Here, we present a protocol to study the molecular mechanism of proton translocation across lipid membranes of single liposomes, using cytochrome bo3 as an example. Combining electrochemistry and fluorescence microscopy, pH changes in the lumen of single vesicles, containing single or multiple enzyme, can be detected and analyzed individually.
Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used for ATP production. The amphiphilic nature of membrane proteins requires particular attention to their handling, and reconstitution into the natural lipid environment is indispensable when studying membrane transport processes like proton translocation. Here, we detail a method that has been used for the investigation of the proton-pumping mechanism of membrane redox enzymes, taking cytochrome bo3 from Escherichia coli as an example. A combination of electrochemistry and fluorescence microscopy is used to control the redox state of the quinone pool and monitor pH changes in the lumen. Due to the spatial resolution of fluorescent microscopy, hundreds of liposomes can be measured simultaneously while the enzyme content can be scaled down to a single enzyme or transporter per liposome. The respective single enzyme analysis can reveal patterns in the enzyme functional dynamics that might be otherwise hidden by the behavior of the whole population. We include a description of a script for automated image analysis.
Information about enzyme mechanisms and kinetics is usually obtained on the ensemble or macroscale level with enzyme population in the thousands to millions of molecules, where measurements represent a statistical average. It is known, however, that complex macromolecules such as enzymes may demonstrate heterogeneity in their behavior and molecular mechanisms observed at the ensemble level are not necessarily valid for every molecule. Such deviations on the individual molecule scale have been extensively confirmed by studies of single enzymes with a variety of methods emerging during the last two decades1. Notably, fluorescence detection of individual enzyme activity has been used to investigate heterogeneity of enzymes activity2,3 or discover the so-called memory effect (periods of high enzymes activity succeeded by periods of low activity and vice versa)4,5.
Many single enzyme studies require that the enzymes are immobilized on the surface or spatially fixed in another way to remain sufficiently long in the field of view for continuous observation. Enzyme encapsulation into liposomes has been shown to enable enzyme immobilization while preventing any negative impact due the surface-enzyme or protein-protein interactions6,7. In addition, liposomes offer a unique possibility to study single membrane proteins in their natural lipid bilayer environment8,9,10.
A class of membrane proteins, transporters, exercises a directional translocation of substances across the cell membrane, a behavior that can only be studied when proteins are reconstituted into the lipid bilayers (e.g., liposomes)11,12,13. For example, proton translocation, exhibited by several enzymes of prokaryotic and eukaryotic electron transport chains, plays an important role in cellular respiration by creating a proton-motive force used for ATP synthesis. In this case, the proton pumping activity is coupled to the electron transfer, although the detailed mechanism of this process often remains elusive.
Recently, we demonstrated the possibility to couple fluorescent detection with electrochemistry to study proton pumping activity of single enzymes of the terminal ubiquinol oxidase of Escherichia coli (cytochrome bo3) reconstituted in the liposomes14. This was achieved by encapsulation of a pH-sensitive membrane-impermeable fluorescent dye into the lumen of liposomes prepared from E. coli polar lipids (Figure 1A). The protein amount was optimized so that most liposomes either contained no or only one reconstituted enzyme molecule (according to Poisson distribution). The two substrates of cytochrome bo3 were provided by adding ubiquinone to the lipid mix that formed the liposomes and (ambient) oxygen in solution. The liposomes are then sparsely adsorbed on a semi-transparent ultra-smooth gold electrode, covered with a self-assembled monolayer of 6-mercaptohexanol. Finally, the electrode is mounted on the bottom of a simple spectroelectrochemical cell (Figure 1B). Electrochemical control of the quinone pool redox state allows one to flexibly trigger or to stop the enzymatic reaction at any moment, while the pH-sensitive dye is used to monitor pH changes inside the lumen of the liposomes as a result of proton translocation by the enzymes. By using the fluorescence intensity of a second, lipid-bound fluorescent dye, the size and volume of individual liposomes can be determined and thus the quantification of enzyme proton pumping activity. Using this technique, we notably found that cytochrome bo3 molecules are able to enter into a spontaneous leak state that rapidly dissipates the proton motive force. The goal of this article is to introduce the technique of single liposome measurements in detail.
1. Preparation of a Lipid/UQ-10/FDLL Mix
NOTE: E. coli lipids used for the liposomes’ preparation should be aliquoted and thoroughly mixed with ubiquinone-10 (enzyme substrate) and long-wavelength fluorescent dye-labeled lipids (for liposomes size determination) prior to the reconstitution.
2. Reconstitution of Cytochrome bo3
NOTE: For the purification of cytochrome bo3 from E. coli, follow the protocol from Rumbley et al.15 To ensure high purity of natively-folded enzyme samples, add size-exclusion chromatography after the affinity purification step described by Rumbley et al. 15
3. Fabrication of Semi-Transparent Gold Electrodes
NOTE: The smooth gold surface is obtained by a template stripping method of a 30 nm-thick layer of 99.99% gold from an atomically smooth silicon wafer. The small thickness of the gold layer is important since it must be semi-transparent to permit fluorescence observation. Details of gold evaporation (physical vapor deposition, PVD) can be find elsewhere16 and only template-stripping is covered here. Alternatively, ultra-smooth gold chips can be purchased elsewhere (see Table of Materials).
4. Modification of the Gold Surface with Self-Assembled Monolayer (SAM)
5. Electrochemical Testing of Proteoliposomes Activity
NOTE: The electrochemical activity of enzyme is first verified on a closely packed liposomes layer (step 5) and lower vesicle coverages are used in single vesicles experiment to measure pH changes in the lumen of the liposomes (step 6).
6. Detection of Enzymatic Proton Pumping by Fluorescence Microscopy
7. Analysis of Fluorescence Images
NOTE: A typical experiment produces a set of images with a time step (e.g., 2.6 s) for each of the two channels, i.e., (duration * 2 / 2.6) images. An example of such image set recorded during a single enzyme experiment can be accessed via the Research Data Leeds repository21. An image treatment consists of several steps by using Fiji (ImageJ) and high-level mathematical analysis programming language software (henceforth referred to as scripting software, see Table of Materials for details).
8. Performing a Calibration Curve of HPTS Fluorescence
NOTE: To convert HPTS fluorescence ratio to an intravesicular pH, a calibration curve must be first established that would take into account particular conditions of experiments such as gold transmittance, filters quality, etc. This step has to be performed only once or twice and the calibration data can be used as long as the setup and measurement parameters remain the same in step 6.
The quality of the gold-modified cover slip (the electrode) with a SAM of 6MH is checked before each experiment with electrochemical impedance spectroscopy. Figure 2A shows representative Cole-Cole plots measured using electrochemical impedance spectroscopy before and after liposomes are adsorbed. If the quality of SAM is sufficient, impedance spectroscopy should demonstrate an almost pure capacitive behavior resulting in a semi-circle Cole-Cole plot. The diameter of the semi-circle in a Cole-Cole plot equals the double layer capacitance of the electrode surface, which should be in the 2.5-3.0 µF/cm2 range. Note that the capacitance should not change significantly upon liposomes adsorption, although a slight increase in capacitance might be observed (high liposomes coverage, Figure 2A, bottom) or a slight shift in impedance can be observed at low frequencies (low liposome coverages, Figure 2A, top).
Electrochemistry can be used to test the catalytic activity of cytochrome bo3 in the proteoliposomes, where catalytic currents measured with cyclic voltammetry reflect the ubiquinol-oxygen oxidoreductase activity. However, significant catalytic currents can only be detected at high quantities of adsorbed proteoliposomes and a closely packed layer of proteoliposomes on the gold-modified cover slip is needed. Figure 2B demonstrates representative cyclic voltammograms (CVs) of the electrode before and after liposomes adsorption with either low or high liposome coverage. No faradaic current is observed on the blank CV because oxygen reduction by the bare gold electrode is blocked by the SAM. When the surface is saturated with liposomes with high cytochrome bo3 content (1.3% (w/w) in this case), a clear catalytic wave due to oxygen reduction is observed with an onset potential of 0 V, i.e., the potential of ubiquinone reduction (Figure 2B, blue line). The ubiquinone pool acts both as the natural substrate for the enzyme and as an electron mediator, transferring electrons from the enzyme to the electrode surface. We note that under high liposome coverage, no distinction of individual vesicles is possible by microscopy and lower coverage is needed for single liposome studies. At low liposome coverage (but high protein to lipid ratio), the catalytic current is significantly reduced, barely distinguishable from background (Figure 2B, red line). Note that under single enzyme conditions (low protein to lipid ratio), the catalytic current is even lower and cannot be measured reliably.
Figure 3 shows fluorescence images of liposomes adsorbed on the electrodes at three different coverages. All images were taken in identical light and exposure conditions and their brightness was adjusted equally to enable direct comparison. The dye-containing-liposomes are visible on the images as bright spots. The central part of the image was photobleached for a couple of minutes to reveal background fluorescence level (we note that FDLL is relatively photostable and is not photobleached completely in Figure 3). The images on two HPTS channels are superimposable, where the ratio between the two channels (410/535 and 470/535) corresponds to pH 7.4 used in this experiment. A larger number of liposomes are visible with the FDLL channel, which indicates the presence of liposomes that have no HPTS encapsulated. The difference between the HPTS and FDLL channels is more pronounced at higher coverages, possibly because at high liposome coverage, liposomes are more likely to burst or fuse on the surface.
The image analysis requires the alignment of frames (against the first frame) using the ImageJ, plugin StackReg. The alignment is indispensable in most situations since a slight thermal drift of the sample usually occurs within the timescale of the experiment, changing the liposome coordinates. All further analysis is performed using a scripting software code. This code performs automatic liposome identification. As the size of liposomes are below the Abbe diffraction limit, their fluorescence is seen as a point spread function much larger than the actual liposome diameter (Figure 4A). The code fits the fluorescence intensity of each vesicle on two channels to a 2D-Gaussian function (Figure 4A) and calculates their volumetric intensity ratio that can be converted to pH using a calibration curve. By performing these actions on all time frames, the pH is obtained inside every vesicle within timescale of the experiment (Movie 1). Figure 4B shows the medians of all vesicle pH changes within a single image when cytochrome bo3 content is 1.3%. An increase in pH (proton pumping) is clearly visible when a potential between -0.1 and -0.3 V is applied, but not for 0 V since the latter is not sufficient to reduce the quinone pool. When cytochrome bo3 content is much lower (protein-to-lipid ratio of 0.1%, Figure 4C), the median curves become almost indistinguishable from those of empty liposomes (Figure 4D). The difference becomes evident when considering individual pH traces of random liposomes (Figure 5, 6 and 7). While liposomes without cytochrome bo3 display no significant pH changes with respect to noise (Figure 7), a selection of liposomes show an increase (dominantly) or a decrease of pH when a potential is applied that actives cytochrome bo3 (Figures 6, grey zone). The obvious difference in pH-traces between liposomes is consistent with the low protein-to-lipid ratio, where cytochrome bo3 is only present in a small subset of liposomes activity. The prevalence of a pH increase over decrease is explained by the reconstitution method that favors an "outward" orientation of enzyme molecules (ca. 75%). The fraction of liposomes that display a pH change increases when the cytochrome bo3 to lipid ratio is higher (Figure 5). Note also that some liposomes stop proton pumping and enter into proton dissipation mode before the end of the potential application. We attribute this behavior to the cytochrome bo3 molecules entering a "leak state", which allows protons to flow back into the liposome lumen14.
The further analysis of the pH traces including their fitting and determination of proton pumping/leaking rates can be done using a script we have published previously14,22 and can be obtained from the Research Data Leeds Repository23,24.
Figure 1: Principle of the method. (A) Principle of single enzyme activity monitoring and (B) the scheme of the experimental setup used in this work with a cutaway to demonstrate an internal part of the cell. Please click here to view a larger version of this figure.
Figure 2: Electrochemical response of liposomes. (A) Cole-Cole plots measured at OCP (0.22 V vs SHE; black squared line) before and after liposomes (1.3% w/w cytochrome bo3) adsorption from solutions at concentrations of 5 µg/mL (red circle line) and 500 µg/mL (blue circle line). (B) Cyclic voltammograms at 100 mV/s (left panel) and 10 mV/s (right panel) of Au-SAM (black dashed line) before and after liposomes (1.3% w/w cytochrome bo3) adsorption from solutions at concentrations of 5 µg/mL (red line) and 500 µg/mL (blue line). Please click here to view a larger version of this figure.
Figure 3: Fluorescence microscopy of liposomes. Fluorescence images of liposomes at different surface coverages: surfaces incubated for 30 min with a liposomes solution of 5 µg/mL (top row), 20 µg/mL (middle row) and 500 µg/mL (bottom row). Left column corresponds to 470/535 nm excitation/emission filter setup (1st HPTS channel); middle column – 410/535 nm (2nd HPTS channel); right column – 560/645 nm (FDLL channel). The images were taken at magnification 90X (60X objective and 1.5X magnification by the microscope prior to the camera), 1 s exposure and similar light intensity. Scale bars correspond to 50 µm. Please click here to view a larger version of this figure.
Figure 4: Liposome identification and pH change. (A) 3D-view of a part of fluorescence image containing a typical single liposome. Its fluorescence intensity is seen as a point spread function (color surface) that is fitted to a 2D-Gaussian function (black mesh). (B-D) pH, displayed as the median of all vesicles within single image area (typically several hundred) at different applied potentials. From 0-60 s, no potential is applied (OCP). Between 60 and 180 s (grey zone) different potentials were applied: 0 V (black), -0.1 V (red), -0.2 V (green), -0.3 V (blue). Between 180 and 300 s, 0.4 V vs. SHE was applied. The cytochrome bo3 to lipid ratios were (B) 1.3%, (C) 0.1%, (D) 0%. The traces are offset for clarity. Please click here to view a larger version of this figure.
Figure 5: pH traces of liposomes containing 1.3% of enzyme. Traces of pH change of 72 single liposomes, randomly selected, containing cytochrome bo3 (1.3% w/w) measured and analyzed during one experiment. From 0-60 s, no potential is applied (OCP); between 60 and 180 s (grey zone) -0.2 V vs. SHE; between 180 and 300 s, 0.4 V vs. SHE was applied. Please click here to view a larger version of this figure.
Figure 6: pH traces of liposomes containing 0.1% of enzyme. Traces of pH change of 72 single liposomes, randomly selected, containing cytochrome bo3 (0.1% w/w) measured and analyzed during one experiment. From 0-60 s, no potential is applied (OCP); between 60 and 180 s (grey zone) -0.2 V vs. SHE; between 180 and 300 s, 0.4 V vs. SHE was applied. Please click here to view a larger version of this figure.
Figure 7: pH traces of liposomes without enzyme. Traces of pH change of 72 single liposomes, randomly selected, without cytochrome bo3 measured and analyzed during one experiment. From 0-60 s: no potential is applied (OCP); between 60 and 180 s (grey zone) -0.2 V vs. SHE; between 180 and 300 s, 0.4 V vs. SHE was applied. Please click here to view a larger version of this figure.
Movie 1: Animation of single liposome pH change. (top panel) Change of a liposome fluorescence on two HPTS channels (shown as 3D-surface plot of the corresponding area) during 300 s of the experiment. The reductive potential was applied between 60 s and 180 s. (bottom panel) Corresponding plot of volumetric intensity ratio of two HPTS channels versus time. The zone of reductive potential application is shaded in grey. Please click here to view this video. (Right-click to download.)
Supplemental Files. Please click here to download the files.
The method described is suitable to study proton pumping by respiratory membrane proteins that can be reconstituted into liposomes and are able to exchange electrons with the quinone pool. Proton pumping activity can be monitored at the single-enzyme level using pH-sensitive (ratiometric) dyes encapsulated in the liposome lumen (Figure 1A).
The method relies on the ability of ubiquinone (or other quinones), incorporated into the lipid bilayer, to exchange electrons with electrodes modified with a SAM18. The properties of the gold electrode, modified with a SAM, are very specific. Liposomes need to adsorb onto the SAM and the SAM needs to be thin enough to enable rapid electrochemical oxidation and reduction of the quinone pool in the liposomes. Importantly, the interaction between the electrode and liposome needs to be weak enough not to impair the integrity of the lipid membrane. Furthermore, depending on the details of the experimental system, it is desirable if the SAM prevents gold-catalyzed side reactions, such as oxygen reduction. Finally, the SAM needs to be stable within the potential window used. The use of ultra-flat gold electrodes modified with high-quality SAM of 6MH fulfills these requirements in the case of cytochrome bo3. Stripping gold electrodes from atomically-flat silicon wafers creates an ultra-smooth gold surface with a near ideal SAM as indicated by the impedance spectra. Please note that we form the SAM from 6MH in water. We previously reported on SAMs with 6MH in isopropanol16, but these SAMs exhibit higher double-layer capacitance values and impedance spectra that indicate a more heterogeneous surface (i.e., more defects) with a lower resistance. Furthermore, when SAMs from 6MH are prepared from a water solution, less background oxygen reduction (due to gold catalyzed oxygen reduction) is observed compared to SAMs formed from an ethanol/isopropanol solution. All these observations indicate that SAMs from 6MH in water solutions have a higher quality with less defects. Higher-quality SAMs can also be formed from thiol-compounds with longer alkane chains (e.g., 1-hydroxy-decane-thiol), but the electron transfer kinetics become slower as the SAM thickness increases.
During (spectro)electrochemistry, chloride ions should be avoided in the buffer solution since they may chemosorb on the gold surface at high potentials, possibly deteriorating the SAM quality. For the same reason, a chloride-free reference electrode should be preferred.
Another important point is the background permeability of the liposomes to protons, which should be low enough to allow proton accumulation as a result of enzymatic proton translocation on the timescale of experiment. The exact lipid composition may influence this permeability. In our work, we use polar E. coli lipid extracts supplemented with ubiquinone-10. Proton permeability has been shown to depend strongly on the fatty-acid chain length25, although this has been contradicted in studies with more complex lipid compositions26. Interestingly, ubiquinone has recently been shown to enhance membrane stability27. Finally, residual detergents from the protein reconstitution procedure can influence proton leakage and hence it is important to remove detergents as completely as possible using hydrophobic polystyrene microbeads, dilution followed by centrifugation and thorough rinsing of the electrochemical cell after the proteoliposomes are adsorbed on surface. If doubtful, liposome permeability can be verified by following the lumen pH change (via HPTS) in a response to an external pH jump28.
Single enzyme measurement require unilamelar liposomes and smaller liposomes are expected to show faster or higher pH changes as the lumen's volume decreases. To achieve this, polystyrene microbeads are added gradually in small amounts leading to slow rate of liposomes formation. In such conditions, small liposomes of homogeneous size are formed with an average diameter of 70 nm and a polydispersity index of 0.24 in our case14. HPTS also adsorbs on the polystyrene microbeads, reducing the capacity of polystyrene microbeads to adsorb detergent. Hence, excess polystyrene microbeads should be added to compensate for its loss. Treatment with polystyrene microbeads was observed to reduce the HPTS concentration in the lipid-protein suspension by about a quarter (from 5 mM to 3 – 4 mM). However, the actual HPTS concentration in the liposome's lumen might be higher since the liposomes are formed early on the treatment with polystyrene microbeads. Instead of HPTS, other pH-sensitive dyes can be used. However, it is important that the dye is membrane-impermeable and ratiometric. The latter avoids experimental errors due to photobleaching and varying amounts of encapsulated dye.
Simple calculations can be made to estimate whether, stochastically, most of the non-empty liposomes contain only one enzyme molecule. Assuming an average liposome diameter of 70 nm, a lipid molecular weight of 750 Da and a lipid area of 0.65 nm2 gives us a liposome mass of 5.2×10-17 g, i.e., 9.6×1013 liposomes per preparation (5 mg of lipid). At 0.1% of cytochrome bo3 (MW = 144 kDa), 2×1013 enzyme molecules are present, corresponding to a 0.2 protein:liposome ratio. Using a Poisson distribution, one may further calculate that the probability to find one enzyme molecule (17%) per liposome is ten times higher than probability to find more than one molecule (1.7%). More precise calculations can be made if the losses of enzyme and lipids during the reconstitution determined from corresponding assays are taken into account. The latter can be estimated from a protein assay and by measuring FDLL fluorescence of prepared liposomes.
Once the protocol is established and single enzyme traces are recorded, further modifications of the method is feasible depending on the aim. One might think about an enzyme inhibitor addition or introduction of an ionophore into the system. Care should be taken to verify that addition of solvents used to prepare ionophore or inhibitor stock do not influence the state of SAM or permeability of the liposomes.
The technique as described here is limited to a particular group of enzymes that are both membrane proton transporters and quinone-converting. The equipment and experimental conditions as used here were optimized for the enzymatic activity of cytochrome bo3. Here, the time resolution is 2-3 seconds, determined by the exposure time and the time it takes for the turret to change filters. The duration of the experiment is limited by photobleaching of fluorescent dye and, thus, by the light intensity. We have previously found average proton translocation rate of cytochrome bo3 to be 73 ± 2.2 protons/s using this technique, although activities down to 20 protons/s were detected. To use these techniques for enzymes with either more or less activity, image acquisition parameters need to be adapted (i.e., light intensity, exposure time and duration of experiment). In the future, this method can be extended towards other electron transport driving proton pumping enzymes, an example being mitochondrial complex I. Other enzymes might require different reconstitution protocols, and this would need to be optimized for each different transporter. Proton pumps that are not quinone-converting enzymes can also be studied, e.g., ATP-driven29, although in this case proton translocation cannot be triggered electrochemically, but addition of an initiator (e.g., ATP) is required. In the latter case, there is no need to use a gold-modified cover slip. Moreover, provided that a suitable membrane-impermeable and ion-sensitive fluorescent dye is used, this method can be extended to other ionic pumps, e.g., sodium and potassium.
The authors have nothing to disclose.
The authors acknowledge the BBSRC (BB/P005454/1) for financial support. NH was funded by the VILLUM Foundation Young Investigator Program.
6-Mercapto-1-hexanol (6MH) | Sigma | 451088 | 97% |
8-hydroxypyrene-1,3,6-trisulphonic acid (HPTS) | BioChemika | 56360 | |
Aluminium holder (Electrochemical cell) | Custom-made | 30x30x7 mm; inner diameter: 26 mm; hole diameter: 15 mm | |
Auxiliary electrode | platinum wire | ||
Chloroform | VWR Chemicals | 83627 | |
E.coli polar lipids | Avanti | 100600C | 25 mg/mL in chloroform |
Epoxy | EPO-TEK | 301-2FL | low fluorescence epoxy |
Fiji (ImageJ 1.52d) | Required plugins: StackReg and TurboReg (http://bigwww.epfl.ch/thevenaz/stackreg/) | ||
Filter cube ("ATTO633") | Chroma Technology Corporation | Ex: 620/60 nm; DM: 660 nm; Em: 700/75 nm | |
Filter cube ("HPTS1") | Chroma Technology Corporation | Ex: 470/20 nm; DM: 500 nm; Em: 535/48 nm | |
Filter cube ("HPTS2") | Chroma Technology Corporation | Ex: 410/300 nm; DM: 500 nm; Em: 535/48 nm | |
Filter cube ("Texas Red") | Chroma Technology Corporation | Ex: 560/55 nm; DM: 595 nm; Em: 645/75 nm | |
Fluorescent dye-labelled lipids (FDLL) | ThermoFisher Scientific | T1395MP | TexasRed-DHPC was used in this work (λexc 595 nm; λem 615 nm) |
Fluorescent dye-labelled lipids (FDLL) (alternative) | ATTO-TEC | AD 633-161 | ATTO633-DOPE can be used as alternative (λexc 630 nm; λem 651 nm) |
Gel filtration column | GE Healthcare | 28-9893-33 | HiLoad 16/600 Superdex 75 pg, for additional protein purification |
Glass coverslips | VWR International | 631-0172 | No 1.5 |
Glass syringe | Hamilton | 1725 RNR | 250 µL |
Glass vials | Scientific Glass Laboratories Ltd | T101/V1 | 1.75 mL capacity |
Gold | GoodFellow | 99.99% | |
Gramacidin | Sigma | G5002 | |
Mercury sulfate reference electrode | Radiometer (Hash) | E21M012 | |
Microcentrifuge | Eppendorf | Minispin NL040 | |
Microscope | Nikon | Eclipse Ti | |
Microscope Camera | Andor | Zyla 5.5 sCMOS | |
Microscope Lamp | Nikon | Intensilight C-HGFI | |
NIS-Elements AR 5.0.2 | Nikon | Microscope acquisition software | |
n-Octyl β-D-glucopyranoside | Melford Laboratories | B2007 | |
Nova 1.10 | Metrohm | Potentiostat control software | |
Objective | Nikon | Plan Apo λ 60x/1.4 oil | |
OriginPro 2017 | OriginLab | Plotting software | |
O-ring (Electrochemical cell) | Orinoko | Inner diameter: 16 mm; cross section: 1.5 mm | |
Plastic tubes | Eppendorf | 3810X | 1.5 mL |
Polystyrene microbeads | Bio-RAD | 152-3920 | Biobeads, 20-50 mesh |
Potentiostat | Metrohm Autolab | PGSTAT 128N | |
Potentiostat | CH Instruments | CHI604C | |
Scripting software | Matlab | R2017a | Required toolboxes: 'Image Processing Toolbox', 'Parallel Computing Toolbox', 'Curve Fitting Toolbox', 'System Identification Toolbox', 'Optimization Toolbox' |
Silicon wafers | IDB Technologies LTD | Si-C2 (N<100>P) | Ø 25 mm, 525 um thick |
Teflon cell (Electrochemical cell) | Custom-made | Outer diameter: 26 mm; inner diameter: 13.5 mm | |
Temple-Stripped Ultra-Flat Gold Surfaces | Platypus Technologies | AU.1000.SWTSG | Alternative ready-to-use ultra-flat gold surfaces (Thickness below 100 nm on demand) |
Thin micropipette tips | Sarstedt | 70.1190.100 | or similar gelloader tips 200 µL |
Ubiquinone-10 | Sigma | C-9538 | |
Ultracentrifuge | Beckman-Coulter | L-80XP | with Ti 45 rotor |
Ultrasonic bath | Fisher Scientific | FB15063 |