The authors acknowledge Asifa Akhtar and Angelika Rambold (both Max Planck Institute for Immunobiology and Epigenetics) for providing HeLa cells.

NOTE: All the cell lines mentioned here were cultured in high glucose Dulbecco&#39;s Modified Eagle&#39;s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), glutamine, penicillin/streptomycin, and pyruvate. Equilibrate all media and supplements to be used on the day of labeling and imaging by warming them up to 37 &#176;C in an incubator set to 5% CO2. All cell culture work including labeling takes place in sterile conditions under a laminar flow hood.
1. Live Cell Labeling
<ol>
	<li>One day before the labeling procedure, culture 4 x 105 HeLa cells in 2 mL of medium in a 35 mm Petri dish with #1.5 glass bottom. Multi-well chambered slides can be used, but volumes should be adjusted proportionally to the volume of the well. Dilute the cell suspension to have approximately 50,000 cells/mL and seed 2 mL of the cell suspension in a 35 mm Petri dish.</li>
	<li>Prepare the following dilutions of SYBR Gold (SG) commercial stock solution: 1:5,000. Prepare Mitotracker Deep Red (far red stain) or Mitotracker CMXRos Red (red stain) (Table of Materials; commercial stock diluted 1:2,000) in phenol red-free culture medium. Prepare the same set dilutions of picoGreen as for SG. 
	NOTE: The solutions described in step 1.2 are the 2x labeling solutions. Protect all the solutions containing fluorescent dyes from light as much as possible.</li>
	<li>Wash the adherent cells with 2 mL PBS once. Add 1 ml phenol red-free culture medium to the 35 mm Petri dish. For an 8-well chambered slide, use a volume equal to 1/2 of the total well capacity (e.g., add 125 &#181;L if the total well capacity is 250 &#181;L).</li>
	<li>Add 1 mLof the 2x labeling solution prepared in step 1.2 to 35 mm Petri dish. For an 8-well chambered slide, use a volume equal to 1/2 of the total well capacity. Incubate the cells for 30 min at 37 &#176;C under 5% CO2. 
	NOTE: Do not incubate the cells with SG labeling solution for longer than 1 h, because longer incubation will cause labeling of nuclear DNA.</li>
	<li>Carefully aspirate the medium containing the fluorescent dyes and wash the cells once with PBS.</li>
	<li>Add phenol red-free cell culture medium and keep the cells in the dark in a CO2 incubator at 37 &#176;C until imaging.</li>
	<li>Image living cells on an SR-SIM microscope equipped with an incubation unit (see below).</li>
</ol>
2. SR-SIM Image Acquisition
<ol>
	<li>Install a stage-top incubator on the microscope stage. Set the desired temperature and CO2 concentration (e.g., 37 &#176;C and 5% for mammalian cells) and keep warm for at least 1 h before starting image acquisition.</li>
	<li>Switch on all the components of the SR-SIM microscope, including the lasers, and leave them to warm up for at least 1 h.</li>
	<li>Choose a high magnification, high numerical aperture (NA) immersion objective (e.g., 100x 1.46 NA oil) recommended for SR-SIM by the microscope manufacturer.</li>
	<li>Install a chambered slide or 35 mm dish with labeled cells (from step 1.6) on the incubated microscope stage.</li>
	<li>Locate the area of interest on the sample, preferably with oculars. To achieve the best SR-SIM imaging quality, choose the cells that are well attached to the glass.</li>
	<li>Use a back-tinned high-end electron multiplying charge-coupled device (EM-CCD) camera to acquire SR-SIM images.</li>
	<li>Using the image acquisition software, set a high EM gain recommended for the camera used (e.g., 300).</li>
	<li>Before acquiring the time lapse series for nucleoid tracking, acquire a two-color SR-SIM image of the same field of view: one channel for mitochondrial staining and another one for SG. Set the first color channel appropriate to the mitochondrial stain used (e.g., for a far red mitochondrial stain, set the excitation at 633 nm or longer and a 650 nm longpass emission filter). For the SG signal, set the excitation at 488 nm and a 500&#8211;550 nm bandpass emission filter.</li>
	<li>Using the software, set the lowest laser power possible for both 488 nm and far red stain lines. 
	NOTE: A typical setting is 1% laser output power adjusted by an acousto-optical tunable filter (AOTF).</li>
	<li>If the SIM microscope acquires channels only sequentially (single-camera setup), then switch off the far-red stain detection channel.</li>
	<li>Set the acquisition of a single focal plane by switching off the Z-stack acquisition by unticking the Z-stack box in the software.</li>
	<li>Set the shortest possible EM-CCD camera exposure time.</li>
	<li>Set three rotations of the grid rather than five rotations.</li>
	<li>To increase the frame rate, in the Acquisition tab of the software, set the camera to read the data only from the central area of camera sensor instead of &#34;Full Chip&#34;. 
	NOTE: For example, switching from reading an area of 1,000 x 1,000 pixels full sensor to reading only 256 x 256 pixels allows a reduction in camera exposure time from 50 ms to 13.4 ms and the total frame time from 1.8 s to 1.2 s. If only a central part of the camera sensor is read, then the field of view will be very small for a 63x or 100x objective typically used for SR-SIM. A shorter exposure time will reduce the signal-to-noise ratio of the images.</li>
	<li>Optimize laser power and camera exposure time: acquire SR-SIM two-dimensional (2D) images of labeled cells at several laser power values (e.g., 0.5%, 1%, 1.5%, 2%) and several exposure times (e.g., 13.4 ms, 25 ms, 50 ms).</li>
	<li>Process the raw SIM datasets (see step 3.1).</li>
	<li>Choose laser power and camera exposure times that yield SIM images with bright spots in the mitochondria (i.e., the nucleoids) with little or no artifacts generated by SIM processing. Inspect the areas outside of the mitochondria (e.g., 1% power of the 488 nm solid-state laser and 25 ms exposure of EM-CCD camera).</li>
	<li>Start acquisition of the time lapse series using the settings optimized in steps 2.15&#8211;2.17.</li>
</ol>
3. Data Processing and Analysis
<ol>
	<li>Process raw SIM datasets with the structured illumination module of a suitable software (Table of Materials): choose the Auto checkbox for SIM processing parameters. For automatic processing of multiple files, use the batch processing tool, deselect Use Current for Batch, click Run Batch and select multiple files for SIM processing.</li>
	<li>Convert SIM-processed time series datasets to ims format with Imaris file converter software (i.e., the version corresponding to the version of Imaris software).</li>
	<li>Open a converted file in the software (version 8.4.1 or later) that has a license for the Lineage (or Track) module.</li>
	<li>Start the &#34;Spots&#34; creation wizard by clicking Add New Spots icon. A wizard for creation will be launched.</li>
	<li>Choose the Create tab of the wizard.</li>
	<li>On the first step of the wizard, click Track Spots (over time) and proceed to the second step of the wizard.</li>
	<li>Set Estimated XY Diameter to 0.1-0.15 &#181;m, click Background Subtraction and proceed to the third step.</li>
	<li>Adjust the threshold in the filter &#34;Quality&#34; by dragging the vertical line in the histogram so that the majority of the nucleoids are detected as &#34;spots&#34;, while the artifacts are not detected (i.e., the detected spots are marked as balls overlaid on the image). Check if this is the case on each frame and readjust the threshold if necessary. Proceed to the fourth and then to the fifth step of the wizard.</li>
	<li>Choose the Autoregressive Motion algorithm. Set Max Distance to 0.5 &#181;m. Set Max Gap Size to 0.</li>
	<li>Look at each frame in the time series and check if any false tracks are drawn and if any gaps between tracks are introduced (tracks built by the wizard with current settings are instantly marked as lines overlaid on the image). Adjust &#34;Max Distance&#34; if necessary. Proceed to the sixth step of the wizard. Choose the Track Duration filter and set a threshold of 3&#8211;5 s.</li>
	<li>If the currently detected spots or tracks are not optimal, go back to the preceding steps of the wizard using navigation buttons (in the bottom of wizard window) needed to fine-tune any parameter for spots or tracks creation.</li>
	<li>When the fine-tuned parameters allow the software to detect all the spots and build tracks correctly, click the green arrow navigation button in the wizard to confirm creation of the tracks.</li>
	<li>Extract the statistics of the tracks by clicking the Statistics icon window. Choose the needed statistics parameters (Detailed and Average Values tabs under Statistics) and export the values as csv files by clicking the Floppy Drive icon for quantification and visualization. 
	NOTE: Maximum track speed, mean track speed, track length, and track displacement are examples of parameters which are important.</li>
	<li>If the image dataset needs to be presented or published, then create snapshots and/or video files representing the time lapse series with optionally overlaid tracks using the Snapshot or Animation tools.</li>
</ol>
4. Confocal Image Acquisition
<ol>
	<li>Install a stage-top incubator on the microscope stage, set desired temperature and CO2 concentration (i.e., 37 &#176;C and 5% for mammalian cells) and keep warm for at least 1 h before starting image acquisition.</li>
	<li>Switch on all the components of a confocal laser scanning microscope, including the lasers, and leave them to warm up for at least 1 h.</li>
	<li>Choose a desired middle to high magnification objective and spatial sampling according to Nyquist criteria.</li>
	<li>For the SG channel, set the excitation at 488 nm and a detection range of 500&#8211;550 nm. For the mitochondrial stain channel, set to 561 nm and 570&#8211;630 nm emission for a red dye, or 633 nm excitation and &#62;650 nm emission for a far red dye.</li>
	<li>Set the lowest possible laser power (typically 1% or less) that permits acquisition of the signals with PMT detector gains of 700 mV.</li>
	<li>Acquire two-color confocal images of the cells, where the mitochondrial nucleoids are detected in the &#34;SG channel&#34; and mitochondria are detected in the &#34;red&#34; or &#34;far red&#34; channel.</li>
</ol>

<ol>
	<li>Kaufman, B. A., 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=The+mitochondrial+transcription+factor+TFAM+coordinates+the+assembly+of+multiple+DNA+molecules+into+nucleoid-like+structures.">The mitochondrial transcription factor TFAM coordinates the assembly of multiple DNA molecules into nucleoid-like structures.</a> Molecular Biology of the Cell. 18 (9), 3225-3236 (2007).</li><li>Farge, G., 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=Protein+sliding+and+DNA+denaturation+are+essential+for+DNA+organization+by+human+mitochondrial+transcription+factor+A.">Protein sliding and DNA denaturation are essential for DNA organization by human mitochondrial transcription factor A.</a> Nature Communications. 3, 1013 (2012).</li><li>Bogenhagen, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+DNA+nucleoid+structure.">Mitochondrial DNA nucleoid structure.</a> Biochimica et Biophysica Acta. 1819 (9-10), 914-920 (2012).</li><li>Kang, D., Kim, S. H., Hamasaki, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+transcription+factor+A+(TFAM):+roles+in+maintenance+of+mtDNA+and+cellular+functions.">Mitochondrial transcription factor A (TFAM): roles in maintenance of mtDNA and cellular functions.</a> Mitochondrion. 7 (1-2), 39-44 (2007).</li><li>Tauber, J., 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=Distribution+of+mitochondrial+nucleoids+upon+mitochondrial+network+fragmentation+and+network+reintegration+in+HEPG2+cells.">Distribution of mitochondrial nucleoids upon mitochondrial network fragmentation and network reintegration in HEPG2 cells.</a> International Journal of Biochemistry &amp; Cell Biology. 45 (3), 593-603 (2013).</li><li>Garrido, N., 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=Composition+and+dynamics+of+human+mitochondrial+nucleoids.">Composition and dynamics of human mitochondrial nucleoids.</a> Molecular Biology of the Cell. 14 (4), 1583-1596 (2003).</li><li>Pernas, L., Scorrano, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mito-Morphosis:+Mitochondrial+Fusion,+Fission,+and+Cristae+Remodeling+as+Key+Mediators+of+Cellular+Function.">Mito-Morphosis: Mitochondrial Fusion, Fission, and Cristae Remodeling as Key Mediators of Cellular Function.</a> Annual Review of Physiology. 78, 505-531 (2016).</li><li>Caielli, 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=Oxidized+mitochondrial+nucleoids+released+by+neutrophils+drive+type+I+interferon+production+in+human+lupus.">Oxidized mitochondrial nucleoids released by neutrophils drive type I interferon production in human lupus.</a> Journal of Experimental Medicine. 213 (5), 697-713 (2016).</li><li>Okayama, S., Ohta, K., Higashi, R., Nakamura, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correlative+light+and+electron+microscopic+observation+of+mitochondrial+DNA+in+mammalian+cells+by+using+focused-ion+beam+scanning+electron+microscopy.">Correlative light and electron microscopic observation of mitochondrial DNA in mammalian cells by using focused-ion beam scanning electron microscopy.</a> Microscopy (Oxf). 63 (Suppl 1), i35 (2014).</li><li>Alan, L., Spacek, T., Jezek, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Delaunay+algorithm+and+principal+component+analysis+for+3D+visualization+of+mitochondrial+DNA+nucleoids+by+Biplane+FPALM/dSTORM.">Delaunay algorithm and principal component analysis for 3D visualization of mitochondrial DNA nucleoids by Biplane FPALM/dSTORM.</a> European Biophysics Journal. 45 (5), 443-461 (2016).</li><li>Kukat, C., 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=Super-resolution+microscopy+reveals+that+mammalian+mitochondrial+nucleoids+have+a+uniform+size+and+frequently+contain+a+single+copy+of+mtDNA.">Super-resolution microscopy reveals that mammalian mitochondrial nucleoids have a uniform size and frequently contain a single copy of mtDNA.</a> Proceedings of the National Academy of Sciences of the United States of America. 108 (33), 13534-13539 (2011).</li><li>Kukat, C., Larsson, N. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=mtDNA+makes+a+U-turn+for+the+mitochondrial+nucleoid.">mtDNA makes a U-turn for the mitochondrial nucleoid.</a> Trends in Cell Biology. 23 (9), 457-463 (2013).</li><li>Betzig, E., 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=Imaging+intracellular+fluorescent+proteins+at+nanometer+resolution.">Imaging intracellular fluorescent proteins at nanometer resolution.</a> Science. 313 (5793), 1642-1645 (2006).</li><li>Rust, M. 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A., 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=Superresolution+fluorescence+imaging+of+mitochondrial+nucleoids+reveals+their+spatial+range,+limits,+and+membrane+interaction.">Superresolution fluorescence imaging of mitochondrial nucleoids reveals their spatial range, limits, and membrane interaction.</a> Molecular and Cellular Biology. 31 (24), 4994-5010 (2011).</li><li>Lewis, S. C., Uchiyama, L. F., Nunnari, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ER-mitochondria+contacts+couple+mtDNA+synthesis+with+mitochondrial+division+in+human+cells.">ER-mitochondria contacts couple mtDNA synthesis with mitochondrial division in human cells.</a> Science. 353 (6296), aaf5549 (2016).</li><li>Dellinger, M., Geze, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+mitochondrial+DNA+in+living+animal+cells+with+fluorescence+microscopy.">Detection of mitochondrial DNA in living animal cells with fluorescence microscopy.</a> Journal of Microscopy. 204 (Pt 3), 196-202 (2001).</li><li>Ban-Ishihara, R., Ishihara, T., Sasaki, N., Mihara, K., Ishihara, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+nucleoid+structure+regulated+by+mitochondrial+fission+contributes+to+cristae+reformation+and+release+of+cytochrome+c.">Dynamics of nucleoid structure regulated by mitochondrial fission contributes to cristae reformation and release of cytochrome c.</a> Proceedings of the National Academy of Sciences of the United States of America. 110 (29), 11863-11868 (2013).</li><li>Zurek-Biesiada, D., 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=Localization+microscopy+of+DNA+in+situ+using+Vybrant((R))+DyeCycle+Violet+fluorescent+probe:+A+new+approach+to+study+nuclear+nanostructure+at+single+molecule+resolution.">Localization microscopy of DNA in situ using Vybrant((R)) DyeCycle Violet fluorescent probe: A new approach to study nuclear nanostructure at single molecule resolution.</a> Experimental Cell Research. 343 (2), 97-106 (2016).</li><li>He, J. Y., 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=The+AAA(+)+protein+ATAD3+has+displacement+loop+binding+properties+and+is+involved+in+mitochondrial+nucleoid+organization.">The AAA(+) protein ATAD3 has displacement loop binding properties and is involved in mitochondrial nucleoid organization.</a> Journal of Cell Biology. 176 (2), 141-146 (2007).</li><li>Ashley, N., Harris, D., Poulton, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+mitochondrial+DNA+depletion+in+living+human+cells+using+PicoGreen+staining.">Detection of mitochondrial DNA depletion in living human cells using PicoGreen staining.</a> Experimental Cell Research. 303 (2), 432-446 (2005).</li><li>Jevtic, V., Kindle, P., Avilov, S. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SYBR+Gold+dye+enables+preferential+labeling+of+mitochondrial+nucleoids+and+their+time-lapse+imaging+by+structured+illumination+microscopy.">SYBR Gold dye enables preferential labeling of mitochondrial nucleoids and their time-lapse imaging by structured illumination microscopy.</a> PLoS One. 13 (9), e0203956 (2018).</li></ol>

The authors have nothing to disclose.

There are several critical components to the protocol: To achieve preferential labeling of mitochondrial DNA, the concentration of the DNA binding dye during incubation should be kept very low (e.g., a 1:10,000 dilution of a typical commercial stock), and the incubation time should be 30 min. The incubation time should never exceed 1 h. SYBR Gold dye should be used; other DNA-binding dyes are not bright enough to generate a strong signal upon labeling at a low concentration.
The limitation of ...

Circular 16.5 kbp DNA molecules constitute the genetic material of mitochondria, encoding 22 tRNAs, 2 rRNAs, and 13 polypeptides needed for mitochondrial oxidative phosphorylation complexes. Mitochondrial DNA bound to mitochondrial transcription factor a (TFAM) and several other proteins form the mitochondrial nucleoids1,2,3,4. Mitochondrial nucleoids moveand redistribute between the components of the mitochondrial network5,6 during its morphological remodeling, fission or fusion depending on cell cycle phase, stress, and other factors (reviewed in Pernas et al.7). In addition, the motion of mitochondrial nucleoids, is implicated in systemic lupus erythematosus disease8 and may play a role in other diseases. Fluorescence microscopy is a straightforward technique for live-cell studies of organelles, but the technique has a resolution of &#62;200 nm, which is larger than the size of mitochondrial nucleoids (~100 nm9,10,11,12). This limit has been circumvented by so called &#34;super-resolution&#34; techniques, such as stimulated emission depletion (STED) and single molecule localization microscopy (SMLM)13,14. So far, mitochondrial nucleoids and other DNAs were imaged in live cells by direct stochastic optical reconstruction microscopy (dSTORM)15. Fine sub-mitochondrial structures with positions correlating with mtDNA were observed by STED in live cells16. However, these super-resolution techniques require high illumination intensity, which causes phototoxic effects on living cells17. Therefore, time lapse imaging of mitochondrial nucleoids with resolution beyond diffraction limit is challenging. To address this, we used super-resolution structured illumination microscopy (SR-SIM)18. SIM requires a much lower illumination power dose than STED and SMLM19. Furthermore, in contrast to STED and SMLM techniques, SIM permits straightforward multicolor three-dimensional (3D) imaging, and it does not require particular photophysical properties of the fluorophores or imaging buffer composition19.
The conventional strategy for labeling mitochondrial nucleoids in live cells is fluorescent tagging of a mitochondrial nucleoid protein, such as TFAM20. However, in many cases, this strategy is not suitable. Moreover, overexpression of fluorescent protein-tagged TFAM produces a serious artifact21. Labeling of DNA with organic dyes has advantages over a fluorescent protein (FP)-based strategy. Organic dyes are free of constrains related to FP tagging: they can be used for any type of cells or tissueand can beapplied at any time point of an experiment. Live cell imaging of mitochondrial nucleoids has been reported with several DNA-binding dyes: DAPI22, SYBR Green23, Vybrant DyeCycle24, and picoGreen15,25,26. A substantial drawback of most DNA-binding dyes for nucleoid labeling is that they stain all DNA within the cell. Targeting a dye solely to mitochondrial DNA is highly desirable. To achieve that, careful selection of a dye possessing suitable physico-chemical properties is necessary. Lipophilic dyes possessing delocalized positive charge, such as rhodamine 123, are known to accumulate in live mitochondria, which preserve their negative membrane potential. In addition, an ideal dye for specific labeling of mitochondrial nucleoids should bind DNA with high affinity and emit bright fluorescence upon DNA binding. Considering these requirements, certain cyanines are promising (e.g., picoGreen), but nuclear DNA is abundantly stained by these dyes simultaneously with mitochondrial DNA15,25,26. The present protocol describes specific labeling of mitochondrial nucleoids in live cells with another cyanine dye, SYBR Gold (SG), and tracking of the nucleoids in time lapse super-resolution SIM videos. Moreover, SG-stained live cells can be imaged by any type of inverted fluorescent microscope (confocal, spinning disk, epifluorescence, etc.) suitable for living cells and equipped with a 488 nm light source.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Elyra PS1</td>
			<td>Carl Zeiss</td>
			<td></td>
			<td>multi-modal super-resolution microscope containing module for super-resolution structured illumination microscopy (SR-SIM)</td>
		</tr>
		<tr>
			<td>high glucose DMEM</td>
			<td>GIBCO/ThermoFisher</td>
			<td>31966021</td>
			<td></td>
		</tr>
		<tr>
			<td>ibidi 35-mm dish, glass bottom</td>
			<td>Ibidi Gmbh</td>
			<td>81158</td>
			<td></td>
		</tr>
		<tr>
			<td>ibidi 8-well microSlide, glass bottom</td>
			<td>Ibidi Gmbh</td>
			<td>80827</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaris 8.4.1</td>
			<td>Bitplane/Oxford Instruments</td>
			<td></td>
			<td>image porcessing and visualisation software package</td>
		</tr>
		<tr>
			<td>iXon 885</td>
			<td>Andor Technologies</td>
			<td></td>
			<td>EMCCD camera with back-illuminated sensor</td>
		</tr>
		<tr>
			<td>LSM880 Airyscan</td>
			<td>Carl Zeiss</td>
			<td></td>
			<td>laser scanning confocal microscope with array detector</td>
		</tr>
		<tr>
			<td>Mitotracker CMXRos Red</td>
			<td>ThermoFischer</td>
			<td>M7512</td>
			<td>red live cell mitochondrial stain</td>
		</tr>
		<tr>
			<td>Mitotracker Deep Red FM</td>
			<td>ThermoFischer</td>
			<td>M22426</td>
			<td>far red live cell mitochondrial stain</td>
		</tr>
		<tr>
			<td>picoGreen</td>
			<td>ThermoFischer</td>
			<td>P7581</td>
			<td>cell permeant DNA stain</td>
		</tr>
		<tr>
			<td>Plan Apochromat 100x/1.46 Oil objective</td>
			<td>Carl Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SYBR Gold</td>
			<td>ThermoFischer</td>
			<td>S11494</td>
			<td>cell permeant DNA stain</td>
		</tr>
		<tr>
			<td>Zen Black 2012 software</td>
			<td>Carl Zeiss</td>
			<td></td>
			<td>image acquisition and processing software</td>
		</tr>
	</tbody>
</table>

specific labeling of mitochondrial nucleoids for time-lapse structured illumination microscopy

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and several essential mitochondrial polypeptides. Mitochondrial nucleoids divide and distribute within the dynamic mitochondrial network that undergoes fission/fusion and other morphological changes. High resolution live fluorescence microscopy is a straightforward technique to characterize a nucleoid&#39;s position and motion. For this technique, nucleoids are commonly labeled through fluorescent tags of their protein components, namely transcription factor a (TFAM). However, this strategy needs overexpression of a fluorescent protein-tagged construct, which may cause artifacts (reported for TFAM), and is not feasible in many cases. Organic DNA-binding dyes do not have these disadvantages. However, they always show staining of both nuclear and mitochondrial DNAs, thus lacking specificity to mitochondrial nucleoids. By taking into account the physico-chemical properties of such dyes, we selected a nucleic acid gel stain (SYBR Gold) and achieved preferential labeling of mitochondrial nucleoids in live cells. Properties of the dye, particularly its high brightness upon binding to DNA, permit subsequent quantification of mitochondrial nucleoid motion using time series of super-resolution structured illumination images.

Characterization of live cell labeling with SG 
First, the distribution of SG in the cells upon incubation with the dye at various dilutions was characterized by confocal microscopy. After incubation with high concentrations of SG or picoGreen, both dyes mostly labeled the nuclei and showed a punctate staining in the cytoplasm (Figure 1), similarly to published data for another positively charged cyanine dye (i.e., picoGreen)<sup class="...

Watch this Scientific Journal Video about Specific Labeling of Mitochondrial Nucleoids for Time-lapse Structured Illumination Microscopy at JoVE.com

Specific Labeling of Mitochondrial Nucleoids for Time-lapse Structured Illumination Microscopy

The protocol describes specific labeling of mitochondrial nucleoids with a commercially available DNA gel stain, acquisition of time lapse series of live labeled cells by super-resolution structured illumination microscopy (SR-SIM), and automatic tracking of nucleoid motion.

Mitochondrial nucleoids are compact particles formed by mitochondrial DNA molecules coated with proteins. Mitochondrial DNA encodes tRNAs, rRNAs, and ...

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7.1K Views.  Max Planck Institute of Immunobiology and Epigenetics. The protocol describes specific labeling of mitochondrial nucleoids with a commercially available DNA gel stain, acquisition of time lapse series of live labeled cells by super-resolution structured illumination microscopy (SR-SIM), and automatic tracking of nucleoid motion.

Video: Specific Labeling of Mitochondrial Nucleoids for Time-lapse Structured Illumination Microscopy

Monitoring Actin Disassembly with Time-lapse Microscopy

Okay, so what I'd like to do today is to show you how to construct a flow chamber, a fusion chamber for imaging on an upright microscope. What then I then do is I create two spaces using film. So what I do is I take the para film, I cut it into small strips.Next I'm going to, this will basically form the channels for the, the walls, and I dry them with a tilted air. I put the cover slip on top of the para, And here's the finished. Then I heat it on a heat block to melt the param and basically create a seal.So what this is, is the, the cover glass here, and then on top of the cover glass is two layers of phim as you see right here. And then on top of this, again, there's a, it's a 22 by two two mil meter cover slip. This one right here, which I pressed against the param to fall the seal.Okay, so the liquid goes in in one direction here and I can suck it out with a filter paper here in this direction. And I'll show you how to do that in a bit. So what I like to do is I like to put it in a humidified chamber, which is very simple.Take two kind of elevated strips and have a filter of paper underneath. And to keep it noisy as I, I put the cover, cover slip on the flow chamber on. Now the way you introduce a solution to the chamber is that you just put the pipet tip at the entrance of the chamber and you just split.Now what I do is I incubate the chamber with the lid closed for 10 minutes. What I usually do to suck the liquid out from the other side is to use it, get a filter paper and cut it into thin strips like this. And I take one of these thin strips and I simultaneously apply the solution into the entrance while putting the filter paper at the exit to suck the exchange, the solution.So you'll see the liquid flowing through the solution. And then I wait for five minutes for the mid close. Now what I'm ready to do is to actually put that the actin it and the actin, I actually have to polymerize, I polymerize inside the flow chamber and it's just the same procedures before.So because like this, and then now that the action is polymerized, I wash out all the un excess un polymerized action using a large volume of just buffer. So here I have about a hundred microliters and it's a few reaction chamber volumes worth of buffer. Just, you know, when the filter paper becomes saturated, I have to change the direction.Okay?And so I'm ready to do imaging with this. So perfusion chamber with actin attached to it. Now I take the slide, mount it under the microscope, and since I'm imaging with an oil objective, I'm gonna put some immersion oil on the top surface of the cover, grass touch, such so assess solution or protein or anything on acting inside.So what I do is I can actually exchange the solution side while I'm doing the time lapse imaging. As such.

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 ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During directional migration, cells move rapidly in response to an extracellular chemotactic signal, or in response to intrinsic cues 3 provided by the basic motility machinery. Random migration occurs when a cell possesses low intrinsic directionality, allowing the cells to explore their local environment.
Cell migration is a complex process, in the initial response cell undergoes polarization and extends protrusions in the direction of migration 2. Traditional methods to measure migration such as the Boyden chamber migration assay is an easy method to measure chemotaxis in vitro, which allows measuring migration as an end point result. However, this approach neither allows measurement of individual migration parameters, nor does it allow to visualization of morphological changes that cell undergoes during migration.
Here, we present a method that allows us to monitor migrating cells in real time using video - time lapse microscopy. Since cell migration and invasion are hallmarks of cancer, this method will be applicable in studying cancer cell migration and invasion in vitro. Random migration of platelets has been considered as one of the parameters of platelet function 4, hence this method could also be helpful in studying platelet functions. This assay has the advantage of being rapid, reliable, reproducible, and does not require optimization of cell numbers. In order to maintain physiologically suitable conditions for cells, the microscope is equipped with CO2 supply and temperature thermostat. Cell movement is monitored by taking pictures using a camera fitted to the microscope at regular intervals. Cell migration can be calculated by measuring average speed and average displacement, which is calculated by Slidebook software.

This method allows monitoring of cells in real time and quantitative measurements of different cell migration parameters such as speed, displacement, and velocity. Unlike the traditional methods, this real time approach is not based on endpoint quantitative migration measurements; instead it allows monitoring and calculating different parameters continuously.

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During ...

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages as well as by the availability of genetic mutants and transgenic strains expressing a myriad of fusion proteins1-4. In addition, dynamic processes such as cell division can be viewed using fluorescently labeled reporter proteins. The study of mitosis can be facilitated through the use of time-lapse experiments in various systems including intact organisms; thus the early C. elegans embryo is well suited for this study. Presented here is a technique by which in vivo imaging of sub-cellular structures in response to anoxic (99.999% N2; &lt;2 ppm O2) stress is possible using a simple gas flow through setup on a high-powered microscope. A microincubation chamber is used in conjunction with nitrogen gas flow through and a spinning disc confocal microscope to create a controlled environment in which animals can be imaged in vivo. Using GFP-tagged gamma tubulin and histone, the dynamics and arrest of cell division can be monitored before, during and after exposure to an oxygen-deprived environment. The results of this technique are high resolution, detailed videos and images of cellular structures within blastomeres of embryos exposed to oxygen deprivation.

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Described here is an in vivo technique to image sub-cellular structures in animals exposed to anoxia using a gas flow through microincubation chamber in conjunction with a spinning disc confocal microscope. This method is straightforward and flexible enough to suit a variety of experimental parameters and model systems.

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages ...

Visualization of Craniofacial Development in the sox10: kaede Transgenic Zebrafish Line Using Time-lapse Confocal Microscopy

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, convergence and extension of facial prominences, and maturation of the craniofacial skeleton. To study the contribution of the cranial neural crest to specific regions of the zebrafish palate a sox10: kaede transgenic zebrafish line was generated. Sox10 provides lineage restriction of the kaede reporter protein to the neural crest, thereby making the cell labeling a more precise process than traditional dye or reporter mRNA injection. Kaede is a photo-convertible protein that turns from green to red after photo activation and makes it possible to follow cells precisely. The sox10: kaede transgenic line was used to perform lineage analysis to delineate CNC cell populations that give rise to maxillary versus mandibular elements and illustrate homology of facial prominences to amniotes. This protocol describes the steps to generate a live time-lapse video of a sox10: kaede zebrafish embryo. Development of the ethmoid plate will serve as a practical example. This protocol can be applied to making a time-lapse confocal recording of any kaede or similar photoconvertible reporter protein in transgenic zebrafish. Furthermore, it can be used to capture not only normal, but also abnormal development of craniofacial structures in the zebrafish mutants.

Visualization of experimental data has become a key element in presenting results to the scientific community. Generation of live time-lapse recording of growing embryos contributes to better presentation and understanding of complex developmental processes. This protocol is a step-by-step guide to cell labeling via photoconversion of kaede protein in zebrafish.

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, ...

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques &#8211; stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 &#181;m from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy f3D-SIM

Spatiotemporal information about dynamic proteins inside live cells is crucial for understanding biology. A type of super-resolution microscopy called fast 3D-structured illumination microscopy (f3D-SIM) reveals unique information about the cytokinetic Z ring in bacteria: both its bead-like appearance and the rapid dynamics of FtsZ within the ring.

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the ...

Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time points. Time intervals can be set through a computer interface that controls the microscope-integrated camera. This kind of microscopy requires both the ability to acquire very rapid events and the signal generated by the observed cellular structure during these events. After the images have been collected, a movie of the entire experiment is assembled to show the dynamic of the molecular events of interest. Time-lapse video microscopy has a broad range of applications in the biomedical research field and is a powerful and unique tool for following the dynamics of the cellular events in real time. Through this technique, we can assess cellular events such as migration, division, signal transduction, growth, and death. Moreover, using fluorescent molecular probes we are able to mark specific molecules, such as DNA, RNA or proteins and follow them through their molecular pathways and functions. Time-lapse video microscopy has multiple advantages, the major one being the ability to collect data at the single-cell level, that make it a unique technology for investigation in the field of cell biology. However, time-lapse video microscopy has limitations that can interfere with the acquisition of high quality images. Images can be compromised by both external factors; temperature fluctuations, vibrations, humidity and internal factors; pH, cell motility. Herein, we describe a protocol for the dynamic acquisition of a specific protein, Parkin, fused with the enhanced yellow fluorescent protein (EYFP) in order to track the selective removal of damaged mitochondria, using a time-lapse video microscopy approach.

Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

The plant immune response associated with a genome-wide transcriptional reprogramming is initiated at the site of infection. Thus, the immune response is regulated spatially and temporally. The use of a fluorescent gene under the control of an immunity-related promoter in combination with an automated fluorescence microscopy is a simple way to understand spatiotemporal regulation of plant immunity. In contrast to the root tissues that have been used for a number of various intravital fluorescent imaging experiments, there exist few fluorescent live-imaging examples for the leaf tissues that encounter an array of airborne microbial infections. Therefore, we developed a simple method to mount leaves of Arabidopsis thaliana plants for live-cell imaging over an extended period of time. We used transgenic Arabidopsis plants expressing the yellow fluorescent protein (YFP) genefused to the nuclear localization signal (NLS) under the control of the promoter of a defense-related marker gene, Pathogenesis-Related 1 (PR1). We infiltrated a transgenic leaf with Pseudomonas syringae pv. tomato DC3000 (avrRpt2) strain (Pst_a2) and performed in vivo time-lapse imaging of the YFP signal for a total of 40 h using an automated fluorescence stereomicroscope. This method can be utilized not only for studies on plant immune responses but also for analyses of various developmental events and environmental responses occurring in leaf tissues.

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging

We report a simple and versatile method for performing fluorescent live-imaging of Arabidopsis thaliana leaves over an extended period of time. We use a transgenic Arabidopsis plant expressing a fluorescent reporter gene under the control of an immunity-related promoter as an example for gaining spatiotemporal understanding of plant immune responses.

The plant immune response associated with a genome-wide transcriptional reprogramming is initiated at the site of infection. Thus, the immune response is ...

Specific Labeling of Mitochondrial Nucleoids for Time-lapse Structured Illumination Microscopy

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Chapters in this video

Monitoring Actin Disassembly with Time-lapse Microscopy

Time-lapse Imaging of Mitosis After siRNA Transfection

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Visualization of Craniofacial Development in the sox10: kaede Transgenic Zebrafish Line Using Time-lapse Confocal Microscopy

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

A Versatile Method for Mounting Arabidopsis Leaves for Intravital Time-lapse Imaging