Analytical and Bioanalytical Chemistry

, Volume 410, Issue 3, pp 853–862 | Cite as

Continuous purification of reaction products by micro free-flow electrophoresis enabled by large area deep-UV fluorescence imaging

  • Simon A. Pfeiffer
  • Benjamin M. Rudisch
  • Petra Glaeser
  • Matthias Spanka
  • Felix Nitschke
  • Andrea A. Robitzki
  • Christoph Schneider
  • Stefan Nagl
  • Detlev Belder
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Microreactors have gained increasing attention in their application toward continuous micro flow synthesis. An unsolved problem of continuous flow synthesis is the lack of techniques for continuous product purification. Herein, we present a micro free-flow electrophoresis device and accompanying setup that enables the continuous separation and purification of unlabeled organic synthesis products. The system is applied to the separation and purification of triarylmethanes. For imaging of the unlabeled analytes on-chip a novel setup for large area (3.6 cm2) deep ultra violet excitation fluorescence detection was developed. Suitable separation conditions based on low conductivity electrophoresis buffers were devised to purify the product. With the optimized conditions, starting materials and product of the synthesis were well separated (R > 1.2). The separation was found to be very stable with relative standard deviations of the peak positions smaller than 3.5% over 15 min. The stable conditions enabled collection of the separated compounds, and purity of the product fraction was confirmed using capillary electrophoresis and mass spectrometry. This result demonstrates the great potential of free-flow electrophoresis as a technique for product purification or continuous clean-up in flow synthesis.

Graphical Abstract

Micro free-flow electrophoresis (μFFE) allows continuous separation and purification of small organic synthesis products. Enabled by a novel deep-UV imaging setup starting materials and product of a recently developed synthesis for triarylmethanes could be purified. Thereby demonstrating the potential of μFFE as continuous purification technique for micro flow synthesis.


Continuous flow Free-flow separation Flow microreactor synthesis Ultraviolet fluorescence 


Micro flow synthesis has gained a lot of attention recently because of its many advantages in reaction optimization and synthesis automation [1, 2, 3, 4]. The main strength of microreactors are that many commonly performed reaction steps like dosing of reagents, heating, cooling, and mixing can be conveniently implemented in a continuous fashion. However, other crucial operations in multi-step synthesis are thus far not easily transferred to continuous flow. For example, intermediate solvent exchanges, purification of intermediates, and most importantly the purification of the product are often performed in a discontinuous off-chip manner.

Research into continuous purification systems has only taken off in recent years. Among others, the Jensen group has been developing multistep microreactor networks with intermediate phase separations [5] and distillation [6] devices. Other continuous work-up strategies are micro evaporation [7], spray drying [8], and liquid–liquid extraction [9].

Micro free-flow electrophoresis (μFFE) has been identified as a promising technique for purification of reaction products emerging from microreactors [10, 11]. Yet, only few publications have demonstrated work in this direction. Most research on μFFE is concerned with separating biologically relevant molecules like peptides or proteins [11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21]. We have previously shown the feasibility of integrating a microreactor with subsequent separation by free zone electrophoresis [10] and isoelectric focusing [22]. In those publications, fluorescent labeling of amino acids and peptides was performed and the reaction mixtures were continuously separated. However, recovery of the analytes after separation was not a focus in those reports.

Separation and purification of more conventional organic synthesis products has not been shown yet. This is a result of various technical challenges arising when the effluent of an organic synthesis is to be separated. Among others, the solubility of synthesis products, the abundance of neutral species, and compatibility issues of synthesis solvent and separation buffer can be challenging [11].

Furthermore, the observation of the separation, which is necessary for optimizing separations on the device, is challenging. The prevailing detection technique in μFFE is fluorescence imaging in the visible light (VIS) range [23, 24]. Obviously, this requires intrinsically fluorescent molecules or necessitates a labeling reaction. Unfortunately, the majority of synthesis products are not fluorescent in the VIS range nor is labeling of the products usually desired. Great progress has been made in recent years with regards to novel detection techniques for μFFE. Among others, detection systems were devised based on mass spectrometry [25, 26, 27, 28], deep ultra violet (deep-UV) fluorescence excitation [29, 30], surface enhanced Raman spectroscopy [31], cell-based signaling [32], and detection via saccharide specific fluorescent probes [33].

Despite this progress with regards to detection techniques, to date a continuous separation and purification of organic synthesis products using μFFE has not been shown. Herein, we present our most recent work with μFFE, including the first continuous separation and fractionation of unlabeled organic synthesis products. For on-line observation of the separation, a novel detection setup for large area deep-UV excited fluorescence detection was developed. The detection system was instrumental for optimization of the chip system and the separation conditions.



All substances were used as received without further purification. Polyethyleneglycol diacrylate (PEG-DA, MWavg 250, and 575 g*mol–1), 3-(trimethoxysilyl)propyl methacrylate (TPM), 2,2-dimethoxy-2-phenylacetophenone (DMPA), 1-napthol, and 3-(cyclohexylamino)-1-propanesulfonic acid (CAPS) were purchased from Sigma-Aldrich (Darmstadt, Germany). Chloroform, n-heptane, and acetonitrile (ACN) were purchased from VWR (Dresden, Germany). Na2CO3, NaHCO3, NaCl, H2SO4, and H2O2 were from Carl Roth (Karlsruhe, Germany). HPMC was from Ferak Laborat (Berlin, Germany), and Triton X-100 from Riedel de Haen (Seelze, Germany). The other synthesis materials as shown later were synthesized in the work group of Professor Schneider (University of Leipzig).

Chip system

μFFE chips were produced largely following our previously published procedures using liquid-phase lithography between two glass slides [34]. For the lid of the device, access holes for fluidic and electrical contacting were powder blasted (Barth, Königstein, Germany) in quartz microscopy glass slides (Science Services, Munich, Germany). The bottom quartz glass slide was sputtered with platinum to form electrodes for electrical contacting. Platinum deposition was performed in two steps: First, the slides were thoroughly cleaned using acetone, isopropanol, ultrapure water, and piranha solution (H2SO4, H2O2, 2:1). As a positive photo resist AR-P 3510 (ALLRESIST, Strausberg, Germany) was applied to the glass slide using a spin coater following the manufacturer’s recommendations. After a prebake of the substrate, a photomask was applied and exposed (18 s) using a mask aligner (MA-6; SÜSS MicroTec, Munich, Germany). The photoresist was developed using AR 300-26 developer agent (ALLRESIST). In the second step, the glass slides were transferred to a sputter coater (BAE 250, Bal-Tec). A layer of chromium (60 nm) and platinum (350 nm) were deposited in sequence. After stripping of the photoresist using acetone, the platinum electrodes remained on the glass slide.

Both the lid and the bottom glass slide were pretreated with 3-(trimethoxysilyl)propyl methacrylate (4% v/v) in a heptane:chloroform (4:1) solution for 5 min and rinsed with heptane and toluene. Adhesive copper foil (Conrad Electronic, Hirschau, Germany) was placed between the lid and the bottom glass slide to obtain a defined channel height of 50 μm. A solution of 1% (w/w) DMPA in PEG-DA (Mavg 250 g*mol–1) pipetted between the two glass slides and a photomask with the fluidic structure was aligned on top of the assembly. After UV exposure (1.3 s) using a flood illuminator (4″ FE5; SÜSS MicroTec) the unpolymerized oligomer was removed by aspiration through the access holes. The devices were thoroughly rinsed with ethanol to remove residual photopolymer and exposed two more times for a total 4 s using the flood source. A schematic outline of the finished μFFE chip is shown in Fig. 1a. The layout of the photomasks used throughout is shown in Fig. 1b. To ease electrical contact with the power supply, short sections of copper wire were soldered on the exposed platinum electrodes before further processing as shown in Fig. 1c.
Fig. 1

Overview of the fabricated chip systems. (a) Exploded view of the μFFE-chip and its layers. Channels are not to scale for clarity. (b) Photomask used for polymerization of the PEG layer. Channel width was 100 μm for the meandering inlet and 300 μm for all others. Scale bar 1 cm. (c) Photograph of an assembled chip showing the copper wires used for electrical contacting and tubing sections for fluidic contact of the electrode channels

To separate the electrode compartments from the separation bed, conductive hydrogel walls were polymerized inside the assembled chips following a previously published procedure [26]. Figure 2 shows the position of the hydrogel walls inside the chip. As the photopolymer, a mixture of 40% water and 60% (w/w) PEG-DA (Mavg 575 g*mol-1) was prepared and doped with 1% (w/w) DMPA. For polymerization, an inverted epi-fluorescence microscope was equipped with a UV-LED (M365L; Thorlabs, Dachau, Germany) and a 20× objective (NA 0.5, UplanFL N; Olympus, Hamburg, Germany). The chips were filled with the prepolymer and placed on the specimen stage. The LED was switched on and the chip slowly moved over the objective from end-to-end polymerizing the hydrogel wall. After rinsing with ethanol and water, the chips were stored under water until use to prevent drying of the hydrogel.
Fig. 2

Frontal view of the separation bed. Arrows highlight the different liquid flows and the respective inlets. Hydrogel walls separate the electrodes and the separation bed. Channel dimensions are not to scale for better visibility

μFFE device operation

The μFFE chip was connected to Nemesys syringe pumps (Cetoni, Korbussen, Germany) equipped with glass syringes from ILS (Stützerbach, Germany). All but the electrode channels were connected by capillary tubing (i.d. 150 μm, o.d. 360 μm; Machery-Nagel, Düren, Germany). The electrode channels were connected using Teflon tubing (i.d. 500 μm, o.d. 1.58 mm; ESKA, Hamburg, Germany). The analyte inlet was further connected to a HPLC injection valve (Knauer, Berlin, Germany) equipped with a 20 μL sample injection loop.

Figure 2 shows a frontal view of the separation bed and highlights the different inlets; namely, analyte, buffer, and electrode stream. The separation buffer consisted of 20 mM CAPS (pH 10.0) containing 0.1% HPMC. The electrode channels were flushed with the same CAPS buffer but supplemented with 1 M NaCl and 1 mM Triton X-100.

Optical setup

The deep-UV fluorescence detection setup was based on a confocal fluorescence lifetime microscope MicroTime200 (PicoQuant, Berlin, Germany). The system was additionally equipped with a deep-UV extension consisting of a deep-UV capable photo multiplier tube (PMA 165-N-M; PicoQuant) and a 40× quartz objective (Partec, Münster, Germany). As the excitation source, the fourth harmonic of a Nd:YVO4 laser (Cougar, Time-bandwidth) was coupled to the auxiliary light path. The beam was reflected by dichroic mirrors (266 RazorEdge, Semrock) and guided into the objective of the modified epifluorescence microscope (iX 71; Olympus). The fluorescence emission was collected with the same objective and guided through a quartz tube lens, a 100 μm confocal pin-hole, and an emission filter (short pass 532 nm, SP532-RS; Semrock) onto the photo multiplier. The laser and detection unit were synchronized using the time correlated single photon counting electronics provided by the MicroTime200. Data acquisition was performed using the accompanying SymphoTime32 software (PicoQuant). To enable large area scanning, the microscope was further fitted with a motorized specimen stage (TANGO 2 Desktop; Märzhäuser Wetzlar, Wetzlar, Germany), which was controlled by the accompanying computer software (SwitchBoard; Märzhäuser Wetzlar). A photograph of the optical setup and its components is provided in the Electronic Supplementary Material (Fig. S1).


The software solution translating fluorescence intensity over time to pseudo colored intensity images as described in the Results section was implemented in Python ( Libraries used for implementation of the necessary transformations were numpy (, scipy (, photon-tools (, PyQT4 (Riverbank Computing, Wimborne, UK), and plotly ( For more general data analysis and plotting, OriginPro 8.5 (OriginLab, Northampton, MA, USA) was used. pKa values of the different synthetic compounds were calculated using ChemDraw 15.0 Professional (Perkin Elmer, Hamburg, Germany).

Analytical methods

Capillary electrophoresis experiments were conducted on a commercial instrument (P/ACE MDQ; Beckman Coulter, Krefeld, Germany). The instrument was equipped with a fused silica capillary (i.d. 50 μm) with a total length of 60 cm (50 cm effective) and a diode array absorbance detector. Injections were performed by applying pressure (35 mbar) for 15 s to the sample vial. For separation, a potential of 20 kV was applied to the thermostatted capillary (25 °C). Samples from the reaction mixture were diluted in separation buffer prior to separation. Samples from the fractions collected at the outlets of the μFFE chip were injected carefully. The starting materials and reaction products of the triarylmethane synthesis were spectroscopically characterized using absorbance and fluorescence spectrometers (V-650 and FP-6200; Jasco, Groß-Umstadt, Germany).

Results and Discussion

To demonstrate and assess the capabilities of μFFE in continuous purification of reaction mixtures, we chose a recently devised synthesis of triarylmethanes as the target reaction [35]. Triarylmethanes are important lead structures in a variety of applications ranging from dye precursors to drug candidates, and various synthesis strategies have been pursued thus far [35]. The synthesis scheme for the Brønsted acid catalyzed Friedel-Crafts alkylation is shown in Fig. 3a. The reaction employs ortho-hydroxylbenzhydryl alcohols 1 and 1-naphthol (2), which under phosphoric acid catalysis yields triarylmethanes 3 via in situ-prepared ortho-quinone methides. High yields and a wide product scope have been demonstrated with this approach in the batch synthesis [35]. Herein, diphenyl phosphate (4) was employed as catalyst and racemic mixtures of triarylmethanes were analyzed.
Fig. 3

(a) Reaction scheme for the synthesis of triarylmethanes 3 from benzhydryl alcohol 1 and 1-naphthol (2) under acid catalysis (4). (b) Electropherogram of a reaction mixture allowed to react for 45 min in batch. Conditions: Absorbance detection at 250 nm, 20 mM carbonate buffer pH 10, 20 kV, i.d. 50 μm, 50 cm eff. length. (c) Emission spectra of the substances 1-3 under deep-UV excitation (266 nm, substances: 100 μM in acetonitrile). Diphenyl phosphate (4) does not fluoresce under these conditions

For purification of these products and separation from catalysts and starting materials, Saha et al. employed column chromatography [35]. For a successful transfer of the purification step to continuous free-flow (zone) electrophoresis the molecules should carry a charge or be ionizable in a suitable buffer. Therefore, the pKa values of the hydroxyl groups of the reaction partners (1-3) were calculated. The pKa values were in the range of 9–10. This suggested that deprotonation should occur at a buffer pH of about 10 or higher. To confirm the pKa values and to prototype possible buffers for μFFE separations, capillary electrophoresis (CE) experiments were conducted. A carbonate buffer (20 mM, pH 10) was selected as a starting point. Figure 3b shows the electropherogram of a reaction mixture containing 100 mM each of the starting materials and 5 mM of diphenyl phosphate catalyst after 45 min reaction time in a vial. All substances were found to migrate after the electroosmotic flow (EOF) and therefore carried negative charge as expected. The peak identity was confirmed using standards and the obtained migration time matched observed migration times of standards. Full baseline separation of the three compounds (1-3) was achieved in less than 7 min with this protocol (Fig. 3b). Only the acid catalyst (4) was not fully separated from 1-naphthol (2).

The analytes were then screened with regard to their spectroscopic behavior. Macroscopically, solutions of the compounds in acetonitrile were optically clear, suggesting no appreciable absorbance in the VIS range, which was confirmed by absorbance measurements (data not shown). The analytes only showed absorption below 350 nm with strong absorption maxima at or below 250 nm. The substances were further characterized with regard to their fluorescent properties and specifically with respect to deep-UV (<300 nm) excitation. Figure 3c shows the fluorescence emission spectra of the starting materials and one representative triarylmethane product. All species involved, with the exception of the diphenyl phosphate, displayed fluorescence emission in the region of 300–450 nm upon excitation with 266 nm.

Large area deep-UV fluorescence imaging

To visualize the separation on the μFFE device, we opted to employ deep UV fluorescence detection. It has been previously shown that deep-UV excitation fluorescence detection is a valuable tool to detect unlabeled analytes in microfluidic systems in general [36, 37] and to visualize μFFE separations in particular [29, 30]. However, these instrumental developments were not applicable for the current approach because they only enabled visualization of a small portion of the separation bed. Herein, we employed μFFE devices with a separation bed area of up to 3.6 cm2.

Therefore, to enable large area deep-UV excitation fluorescence imaging, a microscope was fitted with an electronic specimen stage. The specimen stage enabled computer controlled movement of the chip over the microscope’s objective. To obtain spatially resolved data over the whole separation bed, the chip was moved over the objective on a serpentine trajectory as depicted in Fig. 4a. By continuously collecting fluorescence intensity during the scan process, an intensity trace was acquired over the whole area (Fig. 4b). The data were then processed with a software developed in this study (Fig. 4c). The software translates from the intensity time trace (Fig. 4b) to a pseudo-colored intensity image of the separation bed as shown in Fig. 4d. This was possible because during the scan process the system also detects the autofluorescence originating from edges of the separation bed. This leads to distinct peaks in the time trace (marked red in Fig. 4b), which were used as markers for the end of the separation bed.
Fig. 4

Schematic outline over the image generation process using the devised scanning approach. (a) The chip is scanned over the objective on a serpentine trajectory using a computer controlled specimen stage. (b) Example of a time trace acquired during the scan. Asterisk denotes the sample peak. Peaks marked in red are edges of the separation chamber used for slicing the time trace into individual line scans. (c) Custom software is employed to slice the acquired data and generate images. (d) Pseudo-colored intensity image of a reaction mixture sample flowing through the separation bed

The resolution of the resulting images depends on the scanning speed of the electronic specimen stage, the number of lines scans along the bed, and the detection volume of the detector system. In a typical experiment, as shown herein, 20 scans across the separation bed were performed equally distributed from the inlets to the outlets of the chip (~8 lines per cm). Every line scan took about 2.5 s to complete; thus a complete image was acquired after 50 s. While this might be slow in comparison to directly imaging using, e.g., a CCD-camera [30], it is more than sufficient for μFFE separations, which ideally show a steady positioning of analyte bands. For comparison, the residence time in the separation shown below was 55 s. In essence, as long as the sample is continuously introduced and the residence time of the sample exceeds the acquisition time, the resulting images accurately reflect the band position in the separation bed. The distinct advantage of the developed setup is the large area that can be scanned and imaged. Previous deep UV imaging setups for μFFE were restricted to imaging of 0.12 cm2 [29] and 0.57 cm2 [30]. The newly devised approach is able to image the whole separation bed of devices herein, which covered an area of 3.6 cm2 (1.4 cm × 2.6 cm, W × L). Furthermore, the maximum translation of the electronic stage is even bigger and might allow imaging of large areas or multiple sites on larger integrated devices in the future. Additionally, because the underlying detector system of the microscope was a commercially available time correlated single photon counting system, fluorescence lifetimes of the analytes bands could be obtained as well.

μFFE separation and purification

The starting point for optimizing the μFFE separation conditions was the carbonate buffer (20 mM, pH 10) employed in CE measurements as shown in Fig. 3b. While this buffer enabled electrophoretic baseline separations in CE using 50 μm i.d. capillaries, it proved to be less suitable in μFFE. At higher electric fields, the μFFE separations were poorly reproducible due to excessive joule heating. Therefore, lower conductivity buffers were investigated as separation media. Specifically, CAPS (10 mM, pH 10) was found to be a viable alternative. The prepared CAPS buffered had a conductivity of 0.6 mS*cm–1 (Carbonate buffer 2.7 mS*cm–1) and performed otherwise similarly in CE separations. It was found that supplementing the separation buffer with HPMC (0.1% w/w) to suppress residual EOF helped to stabilize the separation conditions. The separation electrolyte with the high pH did not damage the PEG walls of the microfluidic system.

Figure 5a shows the successful separation of the analytes (1-3) on the μFFE platform using the low conductivity buffer. The separation potential applied was 66 V (47 V*cm–1) at an electrophoretic current of 700 μA and sample residence time of 55s (0.5 mm*s–1). The resolution between the benzhydryl alcohol 1 and the synthesis product 3 was 1.4 (R 1-3 ) and between the product and 1-naphthol 1.2 (R 2-3 ). Peak identity was confirmed by individually injecting the analytes and recording the position in the separation bed. The acid catalysts show no fluorescence upon deep-UV excitation and were therefore not detected.
Fig. 5

μFFE separation of a reaction mixture containing analytes 1-3. (a) Digital render of the chip with correct layout (channels not to scale). Crosses denoted sealed channels. Inset: Pseudo colored fluorescence intensity image of the separation bed during separation. (b) Intensity cross section close to the outlets of the separation bed (red marker in a). Peaks were fitted using Gaussian functions. Conditions: analytes 1–3, 1 mM in CAPS (10 mM, pH 10, 10% ACN). Separation buffer: CAPS (10 mM, pH 10, 0.1% HPMC). Sample residence time: 55 s (0.5 mm*s-1). Separation potential: 66 V (700 μA)

It was found that the electrophoretic separation was very stable and only minimal fluctuations occurred. Figure 6 shows consecutive scans across the separation bed near the outlet over a period of 15 min. The peak positions showed excellent stability with relative standard deviation less than 3.5% (n = 20, consecutive scans) demonstrating the good stability of the band positions during separation. We did not find significant differences in band position between separations and thus did not investigate this in detail. Similarly, we have not yet investigated the reproducibility between individual chips in detail.
Fig. 6

Intensity cross-sections of the separation bed close to the outlet recorded for 15 min. Conditions and peak identity as in Fig. 5

The developed electrolyte system was then employed to separate a mixture of analytes resembling a reaction mixture. The sample contained 1 mM each of the analytes (1-3) in a buffer acetonitrile mixture (10 mM CAPS pH 10, 10% ACN). A 20 μL sample was injected and the effluents of the different outlets of the separation bed were collected. Over a period of 20 min, fractions of about 70 μL were obtained from each individual outlet. The separation was monitored simultaneously to ensure stable band positioning in front of the outlets.

In order to confirm the successful preparative separation of the products with μFFE, the collected samples were then analyzed using CE. Figure 7 shows the electropherograms obtained from the fractions 1-4 (FR 1-4, anode to cathode). It was found that the fractions FR 1-4 contained 1-naphthol (2), triarylmethane 3, benzhydryl alcohol 1, and only buffer. In every fraction only one component could be detected using the qualitative CE-method. The migration times of the peaks present in the data were consistent with separations of standards. Additionally, the individual fractions were spiked with the respective compounds to confirm peak identity (Fig. 7 blue traces). To further confirm the purity of the fraction containing the triarylmethane product, direct infusion electrospray ionization mass spectrometry was performed. In the mass spectra the only sample that showed the ion for the triarylmethane derivative (355 m/z, [M–H]) was FR 2 (see ESM, Fig. S2).
Fig. 7

Electropherograms of the fraction (FR 1-4, order: anode to cathode) collected from the μFFE chip after separation of the components. Analyte peaks are marked with an asterisk. Black traces are the pure fractions. Blue traces are spiked aliquots of the fraction with: FR1: 1-Naphthol (2); FR2: Triarylmethane product (3); FR3: Benzyhydryl alcohol 1. CE conditions:10 mM CAPS buffer (pH 10.0). Otherwise conditions as in Figure 3c


It could be shown that μFFE is capable of separating and resolving native unlabeled synthesis products. This was demonstrated for a synthesis of triarylmethanes as a target reaction. To enable observation of the separation, a novel deep-UV excitation fluorescence detection system was developed. In contrast to previous deep-UV imaging approaches, the developed system enabled large area imaging of the separation bed. The chip system and separation conditions were optimized to separate the components of a synthesis mixture. It was not only possible to achieve separation but also to collect the separated bands from an artificial sample. The purity of the collected fractions was confirmed using capillary electrophoresis and mass spectrometry.

To the best of our knowledge, this is the first demonstration of the separation of small synthetic molecules separated and individually collected using μFFE. The next step could be the direct connection of a micro flow reactor and μFFE to perform synthesis and separation on a single chip. The presented approach toward product purification is currently limited to synthesis products that show appreciable fluorescence under deep-UV excitation, which is however, more broadly applicable then, e.g., detection in the VIS region. While we used a sophisticated research laser microscope in the present set-up, the dramatic developments in photonic technology such as deep-UV LEDs and image sensors should allow realizing more economic detection set-ups in the future, thereby facilitating more widespread use of the approach. Especially for further automation in micro flow synthesis where continuous separation techniques are needed, we believe that the application of μFFE has a great potential to supplement the chemist’s toolbox.



The authors gratefully acknowledge funding by the Deutsche Forschungsgemeinschaft (DFG) through grant FOR 2177.

Compliance with ethical standards

Conflict of interest

The authors declare no conflict of interest.

Supplementary material

216_2017_697_MOESM1_ESM.pdf (282 kb)
ESM 1 (PDF 281 kb)


  1. 1.
    Jensen KF. Microreaction engineering — is small better? Chem Eng Sci. 2001;56:293–303.CrossRefGoogle Scholar
  2. 2.
    Jähnisch K, Hessel V, Löwe H, Baerns M. Chemistry in Microstructured Reactors. Angew Chem Int Ed. 2004;43:406–46.CrossRefGoogle Scholar
  3. 3.
    McMullen JP, Jensen KF. Integrated microreactors for reaction automation: new approaches to reaction development. Annu Rev Anal Chem. 2010;3:19–42.CrossRefGoogle Scholar
  4. 4.
    Plutschack MB, Pieber B, Gilmore K, Seeberger PH. The Hitchhiker’s Guide to Flow Chemistry. Chem Rev. 2017;
  5. 5.
    Sahoo HR, Kralj JG, Jensen KF. Multistep Continuous-flow microchemical synthesis involving multiple reactions and separations. Angew Chem. 2007;119:5806–10.CrossRefGoogle Scholar
  6. 6.
    Hartman RL, Naber JR, Buchwald SL, Jensen KF. Multistep microchemical synthesis enabled by microfluidic distillation. Angew Chem Int Ed. 2010;49:899–903.CrossRefGoogle Scholar
  7. 7.
    Timmer BH, van Delft KM, Olthuis W, Bergveld P, van den Berg A. Micro-evaporation electrolyte concentrator. Sensors Actuators B Chem. 2003;91:342–6.CrossRefGoogle Scholar
  8. 8.
    Ley SV, Fitzpatrick DE, Ingham RJ, Myers RM. Organic synthesis: march of the machines. Angew Chem Int Ed. 2015;54:3449–64.CrossRefGoogle Scholar
  9. 9.
    Kralj JG, Sahoo HR, Jensen KF. Integrated continuous microfluidic liquid–liquid extraction. Lab Chip. 2007;7:256–63.CrossRefGoogle Scholar
  10. 10.
    Jezierski S, Tehsmer V, Nagl S, Belder D. Integrating continuous microflow reactions with subsequent micropreparative separations on a single microfluidic chip. Chem Commun. 2013;49:11644–6.CrossRefGoogle Scholar
  11. 11.
    Agostino FJ, Krylov SN. Advances in steady-state continuous-flow purification by small-scale free-flow electrophoresis. TrAC Trends Anal Chem. 2015;72:68–79.CrossRefGoogle Scholar
  12. 12.
    Turgeon RT, Bowser MT. Micro free-flow electrophoresis: theory and applications. Anal Bioanal Chem. 2009;394:187–98.CrossRefGoogle Scholar
  13. 13.
    Köhler S, Weilbeer C, Howitz S, Becker H, Beushausen V, Belder D. PDMS free-flow electrophoresis chips with integrated partitioning bars for bubble segregation. Lab Chip. 2011;11:309–14.CrossRefGoogle Scholar
  14. 14.
    Köhler S, Benz C, Becker H, Beckert E, Beushausen V, Belder D. Micro free-flow electrophoresis with injection molded chips. RSC Adv. 2012;2:520–5.CrossRefGoogle Scholar
  15. 15.
    Geiger M, Frost NW, Bowser MT. Comprehensive multidimensional separations of peptides using nano-liquid chromatography coupled with micro free-flow electrophoresis. Anal Chem. 2014;86:5136–42.CrossRefGoogle Scholar
  16. 16.
    Herzog C, Beckert E, Nagl S. Rapid isoelectric point determination in a miniaturized preparative separation using jet-dispensed optical pH sensors and micro free-flow electrophoresis. Anal Chem. 2014;86:9533–9.CrossRefGoogle Scholar
  17. 17.
    Geiger M, Harstad RK, Bowser MT. Effect of surface adsorption on temporal and spatial broadening in micro free-flow electrophoresis. Anal Chem. 2015;87:11682–90.CrossRefGoogle Scholar
  18. 18.
    Poehler E, Herzog C, Suendermann M, Pfeiffer SA, Nagl S. Development of microscopic time-domain dual lifetime referencing luminescence detection for pH monitoring in microfluidic free-flow isoelectric focusing. Eng Life Sci. 2015;15:276–85.CrossRefGoogle Scholar
  19. 19.
    Anciaux SK, Geiger M, Bowser MT. 3D printed micro free-flow electrophoresis device. Anal Chem. 2016;88:7675–82.CrossRefGoogle Scholar
  20. 20.
    Novo P, Jender M, Dell’Aica M, Zahedi RP, Janasek D. Free-flow electrophoresis separation of proteins and DNA using microfluidics and polycarbonate membranes. Procedia Eng. 2016;168:1382–5.CrossRefGoogle Scholar
  21. 21.
    Johnson AC, Bowser MT. High-speed, comprehensive, two dimensional separations of peptides and small molecule biological amines using capillary electrophoresis coupled with micro free-flow electrophoresis. Anal Chem. 2017;89:1665–73.CrossRefGoogle Scholar
  22. 22.
    Herzog C, Poehler E, Peretzki AJ, Borisov SM, Aigner D, Mayr T, Nagl S. Continuous on-chip fluorescence labelling, free-flow isoelectric focusing and marker-free isoelectric point determination of proteins and peptides. Lab Chip. 2016;16:1565–72.Google Scholar
  23. 23.
    Kochmann S, Krylov S. Image processing and analysis system for development and use of free-flow electrophoresis chips. Lab Chip. 2017;17:256–66.CrossRefGoogle Scholar
  24. 24.
    Novo P, Janasek D. Current advances and challenges in microfluidic free-flow electrophoresis – a critical review. Anal Chim Acta. 2017;
  25. 25.
    Chartogne A, Tjaden UR, Van der Greef J. A free-flow electrophoresis chip device for interfacing capillary isoelectric focusing on-line with electrospray mass spectrometry. Rapid Commun Mass Spectrom. 2000;14:1269–74.CrossRefGoogle Scholar
  26. 26.
    Benz C, Boomhoff M, Appun J, Schneider C, Belder D. Chip-based free-flow electrophoresis with integrated nanospray mass-spectrometry. Angew Chem Int Ed. 2015;54:2766–70.CrossRefGoogle Scholar
  27. 27.
    Park JK, Campos CDM, Neužil P, Abelmann L, Guijt RM, Manz A. Direct coupling of a free-flow isotachophoresis (FFITP) device with electrospray ionization mass spectrometry (ESI-MS). Lab Chip. 2015;15:3495–502.CrossRefGoogle Scholar
  28. 28.
    Kochmann S, Agostino FJ, LeBlanc JCY, Krylov SN. Hyphenation of production-scale free-flow electrophoresis to electrospray ionization mass spectrometry using a highly conductive background electrolyte. Anal Chem. 2016;88:8415–20.CrossRefGoogle Scholar
  29. 29.
    Köhler S, Nagl S, Fritzsche S, Belder D. Label-free real-time imaging in microchip free-flow electrophoresis applying high speed deep UV fluorescence scanning. Lab Chip. 2012;12:458–63.CrossRefGoogle Scholar
  30. 30.
    Poehler E, Herzog C, Lotter C, Pfeiffer SA, Aigner D, Mayr T, Nagl S. Label-free microfluidic free-flow isoelectric focusing, pH gradient sensing and near real-time isoelectric point determination of biomolecules and blood plasma fractions. Analyst. 2015;140:7496–502.Google Scholar
  31. 31.
    Becker M, Budich C, Deckert V, Janasek D. Isotachophoretic free-flow electrophoretic focusing and SERS detection of myoglobin inside a miniaturized device. Analyst. 2008;134:38–40.CrossRefGoogle Scholar
  32. 32.
    Jezierski S, Klein AS, Benz C, Schaefer M, Nagl S, Belder D. Towards an integrated device that utilizes adherent cells in a micro-free-flow electrophoresis chip to achieve separation and biosensing. Anal Bioanal Chem. 2013;405:5381–6.CrossRefGoogle Scholar
  33. 33.
    Yin X-Y, Dong J-Y, Wang H-Y, Li S, Fan L-Y, Cao C-X. A simple chip free-flow electrophoresis for monosaccharide sensing via supermolecule interaction of boronic acid functionalized quencher and fluorescent dye. Electrophoresis. 2013;34:2185–92.CrossRefGoogle Scholar
  34. 34.
    Jezierski S, Gitlin L, Nagl S, Belder D. Multistep liquid-phase lithography for fast prototyping of microfluidic free-flow-electrophoresis chips. Anal Bioanal Chem. 2011;401:2651–6.CrossRefGoogle Scholar
  35. 35.
    Saha S, Alamsetti SK, Schneider C. Chiral Brønsted acid-catalyzed Friedel–Crafts alkylation of electron-rich arenes with in situ-generated ortho-quinone methides: highly enantioselective synthesis of diarylindolylmethanes and triarylmethanes. Chem Commun. 2015;51:1461–4.CrossRefGoogle Scholar
  36. 36.
    Schulze P, Ludwig M, Kohler F, Belder D. Deep UV laser-induced fluorescence detection of unlabeled drugs and proteins in microchip electrophoresis. Anal Chem. 2005;77:1325–9.CrossRefGoogle Scholar
  37. 37.
    Ohla S, Schulze P, Fritzsche S, Belder D. Chip electrophoresis of active banana ingredients with label-free detection utilizing deep UV native fluorescence and mass spectrometry. Anal Bioanal Chem. 2011;399:1853–7.CrossRefGoogle Scholar

Copyright information

© Springer-Verlag GmbH Germany 2017

Authors and Affiliations

  • Simon A. Pfeiffer
    • 1
  • Benjamin M. Rudisch
    • 1
  • Petra Glaeser
    • 1
  • Matthias Spanka
    • 2
  • Felix Nitschke
    • 3
  • Andrea A. Robitzki
    • 3
  • Christoph Schneider
    • 2
  • Stefan Nagl
    • 1
    • 4
  • Detlev Belder
    • 1
  1. 1.Institut für Analytische ChemieUniversität LeipzigLeipzigGermany
  2. 2.Institut für Organische ChemieUniversität LeipzigLeipzigGermany
  3. 3.Center for Biotechnology and BiomedicineUniversität LeipzigLeipzigGermany
  4. 4.Department of ChemistryThe Hong Kong University of Science and TechnologyKowloonChina

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