This work was supported by the Slovenian Research Agency (project J2-9764 and programme P2-0249). This video represents supplementary material for the "Electroporation-based Technologies and Treatments" scientific workshop and postgraduate course, organized by the Faculty of Electrical Engineering at the University of Ljubljana, Slovenia.

I. Loading the cells with Cell trackers CMFDA and CMRA
<ol>
 <li>Experiments were performed on previously prepared cells of mouse melanoma cell line (B16-F1). Cells are grown in two separated 25 cm2 culture flasks (TPP, ZDA) to 70-80 % confluence in DMEM culture medium (Dulbecco's modified Eagle's medium) supplemented with 10 % fetal bovine serum, 0.15 mg/ml L-glutamine, 16 mg/ml gentamicin (all from Sigma-Aldrich, Germany), 200 units/ml crystacillin (Pliva, Croatia), and incubated in 5% CO2 at 37&deg;C.</li>
 <li>Prepare two 10 mM stock solutions of Cell trackers (Invitrogen, USA) by adding 10.76 &mu;l and 9 &mu;l (for Green CMFDA and for Orange CMRA, respectively) of DMSO (Sigma-Aldrich, Germany) to 50 &mu;g of the dye in the original Invitrogen vial. The stock solution can be stored in a refrigerator at 4&deg;C for few months. Before starting the experiments, warm up the solution until the crystals of DMSO dissolve.</li>
 <li>Prepare bicarbonate-free Krebs-Hepes buffer (130 mM NaCl, 4.7 mM KCl, 1.2 mM MgSO4 , 1.2 mM KH2PO4, 11.7 mM D-glucose, 1.3 mM CaCl2, 10 mM HEPES, pH 7.4). In two 15 ml Eppendorf tubes separately mix 2.1 &mu;l of each stock solution (10 mM CMFDA or CMRA, respectively) in 3 ml of bicarbonate-free Krebs-Hepes buffer. This yields a "loading solution" containing approximately 7 &mu;M CMFDA (or CMRA).</li>
 <li>Rinse cells twice with bicarbonate-free Krebs-Hepes buffer and then insert loading solutions into the flasks. Incubate cells for 30 minutes in 5% CO2 at 37 &deg;C. During this first incubation reagents pass freely through cell membranes, but once inside the cell, the reagents are transformed into cell-impermeant fluorescent reaction products.</li>
 <li>After first incubation, rinse and incubate cells with culture medium for another two hours in 5% CO2 at 37&deg;C.</li>
 <li>Trypsinize cells in both flasks (loaded with CMFDA and CMRA) and mix red and green cells together in a ratio 1:1 in a 50 ml centrifuge tube (TPP, ZDA). Adjust cell concentration to 5 x 106 cells/ml by dilution with DMEM medium or by concentration with centrifuge. Place a 20 &mu;l drop of cell suspension in each well of 24 multiwell plate (TPP, ZDA). Incubate cells in 5 % CO2 at 37 &deg;C for 20 min to allow them to slightly attach to the surface of the well and establish cell contacts.</li>
</ol>
II. Electrofusion
<ol>
 <li>Prepare isoosomolar potassium phosphate buffer (10 mM KH2PO4, 10 mM K2HPO4, 1mM MgCl2, 250 mM sucrose) and hypoosmolar potassium phosphate buffer (10 mM KH2PO4, 10 mM K2HPO4, 1mM MgCl2, 75 mM sucrose).</li>
 <li>Place the multiwell with cells onto the microscope stage, position the electrodes at the bottom of the well and connect them to the pulse generator.</li>
 <li>Wash the cells with 1 ml of isoosomolar potassium phosphate buffer. Add 350 &mu;l of hypoosmolar potassium phosphate buffer in order to induce cell swelling. The buffer should cover the electrodes.</li>
 <li>Leave cells in hypoosmolar buffer for 2 minutes before applying electric pulses. During this time influx of water molecules into the cells due to osmotic imbalance between the interior and the exterior of the cells cause an increase of cell volume. Electric pulses should be applied when cells are close to their maximum volumes, before regulatory volume decrease start.</li>
 <li>To achieve optimal electrofusion and maintain cell viability, optimal parameters of electric pulses should be used. These depend on cell line used [1]. In this experiment a train of 8 rectangular pulses (each with duration of 100 &mu;s at 1 Hz) is applied to each sample, using electroporation device (in our case Cliniporator, IGEA, Italy). The pulses are delivered to two parallel Pt/Ir wire electrodes with 0.8 mm diameter and 5 mm distance between them, creating an electric field of approximately 1200 V/cm between the electrodes in each well, except for the control well.</li>
 <li>Leave the cells undisturbed for 10 minutes after pulse delivery. Determine the fusion yield by means of fluorescent and phase contrast microscopy.</li>
</ol>
III. Image acquisition and determination of the fusion yield
<ol>
 <li>The cells are observed using a fluorescence microscope (in our case Zeiss AxioVert 200, Zeiss, Germany) equipped with an objective (x20) and a cooled CCD camera (VisiCam 1280, Visitron, Germany). The images are acquired in MetaMorph 7.1.1 (Molecular Devices, USA), but other similar acquisition software can also be used. CMFDA is excited with a monochromator (Polychrome IV, Visitron, Germany) at 492 nm and CMRA at 548 nm. The fluorescence of CMFDA and CMRA is acquired using two emission filters, one centered at 535 nm (HQ535/30m, for CMFDA) and the other centered at 510 nm (D605/55m, for CMRA, both Chroma, USA). The use of dichroic mirror (Q515LP) prevented the channel cross talk.</li>
 <li>Acquire three images (phase contrast, red and green fluorescence) for five randomly chosen fields in each well. Create three channel images from each image triplet. In such image fluorescing cytoplasm can be seen together with the cell membranes. Fused cells can thus be easily determined [Figure 1].</li>
 <li>Count all three types of cells (red, green and dually fluorescent) in each three channel image. Determine the percentage of dually fluorescent cells by dividing the number of dually fluorescent cells with the number of all cells in each image. Fusion yield is defined as the percentage of dually fluorescent cells multiplied by 2 since half of fused cells are not detected (when cells of the same color fuse). </li>
</ol>
Representative Results
<img src="/files/ftp_upload/1991/1991fig1.jpg" alt="Figure 1" /> Figure 1. Three channel microscopy image of B16F1 cells after electrofusion: phase contrast, CMRA fluorescence (excitation at 548 nm) and CMFDA fluorescence (excitation at 492 nm), objective magnification 20x

<ol>
	<li>&#268;emazar, M., Jarm, T., Miklav&#269;i&#269;, D., Ma&#269;ek-Lebar, A., Ihan, A., Kopitar, N. A., Ser&#353;a, G. <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+electric-field+intensity+on+electropermeabilization+and+electrosensitivity+of+various+tumor-cell+lines+in+vitro.">Effect of electric-field intensity on electropermeabilization and electrosensitivity of various tumor-cell lines in vitro.</a> Electro. Magnetobiol. 17, 263-272 (1998).</li><li>Trontelj, K., Reber&#353;ek, M., Kandu&#353;er, M., &#269;urin &#353;erbec, V., Miklav&#269;i&#269;, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+bulk+cell+electrofusion+in+vitro+for+production+of+human-mouse+heterohybridoma+cells.">Optimization of bulk cell electrofusion in vitro for production of human-mouse heterohybridoma cells.</a> Bioelectrochemistry. 74, 124-129 (2008).</li><li>Rols, M. -. P., Teissi&#233;, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulation+of+electrically+induced+permeabilization+and+fusion+of+Chinese+hamster+ovary+cells+by+osmotic+pressure.">Modulation of electrically induced permeabilization and fusion of Chinese hamster ovary cells by osmotic pressure.</a> Biochemistry. 29, 4561-4567 (1990).</li><li>Neil, G., Zimmermann, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrofusion.">Electrofusion.</a> Methods in Enzymology. 220, 174-196 .</li><li>Abidor, I. G., Li, L. -. H., Hui, S. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Studies+of+cell+pellets:+II.+Osmotic+properties,+electroporation,+and+related+phenomena:+Membrane+interactions.">Studies of cell pellets: II. Osmotic properties, electroporation, and related phenomena: Membrane interactions.</a> Biophysical journal. 67, 427-435 (1994).</li><li>Jaroszeski, M. J., Gilbert, R., Fallon, P. G., Heller, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanically+facilitated+cell-cell+electrofusion.">Mechanically facilitated cell-cell electrofusion.</a> Biophys J. 67 (4), 1574-1581 (1994).</li><li>Ramos, C., Bonefant, D., Teissi&#233;, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+hybridization+by+electrofusion+on+filters.">Cell hybridization by electrofusion on filters.</a> Analytical biochemistry. 302, 213-219 (2002).</li><li>Cao, Y., Yang, J., Yin, Z. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Study+of+high-throughput+cell+electrofusion+in+a+microelectrode-array+chip.">Study of high-throughput cell electrofusion in a microelectrode-array chip.</a> Microfluid Nanofluid. 5, 669-675 (2008).</li><li>U&#353;aj, M., Trontelj, M., Kandu&#353;er, M., Miklav&#269;i&#269;, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+size+dynamics+and+viability+of+cells+exposed+to+hypotonic+treatment+and+electroporation+for+electrofusion+optimization.">Cell size dynamics and viability of cells exposed to hypotonic treatment and electroporation for electrofusion optimization.</a> Radiol. Oncol. 43, 108-119 (2009).</li><li>Teissi&#233;, J., Ramos, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlation+between+electric+field+pulse+induced+long-lived+permeabilization+and+fusogenicity+in+cell+membranes.">Correlation between electric field pulse induced long-lived permeabilization and fusogenicity in cell membranes.</a> Biophys. J. 74, 1889-1898 (1998).</li></ol>

No conflicts of interest declared.

The ability of cell membranes to fuse non-specifically, e.g., by external electric fields, is important for biotechnology, medicine and research in biology. Such nonspecific fusion enables production of highly valuable hybrid cells and their products, such as monoclonal antibodies, and provides information about fundamental mechanisms of fusion [2]. Electrofusion is a potentially very effective method since it can be properly adjusted to different types of cells. Electrofusion is achieved when cells in close physical contact are brought into their fusogenic state (prone to fusion) by means of high-voltage electric pulses. The efficiency of electrofusion depends on various parameters that affect two parts of the electrofusion process. First part of the electrofusion process is achievement of the close physical contact between cells, which can be obtained with different methods [3-8]. Adherence method (growing cells to confluence) can be used efficiently due to spontaneously established cell contacts in large zones between cells; however, it produces very large fused cells with many nuclei. We are using the modified adherence method, where smaller cells (with 2 to 5 nuclei), which are more likely to survive and proliferate, are obtained (Figure 1). Contact between cells also benefit from osmotic swelling of cells, due to osmotic treatment used in the experiment [9]. Second part of the electrofusion process is the achievement of the fusogenic state of the cell membranes. Fusogenic state correlates well with electropermeabilized state of the membrane (cells are non-specifically permeabilized to molecules that normally cannot pass through intact membrane) and is governed by the same parameters of the electric pulses (amplitude, length, number and frequency) [10]. The values of electrical parameters needed for optimal electroporation [1] and electrofusion differ between different cells and depend on cells size and their biological properties. Electrical parameters thus need to be optimized for different cell lines, which are used as fusion partners, to obtain fusion.

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		<th>Comment</th>
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		<tbody>
		
<tr>
<td>CMRA</td>
<td>Reagent</td>
<td>Invitrogen</td>
<td>C34551</td>
<td>Cytosolic fluorescent dye</td>
</tr>
<tr>
<td>CMFDA</td>
<td>Reagent</td>
<td>Invitrogen</td>
<td>C7025</td>
<td>Cytosolic fluorescent dye</td>
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<td>DMSO</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>D2650</td>
<td>&nbsp;</td>
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<td>DMEM </td>
<td>Reagent</td>
<td>Sigma-Aldrich </td>
<td>D5671</td>
<td>Dulbecco's modified Eagle's medium</td>
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<td>Fetal calf serum</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>F4135</td>
<td>&nbsp;</td>
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<td>L-glutamine</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>G7513</td>
<td>&nbsp;</td>
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<td>crystacillin</td>
<td>Reagent</td>
<td>Pliva</td>
<td>625110</td>
<td>antibiotic</td>
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<td>gentamicin</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>G1397</td>
<td>antibiotic</td>
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<td>Hepes</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>H0887</td>
<td>&nbsp;</td>
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<td>KH2PO4</td>
<td>Reagent</td>
<td>Merck </td>
<td>A124873 927</td>
<td>&nbsp;</td>
<tr>
<td>KH2PO4</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>4248</td>
<td>&nbsp;</td>
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<td>MgCl2</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>M-8266</td>
<td>&nbsp;</td>
<tr>
<td>NaCl</td>
<td>Reagent</td>
<td>Fluka</td>
<td>71382</td>
<td>&nbsp;</td>
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<td>KCl</td>
<td>Reagent</td>
<td>Merck</td>
<td>A154336 908</td>
<td>&nbsp;</td>
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<td>MgSO4</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>M2643</td>
<td>&nbsp;</td>
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<td>D-glucose</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>G8270</td>
<td>&nbsp;</td>
<tr>
<td>CaCl2</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>C4901</td>
<td>&nbsp;</td>
<tr>
<td>sucrose</td>
<td>Reagent</td>
<td>Sigma-Aldrich</td>
<td>16104</td>
<td>&nbsp;</td>
<tr>
<td>Electric pulse generator</td>
<td>Tool</td>
<td>Igea</td>
<td>&nbsp;</td>
<td>Cliniporator VITAE</td>
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<td>Multiwell plate</td>
<td>Tool</td>
<td>TPP</td>
<td>92424</td>
<td>&nbsp;</td>
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<td>50 ml centrifuge tube</td>
<td>Tool</td>
<td>TPP</td>
<td>91050</td>
<td>&nbsp;</td>
<tr>
<td>15 ml centrifuge tube</td>
<td>Tool</td>
<td>TPP</td>
<td>91015</td>
<td>&nbsp;</td>
<tr>
<td>25 cm2 culture flask</td>
<td>Tool</td>
<td>TPP</td>
<td>90026</td>
<td>&nbsp;</td>
<tr>
<td>Electrodes</td>
<td>Tool</td>
<td>Custom made</td>
<td>&nbsp;</td>
<td>Pt/Ir</td>
</tr>		</tbody>
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cell electrofusion visualized with fluorescence microscopy

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves application of short high-voltage electric pulses to cells that are in close contact. Application of short, high-voltage electric pulses causes destabilization of cell plasma membranes. Destabilized membranes are more permeable for different molecules and also prone to fusion with any neighboring destabilized membranes. Electrofusion is thus a convenient method to achieve a non-specific fusion of very different cells in vitro. In order to obtain fusion, cell membranes, destabilized by electric field, must be in a close contact to allow merging of their lipid bilayers and consequently their cytoplasm. In this video, we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method. In this method, cells are allowed to attach only slightly to the surface of the well, so that medium can be exchanged and cells still preserve their spherical shape. Fusion visualization is assessed by pre-labeling of the cytoplasm of cells with different fluorescent cell tracker dyes; half of the cells are labeled with orange CMRA and the other half with green CMFDA. Fusion yield is determined as the number of dually fluorescent cells divided with the number of all cells multiplied by two.

Watch this Scientific Journal Video about Cell Electrofusion Visualized with Fluorescence Microscopy at JoVE.com

Cell Electrofusion Visualized with Fluorescence Microscopy

In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves ...

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14.0K Views.  University of Ljubljana. In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Video: Cell Electrofusion Visualized with Fluorescence Microscopy

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Dopamine Release at Individual Presynaptic Terminals Visualized with FFNs

The nervous system transmits signals between neurons via neurotransmitter release during synaptic vesicle fusion. To observe neurotransmitter uptake and release from individual presynaptic terminals directly, we designed fluorescent false neurotransmitters as substrates for the synaptic vesicle monoamine transporter. Using these probes to image dopamine release in the striatum, we made several observations pertinent to synaptic plasticity. We found that the fraction of synaptic vesicles releasing neurotransmitter per stimulus was dependent on the stimulus frequency. A kinetically distinct "reserve" synaptic vesicle population was not observed under these experimental conditions. A frequency-dependent heterogeneity of presynaptic terminals was revealed that was dependent in part on D2 dopamine receptors, indicating a mechanism for frequency-dependent coding of presynaptic selection.

Hui Zhang and Niko G. Gubernator contributed equally to this work.

A new means to measure neurotransmission optically using fluorescent dopamine analogs.

The nervous system transmits signals between neurons via neurotransmitter release during synaptic vesicle fusion. To observe neurotransmitter uptake and ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Protein-protein Interactions Visualized by Bimolecular Fluorescence Complementation in Tobacco Protoplasts and Leaves

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular fluorescence complementation (BiFC) is an in vivo method to monitor such interactions in plant cells. In the presented protocol the investigated candidate proteins are fused to complementary halves of fluorescent proteins and the respective constructs are introduced into plant cells via agrobacterium-mediated transformation. Subsequently, the proteins are transiently expressed in tobacco leaves and the restored fluorescent signals can be detected with a confocal laser scanning microscope in the intact cells. This allows not only visualization of the interaction itself, but also the subcellular localization of the protein complexes can be determined. For this purpose, marker genes containing a fluorescent tag can be coexpressed along with the BiFC constructs, thus visualizing cellular structures such as the endoplasmic reticulum, mitochondria, the Golgi apparatus or the plasma membrane. The fluorescent signal can be monitored either directly in epidermal leaf cells or in single protoplasts, which can be easily isolated from the transformed tobacco leaves. BiFC is ideally suited to study protein-protein interactions in their natural surroundings within the living cell. However, it has to be considered that the expression has to be driven by strong promoters and that the interaction partners are modified due to fusion of the relatively large fluorescence tags, which might interfere with the interaction mechanism. Nevertheless, BiFC is an excellent complementary approach to other commonly applied methods investigating protein-protein interactions, such as coimmunoprecipitation, in vitro pull-down assays or yeast-two-hybrid experiments.

Formation of protein complexes in vivo can be visualized by bimolecular fluorescence complementation. Interaction partners are fused to complementary parts of fluorescent tags and transiently expressed in tobacco leaves, resulting in a reconstituted fluorescent signal upon close proximity of the two proteins.

Many proteins interact transiently with other proteins or are integrated into multi-protein complexes to perform their biological function. Bimolecular ...

Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule imaging assays which allows adhesion of single virions to glass surfaces with specificity. This preparation is based on grafting the surface of the glass with a mixture of PLL-g-PEG and PLL-g-PEG-Biotin, adding a layer of avidin, and finally creating virion anchors through attachment of biotinylated virus specific antibodies. We have applied this technique across a range of experiments including atomic force microscopy (AFM) and super-resolution fluorescence imaging. This sample preparation method results in a control adhesion of the virions to the surface.

The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

Light Sheet Fluorescence Microscopy of Plant Roots Growing on the Surface of a Gel

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the observation of the whole organ with a high spatial- as well as temporal resolution over prolonged periods of time, which may cause photo-toxic effects. This protocol shows a plant sample preparation method for light-sheet microscopy, which is characterized by mounting the plant vertically on the surface of a gel. The plant is mounted in such a way that the roots are submerged in a liquid medium while the leaves remain in the air. In order to ensure photosynthetic activity of the plant, a custom-made lighting system illuminates the leaves. To keep the roots in darkness the water surface is covered with sheets of black plastic foil. This method allows long-term imaging of plant organ development in standardized conditions.

This protocol shows a plant sample preparation method for light-sheet microscopy. The setup is characterized by mounting the plant vertically on the surface of a gel and letting it grow in controlled bright conditions. This allows long-term observation of plant organ development in standardized conditions.

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the ...

Open-source Single-particle Analysis for Super-resolution Microscopy with VirusMapper

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm allows for the routine imaging of nanoscale biological complexes and processes. This increase in resolution also means that methods popular in electron microscopy, such as single-particle analysis, can readily be applied to super-resolution fluorescence microscopy. By combining this analytical approach with super-resolution optical imaging, it becomes possible to take advantage of the molecule-specific labeling capacity of fluorescence microscopy to generate structural maps of molecular elements within a metastable structure. To this end, we have developed a novel algorithm &#8212; VirusMapper &#8212; packaged as an easy-to-use, high-performance, and high-throughput ImageJ plugin. This article presents an in-depth guide to this software, showcasing its ability to uncover novel structural features in biological molecular complexes. Here, we present how to assemble compatible data and provide a step-by-step protocol on how to use this algorithm to apply single-particle analysis to super-resolution images.

This manuscript uses the Fiji-based open-source software package VirusMapper to apply single-particle analysis to super-resolution microscopy images in order to generate precise models of nanoscale structure.

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm ...