The authors acknowledge the BBSRC (BB/P005454/1) for financial support. NH was funded by the VILLUM Foundation Young Investigator Program.

1. Preparation of a Lipid/UQ-10/FDLL Mix
NOTE: E. coli lipids used for the liposomes&#8217; 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.
<ol>
	<li>Using a glass syringe, transfer 200 &#181;L of chloroform stock of lipid polar extract from Escherichia coli (25 mg/mL) into glass vials to make 5 mg aliquots.</li>
	<li>Add 50 &#181;L of 1 mg/mL ubiquinone-10 (UQ-10; in chloroform) to the lipids to make the final ratio of UQ-10: lipids 1:100 (1% w/w).</li>
	<li>Add 20 &#181;L of 1 mg/mL (0.4% w/w) of a long-wavelength fluorescent dye-labeled lipid (FDLL) to the lipids/UQ-10 mix.</li>
	<li>Homogenize the chloroform solution by short vortexing and evaporate most of the chloroform under a gentle nitrogen or argon flow. Remove the chloroform traces entirely by further evaporation under vacuum for at least 1 h. 
	NOTE: Lipid aliquots can be stored under an inert atmosphere at -20 &#176;C for several months.</li>
</ol>
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
<ol>
	<li>Add 312.5 &#181;L of 40 mM MOPS-KOH/60 mM K2SO4, pH 7.4, to one aliquot of lipids/UQ-10/FDLL dry mix (5 mg, step 1.4) and mix with vortex till the lipid film is fully resuspended, followed by 2 min of treatment in an ultrasonic bath.</li>
	<li>Add 125 &#181;L of 25 mM 8-hydroxypyrene-1,3,6-trisulphonic acid (HPTS), a pH-sensitive fluorescent dye that needs to be encapsulated inside the liposomes.</li>
	<li>Add 137.5 &#181;L of 250 mM n-octyl &#946;-D-glucopyranoside (OGP) surfactant, mix using vortex and sonicate in an ultrasonic water bath for 10 min to ensure all lipids are solubilized into surfactant micelles. Transfer the dispersion into a 1.5 mL plastic tube. 
	NOTE: Cloudy suspension of lipids should become transparent after the solubilization with surfactant.</li>
	<li>Add the required amount of cytochrome bo3 (see the note below) and add ultrapure water to make a total volume of 50 &#181;L (cytochrome bo3 solution plus water). Incubate at 4 &#176;C for 10 min on a roller mixer. 
	NOTE: Typical amount for single enzyme conditions is 0.1 &#8211; 0.2% (w/w protein-to-lipid, i.e. 5 &#8211; 10 &#181;g of protein), although it can be increased till 1 &#8211; 2% (50 &#8211; 100 &#181;g of protein) if the goal is to observe only the electrochemical activity (see below). As a negative control, liposomes without cytochrome bo3 can be prepared.</li>
	<li>Weigh 2x 50 mg and 2x 100 mg of polystyrene microbeads into four 1.5 mL-tube caps and close with paraffin film to prevent drying. 
	NOTE: Before use, polystyrene microbeads should be washed with methanol, water and stored in water according to the manufacturer&#8217;s manual. 
	NOTE: If a water/microbead mixture is used, polystyrene microbeads can conveniently be transferred to the cap with a micropipette with a wide tip. Water can then be removed from the microbeads using a micropipette with a thin tip.</li>
	<li>Add the 1st 50 mg of polystyrene microbeads into the reconstitution mixture (step 2.4) by putting the cap with polystyrene microbeads on the 1.5 mL-plastic tube with dispersion and performing a short spin for a couple of seconds. Incubate at 4 &#176;C on a roller mixer for polystyrene microbeads to adsorb the surfactant for 30 min.
	<ol>
		<li>Repeat the additions of polystyrene microbeads and incubations as follows: add 50 mg of microbeads for 60 min of incubation; add 100 mg of microbeads for 60 min of incubation; and add 100 mg of microbeads for 120 min of incubation. 
		NOTE: The solution above the settled microbeads will turn translucent during step 2.6 as proteoliposomes are formed.</li>
	</ol>
	</li>
	<li>Separate the proteoliposome solution from the polystyrene microbeads using a micropipette with a thin tip. Dilute the dispersion in 90 mL of 20 mM MOPS-KOH/30 mM K2SO4, pH 7.4 (MOPS buffer) and transfer in a Ti45 ultracentrifuge tube.</li>
	<li>Ultracentrifuge the dispersion using Type 45 Ti rotor at 125,000 x g (at rmax) for 1 h to pellet the proteoliposomes. 
	NOTE: Smaller centrifuge tubes can be used although the dilution in large buffer volume helps to reduce the concentration of the non-encapsulated HPTS in the final suspension.</li>
	<li>Discard the supernatant, rinse the pellets with 20 mM MOPS-KOH/30 mM K2SO4, pH 7.4 buffer (without resuspending the pellet). After discarding the rinsing buffer, re-suspend the proteoliposomes in 500 &#181;L of MOPS buffer by pipetting it back and forth with thin tip micropipette. Then transfer to a 1.5 mL plastic tube.</li>
	<li>Centrifuge the suspension for 5 min at 12,000 x g to remove the debris. Transfer the supernatant (reconstituted proteoliposomes) into a new vial.</li>
	<li>Store the reconstituted proteoliposomes dispersion at 4 &#176;C overnight and use within 2 days.</li>
</ol>
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).
<ol>
	<li>Glue up to 9 glass cover slips (0.17 mm thick) onto the evaporated gold surface using bi-component low-fluorescence epoxy. Cure the glue at 80 &#176;C for 4 h.</li>
	<li>Just before modification with a self-assembled monolayer (step 4), detach the glass cover slips from the silicon wafers with a blade. Due to the thinness of the cover slips, take care when detaching the cover slips to not crack or break the glass slides.</li>
</ol>
4. Modification of the Gold Surface with Self-Assembled Monolayer (SAM)
<ol>
	<li>Prepare 5 mL of water solution of 1 mM 6-mercaptohexanol (6MH).</li>
	<li>Dip freshly detached gold-coated cover slips (step 3.1) into 6MH solution and leave at 20 &#8211; 25 &#176;C overnight (&#62;16 h) to form the SAM. 
	NOTE: Thiol solutions have an unpleasant smell, so a closed vessel should be used when incubating the gold-coated cover slip.</li>
	<li>The next day, remove the gold-coated cover slip from the 6MH solution, wash briefly with water or methanol and then with isopropanol. Dry under a gentle gas flow.</li>
</ol>
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).
<ol>
	<li>Assemble the gold-coated cover slip in a spectro-electrochemical cell (see Figure 1). Make contact to the gold with a flat wire outside an area defined by a rubber O-ring.</li>
	<li>Add 2 mL of the electrolyte buffer solution and place the reference and auxiliary electrodes in the cell. 
	NOTE: As further discussed in the Discussion section, references electrodes without chloride are preferred to prevent formation of Au(I)Cl during electrochemistry. Here, a Hg/Hg2SO4 (sat. K2SO4) reference electrode is used and potentials are given versus Standard Hydrogen Electrode (SHE) using 0.658 V vs SHE for the Hg/Hg2SO4 (sat. K2SO4).</li>
	<li>Run electrochemical impedance spectroscopy (0.1 Hz &#8211; 100 kHz) at open cell potential (OCP) potential to assess the quality of SAM. Convert impedance values to admittance and divide by 2&#960;&#969; to plot a Cole-Cole plot, where &#969; is the frequency (see the following references16,17,18 for details on converting and interpreting impedance spectra). 
	NOTE: Compact and dense SAMs of 6MH should give a close to semi-circle Cole-Cole plot and give capacitance values in the range 2.5-3.0 &#181;F/cm2 (Figure 2A). If significant deviation from semi-circle shape or out-of-range capacitance values are obtained, change the electrode.</li>
	<li>Run blank cyclic voltammograms (CVs) with scan rates 100 and 10 mV/s in the potential region -0.3 &#8211; 0.8 V. A typical CV is shown on (Figure 2B, dashed line). 
	NOTE: This should demonstrate almost pure capacitive behavior and an absence of significant faradaic current, even under ambient oxygen conditions as used here.</li>
	<li>Add proteoliposomes (0.5 mg/mL final lipids concentration, 1-2% (w/w) ratio of cytochrome bo3 to lipid) to the electrochemical cell and mix slightly with a pipette. Wait until the adsorption of proteoliposomes on the electrode surface is finished (30-60 min at room temperature). 
	NOTE: Cyclic voltammograms (CVs) at 10 mV/s can be run during the proteoliposomes adsorption to follow the process. The adsorption is finished when consecutive CVs stop changing. Information about CV techniques can be found in the following textbooks.19,20</li>
	<li>Wash the cell by changing the buffer solution at least 10 times but avoid leaving the electrode surface completely dry.</li>
	<li>Run the electrochemical impedance spectroscopy at OCP (Figure 2A, blue line) to confirm the SAM on the gold electrode remains unchanged and CVs with scan rates 10 and 100 mV/s to observe catalytic ubiquinol oxidation (and oxygen reduction) by cytochrome bo3 at onset potentials of electrochemical quinone reduction about 0 V vs SHE) (Figure 2B).</li>
</ol>
6. Detection of Enzymatic Proton Pumping by Fluorescence Microscopy
<ol>
	<li>Modify the gold electrode as in step 5 but using 100x less proteoliposomes compared to step 5.5 (i.e., 5 &#181;g/mL). For single enzyme studies, reduce the cytochrome bo3 to lipid ratio to 0.1-0.2% (w/w). 
	NOTE: Liposomes will sparsely adsorb on the electrode surface enabling single vesicle monitoring by fluorescence microscopy. Under these single-enzyme conditions, the amount of enzyme immobilized on the electrode surface is insufficient for the observation of a catalytic current.</li>
	<li>Place the electrochemical cell on the oil objective (60X) of an inverted fluorescence microscope with a drop of immersion oil. Using appropriate filters for FDLL fluorescence, focus on the electrode surface. Single liposomes should appear as bright spots at the diffraction limit of the microscope/objective. Take an image of FDLL fluorescence.</li>
	<li>Switch to the one of HPTS fluorescence filter sets on the microscope to verify that HPTS fluorescence is clearly visible and distinguishable from the background (at the chosen exposure time, see step 6.4). Increase the light intensity if it is&#160;not the case.</li>
	<li>Program the microscope software to perform a timed image acquisition by alternating two HPTS filter sets (menu Applications | Define/Run ND Acquisition). Set the delay between image acquisitions at minimum. In this experiment, use a 1 s exposure and 0.3 s delay (due to the turret movement). 
	NOTE: The ratio between fluorescence intensities at these two channels will be later converted to pH inside the liposomes at each time point. The duration of the acquisition can vary according to the expected pH change rate; 5 min is used in this article.</li>
	<li>Adjust the settings of the potentiostat to change the potential during the image acquisition. For example, in this experiment, use the following sequence: 0 &#8211; 60 s: no potential applied (i.e. OCP); 60 &#8211; 180 s: -0.2 V (vs SHE); 180 &#8211; 300 s: 0.4 V (vs SHE). In this case, only the applied potential in the second phase (-0.2 V vs SHE) is sufficient to efficiently reduce the quinone pool.</li>
	<li>Run simultaneously the timed images acquisition (microscope) and the potential sequence (potentiostat) by manually starting both measurements at the same time. 
	NOTE: The experiment can be repeated on the same electrode several time by moving the microscope stage to a different area on the surface. The delay between the acquisitions should be at least 5-10 min to insure complete pH equilibration of liposomes on the surface. Different durations and potential patterns can be applied depending on the need, although imaging time is limited due to photobleaching of HPTS during acquisition.</li>
</ol>
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).
<ol>
	<li>Use Fiji to separate time lapse file, align images and save as separate channels and timeframes.
	<ol>
		<li>Open time lapse file using the Bioformats importer provided within Fiji (macro command: run (&#34;Bio-Formats Importer&#34;)).</li>
		<li>Use the plugin StackReg to align each frame to the first one to account for possible stage movements or thermal drift occurred during the acquisition (macro command: run(&#34;StackReg &#34;, &#34;transformation=Translation&#34;)).</li>
		<li>Save separate uncompressed image files for each channel and each time step in TIFF format into a single folder.</li>
		<li>Extract the exact time stamps for each frame from the time lapse file metadata and save them as a CSV-file in the same folder as the images. Alternatively, extract time stamps using the acquisition software and save manually into a CSV-file using a spreadsheet software. 
		NOTE: The folder with images and time stamps is now ready to be analyzed by the scripting software. Steps 7.1.1. &#8211; 7.1.4. can be automatized using a Fiji script (a script written in Python for batch processing of time lapse files is provided, use &#8220;Ctrl-Shift-N&#8221; (in Windows), then File | Open to load the script).</li>
	</ol>
	</li>
	<li>Use scripting software for automatized processing of the images. Load the provided code to the software and click Run to end. When prompted, select the folder containing the images from step 7.1. 
	NOTE: A script for analysis is provided as mlx-format live script, with extensive comments, to identify single liposomes, fit them to 2D-Gaussian function, filter them and quantify the pH values at each point of time. The following sub-steps are executed automatically by the script.
	<ol>
		<li>Load all time frames images for a single experiment into the memory.</li>
		<li>Average all images for a single channel (select the channel with the highest fluorescence intensity) and use the averaged image to identify all maximums that might correspond to the single liposomes.</li>
		<li>Fit the identified maximums on the averaged image to the 2D-Gaussian function and save the resulted fit parameters for each liposome (see Figure 4A for fitted liposome example).</li>
		<li>Filter the maximums according to the expected single liposomes criteria, such as size, circularity and intensity. Reject liposomes that are poorly fitted due to low signal to noise, closely neighboring liposome or being too close to the image edge.</li>
		<li>Load time stamps from the external file and fit each filtered liposome to the 2D-gaussian function on each time frame image separately. 
		NOTE: The use of parallel programming and multicore CPU can significantly enhance the calculation speed at this stage.</li>
		<li>Filter the liposomes again according to the similar criteria as in step 7.2.4 but applied to each time frame.</li>
		<li>On each time step, define the fluorescence intensity ratio of a liposome as the ratio of volumes enclosed by fitted 2D-Gaussian function at two channels. Calculate the pH values from the ratio of intensities determined from the two HPTS channels and using a calibration curve (step 8). Plot resulting pH-time curves for the liposomes and observe their pH change when the potential is applied.</li>
	</ol>
	</li>
</ol>
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.
<ol>
	<li>Prepare an electrode with sparsely adsorbed liposomes (without cytochrome bo3 as described in Steps 5 and 6.1).</li>
	<li>Add 2 &#181;L of 0.1 mg/mL gramicidin solution in ethanol to 2 mL of buffer to create the concentration 100 ng/mL.</li>
	<li>Capture two fluorescence images for the two HPTS channels.</li>
	<li>Change the pH of the cell by addition of small aliquots of 1 M HCl or 1 M H2SO4. Measure the pH of the buffer with a standard pH meter and capture two fluorescence images for the two HPTS channels.</li>
	<li>Repeat steps 8.4 for the pH range 6 to 9.</li>
	<li>Using the algorithm from 7.2, fit, filter and calculate the average HPTS fluorescence ratio of individual liposomes at every pH. Take the HPTS ratio average over all liposomes.</li>
	<li>Fit the resulting pH-ratio dependence to the following equation: 
	<img alt="Equation" src="/files/ftp_upload/58896/58896eq1.jpg" />, where pKa, Ra and Rb are fitting parameters.</li>
	<li>Use pKa, Ra and Rb to convert the HPTS ratio of individual liposomes in step 7.2.7. to pH values.</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> MATLAB code for the analysis of proton transport activity in single liposomes. , (2017).</li><li>Li, M., Tuma, R., Jeuken, L. J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Time-resolved fluorescence microscopic data (traces) of individual lipid vesicles with proton transport activity. , (2017).</li><li>Paula, S., Volkov, A. G., Van Hoek, A. N., Haines, T. H., Deamer, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Permeation+of+protons,+potassium+ions,+and+small+polar+molecules+through+phospholipid+bilayers+as+a+function+of+membrane+thickness.">Permeation of protons, potassium ions, and small polar molecules through phospholipid bilayers as a function of membrane thickness.</a> Biophysical Journal. 70 (1), 339-348 (1996).</li><li>Brookes, P. S., Hulbert, A. J., Brand, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+proton+permeability+of+liposomes+made+from+mitochondria)+inner+membrane+phospholipids:+No+effect+of+fatty+acid+composition.">The proton permeability of liposomes made from mitochondria) inner membrane phospholipids: No effect of fatty acid composition.</a> Biochimica et Biophysica Acta (BBA) - Biomembranes. 1330 (2), 157-164 (1997).</li><li>Eriksson, E. K., Agmo Hern&#225;ndez, V., Edwards, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+ubiquinone-10+on+the+stability+of+biomimetic+membranes+of+relevance+for+the+inner+mitochondrial+membrane.">Effect of ubiquinone-10 on the stability of biomimetic membranes of relevance for the inner mitochondrial membrane.</a> Biochimica et Biophysica Acta - Biomembranes. 1860 (5), 1205-1215 (2018).</li><li>Seneviratne, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+reconstitution+method+for+integral+membrane+proteins+in+hybrid+lipid-polymer+vesicles+for+enhanced+functional+durability.">A reconstitution method for integral membrane proteins in hybrid lipid-polymer vesicles for enhanced functional durability.</a> Methods. , 1-8 (2018).</li><li>Veshaguri, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+observation+of+proton+pumping+by+a+eukaryote+P-type+ATPase.">Direct observation of proton pumping by a eukaryote P-type ATPase.</a> Science. 351 (6280), 1-6 (2016).</li></ol>

The authors have nothing to disclose.

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 electron...

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.

<table>
	<tbody>
		<tr>
			<td>6-Mercapto-1-hexanol (6MH)</td>
			<td>Sigma</td>
			<td>451088</td>
			<td>97%</td>
		</tr>
		<tr>
			<td>8-hydroxypyrene-1,3,6-trisulphonic acid (HPTS)</td>
			<td>BioChemika</td>
			<td>56360</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminium holder (Electrochemical cell)</td>
			<td>Custom-made</td>
			<td></td>
			<td>30x30x7 mm; inner diameter: 26 mm; hole diameter: 15 mm</td>
		</tr>
		<tr>
			<td>Auxiliary electrode</td>
			<td></td>
			<td></td>
			<td>platinum wire</td>
		</tr>
		<tr>
			<td>Chloroform</td>
			<td>VWR Chemicals</td>
			<td>83627</td>
			<td></td>
		</tr>
		<tr>
			<td>E.coli polar lipids</td>
			<td>Avanti</td>
			<td>100600C</td>
			<td>25 mg/mL in chloroform</td>
		</tr>
		<tr>
			<td>Epoxy</td>
			<td>EPO-TEK</td>
			<td>301-2FL</td>
			<td>low fluorescence epoxy</td>
		</tr>
		<tr>
			<td>Fiji (ImageJ 1.52d)</td>
			<td></td>
			<td></td>
			<td>Required plugins: StackReg and TurboReg (http://bigwww.epfl.ch/thevenaz/stackreg/)</td>
		</tr>
		<tr>
			<td>Filter cube (&#34;ATTO633&#34;)</td>
			<td>Chroma Technology Corporation</td>
			<td></td>
			<td>Ex: 620/60 nm; DM: 660 nm; Em: 700/75 nm</td>
		</tr>
		<tr>
			<td>Filter cube (&#34;HPTS1&#34;)</td>
			<td>Chroma Technology Corporation</td>
			<td></td>
			<td>Ex: 470/20 nm; DM: 500 nm; Em: 535/48 nm</td>
		</tr>
		<tr>
			<td>Filter cube (&#34;HPTS2&#34;)</td>
			<td>Chroma Technology Corporation</td>
			<td></td>
			<td>Ex: 410/300 nm; DM: 500 nm; Em: 535/48 nm</td>
		</tr>
		<tr>
			<td>Filter cube (&#34;Texas Red&#34;)</td>
			<td>Chroma Technology Corporation</td>
			<td></td>
			<td>Ex: 560/55 nm; DM: 595 nm; Em: 645/75 nm</td>
		</tr>
		<tr>
			<td>Fluorescent dye-labelled lipids (FDLL)</td>
			<td>ThermoFisher Scientific</td>
			<td>T1395MP</td>
			<td>TexasRed-DHPC was used in this work (&#955;exc 595 nm; &#955;em 615 nm)</td>
		</tr>
		<tr>
			<td>Fluorescent dye-labelled lipids (FDLL) (alternative)</td>
			<td>ATTO-TEC</td>
			<td>AD 633-161</td>
			<td>ATTO633-DOPE can be used as alternative (&#955;exc 630 nm; &#955;em 651 nm)</td>
		</tr>
		<tr>
			<td>Gel filtration column</td>
			<td>GE Healthcare</td>
			<td>28-9893-33</td>
			<td>HiLoad 16/600 Superdex 75 pg, for additional protein purification</td>
		</tr>
		<tr>
			<td>Glass coverslips</td>
			<td>VWR International</td>
			<td>631-0172</td>
			<td>No 1.5</td>
		</tr>
		<tr>
			<td>Glass syringe</td>
			<td>Hamilton</td>
			<td>1725 RNR</td>
			<td>250 &#181;L</td>
		</tr>
		<tr>
			<td>Glass vials</td>
			<td>Scientific Glass Laboratories Ltd</td>
			<td>T101/V1</td>
			<td>1.75 mL capacity</td>
		</tr>
		<tr>
			<td>Gold</td>
			<td>GoodFellow</td>
			<td></td>
			<td>99.99%</td>
		</tr>
		<tr>
			<td>Gramacidin</td>
			<td>Sigma</td>
			<td>G5002</td>
			<td></td>
		</tr>
		<tr>
			<td>Mercury sulfate reference electrode</td>
			<td>Radiometer (Hash)</td>
			<td>E21M012</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge</td>
			<td>Eppendorf</td>
			<td>Minispin NL040</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Nikon</td>
			<td>Eclipse Ti</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Camera</td>
			<td>Andor</td>
			<td>Zyla 5.5 sCMOS</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Lamp</td>
			<td>Nikon</td>
			<td>Intensilight C-HGFI</td>
			<td></td>
		</tr>
		<tr>
			<td>NIS-Elements AR 5.0.2</td>
			<td>Nikon</td>
			<td></td>
			<td>Microscope acquisition software</td>
		</tr>
		<tr>
			<td>n-Octyl &#946;-D-glucopyranoside</td>
			<td>Melford Laboratories</td>
			<td>B2007</td>
			<td></td>
		</tr>
		<tr>
			<td>Nova 1.10</td>
			<td>Metrohm</td>
			<td></td>
			<td>Potentiostat control software</td>
		</tr>
		<tr>
			<td>Objective</td>
			<td>Nikon</td>
			<td>Plan Apo &#955; 60x/1.4 oil</td>
			<td></td>
		</tr>
		<tr>
			<td>OriginPro 2017</td>
			<td>OriginLab</td>
			<td></td>
			<td>Plotting software</td>
		</tr>
		<tr>
			<td>O-ring (Electrochemical cell)</td>
			<td>Orinoko</td>
			<td></td>
			<td>Inner diameter: 16 mm; cross section: 1.5 mm</td>
		</tr>
		<tr>
			<td>Plastic tubes</td>
			<td>Eppendorf</td>
			<td>3810X</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td>Polystyrene microbeads</td>
			<td>Bio-RAD</td>
			<td>152-3920</td>
			<td>Biobeads, 20-50 mesh</td>
		</tr>
		<tr>
			<td>Potentiostat</td>
			<td>Metrohm Autolab</td>
			<td>PGSTAT 128N</td>
			<td></td>
		</tr>
		<tr>
			<td>Potentiostat</td>
			<td>CH Instruments</td>
			<td>CHI604C</td>
			<td></td>
		</tr>
		<tr>
			<td>Scripting software</td>
			<td>Matlab</td>
			<td>R2017a</td>
			<td>Required toolboxes: &#39;Image Processing Toolbox&#39;, &#39;Parallel Computing Toolbox&#39;, &#39;Curve Fitting Toolbox&#39;, &#39;System Identification Toolbox&#39;, &#39;Optimization Toolbox&#39;</td>
		</tr>
		<tr>
			<td>Silicon wafers</td>
			<td>IDB Technologies LTD</td>
			<td>Si-C2 (N&#60;100&#62;P)</td>
			<td>&#216; 25 mm, 525 um thick</td>
		</tr>
		<tr>
			<td>Teflon cell (Electrochemical cell)</td>
			<td>Custom-made</td>
			<td></td>
			<td>Outer diameter: 26 mm; inner diameter: 13.5 mm</td>
		</tr>
		<tr>
			<td>Temple-Stripped Ultra-Flat Gold Surfaces</td>
			<td>Platypus Technologies</td>
			<td>AU.1000.SWTSG</td>
			<td>Alternative ready-to-use ultra-flat gold surfaces (Thickness below 100 nm on demand)</td>
		</tr>
		<tr>
			<td>Thin micropipette tips</td>
			<td>Sarstedt</td>
			<td>70.1190.100</td>
			<td>or similar gelloader tips 200 &#181;L</td>
		</tr>
		<tr>
			<td>Ubiquinone-10</td>
			<td>Sigma</td>
			<td>C-9538</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultracentrifuge</td>
			<td>Beckman-Coulter</td>
			<td>L-80XP</td>
			<td>with Ti 45 rotor</td>
		</tr>
		<tr>
			<td>Ultrasonic bath</td>
			<td>Fisher Scientific</td>
			<td>FB15063</td>
			<td></td>
		</tr>
	</tbody>
</table>

single liposome measurements for the study of proton-pumping membrane enzymes using electrochemistry and fluorescent microscopy

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.

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 dia...

Watch this Scientific Journal Video about Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy at JoVE.com

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Mesures de Liposome unique pour l’étude des Enzymes membranaires Proton-pompage, à l’aide de l’électrochimie et microscopie fluorescente

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 ...

single-liposome-measurements-for-study-proton-pumping-membrane

Research

JoVE Journal

Biochemistry

7.3K vues.  University of Leeds. Cette méthode permet l’étude de l’activité de transport des protons des enzymes de la membrane respiratoire comme le cytochrome BO3 dans un environnement naturel de bicposition lipidique. Les principaux avantages de cette technique enzymatique unique est la possibilité de commencer et d’arrêter les réactions enzymatiques à tout moment en utilisant l’électrochimie. À l’aide d’une seringue en verre transférer 200 microlitres de stock chloroforme d’extrait polaire lipidiq...

Vidéo: Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

High-throughput Screening of Carbohydrate-degrading Enzymes Using Novel Insoluble Chromogenic Substrate Assay Kits

Carbohydrates active enzymes (CAZymes) have multiple roles in vivo and are widely used for industrial processing in the biofuel, textile, detergent, paper and food industries. A deeper understanding of CAZymes is important from both fundamental biology and industrial standpoints. Vast numbers of CAZymes exist in nature (especially in microorganisms) and hundreds of thousands have been cataloged and described in the carbohydrate active enzyme database (CAZy). However, the rate of discovery of putative enzymes has outstripped our ability to biochemically characterize their activities. One reason for this is that advances in genome and transcriptome sequencing, together with associated bioinformatics tools allow for rapid identification of candidate CAZymes, but technology for determining an enzyme's biochemical characteristics has advanced more slowly. To address this technology gap, a novel high-throughput assay kit based on insoluble chromogenic substrates is described here. Two distinct substrate types were produced: Chromogenic Polymer Hydrogel (CPH) substrates (made from purified polysaccharides and proteins) and Insoluble Chromogenic Biomass (ICB) substrates (made from complex biomass materials). Both CPH and ICB substrates are provided in a 96-well high-throughput assay system. The CPH substrates can be made in four different colors, enabling them to be mixed together and thus increasing assay throughput. The protocol describes a 96-well plate assay and illustrates how this assay can be used for screening the activities of enzymes, enzyme cocktails, and broths.

A high-throughput assay for enzyme screening is described. This multiplexed ready-to-use assay kit comprises of pre-chosen Chromogenic Polymer Hydrogel (CPH) substrates and complex Insoluble Chromogenic Biomass (ICB) substrates. Target enzymes are polysaccharide degrading endo-enzymes and proteases.

Criblage à haut débit d&#39;Enzymes Glucides dégradant Utilisation Novel Insoluble chromogénique Substrat Assay Kits

Carbohydrates active enzymes (CAZymes) have multiple roles in vivo and are widely used for industrial processing in the biofuel, textile, detergent, paper ...

Recherche

Biochimie

Single-molecule Super-resolution Imaging of Phosphatidylinositol 4,5-bisphosphate in the Plasma Membrane with Novel Fluorescent Probes

Phosphoinositides in the cell membrane are signaling lipids with multiple cellular functions. Phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) is a determinant phosphoinositide of the plasma membrane (PM), and it is required to modulate ion channels, actin dynamics, exocytosis, endocytosis, intracellular signaling, and many other cellular processes. However, the spatial organization of PI(4,5)P2 in the PM is controversial, and its nanoscale distribution is poorly understood due to the technical limitations of research approaches. Here by utilizing single molecule localization microscopy and the Pleckstrin Homology (PH) domain based dual color fluorescent probes, we describe a novel method to visualize the nanoscale distribution of PI(4,5)P2 in the PM in fixed membrane sheets as well as live cells.

PI(4,5)P2 regulates various cellular functions, but its nanoscale organization in the cell plasma membrane is poorly understood. By labeling PI(4,5)P2 with a dual-color fluorescent probe fused to the Pleckstrin Homology domain, we describe a novel approach to study the PI(4,5)P2 spatial distribution in the plasma membrane at the nanometer scale.

Simple-molécule Super-résolution Imaging de phosphatidylinositol 4,5-bisphosphate dans la membrane plasmique avec nouveaux Fluorescent Probes

Phosphoinositides in the cell membrane are signaling lipids with multiple cellular functions. Phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) is a ...

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Utilisation de Scaffold Liposomes à Reconstituer interactions protéine-protéine Lipid-proximal<em&gt; In vitro</em

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

High-resolution Single Particle Analysis from Electron Cryo-microscopy Images Using SPHIRE

SPHIRE (SPARX for High-Resolution Electron Microscopy) is a novel open-source, user-friendly software suite for the semi-automated processing of single particle electron cryo-microscopy (cryo-EM) data. The protocol presented here describes in detail how to obtain a near-atomic resolution structure starting from cryo-EM micrograph movies by guiding users through all steps of the single particle structure determination pipeline. These steps are controlled from the new SPHIRE graphical user interface and require minimum user intervention. Using this protocol, a 3.5 &#197; structure of TcdA1, a Tc toxin complex from Photorhabdus luminescens, was derived from only 9500 single particles. This streamlined approach will help novice users without extensive processing experience and a priori structural information, to obtain noise-free and unbiased atomic models of their purified macromolecular complexes in their native state.

This paper presents a protocol for processing cryo-EM images using the software suite SPHIRE. The present protocol can be applied for nearly all single particle EM projects that target near-atomic resolution.

Analyse de particules à haute résolution à partir de microscopie à électrons à l&#39;aide d&#39;images utilisant SPHIRE

SPHIRE (SPARX for High-Resolution Electron Microscopy) is a novel open-source, user-friendly software suite for the semi-automated processing of single ...

High Precision FRET at Single-molecule Level for Biomolecule Structure Determination

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule level in multiparameter fluorescence detection (MFD) mode is presented here. MFD maximizes the usage of all &#34;dimensions&#34; of fluorescence to reduce photophysical and experimental artifacts and allows for the measurement of interdye distance with an accuracy up to ~1 &#197; in rigid biomolecules. This method was used to identify three conformational states of the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor to explain the activation of the receptor upon ligand binding. When comparing the known crystallographic structures with experimental measurements, they agreed within less than 3 &#197; for more dynamic biomolecules. Gathering a set of distance restraints that covers the entire dimensionality of the biomolecules would make it possible to provide a structural model of dynamic biomolecules.

A protocol for high-precision FRET experiments at the single molecule level is presented here. Additionally, this methodology can be used to identify three conformational states in the ligand-binding domain of the N-methyl-D-aspartate (NMDA) receptor. Determining precise distances is the first step towards building structural models based on FRET experiments.

FRET de haute précision au niveau d&#39;une molécule pour la détermination de la structure biomoléculaire

A protocol on how to perform high-precision interdye distance measurements using F&#246;rster resonance energy transfer (FRET) at the single-molecule ...

Protein Film Infrared Electrochemistry Demonstrated for Study of H2 Oxidation by a [NiFe] Hydrogenase

Understanding the chemistry of redox proteins demands methods that provide precise control over redox centers within the protein. The technique of protein film electrochemistry, in which a protein is immobilized on an electrode surface such that the electrode replaces physiological electron donors or acceptors, has provided functional insight into the redox reactions of a range of different proteins. Full chemical understanding requires electrochemical control to be combined with other techniques that can add additional structural and mechanistic insight. Here we demonstrate a technique, protein film infrared electrochemistry, which combines protein film electrochemistry with infrared spectroscopic sampling of redox proteins. The technique uses a multiple-reflection attenuated total reflectance geometry to probe a redox protein immobilized on a high surface area carbon black electrode. Incorporation of this electrode into a flow cell allows solution pH or solute concentrations to be changed during measurements. This is particularly powerful in addressing redox enzymes, where rapid catalytic turnover can be sustained and controlled at the electrode allowing spectroscopic observation of long-lived intermediate species in the catalytic mechanism. We demonstrate the technique with experiments on E. coli hydrogenase 1 under turnover (H2 oxidation) and non-turnover conditions.

Protein Film Infrared Electrochemistry Demonstrated for Study of H2 Oxidation by a [NiFe] Hydrogenase

Here, we describe a technique, protein film infrared electrochemistry, which allows immobilized redox proteins to be studied spectroscopically under direct electrochemical control at a carbon electrode. Infrared spectra of a single protein sample can be recorded at a range of applied potentials and under a variety of solution conditions.

Protéine Film infrarouge électrochimie témoigne pour l’étude de l’oxydation de2 H une hydrogénase [NiFe]

Understanding the chemistry of redox proteins demands methods that provide precise control over redox centers within the protein. The technique of protein ...

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

Un dosage de spectroscopie par Fluorescence Fluctuation d’Interactions protéine-protéine à Contacts cellule-cellule

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

Production of Dynein and Kinesin Motor Ensembles on DNA Origami Nanostructures for Single Molecule Observation

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and others. Many of these functions require motors to operate in ensembles. Despite a wealth of knowledge about the mechanisms of individual cytoskeletal motors, comparatively less is known about the mechanisms and emergent behaviors of motor ensembles, examples of which include changes to ensemble processivity and velocity with changing motor number, location, and configuration. Structural DNA nanotechnology, and the specific technique of DNA origami, enables the molecular construction of well-defined architectures of motor ensembles. The shape of cargo structures as well as the type, number and placement of motors on the structure can all be controlled. Here, we provide detailed protocols for producing these ensembles and observing them using total internal reflection fluorescence microscopy. Although these techniques have been specifically applied for cytoskeletal motors, the methods are generalizable to other proteins that assemble in complexes to accomplish their tasks. Overall, the DNA origami method for creating well-defined ensembles of motor proteins provides a powerful tool for dissecting the mechanisms that lead to emergent motile behavior.

The goal of this protocol is to form ensembles of molecular motors on DNA origami nanostructures and observe the ensemble motility using total internal reflection fluorescence microscopy.

Production d'ensembles moteurs d'adn et de kinésine sur des nanostructures d'origami d'ADN pour l'observation d'une seule molécule

Cytoskeletal motors are responsible for a wide variety of functions in eukaryotic cells, including mitosis, cargo transport, cellular motility, and ...

Visualizing Diffusional Dynamics of Gold Nanorods on Cell Membrane using Single Nanoparticle Darkfield Microscopy

Analyzing the diffusional dynamics of nanoparticles on cell membrane plays a significant role in better understanding the cellular uptake process and provides a theoretical basis for the rational design of nano-medicine delivery. Single particle tracking (SPT) analysis could probe the position and orientation of individual nanoparticles on cell membrane, and reveal their translational and rotational states. Here, we show how to use traditional dark-field microscopy to monitor the dynamics of gold nanorods (AuNRs) on live cell membrane. We also show how to extract the location and orientation of AuNRs using ImageJ and MATLAB, and how to characterize the diffusive states of AuNRs. Statistical analysis of hundreds of particles show that single AuNRs perform Brownian motion on the surface of U87 MG cell membrane. However, individual long trajectory analysis shows that AuNRs have two distinctly different types of motion states on the membrane, namely long-range transport and limited-area confinement. Our SPT methods can be potentially used to study the surface or intracellular particle diffusion in different biological cells and can become a powerful tool for investigations of complex cellular mechanisms.

Here, we show the use of traditional dark-field microscopy to monitor the dynamics of gold nanorods (AuNRs) on cell membrane. The location and orientation of single AuNRs are detected using ImageJ and MATLAB, and the diffusive states of AuNRs are characterized by single particle tracking analysis.

Visualisation de la dynamique diffusionnelle des nanorods d’or sur la membrane cellulaire à l’aide d’une microscopie darkfield nanoparticule unique

Analyzing the diffusional dynamics of nanoparticles on cell membrane plays a significant role in better understanding the cellular uptake process and ...

Single Cell Analysis Of Transcriptionally Active Alleles By Single Molecule FISH

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk population methods (Northern blot, PCR, and RNAseq) that measure mRNA levels lack the ability to provide information on cell-to-cell variation in responses. Precise single cell and allelic visualization and quantification is possible via single molecule RNA fluorescence in situ hybridization (smFISH). RNA-FISH is performed by hybridizing target RNAs with labeled oligonucleotide probes. These can be imaged in medium/high throughput modalities, and, through image analysis pipelines, provide quantitative data on both mature and nascent RNAs, all at the single cell level. The fixation, permeabilization, hybridization and imaging steps have been optimized in the lab over many years using the model system described herein, which results in successful and robust single cell analysis of smFISH labeling. The main goal with sample preparation and processing is to produce high quality images characterized by a high signal-to-noise ratio to reduce false positives and provide data that are more accurate. Here, we present a protocol describing the pipeline from sample preparation to data analysis in conjunction with suggestions and optimization steps to tailor to specific samples.&#160;

Single molecule RNA fluorescence in situ hybridization (smFISH) is a method to accurately quantify levels and localization of specific RNAs at the single cell level. Here, we report our validated lab protocols for wet-bench processing, imaging and image analysis for single cell quantification of specific RNAs.

Analyse unicellulaire d’allèles transcriptionnellement actifs par molécule unique FISH

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk ...