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>

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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. 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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. 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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 our protocol is that the dye is washed out from the mitochondrial nucleoids during a permeabilization step. Therefore, the described procedure is not suitable for conventional immunofluorescence protocols. In this case, nucleoids in fixed cells can be efficiently labeled with other techniques, such as antibodies against DNA or mitochondrial transcription factors (TFAM).
Direct labeling of mitochondrial nucleoids with the DNA binding organic dye has advantages over fluorescent protein labeling: any type of cell can be labeled within &#60;1 h, without temporal or other constrains of transient or stable expression of fluorescent protein-tagged constructs. Also, in current protocols, conventional fluorescent protein-tagging of TFAM was reported to cause artifacts. Moreover, SG efficiently stains nucleoids only if the mitochondrial membrane potential is intact. This prevents image acquisition of biologically irrelevant &#34;sick&#34; and dead cells, which cannot be avoided with FP-based staining. Finally, previously published protocols based on DNA binding dyes did not achieve preferential staining of mitochondrial DNA.
The proposed protocol is fast and simple, thus we assume it will be widely used for live imaging of mitochondrial nucleoids by various fluorescence techniques, both diffraction limited (e.g., laser scanning confocal, TIRF, etc.) as well as super-resolution SIM.

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><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)15. However, upon incubation with SG at 1:10,000 dilution, faint staining appeared in the nuclei, while in the cytoplasm, we observed a pattern of bright spots (Figure 1). On the other hand, incubation with picoGreen dye diluted 1:10,000 yielded mostly nuclear staining. The SG signal was much brighter than that of picoGreen at the same concentration. The data showed that SG is more suitable for imaging mitochondrial DNA than other similar DNA-binding dyes.
To confirm that the bright dots are localized in the mitochondria, we stained living cells simultaneously with SG and far red mitochondrial stain. The latter is a positively charged cell permeable organic dye that accumulates in the mitochondria of living cells. Upon incubation with SG at 1:10,000 and 1:50,000 dilutions, nearly all SG staining occurred in mitochondria, while upon labeling at 1:500 and 1:1,000 dilutions, significant staining of the nuclei and cytoplasm occurred (Figure 2).
Further, we characterized the time course of staining live cells with SG by time lapse microscopy (Figure 3). The plots of SG fluorescence intensity in mitochondria vs. time (Figure 3B) suggested that after 45 min, nucleoid staining was close to saturation. Thus, we recommend incubation times of ~30&#8211;60 min.
We tested how SG intracellular distribution changes upon fixation and/or permeabilization of the cells. Fixation (2% paraformaldehyde [PFA]) of the live-stained cells caused a slight redistribution of the dye to the nucleus (Figure 4A,B). Permeabilization of the fixed cells (0.1% Triton X100) eliminated the SG dotted pattern in mitochondria, and staining of the nuclei was dominant (Figure 4C). If the SG was added to the cells after fixation and permeabilization, it distributed uniformly across the cytoplasm and nuclei (Figure 4D). Thus, SG labeled cells can be fixed if necessary, but the dye is not suitable for protocols that require permeabilization.
Live cell SR-SIM and mitochondrial nucleoids tracking 
Live cells costained with SG and a far red mitochondrial stain (Table of Materials) were 3D imaged by a super-resolution SIM technique. As in the confocal images (Figure 2), mitochondrial nucleoids appeared as bright spots within the mitochondria (Figure 5A). Further, we acquired a time series of 2D SIM images and tracked the positions of the nucleoids at a resolution beyond the diffraction limit. Immediately before starting the time series we acquired SIM 3D stacks in the SG and the mitochondrial stain channels. Then we acquired a time series in the SG channel only and tracked the mitochondrial nucleoids in order to quantify their motions. The track mean speed was 0.042 &#181;m/s, and the maximal instant speed of track&#160;was 0.078 &#177; 0.012 &#181;m/s. The majority of nucleoids did not displace far from their original positions but showed short-distance random motions that were probably confined to the mitochondrial network, as an overlay of tracks on the mitochondrial images suggests (Figure 5B). Few rapid directional displacements occurred during a typical time series (Movie 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60003/60003fig1.jpg" /> 
Figure 1: Comparison of picoGreen and SG live cell labeling. A549 cells were incubated with picoGreen or SG at indicated dilutions and imaged. Representative fields of view of the labeled cells in the green channel. LSM880 Airyscan FAST, 20x/air objective, Scale bar = 50 &#181;m. Single optical sections. White squares mark the areas shown at a higher magnification to the right of the entire 423 x 423 &#181;m fields of view. For 1:1,000 and 1:2,000 dilutions, the same images are shown at two brightness settings: the &#34;default&#34; brightness settings optimized for 1:10,000 dilution, and the &#34;reduced brightness&#34; settings adjusted to avoid detector saturation. This figure has been modified from Jevtic et al.27. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60003/60003fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60003/60003fig2.jpg" /> 
Figure 2: SG localization in live cells upon labeling at different concentrations. HeLa cells were incubated for 30 min with a mixture of 0.25 &#181;M Mitotracker CMXRos Red and the indicated SG dilution. The solution was replaced with DMEM, and the images were acquired on a LSM880 Airyscan microscope, 63x 1.4 oil objective, with a sequential acquisition of color channels. Single optical slices are shown Scale bar = 10 &#181;m). This figure has been modified from Jevtic et al.27. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60003/60003fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60003/60003fig3.jpg" /> 
Figure 3: HeLa cells during labeling with SYBR Gold (SG). First, live HeLa cells were labeled with Mitotracker CMXRos Red and washed. SG (final dilution 1:10,000 in DMEM) was added to the cells and a time lapse series was acquired. LSM880 microscope, 63x 1.4 Oil objective, sequential acquisition. Z-stacks were acquired at each time point. The maximum intensity projections are shown. (A) Mean intensities of SG fluorescence over time in several regions of interest. (B) Representative fields of view showing the regions of interest where SG fluorescence was measured (colored rectangles). (C) A field of view at several time points during incubation with SG. A square region marked with white line is shown at a higher magnification in the right column. This figure has been modified from Jevtic et al.27. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60003/60003fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60003/60003fig4.jpg" /> 
Figure 4: Effect of fixation and permeabilization on SG localization in the cells. HeLa cells were stained with SG (stock diluted 1:10,000) and Mitotracker CMXRos Red (0.25 &#181;M) for 30 min. Single optical slices were acquired on a spinning disk microscope; sequential acquisition; scale bar = 10 &#181;m. (A) Live HeLa cells stained with SG. (B) Live HeLa cells stained with SG and then fixed with 2% PFA in PBS for 30 min. (C) Live HeLa cells stained with SG, then fixed with 2% PFA in PBS for 30 min and permeabilized with 0.1% Triton X100 for 15 min. On the lower panel, the same image is shown, but the brightness in the green channel is set higher. (D) HeLa cells fixed with 2% PFA, permeabilized with 0.1% Triton X100 for 15 min, and then stained with SG and Mitotracker CMXRos Red. To acquire the images shown in (D), the EMCCD gain of the camera for the green channel was reduced by a factor of 12 in comparison to A-C to avoid overexposure. This figure has been modified from Jevtic et al.27. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60003/60003fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60003/60003fig5.jpg" /> 
Figure 5: Nucleoid tracking on live SIM images. Representative images of a field of view of SG stained&#160;cells. Bottom:&#160;the two-color SIM image taken before the acquisition of the time lapse series. Green = SG channel; magenta = mitotracker channel. Top: nucleoid tracks from a 50-frame SIM time series in the SG channel (Movie 1); frame time = 1.8 s; tracking by Imaris 8.4.1 software. Tracks are color-coded by maximal track speed (&#181;m/s); Elyra PS.1, SIM mode, 100x/1.46 Oil objective; Scale bar = 10 &#181;m. This figure has been modified from Jevtic et al.27. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60003/60003fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Movie 1" src="/files/ftp_upload/60003/60003movie1.jpg" /> 
Movie 1: SIM time lapse series showing representative SG-stained cells. Scale bar = 5 &#181;m. Detected mitochondrial nucleoids are marked as white spheres. Tracks are visualized as &#34;Dragon tails&#34; (8 frames length) and color-coded according to their maximal instant speeds (color bar in the bottom right shows speeds in &#181;m/s). Mitochondrial nucleoids tracking and visualization by Imaris 8.4.1 software. This video has been published in Jevtic et al.27. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/60003/60003movie1.avi" target="_blank">Please click here to view this video (Right click to download).</a>

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 ...

specific-labeling-mitochondrial-nucleoids-for-time-lapse-structured

Universidad San Sebastian

Research

JoVE Journal

Biology

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

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Neuroscience

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

Mitochondria are essential organelles of eukaryotic cells capable of aerobic respiration. They contain circular genome and gene expression apparatus. A mitochondrial genome of baker&#8217;s yeast encodes eight proteins: three subunits of the cytochrome c oxidase (Cox1p, Cox2p, and Cox3p), three subunits of the ATP synthase (Atp6p, Atp8p, and Atp9p), a subunit of the ubiquinol-cytochrome c oxidoreductase enzyme, cytochrome b (Cytb), and mitochondrial ribosomal protein Var1p. The purpose of the method described here is to specifically label these proteins with 35S methionine, separate them by electrophoresis and visualize the signals as discrete bands on the screen. The procedure involves several steps. First, yeast cells are cultured in a galactose-containing medium until they reach the late logarithmic growth stage. Next, cycloheximide treatment blocks cytoplasmic translation and allows 35S methionine incorporation only in mitochondrial translation products. Then, all proteins are extracted from yeast cells and separated by polyacrylamide gel electrophoresis. Finally, the gel is dried and incubated with the storage phosphor screen. The screen is scanned on a phosphorimager revealing the bands. The method can be applied to compare the biosynthesis rate of a single polypeptide in the mitochondria of a mutant yeast strain versus the wild type, which is useful for studying mitochondrial gene expression defects. This protocol gives valuable information about the translation rate of all yeast mitochondrial mRNAs. However, it requires several controls and additional experiments to make proper conclusions.

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

Baker&#8217;s yeast mitochondrial genome encodes eight polypeptides. The goal of the current protocol is to label all of them and subsequently visualize them as separate bands.

Mitochondria are essential organelles of eukaryotic cells capable of aerobic respiration. They contain circular genome and gene expression apparatus. A ...

Biochemistry

High-Throughput Quantification of the Synthesis and Distribution of mtDNA

High-Throughput Image-Based Quantification of Mitochondrial DNA Synthesis and Distribution

The vast majority of cellular processes require a continuous supply of energy, the most common carrier of which is the ATP molecule. Eukaryotic cells produce most of their ATP in the mitochondria by oxidative phosphorylation. Mitochondria are unique organelles because they have their own genome that is replicated and passed on to the next generation of cells. In contrast to the nuclear genome, there are multiple copies of the mitochondrial genome in the cell. The detailed study of the mechanisms responsible for the replication, repair, and maintenance of the mitochondrial genome is essential for understanding the proper functioning of mitochondria and whole cells under both normal and disease conditions. Here, a method that allows the high-throughput quantification of the synthesis and distribution of mitochondrial DNA (mtDNA) in human cells cultured in vitro is presented. This approach is based on the immunofluorescence detection of actively synthesized DNA molecules labeled by 5-bromo-2'-deoxyuridine (BrdU) incorporation and the concurrent detection of all the mtDNA molecules with anti-DNA antibodies. Additionally, the mitochondria are visualized with specific dyes or antibodies. The culturing of cells in a multi-well format and the utilization of an automated fluorescence microscope make it easier to study the dynamics of mtDNA and the morphology of mitochondria under a variety of experimental conditions in a relatively short time.

Author Spotlight: High-Throughput Image-Based Quantification of Mitochondrial DNA Synthesis and Distribution  

A procedure for studying the dynamics of mitochondrial DNA (mtDNA) metabolism in cells using a multi-well plate format and automated immunofluorescence imaging to detect and quantify mtDNA synthesis and distribution is described. This can be further used to investigate the effects of various inhibitors, cellular stresses, and gene silencing on mtDNA metabolism.

The vast majority of cellular processes require a continuous supply of energy, the most common carrier of which is the ATP molecule. Eukaryotic cells ...

Three-dimensional Imaging and Analysis of Mitochondria within Human Intraepidermal Nerve Fibers

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to isolate nerve-specific mitochondria and evaluate disease-induced alterations of mitochondria in the distal tip of sensory nerves. The protocol combines fluorescence immunohistochemistry, confocal microscopy and 3D image analysis techniques to visualize and quantify nerve-specific mitochondria. Detailed parameters are defined throughout the procedures in order to provide a concrete example of how to use these techniques to isolate nerve-specific mitochondria. Antibodies were used to label nerve and mitochondrial signals within tissue sections of skin punch biopsies, which was followed by indirect immunofluorescence to visualize nerves and mitochondria with a green and red fluorescent signal respectively. Z-series images were acquired with confocal microscopy and 3D analysis software was used to process and analyze the signals. It is not necessary to follow the exact parameters described within, but it is important to be consistent with the ones chosen throughout the staining, acquisition and analysis steps. The strength of this protocol is that it is applicable to a wide variety of circumstances where one fluorescent signal is used to isolate other signals that would otherwise be impossible to study alone.

This protocol uses three-dimensional (3D) imaging and analysis techniques to visualize and quantify nerve-specific mitochondria. The techniques are applicable to other situations where one fluorescent signal is used to isolate a subset of data from another fluorescent signal.

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to ...

Visualization and Live Imaging of Oligodendrocyte Organelles in Organotypic Brain Slices Using Adeno-associated Virus and Confocal Microscopy

Neurons rely on the electric insulation and trophic support of myelinating oligodendrocytes. Despite the importance of oligodendrocytes, the advanced tools currently used to study neurons, have only partly been taken on by oligodendrocyte researchers. Cell type-specific staining by viral transduction is a useful approach to study live organelle dynamics. This paper describes a protocol for visualizing oligodendrocyte mitochondria in organotypic brain slices by transduction with adeno-associated virus (AAV) carrying genes for mitochondrial targeted fluorescent proteins under the transcriptional control of the myelin basic protein promoter. It includes the protocol for making organotypic coronal mouse brain slices. A procedure for time-lapse imaging of mitochondria then follows. These methods can be transferred to other organelles and may be particularly useful for studying organelles in the myelin sheath. Finally, we describe a readily available technique for visualization of unstained myelin in living slices by Confocal Reflectance microscopy (CoRe). CoRe requires no extra equipment and can be useful to identify the myelin sheath during live imaging.

Myelinating oligodendrocytes promote rapid action potential propagation and neuronal survival. Described here is a protocol for oligodendrocyte-specific expression of fluorescent proteins in organotypic brain slices with subsequent time-lapse imaging. Further, a simple procedure for visualizing unstained myelin is presented.

Neurons rely on the electric insulation and trophic support of myelinating oligodendrocytes. Despite the importance of oligodendrocytes, the advanced ...

Unveiling Mitochondrial Ultrastructure in Neurons Using STED Microscopy

Using Live Cell STED Imaging to Visualize Mitochondrial Inner Membrane Ultrastructure in Neuronal Cell Models

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of reactive oxygen species (ROS). These mitochondria-mediated processes take on specialized roles in neurons, coordinating aerobic metabolism to meet the high energy demands of these cells, modulating Ca2+ signaling, providing lipids for axon growth and regeneration, and tuning ROS production for neuronal development and function. Mitochondrial dysfunction is therefore a central driver in neurodegenerative diseases. Mitochondrial structure and function are inextricably linked. The morphologically complex inner membrane with structural infolds called cristae harbors many molecular systems that perform the signature processes of the mitochondrion. The architectural features of the inner membrane are ultrastructural and therefore, too small to be visualized by traditional diffraction-limited resolved microscopy. Thus, most insights on mitochondrial ultrastructure have come from electron microscopy on fixed samples. However, emerging technologies in super-resolution fluorescence microscopy now provide resolution down to tens of nanometers, allowing visualization of ultrastructural features in live cells. Super-resolution imaging therefore offers an unprecedented ability to directly image fine details of mitochondrial structure, nanoscale protein distributions, and cristae dynamics, providing fundamental new insights that link mitochondria to human health and disease. This protocol presents the use of stimulated emission depletion (STED) super-resolution microscopy to visualize the mitochondrial ultrastructure of live human neuroblastoma cells and primary rat neurons. This procedure is organized into five sections: (1) growth and differentiation of the SH-SY5Y cell line, (2) isolation, plating, and growth of primary rat hippocampal neurons, (3) procedures for staining cells for live STED imaging, (4) procedures for live cell STED experiments using a STED microscope&#160;for reference, and (5) guidance for segmentation and image processing using examples to measure and quantify morphological features of the inner membrane.

Author Spotlight: Decoding Mitochondrial Aging 

This protocol presents a workflow for the propagation, differentiation, and staining of cultured SH-SY5Y cells and primary rat hippocampal neurons for mitochondrial ultrastructure visualization and analysis using stimulated emission depletion (STED) microscopy.

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of ...

MATLAB Optimization for Analyzing Mitochondrial Networks in Neuroscience

Optimized Automated Analysis of Live Neuronal Mitochondria Homeostasis Modulation by Isoform-Specific Retinoic Acid Receptors

The complex mitochondrial network makes it very challenging to segment, follow, and analyze live cells. MATLAB tools allow the analysis of mitochondria in timelapse files, considerably simplifying and speeding up the process of image processing. Nonetheless, existing tools produce a large output volume, requiring individual manual attention, and basic experimental setups have an output of thousands of files, each requiring extensive and time-consuming handling. 
To address these issues, a routine optimization was developed, in both MATLAB code and live-script forms, allowing for swift file analysis and significantly reducing document reading and data processing. With a speed of 100 files/min, the optimization allows an overall rapid analysis. The optimization achieves the results output by averaging frame-specific data for individual mitochondria throughout time frames, analyzing data in a defined manner, consistent with those output from existing tools. Live confocal imaging was performed using the dye tetramethylrhodamine methyl ester, and the routine optimization was validated by treating neuronal cells with retinoic acid receptor (RAR) agonists, whose effects on neuronal mitochondria are established in the literature. The results were consistent with the literature and allowed further characterization of mitochondrial network behavior in response to isoform-specific RAR modulation. 
This new methodology allowed rapid and validated characterization of whole-neuron mitochondria network, but it also allows for differentiation between axon and cell body mitochondria, an essential feature to apply in the neuroscience field. Moreover, this protocol can be applied to experiments using fast-acting treatments, allowing the imaging of the same cells before and after treatments, transcending the field of neuroscience.

Author Spotlight: An Optimized Automated Method for Investigating Retinoic Acid Receptors in Neuronal Mitochondria

The mitochondrial network is extremely complex, making it very challenging to analyze. A novel MATLAB tool analyzes live confocal imaged mitochondria in timelapse images but results in a large output volume requiring individual manual attention. To address this issue, a routine optimization was developed, allowing for speedy file analysis.

The complex mitochondrial network makes it very challenging to segment, follow, and analyze live cells. MATLAB tools allow the analysis of mitochondria in ...

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH allows the measurement of cell-to-cell variability in mRNA copy number of genes of interest, as well as the subcellular location of the transcripts. The main steps involved are fixation of the bacterial cell culture, permeabilization of cell membranes, and hybridization of the target transcripts with sets of commercially available short fluorescently-labeled oligonucleotide probes. smFISH can allow the imaging of the transcripts of multiple genes in the same cell, with limitations imposed by the spectral overlap between different fluorescent markers. Following completion of the protocol illustrated below, cells can be readily imaged using a microscope coupled with a camera suitable for low-intensity fluorescence. These images, together with cell contours obtained from segmentation of phase contrast frames, or from cell membrane staining, allow the calculation of the mRNA copy number distribution of a sample of cells using open-source or custom-written software. The labeling method described here can also be applied to image transcripts with stochastic optical reconstruction microscopy (STORM).

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

This manuscript describes a method for labeling individual messenger RNA (mRNA) transcripts with fluorescently-labeled DNA probes, for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH is a visualization method that allows the simultaneous detection, localization, and quantification of single mRNA molecules in fixed individual cells.

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ ...

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a ...

Correlative Microscopy Reveals Stress-Induced Mitochondria-Lysosome Interactions

Dual-color Correlative Light and Electron Microscopy for the Visualization of Interactions between Mitochondria and Lysosomes

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for structural analysis, light microscopy is essential for the visualization of dynamic organelles by target-specific labeling. The following protocol describes a method that combines dual-color correlative light and electron microscopy (CLEM) to observe the interactions between mitochondria and lysosomes. In this study, mitochondria were labeled with mEosEM (Mito-mEosEM) and lysosomes with TMEM192-V5-APEX2. The results obtained from CLEM images enable us to observe the changes in the interactions between mitochondria and lysosomes under external stress conditions. Treatment with bafilomycin (BFA), which inhibits lysosomal function, resulted in an increase in contact between mitochondria and lysosomes, leading to the formation of fragmented mitochondria trapped inside lysosomes. Conversely, treatment with U18666A, which inhibits cholesterol export from lysosomes, caused lysosomes to be surrounded by mitochondria, indicating a distinct form of interaction. This study presents an effective method for observing the interactions between mitochondria and lysosomes in fixed cells. Furthermore, CLEM imaging with dual-color probes offers a powerful tool for future investigations of organelle dynamics and their implications for cell function and pathology.

This protocol describes a mechanism for using correlative light and electron microscopy to visualize the interaction of mitochondria and lysosomes labeled with mEosEM and APEX2, respectively.

Cellular organelles, such as mitochondria and lysosomes, display dynamic structures. Despite the higher resolution of transmission electron microscopy for ...

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

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Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

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