Up to ∼ 30 µA

Up to ∼ 30

100**

1

PDMS/galss

InBiSn alloy

Microfabrication Anodic bonding Microfabrication Thermal bonding

[57] All outlets merged into one channel

[58]

Injection molding

[59]

Microfabrication Wet etching Thermal bonding

[60]

Microfabrication Soft-lithography Microfabrication Wet etching Lithography Milling Dichloromethane bonding Microfabrication Soft-lithography

[11, 31] Hand-held device with integrated optical detection Triangular shape Comparison with gel electrophoresis No sheath Photopolimerizable glue for channel fabrication Fast prototyping Cost effective Millifluidic range Liquid InBiSn electrodes Integrated pH gradient generator based on bipolar membranes Integrated pH sensor layer Characterization by MS

[61]

[32, 33]

[52]

[62] [63]

40-70 V

360

93

8

Polyurethane PDMS/glass

Ag/AgCl Pt wire

I = 500 µA

Up to 152

NA

5

OEG-DA/glass

NA

Microfabrication Lithography

Up to 50 V

Up to 78

36-52

2

PDMS/glass

Pt wire

Microfabrication Soft-lithography

No sheath

[12]

300-830 V, 20-60 µA

375-1038

Up to 400 V

[1]

[34, 35]

Adhesive foil/glass/ PTFE membrane/ PET foil PDMS/carbon black composite/ glass

NA

Laser cutting Lamination

Gas removal by using a Teflon membrane

[64]

Gallium and PDMS/carbon black composite

Microfabrication Soft-lithography

Liquid gallium electrodes

[15]

82

5

NA

NA

5

NA

20-300

NA

5

OEG-DA/glass

NA

Microfabrication Lithography

Integrated pH sensor and analyte labelling

[36]

NA

Up to 700 V

Up to 450*

90

5

COP PDMS/glass

Ti/Au thin-film

Injection molding Microfabrication

Injection molding using COP

[65, 66]

∼ 1.2 s$

Up to 100 V

100**

100**

4

ABS

3D printing, Solvent vapor bonding

Reduced EOF as compared with glass

[14]

1.2-2 kV, 100-200 µA

520-870

100**

7

14.15 s*

-602 V, -33 mA

NA

NA

3

16.5-112.2 s

Up to 200 V

Up to 60

100

9

>1-10s min

100s-1000s V

-

-

96

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Acrylamide gel

×

Fluorescein

Device

10-100 s of seconds

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25 µm×(55 µm×23.1 mm)

Side channel PDMS/carbon black composite membrane Microfabricated pillars External electrodes

×

100 µm×(36.4×28.7) mm2

Capillary electrophoresis on chip#

Electrodes connected through side channels

×

Other

IEF

ITP

×

Materials

15-50 µl/min

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(h NA)×(1×18) mm2

Fluorescein Acridine orange Labelled aminoacids Mitochondria Nuclei Peroxisomes Rhodamine 110 Fluorescein FITC labelled aminoacids Fluorescein, Eosin G Acetylsalicylic acid Myoglobin Fluorescein Rhodamine 110 & 123

Applied

Electric field Efficiency (V/cm) (%)

Residence time

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Direct integration

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Flow rates

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Analytes

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Small connection channels

Separation modes FSE

Separation bed dimensions (h×(w×l))

ZE

Electrode/separation compartment division

NA

10.35 s*

Pt wire

Microfabrication Wet etching Thermal bonding

Glass/glass

PDMS/ PC membrane/ glass

NA

NA

Pt wire

Microfabrication Soft-lithography

Direct coupling to MS Hydrodynamic complemented focusing Only one outlet effectively used

[25]

[13]

Flexible fabrication design Direct interface to a 96-well plate

[17]

Commercial, BD FFE

[67–71]

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An important challenge in FFE is the introduction of the electric field. While applying a voltage to the electrodes placed in the buffer solution, electrolysis happens, causing the generation of molecular oxygen and hydrogen and the formation of gas bubbles. These bubbles lead to local medium inhomogeneities which affect the efficiency of electric field introduction (eq. 9) as well as distorting the local flow dynamics, a chaotic process that occurs through time and therefore compromise stability of the separation. The electric field introduction efficiency, ηE , is defined as: (9)

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where Ueff is the effective voltage drop within the separation bed and Uapplied is the total applied voltage. These aspects do not represent serious implications in large systems such as gel electrophoresis, since bubbles are automatically degassed due to its different density, and inhomogeneities in the solution not being perceptible due to the large dimensions. However, in microfluidic systems bubbles are a considerable issue due to two aspects: firstly, bubbles are easily trapped, difficult to purge from microfluidic channels [72] and only certain removal strategies are compatible with µFFE as will be discussed hereafter. Secondly, although their size might be insignificant in the macroscale, bubbles in microstructures have a huge size relative to the dimension of the microchannel as it may be quickly assessed by the appropriate scaling laws. Hence, these bubbles are responsible for significant inhomogeneities which, in connection with the aforementioned issues, results in considerable distortion of the electric field introduction. However, the use of microfluidics is advantageous at several levels, namely sample volume and high control of flows in the laminar regime. These, and others discussed throughout this review, drive the motivation of µFFE researchers and developers. However, in order to exploit the advantages of microfluidics, the first task is to solve the bubble issue and to develop a strategy for electrode integration. In commercially available desktop FFE devices normal semipermeable membranes from e.g. cellulose acetate are used to make the isolation of the electrodes from a separation bed [5, 69]. This configuration allows for gas bubble purging and prevents analyte separation distortion by avoiding bubbles to enter the separation bed as well as by ensuring electric field introduction stability throughout time. With the exception of a few examples, which will also be presented hereafter, most of µFFE devices use design analogies to that of the conventional desktop devices. However, the integration of membranes or other analogous strategies are either difficult to implement, lack of separation stability, efficiency and resolution or compromise them. 4.1. Direct Electrode Integration The simplest µFFE device fabrication are those whose design has only direct electrode integration to the sides of the separation bed. However, it comes at a cost, especially at the range of applicability, separation throughput and stability. For instance, Lu et al. [28] developed a µFF-IEF device for the separation of organelles by applying low potential values of 2 V a value low enough to avoid bubble formation, across a channel of only 1 mm in width with direct electrode integration. It is important to note that

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the separation resolution was ultimately limited by the outlet fractionation throughput which is limited in narrow separation beds and therefore rendering the device highly application specific. In order to avoid the complication of bubble formation in a configuration of direct electrode integration, Kohlheyer et al. [73] used quinhydrone as electron acceptor/donor in the separation buffers. The use of this molecule prevents hydrolysis at the electrodes and therefore allows for higher potential values. However, bubble suppression was limited and high flow rates might be necessary to prevent gas formation at higher potentials, a situation that might fall outside regimens of high analyte separation throughput. There are other examples of direct electrode integration in µFFE [12, 14, 47, 59, 64, 74]. However, these often make use of local channel modifications [14, 47] or make use of a membrane [59, 64] and will therefore be presented in other specialized subsections hereafter. Other works made use of external electrodes [25, 34, 36, 64] whilst emphasizing other aspects of the device, application or results. We will not provide a discussion regarding this configuration as its detailed explanation is often missing in the related publications.

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4.2. Isolation of Electrodes by Geometrical Structures Positioning electrodes in reservoirs that are isolated from the separation bed via geometrical structures is the simplest alternative to direct electrode integration: in this case “geometric structures” correspond to local modifications that are exploited in two opposite ways: create a high hydraulic resistance path between electrode reservoirs and the separation bed [16, 23, 24, 27, 40, 57, 60, 75] or accommodate the electrodes into a chamber with lower hydraulic resistance [14, 47, 59, 65, 66, 76] through which bubbles may be easily purged. Obviously these strategies have also opposite consequences, advantages and disadvantages. The first example provides a safer way to avoid bubbles from entering the separation bed. For instance, Xu et al.[27] positioned electrodes into reservoirs isolated from a separation bed via 108 “connection channels”, each 10 µm high, 5 µm wide, 5 mm long (fig. 5). However, due to fabrication constraints, the electrical resistance between the electrode reservoirs and the separation bed is high, thus decreasing the efficiency of electric field generation. Consequently, high potentials are required, posing higher risks to the user and resulting in increased Joule heating. Another serious issue is the possibility for local buffer inhomogeneities, like pH change, caused during operation which will perturb separation stability. In summary, developments using this strategy are conceptually simple but difficult to implement into a working device, especially when trying not to compromise the range of applications and time of operation. It is important to keep in mind that although miniaturization might be beneficial in many aspects, it often compromises the general applicability as offered by desktop systems, and therefore not suitable for application in a diverse experimental environment as in the lab. In the second case, the efficiency of the electric field introduction is maximized due to the high proximity of the electrodes to the separation bed. For instance Anciaux et al.[14] developed a 3D printed µFFE device in acrylonitrile butadiene styrene (ABS) having a 80 µm high separation bed sided by two 345 µm high channels in which 250 µm thick

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Pt wires were positioned as electrodes (fig. 6). The larger channels to the sides allow both for insertion of the relatively large electrode wires, and for easy bubble purging. However, it is important to note that the reduced hydraulic resistance at the sides of the separation bed is a path of least resistance for the sample solution as well. Hence, careful design of the geometry as well as the set of fluids’ viscosities and flowing conditions is fundamental to ensure that the buffer solution is focused at the desired position at the separation bed, and is stable throughout its length [2, 62]. Furthermore, fast deviating analytes might be lost easily into the side channels. Another interesting feature of this device is that a ∼27-fold lower EOF was observed when comparing identical devices fabricated in ABS vs glass. In a similar yet intermediate configuration, K¨ohler et al. [66] developed an injection molded cyclic olefin polymer (COP) three-level µFFE device (fig. 7). A 50 µm high separation bed was divided by a channel constriction from 300 µm high side channels at which electrodes were directly deposited. This configuration aimed at ensuring that bubbles never enter the separation bed, but at the same time it increased the pressure there, increasing the difficulty of solutions flowing there to exit into the side channels. Since the local constriction was relatively small compared to the long channels used in other publications [27], a high electric field introduction efficiency was possible (e.g. 90% [66]). Although it provideh.igher control of the flow, it still required care to avoid loss of separation buffers and sample to the side channels. This issue was addressed in the work by Agostino et al. [62], who demonstrated the fabrication of a milli-FFE device with relatively larger separation bed dimensions (200 µm in height). Channel openings at the separation bed sides, where Pt wire electrodes are positioned, were used as bubble chimneys. Although these facilitate removal of bubbles, it requires additional design developments to ensure flow uniformity. In opposition to the above examples, Agostino et al. [62] created longitudinal grooves parallel to the flow direction and achieved considerable correction of the flow uniformity and further operation validation at separating fluorescent analytes (fig. 8). As we observe, the compartmentalization is a fundamental issue in the development of µFFE devices in general: researchers are often confronted with decisions that result in opposite effects, i.e. high electric field setting efficiency, simplified flow control and general applicability seem to be parameters hard to integrate in a single device. Usually, an isolation of the electrodes and a separation bed is a crucial feature of µFFE devices, ideally posing an infinite hydraulic resistance, for easy adjustment of flow conditions, and null electric resistance, for maximized electric field introduction efficiency. Yet in another variation of a geometrical structure as barrier between electrodes and the separation bed, Song et al. [12] made use of packed microbeads modified with charged hydrogels. Packed microbeads are a simple way to increase the hydraulic resistance significantly. However, the large amount of inter-bead spaces might result in decreased electrical resistance. Furthermore, different beads, modifications and sizes are commercially available allowing for tunning the referred parameters. The authors showed an electric field introduction efficiency up to 52% (see tab. 2). The difficulty is to easily and efficiently pack these beads in microchannels that isolate the electrodes from the separation bed. Another geometrically similar strategy was developed by Fu

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Figure 5: Top view schematics of µFFE devices with “connection channels” dividing electrode reservoirs from a separation bed, designed for the application of isoelectric focusing (a) and zone electrophoretic separation mode (b). Reproduced from Ref. [27] with permission from the Royal Society of Chemistry.

Figure 6: Images of µFFE devices: a) glass, b) bottom and c) top views of a 3D printed ABS device. 1) buffer inlets, 2) sample inlet, 3) electrode connections, and 4) buffer outlets. Reprinted with permission from [14]. Copyright 2016 American Chemical Society.

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Figure 7: Cross-section schematics of an injection molded µFFE device, having a two-level channel configuration and direct electrode integration. Reproduced from Ref. [66] with permission from the Royal Society of Chemistry.

Figure 8: A milli-FFE device with platinum electrodes positioned at side chimneys, for easy bubble removal, and with grooves parallel to the flow direction to ensure uniformity. Top views of steadystate fluorescent analyte separation as a function of time. Reproduced from Ref. [62] with permission. Copyright Wiley-VCH.

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Figure 9: µFFE with packed silica microbeads as electrode-separation bed division. Left) photograph of the µFFE device. Right) microphotograph of the separation bed at the outlet region showing separated analytes. Reprinted with permission from [12]. Copyright 2013 American Chemical Society.

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et al. [15] who developed a µFFE device having a separation bed sided by 100 µm thick polydimethylsiloxane (PDMS)/carbon black composite membranes and gallium electrodes (fig. 10). Authors mentioned that no Faradaic reactions were observed for up to 160–200 V applied potentials. Although no data was discussed regarding the electric field introduction efficiency, experiments showed the separation of fluorescent dyes at an applied potential of 80 V, a value similar to others in literature (tab. 2). In connection to its seemingly simple fabrication procedures, this novel electrode configuration has potential to revolutionize the way µFFE devices are fabricated and actuated, attracting the attention by other working groups on the field. However, additional tests are required to validate its potential. Another issue that needs careful attention is the use of PDMS in the fabrication of the membrane which might not be compatible with other fabrication methods, and most importantly with largescale industrial production. A fundamentally different approach yet making use of a physical barrier was reported by Janasek et al.[58]. In their device, a solid glass membrane acting as a dielectric barrier insulated the separation bed from the electrodes. The underlying principle of the field introduction is the electrostatic induction based on charge displacement caused by dipole orientation in the glass. Since the efficiency of the induction is determined by the material specific relative permittivity r , the use of ceramics with r >1000 as presented by Couceiro et al.[77] would be preferable to glass and polymers with r values of ≈8 and 2–4, respectively. The challenge for this approach is the fabrication of thin, tightly bonded, crack-free and non-porous barrier membranes as the principle only works under these conditions which is extremely difficult to fabricate and stopped the Janasek group from pursuing that path.

4.3. Gels as Salt Bridges The use of gel barriers or bridges in µFFE devices was first introduced by Kohlheyer et al.[11]. Agarose or SDS-PAGE gels are widely used in electrophoresis and several properties are well characterized, such as porosity. The rationale of using a gel bridge is the same as presented before: create a stable barrier to flow between electrodes and the

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Figure 10: µFFE with and without PDMS/carbon black composite membranes. A, B and C are schematic top views of devices having narrow lateral channels, no channels, and PDMS/carbon black membrane to bridge gallium electrodes with a separation bed, respectively. The bottom images are microphotographs during operation of the corresponding device configurations above. The use of PDMS/carbon black membranes enables proper operation without the formation of bubbles. Reproduced from Ref. [15] with permission. Copyright Wiley-VCH.

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separation bed, while its tunable porosity could be adjusted to increase the efficiency of electric field introduction. In opposition to the purely geometrical modifications, the fabrication of devices with gel bridges requires additional procedures such as surface modification and local photopolymerization. The mechanical gel stability inside the microfluidic device is dependent of its bulk stiffness, and importantly to the grafting strength. Kohlheyer et al.[11] functionalized glass chips with a methacrylate ended silane which were then used to graft an acrylamide gel by photopolymerization (fig. 11). Several experiments were demonstrated successfully, but as mentioned in that publication the gel broke apart after several hours of working. Gels are very fragile and the shear stresses felt in microfluidic devices are substantially higher than those in bulk devices. Associated to the lack of detailed information regarding the gel mechanical stability in microfluidics, namely bulk resistance before breakdown and delamination pressure, the use of gels requires extensive trial and error experiments during device development. Another issue to be taken into account is the gel’s sensitivity to atmospheric conditions requiring careful storage methods for increased shelf time, which may ultimately compromise the use of such strategy due to the substantial device fabrication complexity as compared to a traditional gel casting procedure. Several other works using gels as salt bridges are available in literature [1, 13, 15, 29, 31, 33]. Novelties such as gel functionalization have been performed by Albrecht et al.[29], Wen et al.[33], Song et al.[12] and Cheng et al.[1]. The introduced modifications aim at charging the gel, conferring a given pKa to it, a well-known technique in IEF. Albrecht et al.[29] were the first to make use of modified gel bridges in a µFFE device

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Figure 11: Using gels as salt bridges in a glass µFFE device. a) are photographs of the glass device with and without gel bridges. b) microphotograph showing one of the gel bridges in detail. c) SEM image from a cut chip at the gel front. Reproduced from Ref. [11] with permission from the Royal Society of Chemistry.

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Figure 12: µFFE devices with a single inlet and modified salt gel bridges. Comparing the effect of geometry on the formation of a stable pH gradient throughout the device. A) and C) are top view schematics of the devices used. B) and D) are photographs of the corresponding devices during IEF of methylred and bromothymol blue. Reproduced from Ref. [32] with permission. Copyright Wiley-VCH.

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to perform IEF of pI markers. Adding to the concept of using a charged gel bridge, Wen et al.[32, 33] developed a triangular µFFE device, unique in the field. Although ampholytes, used typically in IEF, are highly mobile even at low electric fields strengths, the time to achieve a stable pH gradient might still be considerably high compared to the flow transit time. Using a triangular shape promoted stable pH gradients shortly after the inlet position (fig. 12). In the previous example the pH gradient is arbitrarily confined within a given interval by the pKa values of the modified gels. Cheng et al.[1, 78] reported on a strategy to generate and control pH gradients dynamically. The apparatus developed consists of two devices connected in series: a first for generation of high and low pH values from a neutral solution, and a second where FFE takes place. In the first stage, two bipolar membranes are connected in series and sandwich a common channel tied to ground, through which the neutral solution is supplied and then connected to the second device. Buffer containers placed to the extremities of the membranes are used to close the circuit. This configuration allowed the application of asymmetric potential values increasing dramatically the range of pH gradients realizable. The generated pH gradient is then supplied to the second device at which a sample mixture is injected for separation.

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Figure 13: Dynamic pH gradient generation, based on the use of bipolar membranes, and direct coupling to a µFFE device for IEF separations. a) photographah of the integrated device. b) and c) are schematics of the integrated device when no potential is applied and vice-versa, for ZE or IEF separation modes, respectively. d) schematics of the bipolar membrane arrangement and generation of H+ and OHions when potential is applied. Reproduced from Ref. [1] with permission from the Royal Society of Chemistry.

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The µFFE device also contains polarized gel bridges in a configuration similar to that presented above. The authors demonstrated the separation of pI markers in IEF mode, and of ovalbumin-Alexa 594 and avidin-Alexa 488 in ZE mode. Indeed, the novelty of this work might revolutionize the range of applications a single device can achieve. However, fabrication complexity, a common issue in microfluidics devices in general, is especially challenging. Furthermore, true control needs to be achieved by means of feedback systems for precise formation of pH gradients. The biggest challenge to achieve this is the development and integration of pH sensors that are stable and with high longevity.

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4.4. Integration of Commercial Membranes More recently, the use of commercial membranes has been introduced [17, 59, 64]. The great advantage of commercial membranes is that they are readily available in a series of materials, dimensions, pore sizes, wettability, among others, circumventing issues related to challenging fabrication procedures and prone to large batch-to-batch variations. Although these might seem difficult to integrate in µFFE devices, in fact they are highly suitable for industrial manufacturing as compared to gel salt bridges. For instance, membranes may be easily integrated with other materials by lamination [64] or ultrasonication [59, 79]. Herzog et al.[64] demonstrated the use of PTFE membranes in a µFFE device (fig. 14) using a different configuration: electrodes were connected externally (similar to a direct integration) and the membrane spaned the entire top of device’s footprint. The idea was that bubbles generated at any portion of the device could evacuate through the membrane which was stabilized with adhesive transfer tape and polyethylene terephthalate (PET) foil to avoid bulging and consequent changes in flow profiles. This configuration and device fabrication was achieved by laser cutting and

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Figure 14: a) and b) are schematic cross sections of the laminated device having a PTFE membrane without and with a stabilization layer, respectively. c) image sequence of a gas bubble in the separation area removed through the PTFE membrane. Bottom) top view photograph of the µFFE device having three inlets and three outlets. The remaining ports are for external electrode plugging. The white ring corresponds to the laminated membrane. Reproduced from Ref. [64] with permission of Springer.

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lamination, hence enabling fabrication transfer to an industrial setting. The authors demonstrated the separation of several fluorescent dyes at an electric field of 500 V/cm and tested stability to values up to 1038 V/cm. Stability was compared between devices with and without membrane. For instance, a device with membrane was operated continuously at electric field of 500 V/cm for at least 180 min vs 7 min for a device without membrane. However, the spatial source of bubbles (should occur only at the electrodes assuming a temperature controlled device) and analysis of the electric field introduction are difficult due to the lack of detailed information regarding electrode integration. The use of external electrodes is beneficial both at the fabrication level and for recycling reasons. For instance, the use of microfabricated gold electrodes might be unnecessarily expensive and labor intensive. In contrast, the use of platinum electrodes is a comparatively cost-effective solution (especially through re-use) and commercially available. Novo et al.[17] developed a µFFE device employing polycarbonate membranes to isolate electrode reservoirs from the separation bed. The membrane was effective at preventing bubbles, generated at the platinum wire electrodes in the reservoirs, from entering into the separation bed (fig. 15). Furthermore, only negligible increase of the electric resistance by the use of the membranes was observed, hence allowing for highly efficient electric field introduction (∼100%). Separations of fluorescent dyes and protein pI markers were demonstrated in several operation modes indicating a large range of applications. However, the device was fabricated in PDMS and the membranes were sealed using an intermediate silanization step, features incompatible with industrialization. Nevertheless, the same device design and membranes used might be realizable with other materials such as thermoplastics, and techniques such as lamination and ultrasonication, to bridge with industrial fabrication.

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In most works in µFFE and microfluidics literature in general, material selection and fabrication methods [80] are often determined by the technologies available to the researchers who are typically focused on demonstrating functional aspects and technology potentials. However, the success of microfluidic technologies is highly dependent on the suitability to transfer the production from the lab to the industry, and the usability from the expert to the average user. Furthermore, the quest for industrial interest and investment will only be successful when a considerable value increase by using microfluidics as compared to production costs is demonstrated, a topic that has not yet been thoroughly discussed in the literature. However, there has been great progress to bridge the potential from research in the field and the industry, especially promoted by young companies often established by young entrepreneurs who spin-off from research. In this section the materials used and fabrication techniques employed on the production of µFFE devices over the last two decades will be presented and divided into two sections: a first for the chip body and a second for the electrodes used. These can be easily compared in tab. 2. 5.1. Materials for Chips Several materials have been employed in the fabrication of µFFE devices: silicon [16], amorphous silicon [57, 81], glass [11, 13, 16, 25, 28, 31, 34, 57, 60, 61, 81], quartz [35], SU8 [28], PDMS [10, 12, 15, 17, 23, 27, 32, 33, 63, 75, 82], OEG-DA (oligoethyleneglycol diacrylate) [34, 36], Norland Optical Adhesive 81 (NOA 81) [52], polyuretane [1],

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Figure 16: Effects of electroosmotic flow in the separation of methyl red in a triangular µFFE device. A) no dynamic coating used. B) using 4% PVA. Adapted with permission from [33]. Copyright 2010 American Chemical Society.

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polyimide [83], polymethylmethacrylate (PMMA) [2, 62, 83], ABS [14], COP [65, 66], polystyrene (PS) [59] and PET [64]. The materials used in µFFE chip fabrication and their trends are analogous to the developments in microfluidics in general. In the 1990s and up to early 2000s, silicon and glass were the materials of selection backed by the well established micromachining technologies such as wet etching and lithography. Raymond et al.[16] was the first to make use of silicon and glass for the fabrication of a µFFE device. Although very precise structures can be fabricated, methods are highly time and cost consuming, involve chemicals that pose serious health risks such as HF or KOH, and in the particular case of silicon require for oxide passivation to avoid short circuits. Silicon was abandoned through time, but glass is still the material of choice in recent publications (e.g. [13, 25]). With the introduction of soft-lithography of PDMS [84], a rapid growth in the field of microfluidics and also in the particular case of µFFE was observed, with the first steps being demonstrated by Andreas Manz’s group [10, 27]. The reason is that development cycles were considerably faster being possible to achieve from design to working device in a single day. This is extremely advantageous in a research environment, but alternatives that proportionate the same results should be kept in mind. Examples of features that need to be carefully weighed when migrating the fabrication to other materials are the non-specific molecule adsorption and surface zeta potentials [85]. Higher zeta potentials lead to increasing EOF which hinders or even compromise separation. Besides the possibility for EOF minimization, such as by the use of cellulose, Triton X-100 [52] and polyvinyl alcohol (PVA) (fig. 16), it is advantageous to select materials with low zeta potentials [14]. Thermoplastics and thermosets as well as techniques such as hot embossing and injection molding are some of the most successful examples in industry. However, high master mold prices and long fabrication times, often for the fabrication of a single device at an intermediate development stage, are prohibitive in a typical research setting. Although we critically address the issue of most literature in microfluidic devices, namely in µFFE, regarding bridging the technology to the industry, it is important to note that it is still possible to develop devices and their operation using readily available techniques while ensuring that they are in fact portable. For instance, integration of microvalves is well-established in PDMS microfluidic devices, but may be impracticable

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5.2. Materials for Electrodes The rule on electrode material for µFFE devices is the use of noble metals such as platinum, gold and silver. Most common electrodes used are made of platinum (tab. 2) as it is highly inert and available commercially in several sizes and shapes. Although a good integration of, for instance, platinum wire in µFFE devices can be achieved, higher degree of precision can only be obtained by microfabrication techniques and direct deposition of thin-film electrodes inside the chip. These include platinum [73] and gold [28, 57, 65, 66], the latter being the most popular (see tab. 2). A good gold adhesion to most materials, namely glass, requires for an intermediate layer of other materials such as chromium or titanium. These, on the other hand, are not stable in FFE and its contact with the solution should be avoided, which may occur in films lacking of homogeneity (pinholes) or at the sides of unpassivated film stacks. Ultimately, such thin-films will limit the operation longevity of the device. In the literature, testing the longevity of the electrodes in µFFE devices is not usual. Therefore, critically addressing this subject is not possible. It is, however, important to note that works referring long continuous operation times (3 h [64], 12 h [62]) used platinum wires. Examples making use of silver [90, 91] are very limited, most probably due to difficulties to fabricate and maintain the stability of the electrodes throughout operation. However, with the rapid development of silver ink screen printing technologies [92, 93], the use of silver, when properly conditioned to ensure stability, is a potential cost-effective solution for the direct deposition of electrodes in µFFE devices. The exception to the rule is the use of conductive polymers [14, 59] and liquid metals [14, 58, 63]. These offer higher fabrication flexibility as they can be molded to any desired shape. For instance, Stone et al.[59] used carbon fiber loaded polystyrene and injection molding to fabricate electrodes, enabling even the fabrication of 2.5D structures. Janasek et al. [58] used Woods alloy. Fu et al.[15] used carbon black composite PDMS membranes and liquid gallium, and Herling et al.[63] used an InBiSn alloy liquidized at 79 °C. Since these are casted in the liquid state, they require additional structures to be implemented in the microfluidic channels. However, when having appropriate fluidic structures, the plasticity offered by these electrodes allows for the fabrication of 3D electrodes. Furthermore, no bubble formation was observed at the PDMS/carbon black membrane interface which simplifies the design/fabrication of arbitrary channel configurations, since no special requirements regarding bubble evacuation are necessary.

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Figure 17: Scheme and photograph of the used µFFE chip as well as the customized fluorescence imaging system. Reproduced from Ref. [97] with permission from The Royal Society of Chemistry.

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6.1. Detection Usually, microfluidic devices are mounted on microscopes for simple, real-time monitoring of µFFE experiments. Furthermore, fluorescence microscopy is widely ready to use in laboratories. Thus, the vast selection of commercially available fluorophores, having different charge states and fluorescence properties, offers a simple method to test devices in proof-of-principle experiments. More elaborate experiments using naturally non-fluorescent analytes can still be performed using the same conceptual setups and by labeling the analytes of interest with selected fluorophores [1, 10–16, 23, 24, 27, 29, 31, 33, 34, 40, 51, 57, 60, 73, 82]. Analysis of the recorded results, either as time-lapsed images or full videos, can be performed using freely available softwares such as FIJI [94] and Inkscape [95]. Another software that might be useful in analyzing video data is Tracker [96]. Kochmann et al.[97] presented a cost-effective alternative to the use of sophisticated microscopes, with self-built fluorescence imaging and including a comprehensive customizable software suite for image processing and analysis of the performance of µFFE chips. The portable imaging setup consisted of a micropipette tip box as a lightproof case, a 470 nm LED illumination source, optical filters, a camera objective and electronics (fig. 17). The software suite provides tools for automated identification of chip features, extraction and analysis of stream trajectories, the evaluation of flow profiles and separation quality. Efforts were employed at rendering the publicly made available hardware and software as modular, flexible and adaptable as possible to enable extension or substitution. The use of a custom built high speed deep UV laser scanner for the detection of unlabeled analytes and native proteins by space-resolved intrinsic deep UV fluorescence scanning was demonstrated by K¨ohler et al.[98] The challenge in this approach was the necessity to develop suitable non-fluorescent and UV transparent fused silica FFE chips. Poehler et al.[35] employed a fluorescence microscope equipped with a 660 nm LED as the light source for a pH sensitive near-infrared fluorescence probe used as an integrated

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6.2. Analytes Conventional bench-top devices have been demonstrated to be applicable to the separation and fractionation of a wide range of samples and analytes [19, 49, 100, 101]. As described in the section above, the easy fluorescence detection in microfluidic structures makes fluorescent dyes [10–16, 23, 40, 57, 73, 82] or fluorescently labeled proteins or peptides [1, 11, 12, 23, 24, 27, 29, 31, 33, 34, 40, 51, 60, 82] as the analytes of choice for proof-of-concept experiments. In more advanced devices the labeling was carried out on-chip [36] or label-free proteins were used [17, 35]. Additionally, mixtures of organo-chemicals [13, 25] or antibiotics [35, 36] have been separated. The fast determination of the electrophoretic mobility of mitochondria using µFFE and online laser induced fluoresecence (LIF) detection was demonstrated by Kostal et al.[81]. In their work, results obtained with the µFFE were in good quantitative agreement with those obtained by CE, performed for validation purposes. Lu et al.[28] employed their µFFE device to isoelectrophoretically separate and focus organelles directly from crude cell lysate and intact cells. Examples of separations comparable to those expected in a real-world application are still very limited [17, 28, 33]. As the technology has reached a substantial level of maturity, we assume that research groups devote more efforts in developing methods and protocols. This includes issues related with sample preparation, optimized BGE systems as well as ensuring compatibility with common post-processing methods. To this end, the development and establishment of methods/devices to interface the chip’s output are required. Examples in the literature are rare [17, 33, 102] and, as with many other features of microfluidics developments, lack standardization. The latter, although a difficult subject in the constantly changing environment of research, would greatly benefit the comparison and/or cooperation between R&D groups as well as improve portability to a true wide user community.

FFE is a technique mainly suited for preparative purposes as part of analytical workflows and systems. Hence, besides reproducibility, high separation resolution is a key feature. This is ultimately limited by the number of outlets and fractionation number. This means, even if high separation resolution is achieved within the separation bed, this result will only translate into the collected fractions if the number of outlets is at least equal or higher than the device’s analyte separation resolving power (commonly described as peak capacity). This feature applies both to standalone devices and to integrated systems such as µFFE-MS setups. For instance, commercially available desktop devices

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7.1. Preparative Outlets to the Macro-World Despite the vital importance of the number of outlets on the resulting fractionation, many of the reported µFFE devices lack of real fractionation interfaces and seem to omit its importance. Instead, most of the initial devices concentrate uniquely in internal analytical aspects by performing proof-of-concept experiments. For instance, the chip devices of Raymond et al.[16], Zhang et al.[10], Xu et al.[27] and de Jesus et al.[104] had 125, 72 and 43 outlet channels, respectively, which however did not connect to the macro-world but were unified in one single outlet reservoir/hole for simplicity reasons. Others had only reduced number of outlets: two [12, 74], three [11] and five outlets [31, 34, 36, 51, 64], respectively. 24 outlet reservoirs were created by Wen et al.[33] as they punched holes in the outlet channels of their PDMS device and fitted capillary pipette tips into them. The challenge for the construction of interface arrays lies in maintaining equality of hydrodynamic resistance of all outlets of the µFFE device in order to not compromise the parallel laminar flow of the BGE and sample through the separation bed, and the space needed for tubes and sealings for preparative micro-to-macro-world interfaces. The compliance of these premises often results in interfaces with massive footprints. An example is shown in fig. 18 where a 9- and a 64-outlet port interface is displayed. Due to the outer diameter of the connected tubes and the sealing by O-rings (ca. 2 mm) (see fig. 18 B2 ) the area of the interface of the 64-outlet port is 25-fold larger than the separation bed itself (fig. 18 B1 ) [60, 105]. The equality of hydrodynamic resistance was achieved by equal-distant channels connecting the outlet side of the separation bed and the connection point to the tubes. The very long outlet channels on-chip and the additional length of the tube connections to the fraction sampler result in long analyte residence times and pose massive hydrodynamic resistance. This causes high mechanical stress to the structures isolating the separation bed from the electrode reservoirs, which is a typically fragile region (see section 4). One way to overcome this high-pressure drop problem is demonstrated with the device developed by Kobayashi et al.[102]. The µFFE chip was attached to a micromodule fraction separator (MFS) consisting of 19 capillaries and tubes connected to a multichannel peristaltic pump in order to draw the liquid from the device. At the inlet side, buffer and sample were pumped into the separation bed. Thus, by balancing the push and pull actions of the two pump systems the pressure drop to each system can be minimized. However, it is a very complex setup. In order to estimate the separation efficiency of that device, cytochrome c and myoglobin were separated in FF-ZE mode, and

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the liquid of each outlet tube was collected and analyzed offline by means of RP-HPLC [102]. Novo et al.[17], developed an additional microfluidic device connected to the µFFE by tubing, which ensured homogeneous flow distribution through each of the 9 outlets and was directly coupled to a 96-well plate. Although this novel interface guaranteed high flow stability as well as a simple method to interface with the well plate, it requires for the manual tubing between microfluidic devices—a task that is not user-friendly and not compatible with scalability perspectives, i.e. a high number of outlets requires labor-intensive individual tubing. This issue reinforces the need for the development of commercial chip-world interfaces. It is noteworthy to mention that although some companies specialized in microfluidics already provide some products for chip-world interface, these are still not suitable for µFFE. Ready to use, plugin like modules would be of the high interest and could possibly be based on simple ideas such as the use of pipette tips by Wen et al.[33].

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7.2. Hyphenation Besides the preparative interfaces for fractionating and collecting, and the offline coupling to other analytical methods like RP-HPLC[102], also online hyphenation of µFFE devices to mass spectrometric (MS) detection have been reported [13, 25, 74, 106]. Chartogne et al.[106] interconnected a µFFE device with capillary isoelectric focusing (CIEF) and electrospray ionization (ESI)-MS in order to online remove the ampholyte BGE. The system was optimized in the way that the purified analytes exited the µFFE chip via the central outlet which in turn was connected to a coaxial sheath flow ESI configuration. The feasibility of the system was demonstrated for the three model proteins myoglobin, carbonic anhydrase I and β-lactoglobulin B. Kinde et al.[74] applied a µFF-ZE in order to extract low molecular weight cationic analytes, namely human angiotensin I and the four-peptide MRFA, from a matrix containing sodium dodecyl sulphate (SDS) into an ESI-compatible solvent stream prior analyzing them mass spectrometrically via an ESI interface which was connected to one of the outlets of the chip. Using this approach, they were able to detect human angiotensin I and MRFA even at SDS concentrations in the original sample solution as high as 10 mM—concentrations which otherwise suppress the ESI-MS detection completely. Benz et al.[13] machined a monolithic emitter tip out of the used borosilicate microchip to generate an electrospray (fig. 19A1 ). The emitter tip was connected to the central outlet of the µFFE and intersected by a channel providing make-up flow, flow splitting and electrical contact for the ESI emitter. The separated zones were hydrodynamically directed towards the central outlet by linear variation of the volumetric flow rates of the flanking buffer streams (fig. 19A2 ), and thus the zones were detected by MS in a scanning manner (fig. 19A3 ). The system was evaluated by µFF-ZE of a mixture of three rhodamine dyes, and applied to study a multicomponent organic reaction with non-fluorescence educts, products and side-products. A similar approach was reported by Park et al.[25]. They connected the central outlet of their µFFE glass chip to an ESI emitter via a fused silica capillary (outlet 13 in fig. 19B1 ). By periodical change of the volume flow through the outermost inlets (inlets

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1 and 5), the isotachophoretic window was moved back and forth. Thus, the zones consecutively passed the outlet 13, exited the chip, were sprayed into the MS device and measured there. For proof-of-concept, a sample mixture of fluorescein (as model contaminant), Alexa Fluor 488, citric acid and glycolic acid were separated in µFF-ITP mode, hydrodynamically focused and measured by MS in a scanning manner (from LE to TE and back to LE - fig. 19B2 ). Over the time course of the experiment, selected ion isotachopherograms of glycolic acid, citric acid and fluorescein were recorded revealing that the model contaminant fluorescein was removed from the ITP window (fig. 19B3 ). 8. Conclusion

The miniaturization of free-flow electrophoresis (FFE) over the last decades has proven the scaling laws [37] and demonstrated the capability of the constructed devices, thus raising hopes to apply the technique of FFE to the challenges of personalized analytics where limited amount of sample is crucial. Current limitations as mentioned in this review impede the dissemination of µFFE. Examples of these limitations include is-

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sues related to selection of materials and fabrication methods which typically do not fit industrial manufacturing. Other issues relate to the lack of easy-to-use interfaces which could potentially benefit from future efforts on standardization. For instance, ready-to-use, plug-in like modules would be of high interest, however are not available yet. However, progress has been done in this direction as well as on simplification of device/setup fabrication[14, 64], and on interfacing to sample collection [33, 102] or other downstream analytical methods [13, 25]. Once the challenges are resolved, µFFE has the potential to be a consummate enhancement in both analytical and preparative processes in the micro-scaled world, since many aspects regarding the separation power as well as different modes for the separation of analytes from a broad range of sample sources have already been demonstrated. Acknowledgements

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This study was supported by the Ministerium f¨ ur Innovation, Wissenschaft und Forschung des Landes Nordrhein-Westfalen, the Regierende B¨ urgermeister von Berlin - inkl. Wissenschaft und Forschung, and the Bundesministerium f¨ ur Bildung und Forschung. References

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The review gives an overview on the field of microfluidic free-flow electrophoresis. The research field has developed vast amounts of devices, methods and applications.

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Wide-spread use of µFFE depends on simplicity of use and industrial fabrication.