Monatshefte für Chemie - Chemical Monthly

, Volume 149, Issue 9, pp 1685–1691 | Cite as

Development of a miniaturized injection cell for online electrochemistry–capillary electrophoresis–mass spectrometry

  • Thomas Herl
  • Nicole Heigl
  • Frank-Michael Matysik
Original Paper


The elucidation of oxidation or reduction pathways is important for the electrochemical characterization of compounds of interest. In this context, hyphenation of electrochemistry and mass spectrometry is frequently applied to identify products of electrochemical reactions. In this contribution, the development of a novel miniaturized injection cell for online electrochemistry–capillary electrophoresis–mass spectrometry (EC–CE–MS) is presented. It is based on disposable thin-film electrodes, which allow for high flexibility and fast replacement of electrode materials. Thus, high costs and time-consuming maintenance procedures can be avoided, which makes this approach interesting for routine applications. The cell was designed to be suitable for investigations in aqueous and particularly non-aqueous solutions making it a universal tool for a broad range of analytical problems. EC–CE–MS measurements of different ferrocene derivatives in non-aqueous solutions were carried out to characterize the cell. Oxidation products of ferrocene and ferrocenemethanol were electrochemically generated and could be separated from the decamethylferricenium cation. The importance of fast CE–MS analysis of instable oxidation products was demonstrated by evaluating the signal of the ferriceniummethanol cation depending on the time gap between electrochemical generation and detection.

Graphical abstract


Capillary zone electrophoresis Electrochemistry Mass spectrometry Reaction mechanisms 


Electrochemical methods are of high relevance in many fields of research. They are essential tools for studies in the context of material sciences such as corrosion studies [1], the development of energy carriers [2], microbial fuel cells [3], or electrosynthetic processes [4]. Electrochemistry is also widely applied in bioanalytical studies such as the electrochemical simulation of oxidative stress [5, 6, 7, 8] or metabolic processes [9, 10, 11, 12, 13, 14, 15, 16, 17].

Pure electrochemical investigations such as cyclic voltammetry are well suited for the characterization of redox activities and reversibility of redox processes [18], but lack of qualitative information regarding mechanistic details. Thus, hyphenation to powerful detection techniques is in demand to obtain additional information on processes taking place on the electrode surface. In this context, hyphenation of electrochemistry (EC) to electrospray ionization–mass spectrometry (ESI–MS) is a frequently applied method, as recent reviews point out [16, 19]. ESI–MS offers high sensitivity and the possibility of identifying products of electrochemical reactions by their molecular masses and isotopic patterns with low fragmentation in the ionization process [20]. Thus, this technique can help in the elucidation of possible reaction mechanisms. However, EC and ESI have to be decoupled as electrochemical cells are operated at low voltages, while in ESI high-voltage conditions are applied [21]. This can be achieved using setups with grounded ESI interfaces [22].

Typical approaches to EC–MS comprise the direct coupling of electrochemical flow cells such as coulometric flow-through cells with porous electrodes or thin-layer flow cells with planar electrodes to mass spectrometry [4, 19, 23, 24]. Furthermore, efforts towards miniaturized setups using microfluidic electrochemical cells and nanoscale electrochemical reactors were made [25]. The advantage of such an approach is the simplicity of the experimental setup and the possibility of very fast detection within seconds [26], as the analytes are directly transported to MS via pumps. However, based on the cell type, the dependence of the conversion efficiency on the flow rate and the electrode surface area has to be kept in mind [23, 24]. An innovative approach to EC as online sample preparation technique for ESI–MS was developed by Dytrtová et al. [27], who coupled an electrochemical cell with switchable working electrodes to ESI–MS to ionize even non-polar organic compounds by adduct formation with electrochemically generated reactive metallic ions.

However, there are also some limitations of direct EC–MS. In complex samples, additional separation steps are necessary, as ion suppression effects in the ion source of MS can influence the detection, especially if mixtures of products are formed or if product and educt species show significant differences in their ionization properties. Thus, a quantification of oxidation or reduction products is difficult. Overlapping mass spectra can prevent the clear identification of individual species. Moreover, the separation behavior can give important additional information on the analytes, such as the presence of functional groups or polarity.

Hyphenation to separation techniques is often achieved by coupling EC to HPLC via electrochemical flow cells installed prior to or after the separation column [7, 13, 24]. However, the instrumental setups for this purpose are quite complex as different pumps and valves are needed, if EC is carried out before HPLC [19]. Another disadvantage that can arise is the compatibility between the conditions needed for electrochemical reactions and the separation conditions [13]. Contrary to direct EC–MS, the time gap between formation and detection of products is longer if a separation step is carried out after oxidation or reduction. Typical analysis times are in the range of several minutes [7, 14, 23]. Working with reversed phase HPLC–MS it has to be considered that non-polar analytes are usually favored for HPLC, but polar analytes are better compatible to ESI–MS conditions [28].

Due to the aspects mentioned above, capillary electrophoresis (CE) is the method to be preferred in some cases. It offers fast separation and low solvent consumption [29, 30] and is suitable for separation of charged species. Thus, it is an ideal separation method for many biomolecules that contain functional groups which can be protonated or deprotonated depending on pH. In contrast to HPLC, separations are possible under nearly physiological conditions [5] and can be carried out in the same electrolyte as the oxidation or reduction, so that the migration behavior in CE is representative for the state of charge of the analytes in the electrolyte. Due to the charge-dependent migration behavior, CE allows for differentiation between ions, which are generated by electrochemical processes and ions generated in the ionization prior to MS detection. This information cannot be obtained in direct EC–MS or EC–HPLC–MS.

First online EC–CE approaches were established in 2003 [31, 32] using batch electrolysis cells and classical three-electrode setups. Thus, they had the disadvantage of time-consuming electrode maintenance procedures, which are necessary to avoid electrode fouling and require experienced users. Additionally, a comparably high sample volume is needed. In contrast to that, Palatzky et al. [33] developed a fully automated device for online EC–CE–MS based on disposable screen-printed electrodes (SPEs). Hence, compared to classical cells, significantly lower sample consumption (about 50 mm3 is sufficient), easy replacement of electrodes avoiding time-consuming cleaning and polishing procedures, and high flexibility concerning electrode materials could be achieved. The electrochemical cell consisted of a droplet of solution placed onto the three-electrode structure of the electrode and sample injection into the CE-system was achieved by placing the separation capillary into this droplet directly above the working electrode. However, this system was not compatible to non-aqueous solutions due to the screen-printed electrode materials, which is a major drawback when it comes to the investigation of analytes that are not readily soluble or stable [31] in water.

This contribution presents an instrumental approach to online EC–CE–MS with disposable electrodes, which is applicable under non-aqueous conditions. It is based on the existing fully automated EAI–CE–MS device described in [33]. To allow for investigations in non-aqueous solutions, different problems had to be addressed. As already mentioned above, screen-printed electrodes are attacked by organic solvents, so that alternative electrode types had to be used. Commercially available thin-film electrodes consist of metal electrode materials fabricated on glass substrates and are ideal for this purpose as they are solvent-resistant [26]. However, simply applying droplets of solution onto the electrode surface as it could be done with aqueous solutions [5, 33] was not possible, as organic solvents easily spread due to low surface tension. This can lead to electrical shortcuts and corrosion problems, when the liquid flows into electrical contacts. Therefore, the cell volume had to be delimited physically to prevent spreading of the liquid. To overcome these problems, a novel miniaturized injection cell for online EC–CE–MS with integrated thin-film electrodes was developed, capable of measurements in aqueous and especially non-aqueous media. A model mixture consisting of ferrocene (Fc), ferrocenemethanol (FcMeOH), and decamethylferrocene (dMFc) was used to characterize this injection cell. The importance of short separation times was demonstrated by evaluation of the dependency of the FcMeOH+ signal on the separation time.

Results and discussion

Design and fabrication of the injection cell

The developed injection cell was based on commercial Micrux thin-film electrodes (size 10 mm × 6 mm, working electrode diameter 1 mm). The cell geometry was adapted to the existing EC–CE–MS setup [33] to allow for the usage of thin-film electrodes without changing the injection unit. Instead of a SPE, the injection cell with integrated thin-film electrode was installed in the injection unit. Due to electrode and cell dimensions, small sample volumes of only 10 mm3 or lower were sufficient for EC–CE–MS measurements, which is especially advantageous if only limited amount of sample is available. A schematic illustration of the final injection cell prototype is shown in Fig. 1.
Fig. 1

Illustrations of a the injection cell in the setup at injection position and b exploded view of injection cell. The cell is installed in the bottom part (2) of the EC–CE–MS device next to buffer reservoirs for CE separation. The separation capillary (1) is installed in the top part. The injection cell consists of a bottom piece (4) with electrode slot and a cover piece (3) with electrical contacts. A silicone sealing ring prevents leakage of the sample

Polyether ether ketone (PEEK), a highly chemical resistant and mechanically stable material, was used for fabrication of the cell body. To prevent leakage, a sealing ring was integrated at the bottom of the open cell chamber. Thus, spreading of droplets could be avoided. As commercially available O-ring materials were attacked by organic solvents, a custom silicone sealing ring with appropriate dimensions (inner diameter 2 mm, outer diameter 4 mm, thickness 1 mm) was prepared. The electrical contact to the implemented thin-film electrode was achieved via spring contact probes. To facilitate a fast assembling and disassembling of the cell, magnets were integrated to keep the cell closed. Due to materials and modularity of the cell, it could be cleaned easily and was suitable for measurements in aqueous as well as non-aqueous solutions. Electrodes could easily be exchanged, which allowed for high flexibility regarding electrode materials. When installed in the EC–CE–MS device, a fully automated hydrodynamic injection of sample directly from the working electrode surface was possible by placing the tapered tip of the fused silica separation capillary onto the electrode surface. The overall experimental setup is illustrated in Fig. 5 in the experimental section.

EC–CE–MS experiments

By CE–MS, a fast separation and detection of neutral and particularly cationic species was possible applying a positive high voltage at the injection end of the capillary (detection end of capillary installed in grounded ESI sprayer). Figure 2a shows a CE–MS measurement that was carried out at a separation voltage of 18 kV (2.4 µA) without previous oxidation using a model mixture of Fc, FcMeOH, and dMFc. The three model substances showed different behavior regarding state of charge and detectability. In the case of dMFc, the migration behavior in CE and the respective mass detected in MS indicated that only the cationic dMFc+ (m/z = 326.20) was present in solution even without electrochemical oxidation. This is because in dMFc Fe(II) is easily oxidized to Fe(III) by dissolved oxygen forming a stable cationic complex [32, 34]. This behavior is well known as already reported in 1990 by Bashkin and Kinlen [35]. FcMeOH was migrating with the EOF, showing that it was neutral in solution. The detected mass (m/z = 199.02) indicated a loss of the hydroxyl group during the ionization process, whereas no protonation or oxidation in the ESI source could be observed under the applied conditions. Non-oxidized Fc could not be detected due to its hydrophobicity and thus poor ionization efficiency in ESI. Unlike Dytrtová et al. [36], who used a NaClO4/HClO4 electrolyte, we could not observe a protonation or oxidation of Fc in solution or in the ESI source. In contrast to dMFc, Fc and FcMeOH exhibited no peaks corresponding to cationic species in CE separation without previous electrochemical oxidation. By electrochemical oxidation at 0.5 V for 10 s, the cationic ferricenium (m/z = 186.01) and ferriceniummethanol species (m/z = 216.02) were formed, as could be confirmed by the m/z values and the migration behavior in CE. A representative electropherogram is shown in Fig. 2b. Fc+ and FcMeOH+ were migrating to the cathode faster than dMFc+. The results demonstrated the importance of CE separation: The migration behavior in CE and the comparison of the electropherograms before and after oxidation facilitated the distinction between cationic species that are present in solution and species formed in the ionization process, which is not possible without separation step.
Fig. 2

Electropherograms of the model mixture Fc, FcMeOH, and dMFc without oxidation (a) and after oxidation at 0.5 V for 10 s (b). The inset in b shows an enlarged view of the separated cationic species dMFc+ (m/z = 326.20), FcMeOH+ (m/z = 216.02), and Fc+ (m/z = 186.01). The migration time of FcMeOH marks the EOF. Before oxidation, only dMFc+ and FcMeOH (m/z = 199.02) were visible. After oxidation, additional peaks of FcMeOH+ and Fc+ were present. Separation voltage 18 kV (2.4 µA); capillary: ID = 25 µm, L = 35 cm; separation in ACN/10 mmol/dm3 NH4OAc/1 mol/dm3 HOAc; 2 s hydrodynamic injection

The results showed that electrochemical sample pretreatment and online analysis of oxidation products could be achieved within a short time scale. Very short oxidation times of only 10 s (injection during last 2 s of oxidation) were enough to generate a sufficient amount of product species for detection. The electrochemical generation and detection of Fc+ and FcMeOH+ were feasible within 90 s and both were separated from dMFc+.

Fast online analysis of oxidation products

Besides high-throughput aspects, short analysis times are crucial for the investigation of reactive or instable species to allow for a reliable identification and sensitive detection. To demonstrate that, the detection of FcMeOH+ was investigated depending on the separation conditions. As in the case of FcMeOH both, the cationic and the neutral species were detectable, the peak of neutral FcMeOH could be used as internal standard for characterization of the analytical performance. The time gap between generation and detection of FcMeOH+ was varied by changing the separation voltage and thus the migration time. Representative electropherograms measured after oxidation are illustrated in Fig. 3. As visible in the measurements, the ratio of the FcMeOH+ peak to the FcMeOH peak continuously increased with higher separation voltage meaning faster migration. This indicated the decomposition of the cation over time, as after longer migration less amount of cation could be detected. As depicted in Fig. 3a, the detection of both species took quite long and the signal corresponding to the cationic species was comparably small when applying a separation voltage of 2 kV, while at a separation voltage of 18 kV, a fast detection, narrow peaks and a high intensity of the FcMeOH+ peak were observed as shown in Fig. 3e.
Fig. 3

EC–CE–MS measurements of FcMeOH (m/z = 199.02, 216.02) after oxidation at 0.5 V for 10 s. Separations were carried out at different separation voltages: a 2 kV (0.7 µA), b 6 kV (1.3 µA), c 10 kV (1.8 µA), d 14 kV (2.1 µA), e 18 kV (2.4 µA). Capillary: ID = 25 µm, L = 35 cm; separation in ACN/10 mmol/dm3 NH4OAc/1 mol/dm3 HOAc; 2 s hydrodynamic injection

Further evaluation of the experimental data led to the results illustrated in Fig. 4. The peak area obtained for FcMeOH+ normalized to the peak area corresponding to FcMeOH was plotted versus the migration time of the respective FcMeOH+ peak. Due to the fully automated oxidation and injection procedures, the measurements showed a good reproducibility regarding migration times and amount of cation formed. The time between generation and detection of FcMeOH+ could precisely be controlled by the high voltage. At short migration times, a significantly larger signal for the cationic species was obtained, so that the sensitivity of the system could be enhanced by applying higher separation voltages. These results indicate that this method is promising for sensitive detection of instable oxidation or reduction products due to the possibility of fast online analysis.
Fig. 4

Peak area of FcMeOH+ (normalized to the peak area of the FcMeOH signal) vs. migration time of FcMeOH+. The migration times were controlled by the separation voltages of 18, 14, 10, 6, and 2 kV (from left to right). The standard deviations of migration times and peak ratios (n = 3) are indicated by error bars


A miniaturized injection cell for online EC–CE–MS was developed and characterized. It was capable of handling very small sample volumes of 10 mm3 or lower. Electrodes and injection cell were solvent-resistant, so that online investigations of electrochemical reactions in aqueous and particularly non-aqueous media were possible. The integration of disposable thin-film electrodes leads to a high flexibility in electrode materials and to an easy exchange of electrodes, which is minimizing artifacts due to adsorption or electrode fouling. Time-consuming electrode maintenance procedures that usually need experienced users can be avoided. In online EC–CE–MS using ferrocene derivatives, short analysis times within few minutes from generation to detection of oxidized species were possible. Fc+ and FcMeOH+ could be generated and separated from dMFc+ within less than 90 s. Due to that, the method is suitable for the investigation of instable products, which was demonstrated by evaluating the signal of electrochemically generated FcMeOH+ depending on separation speed. Fully automated oxidation and injection procedures allowed for reproducible measurements and a reliable control of the time gap between formation and detection of oxidized species.

In conclusion, this novel setup extends the applicability of online EC–CE–MS based on disposable electrodes to analytes that are only soluble in organic solvents, which was not possible using screen-printed electrodes. The high flexibility and possibility of fast online analysis make this setup attractive for further applications, such as kinetic studies or electrochemical simulation of metabolic processes with particular focus on reactive species.


Reagents and chemicals

The following chemicals were used, all of analytical grade or higher if not stated otherwise: acetic acid (Sigma Aldrich, MO, USA), acetonitrile, ammonium acetate (both Merck, Darmstadt, Germany), decamethylferrocene (purity 99%, ABCR, Karlsruhe, Germany), formic acid (Merck, Darmstadt, Germany), ferrocene (purity 98%, Riedel-de-Haën, Seelze, Germany), ferrocenemethanol (purity 99%, ABCR, Karlsruhe, Germany), isopropanol (Roth, Karlsruhe, Germany).


For EC–CE–MS measurements, the fully automated CE system developed by Palatzky et al. [33] was used. The setup was installed in a plexiglass box and connected to a high voltage supply (HCN 7E 35000, FuG Elektronik, Schechen, Germany). A micrOTOF time-of-flight mass spectrometer (Bruker Daltonics, Bremen, Germany), equipped with a coaxial sheath liquid ESI interface (Agilent Technologies, Waldbronn, Germany), was used for detection. It was operated in positive ion mode. A mixture of isopropanol:water:formic acid (49.9:49.9:0.2, v:v:v) was added as sheath liquid at a flow rate of 8 mm3/min with a syringe pump (KS Scientific, Holliston, MA, USA). Separations were carried out in fused silica capillaries (Polymicro Technologies, AZ, USA) with an outer diameter of 360 µm, an inner diameter of 25 µm, and a length of 35 cm. The detection end of the capillary was polished to a plane edge while the injection end of the capillary was polished to an angle of 15°. The capillaries were preconditioned by flushing with 0.1 mol/dm3 NaOH for 10 min, followed by water for 5 min and background electrolyte (BGE) for CE separation (ACN/10 mmol/dm3 NH4OAc/1 mol/dm3 HOAc) for at least 30 min. A thin-film electrode with gold working, counter, and quasireference electrode (ED-SE1-Au, Micrux Technologies, Oviedo, Spain) was used for the oxidation. All potentials given are referred to the Au quasireference electrode and all electrochemical measurements were carried out in ACN/10 mmol/dm3 NH4OAc/1 mol/dm3 HOAc as BGE. The electrode was installed in the novel injection cell described in detail in the results and discussion section. A µSTAT 200 potentiostat (DropSens, Llanera, Spain) controlled by Dropview 200 software was used for applying potentials. A schematic illustration of the experimental setup is depicted in Fig. 5.
Fig. 5

Illustration of the instrumental setup used for EC–CE–MS measurements: a injection unit with novel injection cell and electrolyte reservoirs, b potentiostat, c high voltage source, d fused silica capillary, e mass spectrometer, f computer

Experimental procedures

For evaluation of the cell performance, a solution of 1.5 mmol/dm3 Fc, 1 mmol/dm3 FcMeOH, and 40 µmol/dm3 dMFc in BGE was used. Fast detection studies were carried out with a solution of 1 mmol/dm3 FcMeOH in BGE. For the CE protocol, 8 mm3 of sample solution were filled into the cell. The sample was hydrodynamically injected into the CE system by placing the tapered end of the separation capillary onto the working electrode surface for 2 s at a difference in height of 18 cm between the injection end of the capillary and the detection end of the capillary (hydrostatic pressure). After the injection, the capillary was automatically placed into the 2-cm3 BGE reservoir and the separation voltage denoted in detail in the respective measurements in the results section was applied. Measurements were carried out without previous oxidation and after oxidation at 0.5 V for 10 s (injection during the last 2 s of oxidation). For data evaluation the extracted ion signals of Fc (m/z = 186.01), FcMeOH (m/z = 199.02; m/z = 216.02), and dMFc (m/z = 326.20) were used.

The parameters for the MS detection were as follows: acquisition: ion polarity: positive; mass range: m/z = 100–350; spectra rate 5 Hz; source: end plate offset: − 500 V; capillary: − 4000 V; nebulizer: 1.0 bar; dry gas: 4.0 dm3/min; dry temperature: 190 °C; transfer: capillary exit: 75.0 V; skimmer 1: 25.3 V; hexapole 1: 23.0 V; hexapole RF: 65.0 Vpp; skimmer 2: 23.0 V; lens 1 transfer: 38.0 µs; lens 1 pre pulse storage: 6.0 µs.


  1. 1.
    Arjmand F, Adriaens A (2014) J Solid State Electrochem 18:1779CrossRefGoogle Scholar
  2. 2.
    Chandrasekaran S, Chung JS, Kim EJ, Hur SH (2016) J Electrochem Sci Technol 7:1CrossRefGoogle Scholar
  3. 3.
    Tharali AD, Sain N, Osborne WJ (2016) Front Life Sci 9:252CrossRefGoogle Scholar
  4. 4.
    Gul T, Bischoff R, Permentier HP (2015) TrAC Trends Anal Chem 70:58CrossRefGoogle Scholar
  5. 5.
    Scholz R, Palatzky P, Matysik F-M (2014) Anal Bioanal Chem 406:687CrossRefPubMedGoogle Scholar
  6. 6.
    Cindric M, Vojs M, Matysik F-M (2015) Electroanalysis 27:234CrossRefGoogle Scholar
  7. 7.
    Erb R, Plattner S, Pitterl F, Brouwer H-J, Oberacher H (2012) Electrophoresis 33:614CrossRefPubMedPubMedCentralGoogle Scholar
  8. 8.
    Baumann A, Lohmann W, Jahn S, Karst U (2010) Electroanalysis 22:286CrossRefGoogle Scholar
  9. 9.
    Karst U (2004) Angew Chem Int Ed 43:2476CrossRefGoogle Scholar
  10. 10.
    Jurva U, Wikström HV, Bruins AP (2000) Rapid Commun Mass Spectrom 14:529CrossRefPubMedGoogle Scholar
  11. 11.
    Jurva U, Wikström HV, Weidolf L, Bruins AP (2003) Rapid Commun Mass Spectrom 17:800CrossRefPubMedGoogle Scholar
  12. 12.
    Johansson T, Weidolf L, Jurva U (2007) Rapid Commun Mass Spectrom 21:2323CrossRefPubMedGoogle Scholar
  13. 13.
    Faber H, Vogel M, Karst U (2014) Anal Chim Acta 834:9CrossRefPubMedGoogle Scholar
  14. 14.
    Nouri-Nigjeh E, Permentier HP, Bischoff R, Bruins AP (2010) Anal Chem 82:7625CrossRefPubMedGoogle Scholar
  15. 15.
    Yuan T, Permentier H, Bischoff R (2015) TrAC Trends Anal Chem 70:50CrossRefGoogle Scholar
  16. 16.
    Bussy U, Boisseau R, Thobie-Gautier C, Boujtita M (2015) TrAC Trends Anal Chem 70:67CrossRefGoogle Scholar
  17. 17.
    Jurva U, Weidolf L (2015) TrAC Trends Anal Chem 70:92CrossRefGoogle Scholar
  18. 18.
    Bard AJ, Faulkner LR (2001) Electrochemical methods: fundamentals and applications, 2nd edn. Wiley, HobokenGoogle Scholar
  19. 19.
    Portychová L, Schug KA (2017) Anal Chim Acta 993:1CrossRefPubMedGoogle Scholar
  20. 20.
    Fenn JB, Mann M, Meng CK, Wong SF, Whitehouse CM (1989) Science 246:64CrossRefPubMedGoogle Scholar
  21. 21.
    Zhou F, Van Berkel GJ (1995) Anal Chem 67:3643CrossRefGoogle Scholar
  22. 22.
    Liu P, Zheng Q, Dewald HD, Zhou R, Chen H (2015) TrAC Trends Anal Chem 70:20CrossRefGoogle Scholar
  23. 23.
    Lohmann W, Baumann A, Karst U (2010) LCGC N Am 28:470Google Scholar
  24. 24.
    Bruins AP (2015) TrAC Trends Anal Chem 70:14CrossRefGoogle Scholar
  25. 25.
    van den Brink FTG, Olthuis W, van den Berg A, Odijk M (2015) TrAC Trends Anal Chem 70:40CrossRefGoogle Scholar
  26. 26.
    Herl T, Matysik F-M (2017) Tech Mess 84:672CrossRefGoogle Scholar
  27. 27.
    Jaklová Dytrtová J, Jakl M, Navrátil T, Cvačka J, Pačes O (2016) Electrochim Acta 211:787CrossRefGoogle Scholar
  28. 28.
    Diehl G, Liesener A, Karst U (2001) Analyst 126:288CrossRefPubMedGoogle Scholar
  29. 29.
    Wang X, Li K, Adams E, Van Schepdael A (2013) Curr Drug Metab 14:807CrossRefPubMedGoogle Scholar
  30. 30.
    Mark JJP, Piccinelli P, Matysik F-M (2014) Anal Bioanal Chem 406:6069CrossRefPubMedGoogle Scholar
  31. 31.
    Esaka Y, Okumura N, Uno B, Goto M (2003) Electrophoresis 24:1635CrossRefPubMedGoogle Scholar
  32. 32.
    Matysik F (2003) Electrochem Commun 5:1021CrossRefGoogle Scholar
  33. 33.
    Palatzky P, Zöpfl A, Hirsch T, Matysik F-M (2013) Electroanalysis 25:117CrossRefGoogle Scholar
  34. 34.
    Singh A, Chowdhury DR, Paul A (2014) Analyst 139:5747CrossRefPubMedGoogle Scholar
  35. 35.
    Bashkin JK, Kinlen PJ (1990) Inorg Chem 29:4507CrossRefGoogle Scholar
  36. 36.
    Jaklová Dytrtová J, Jakl M, Navrátil T, Schröder D (2013) Int J Electrochem Sci 8:1623Google Scholar

Copyright information

© Springer-Verlag GmbH Austria, part of Springer Nature 2018

Authors and Affiliations

  • Thomas Herl
    • 1
  • Nicole Heigl
    • 1
  • Frank-Michael Matysik
    • 1
  1. 1.Institute of Analytical Chemistry, Chemo- and BiosensorsUniversity of RegensburgRegensburgGermany

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