Name: simultaneous measurement of mitochondrial calcium and mitochondrial membrane potential in live cells by fluorescent microscopy
Uploaded: 2017-01-24T14:00:00.000+00:00
Duration: 8 min 43 s
Description: Watch this Scientific Journal Video about Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy at JoVE.com

We thank Dr Kirstin Elgass and Dr Sarah Creed from Monash Micro Imaging for technical assistance, and the Wellcome Trust and Medical Research Council UK for financial support. MMcK is supported the Australian Research Council Future Fellowship Scheme (FT120100459), the William Buckland Foundation, The Australian Mitochondrial Disease Foundation (AMDF), The Hudson Institute of Medical Research and Monash University. This work was supported by the Victorian Government Operational Infrastructure Support Scheme.

1. Preparation of Cells
<ol>
	<li>Grow cells on 10 cm cell culture dishes or 75 cm2 flasks in culture media [10 ml Dulbecco&#39;s Modified Eagles Medium (DMEM) supplemented with 5% (v/v) fetal bovine serum (FBS) and 1x penicillin/streptomycin (p/s)] at 37 &#176;C/5% CO2.</li>
	<li>To harvest cells, remove media by aspiration, then wash with 5 ml 1x phosphate buffered saline (1x PBS). Remove 1x PBS by aspiration, then add 1.5 ml 0.25% (w/v) Trypsin/0.25% (w/v) ethylenediaminetetraacetic acid (EDTA) and incubate at 37 &#176;C/5% CO2 for 2 min. Tap dish or flask gently to remove cells, then resuspend in 5 ml culture media.</li>
	<li>Count cells by adding 12 &#181;l of resuspended cells onto a hemocytometer.</li>
	<li>Plate cells in dishes or chambered coverslips for confocal imaging. 
	NOTE: These will have a suitable plastic or glass bottom that has been designed for use with inverted microscopes.</li>
	<li>Plate cells so that ~80% confluency is achieved on the day of imaging. For example, approximately 2 x 104 143B osteosarcoma cells can be plated into one well of an 8-well chamber slide one day before imaging.</li>
	<li>Incubate cells overnight at 37 &#176;C/5% CO2 to allow cells to attach and recover.</li>
</ol>
2. Buffers for TMRM and Fluo-4 Imaging
<ol>
	<li>Prepare Record Solution (RS) containing 156 mM NaCl, 3 mM KCl, 2 mM MgSO4, 1.25 mM KH2PO4, 10 mM D-glucose, 2 mM CaCl2 and 10 mM HEPES pH 7.35. Store RS in aliquots at&#160; -20 &#186;C.</li>
	<li>Prepare Intracellular Medium (IM) containing 6 mM NaCl, 130 mM KCl, 7.8 mM MgCl2, 1 mM KH2PO4, 0.4 mM CaCl2, 2 mM EGTA, 10 mM HEDTA, 2 mM malate, 2 mM glutamate, 2 mM ADP and 20 mM HEPES pH 7.1. 200 mM stock solutions of malate, glutamate and ADP can be prepared separately, aliquoted, and stored at -20 &#176;C. These stocks can be added to the IM just before use.</li>
	<li>Ca2+ free Hank&#39;s buffered salt solution (HBSS) can be obtained pre-prepared from commercial suppliers. Add ethylene glycol-bis(&#946;-aminoethyl ether)-N,N,N&#39;,N&#39;-tetraacetic acid (EGTA) to a final concentration of 500 &#181;M.</li>
	<li>Prepare a stock solution of 10 mM tetramethylrhodamine, methyl ester, perchlorate (TMRM) in 100% methanol. From this stock, make a working stock of 2 &#181;M in distilled H2O.</li>
	<li>Prepare a stock solution of 10 mM Verapamil in 100% ethanol.</li>
	<li>Prepare a 1 mg/ml (w/v) stock solution of Fluo-4 acetoxymethyl ester (Fluo-4, AM) in dimethyl sulfoxide (DMSO). 
	NOTE: Fluo-4, AM is a calcium indicator that exhibits increased fluorescence upon binding Ca2+. Fluo-4 is an analog of fluo-3, with the two chlorine substituents replaced by fluorines. This results in increased fluorescence excitation at 488 nm and higher fluorescence signal levels. The AM ester group results in an uncharged Fluo-4 molecule that can permeate cell membranes. Once inside the cell, the AM ester is cleaved by nonspecific esterases, resulting in a charged form of Fluo-4 that is trapped within the cell.</li>
	<li>Prepare a stock solution of 1 mM carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) in 100% ethanol.</li>
	<li>Prepare a stock solution of 25 mg/ml (w/v) digitonin in distilled H2O.</li>
	<li>Prepare a stock solution of 100 &#181;M thapsigargin in DMSO.</li>
	<li>Prepare a stock solution of 40 mM calcium chloride (CaCl2) in distilled H2O.</li>
</ol>
3. Staining of Cells with TMRM and Fluo-4, AM
<ol>
	<li>Prepare RS staining solution with 20 nM TMRM, 5 &#181;g/ml (w/v) Fluo-4, AM and 0.005% surfactant such as Pluronic F-127 in RS. 
	NOTE: This surfactant is a nonionic polyol that facilitates the solubilization of Fluo-4, AM. Note: 10 &#181;M Verapamil can also be added to inhibit TMRM export by the plasma membrane multidrug transporter if it is expressed in the cell type being analyzed.</li>
	<li>Remove culture media from cells by pipette and wash with 100 &#181;l 1x PBS.</li>
	<li>Remove 1x PBS by pipette and incubate cells in 250 &#181;l RS staining solution for 45 min at room temperature. 
	NOTE: Once the Fluo-4, AM enters the cell, the AM ester is cleaved by cellular esterases. Incubating at room temperature inhibits esterase activity, allowing Fluo-4, AM to enter the mitochondria.</li>
	<li>Remove RS staining solution by pipette and wash cells in 100 &#181;l Ca2+ free HBSS to remove excess Fluo-4, AM and TMRM.</li>
	<li>Prepare IM imaging solution with 25 &#181;g/ml (w/v) digitonin, 200 nM TMRM and 1 &#181;M thapsigargin in IM. 
	NOTE: The concentration of digitonin may need to be adjusted to ensure that permeabilization of the mitochondrial inner membrane does not occur. This can be determined empirically by assessing the intensity of the TMRM signal at different concentrations of digitonin.</li>
	<li>Remove the Ca2+ free HBSS by pipette and add 300 &#181;l of IM imaging solution to the cells. Leave cells to equilibrate at room temperature for 10 min. Cells are now ready for imaging with CaCl2 additions.</li>
</ol>
4. Cell Imaging
<ol>
	<li>Place dish or chambered coverslip with cells in IM imaging solution onto an inverted laser scanning confocal microscope.</li>
	<li>Use setting on the microscope lasers for the excitation and emission spectra of TMRM and Fluo-4. For example, use 543 nm He-Ne and 473 nm argon laser lines to excite TMRM and Fluo-4 respectively. Use a low laser power of approximately 5% to minimize any photo-damage to the cells.</li>
	<li>Set microscope to scan images every 25 sec. Scan cells for 10 min to establish baseline readings for the TMRM and Fluo-4 signals. 
	NOTE: The Fluo-4 signal may be weak or undetectable at resting calcium concentrations.</li>
	<li>Add 3 &#956;l of 40 mM CaCl2 stock solution directly to the cells in 300 &#956;l of IM imaging solution (a 1:100 dilution) and mix gently with a pipette. Pause the image scanning while adding and mixing the CaCl2 if necessary. 
	NOTE: The final free Ca2+ ion concentration [Ca2+] in the IM imaging solution can be calculated using software such as Maxchelator WEBMAXC EXTENDED (<a href="http://www.stanford.edu/%7Ecpatton/maxc.html">http://www.stanford.edu/%7Ecpatton/maxc.html</a>) or Chelator11. A detailed description of how to calculate the final free Ca2+ ion concentration [Ca2+] is described in section 6 below.</li>
	<li>Continue image scanning for approximately 4 min, then repeat CaCl2 additions every 4 min until the desired TMRM or Fluo-4 signals are reached, usually around 8 to 10 additions.</li>
	<li>Using a pipette, add a 1:100 dilution of 1 mM FCCP (final concentration of 10 &#956;M) directly to the cells to dissipate &#916;&#936;m. Image cells for a further 5 min. The experiment is now finished.</li>
</ol>
5. Image Analysis
<ol>
	<li>Once imaging is complete, determine the intensity of the Fluo-4 and TMRM signals using image analysis software, for example ImageJ software with the Bio-Formats plugin (download &#39;bioformats_package.jar&#39; from http://downloads.openmicroscopy.org/bio-formats/ and save it in the &#39;C:\Program Files\ImageJ\plugins&#39; folder).</li>
	<li>Open the imaging file (for example an .oif file) in ImageJ. Click on the selection tool and select a region of interest (ROI) that contains mitochondria. Use the initial TMRM signal to select the ROI.</li>
	<li>Open &#39;Analyze&#62;Tools&#62;ROI Manager&#39;, click &#39;Add&#39; to include the selected ROI for intensity measurement. Multiple ROIs can be selected on a single image for measurement. Repeat previous steps until all ROIs have been added to the ROI Manager.</li>
	<li>Click &#39;More&#62;Multi Measure&#8217;, make sure that &#8216;Measure all slices&#8217; is selected and that &#8216;One Row Per Slice&#8217; is deselected, then click &#8216;OK&#8217; to obtain a table of mean fluorescent intensity of the TMRM and Fluo-4 signals for each ROI at each time point.</li>
	<li>Use data from all ROIs to calculate the average intensities and standard deviation for the TMRM and Fluo-4 signals.</li>
</ol>
6. Calculating the Final Free Ca2+ Ion Concentration [Ca2+]
<ol>
	<li>The free Ca2+ ion concentration [Ca2+] in the IM imaging solution can be determined using software such as Maxchelator WEBMAXC EXTENDED (<a href="http://www.stanford.edu/%7Ecpatton/maxc.html">http://www.stanford.edu/%7Ecpatton/maxc.html</a>) or Chelator11. For example, set the variables in WEBMAXC EXTENDED as follows: Temperature = 24 &#176;C, pH = 7.1, Ionic strength = 0.162, EGTA = 0.002 M, HEDTA = 0.01 M, Ca2+ = 0.0004 M and Mg2+ = 0.0078 M. This results in a final free Ca2+ ion concentration [Ca2+] of 62 nM. Note that these programs cannot take into account every variable, such as reagent purity, accuracy of measurements and pH determination. The most accurate method for determining the free Ca2+ concentration is to use a calibrated Ca2+ selective electrode.</li>
</ol>

<ol>
	<li>Szabadkai, G., Duchen, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondria:+the+hub+of+cellular+Ca2++signaling.">Mitochondria: the hub of cellular Ca2+ signaling.</a> Physiology. 23, 84-94 (2008).</li><li>Jacobson, J., Duchen, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interplay+between+mitochondria+and+cellular+calcium+signalling.">Interplay between mitochondria and cellular calcium signalling.</a> Mol. Cell. Biochem. 256-257, 209-218 (2004).</li><li>Bhosale, G., Sharpe, J. A., Sundier, S. Y., Duchen, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+signaling+as+a+mediator+of+cell+energy+demand+and+a+trigger+to+cell.">Calcium signaling as a mediator of cell energy demand and a trigger to cell.</a> Ann. N. Y. Acad. Sci. 1350, 107-116 (2015).</li><li>Duchen, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondria+calcium-dependent+neuronal+death+and+neurodegenerative+disease.">Mitochondria calcium-dependent neuronal death and neurodegenerative disease.</a> Pflugers Arch. 464, 111-121 (2012).</li><li>Drummond, R. M., Mix, T. C., Tuft, R. A., Walsh, J. V., Fay, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+Ca2++homeostasis+during+Ca2++influx+and+Ca2++release+in+gastric+myocytes+from+Bufo+marinus.">Mitochondrial Ca2+ homeostasis during Ca2+ influx and Ca2+ release in gastric myocytes from Bufo marinus.</a> J. Physiol. 522, 375-390 (2000).</li><li>Hajnoczky, G., Robb-Gaspers, L. D., Seitz, M. B., Thomas, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Decoding+of+cytosolic+calcium+oscillations+in+the+mitochondria.">Decoding of cytosolic calcium oscillations in the mitochondria.</a> Cell. 82, 415-424 (1995).</li><li>Davidson, S. M., Duchen, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+mitochondrial+calcium+signalling+with+fluorescent+probes+and+single+or+two+photon+confocal+microscopy.">Imaging mitochondrial calcium signalling with fluorescent probes and single or two photon confocal microscopy.</a> Methods Mol. Biol. 810, 219-234 (2012).</li><li>Pozzan, T., Rudolf, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurements+of+mitochondrial+calcium+in+vivo.">Measurements of mitochondrial calcium in vivo.</a> Biochim. Biophys. Acta. 1787, 1317-1323 (2009).</li><li>Rizzuto, R., Simpson, A. W., Brini, M., Pozzan, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+changes+of+mitochondrial+Ca2++revealed+by+specifically+targeted+recombinant+aequorin.">Rapid changes of mitochondrial Ca2+ revealed by specifically targeted recombinant aequorin.</a> Nature. 358, 325-327 (1992).</li><li>Pitter, J. G., Maechler, P., Wollheim, C. B., Spat, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondria+respond+to+Ca2++already+in+the+submicromolar+range:+correlation+with+redox+state.">Mitochondria respond to Ca2+ already in the submicromolar range: correlation with redox state.</a> Cell Calcium. 31, 97-104 (2002).</li><li>Schoenmakers, T. J., Visser, G. J., Flik, G., Theuvenet, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CHELATOR:+an+improved+method+for+computing+metal+ion+concentrations+in+physiological+solutions.">CHELATOR: an improved method for computing metal ion concentrations in physiological solutions.</a> BioTechniques. 12, 870-879 (1992).</li><li>McKenzie, M., Duchen, M. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impaired+Cellular+Bioenergetics+Causes+Mitochondrial+Calcium+Handling+Defects+in+MT-ND5+Mutant+Cybrids.">Impaired Cellular Bioenergetics Causes Mitochondrial Calcium Handling Defects in MT-ND5 Mutant Cybrids.</a> PLoS One. 11, e0154371 (2016).</li><li>Berridge, M. J., Lipp, P., Bootman, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+versatility+and+universality+of+calcium+signalling.">The versatility and universality of calcium signalling.</a> Nat. Rev. Mol. Cell Biol. 1, 11-21 (2000).</li><li>Homolya, L., Hollo, Z., Germann, U. A., Pastan, I., Gottesman, M. M., Sarkadi, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+cellular+indicators+are+extruded+by+the+multidrug+resistance+protein.">Fluorescent cellular indicators are extruded by the multidrug resistance protein.</a> J. Biol. Chem. 268, 21493-21496 (1993).</li><li>Fujimoto, K., Chen, Y., Polonsky, K. S., Dorn, G. W. . 2. n. d. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeting+cyclophilin+D+and+the+mitochondrial+permeability+transition+enhances+beta-cell+survival+and+prevents+diabetes+in+Pdx1+deficiency.">Targeting cyclophilin D and the mitochondrial permeability transition enhances beta-cell survival and prevents diabetes in Pdx1 deficiency.</a> Proc. Natl. Acad. Sci. U.S.A. 107, 10214-10219 (2010).</li><li>Rao, V. K., Carlson, E. A., Yan, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+permeability+transition+pore+is+a+potential+drug+target+for+neurodegeneration.">Mitochondrial permeability transition pore is a potential drug target for neurodegeneration.</a> Biochim. Biophys. Acta. 1842, 1267-1272 (2014).</li></ol>

The authors declare that they have no competing financial interests.

Calcium plays a critical role in many cell processes, including muscle contraction, neuronal signaling and cell proliferation13. Increases in cell calcium concentrations are often associated with energy demand, with calcium able to directly stimulate mitochondrial oxidative phosphorylation to raise ATP generation3. It is therefore essential that we have the ability to effectively monitor mitochondrial calcium accumulation and to be able to compare how this function is affec...

Mitochondria play an important role in regulating intracellular Ca2+ concentration by acting as local Ca2+ buffers1. Ca2+ enters the mitochondria via the Ca2+ uniporter, a process driven by the electrochemical gradient that exists across the mitochondrial inner membrane (&#916;&#936;m)2. Once inside the mitochondrial matrix, Ca2+ can activate oxidative phosphorylation by stimulating three rate-limiting dehydrogenases of the citric acid cycle3. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix. If the Ca2+ concentration within the mitochondria becomes very high, mitochondrial permeability transition can be initiated, resulting in the dissipation of &#916;&#936;m, the cessation of oxidative phosphorylation and the induction of cell death signaling pathways4.
The important role that mitochondria play in the spatial buffering of cellular calcium makes the accurate monitoring of mitochondrial calcium critical. Various methods have been established to monitor mitochondrial calcium, including the use of rhodamine based dyes. One such dye, Rhod-2, AM, is quite effective at partitioning to the mitochondria to measure mitochondrial Ca2+ levels5,6. However, care must be used as some dye will accumulate in other organelles, such as liposomes, or remain in the cell cytosol. Nevertheless, downstream analyses can be employed to distinguish these signals from those from the mitochondria7.
Another technique to monitor mitochondrial calcium utilizes fluorescent reporter constructs8. The benefit of these genetically encoded probes is that they can be specifically targeted to the mitochondria by using endogenous N-terminal peptides, for example the N-terminal targeting signal of human COX subunit VIII. This system has been employed to generate a mitochondrial-targeted aequorin probe which has proved extremely useful for investigating mitochondrial calcium signaling9. The main drawback of these genetically encoded probes is that they need to be introduced into the cells by transient expression (which is not feasible for certain cell types and can produce variable results) or by creating stable expression systems (which is time consuming).
To circumvent the problems outlined above, we have developed a new protocol to measure mitochondrial Ca2+ and &#916;&#936;m simultaneously. This protocol is based on a previously described method that adds exogenous calcium to permeabilized cells10. Our protocol has three main advantages over other methods: firstly, we use Fluo-4, AM and TMRM to monitor mitochondrial Ca2+ and &#916;&#936;m, two dyes that have very distinct spectral properties; secondly, the cells are permeabilized so that the Fluo-4 signal is only detecting mitochondrial Ca2+ and not Ca2+ localized to other organelles or the cell cytosol; and thirdly, the use of Fluo-4 to detect mitochondrial Ca2+ allows for fast and simple cell staining, negating any cell transfection or transformation issues that exist if using genetically encoded probes.

<table><tbody><tr><td>Dulbecco&#039;s Modified Eagle Medium (DMEM)</td><td>ThermoFisher</td><td>10566016</td><td></td></tr><tr><td>fetal bovine serum (FBS)</td><td>ThermoFisher</td><td>16000044</td><td></td></tr><tr><td>1x phosphate buffered saline (PBS)</td><td>ThermoFisher</td><td>10010023</td><td></td></tr><tr><td>100x penicillin/streptomycin (p/s)</td><td>ThermoFisher</td><td>15140122</td><td></td></tr><tr><td>0.25% Trypsin/0.25% EDTA</td><td>ThermoFisher</td><td>25200056</td><td></td></tr><tr><td>8-well chambered coverslip</td><td>ibidi</td><td>80826</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>793566</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P9541&amp;nbsp;</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>746452</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>795488</td><td></td></tr><tr><td>D-glucose</td><td>Sigma-Aldrich</td><td>G8270&amp;nbsp;</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>746495</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375&amp;nbsp;</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>M2670&amp;nbsp;</td><td></td></tr><tr><td>EGTA</td><td>Sigma-Aldrich</td><td>E4378&amp;nbsp;</td><td></td></tr><tr><td>HEDTA</td><td>Sigma-Aldrich</td><td>H8126&amp;nbsp;</td><td></td></tr><tr><td>malate</td><td>Sigma-Aldrich</td><td>M1000</td><td></td></tr><tr><td>glutamate</td><td>Sigma-Aldrich</td><td>G1626</td><td></td></tr><tr><td>ADP</td><td>Sigma-Aldrich</td><td>A5285</td><td></td></tr><tr><td>Ca&lt;sup&gt;2+&lt;/sup&gt; free Hank&amp;rsquo;s buffered salt solution (HBSS)&amp;nbsp;</td><td>ThermoFisher</td><td>14175-095</td><td></td></tr><tr><td>tetramethylrhodamine, methyl ester, perchlorate (TMRM)</td><td>ThermoFisher</td><td>T668</td><td></td></tr><tr><td>Verapamil</td><td>Sigma-Aldrich</td><td>V4629</td><td></td></tr><tr><td>Fluo-4 acetoxymethyl ester (Fluo-4, AM)</td><td>ThermoFisher</td><td>F14201</td><td></td></tr><tr><td>dimethyl sulfoxide (DMSO)</td><td>ThermoFisher</td><td>D12345</td><td></td></tr><tr><td>carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP)</td><td>Sigma-Aldrich</td><td>C2920</td><td></td></tr><tr><td>digitonin</td><td>Sigma-Aldrich</td><td>D141</td><td></td></tr><tr><td>thapsigargin</td><td>Sigma-Aldrich</td><td>T9033</td><td></td></tr><tr><td>Pluronic F-127&amp;nbsp;</td><td>ThermoFisher</td><td>P3000MP&amp;nbsp;</td><td></td></tr><tr><td>hemacytometer</td><td>VWR</td><td>631-0925</td><td></td></tr><tr><td>10 cm cell culture dishes</td><td>Corning</td><td>COR430167</td><td></td></tr><tr><td>75 cm&lt;sup&gt;2&lt;/sup&gt; cell culture flasks</td><td>Corning</td><td>COR430641</td><td></td></tr></tbody></table>

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.

We have used this protocol to examine the effects of an MT-ND5 mutation on the ability of 143B cell mitochondria to buffer increases in calcium12. In the example shown here, control 143B cells were loaded with TMRM and Fluo-4, AM before permeabilization with digitonin. After 5 min of imaging, eight sequential additions of a 1:100 dilution of 40 mM exogenous CaCl2 were made, with the final free Ca2+ ion concentration [Ca2+] ...

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

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+ ...

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18.9K Views.  Hudson Institute of Medical Research.  Mitochondriën kan de elektrochemische potentiaal over hun binnenste membraan (Δ Ψm) te gebruiken om calcium sekwestreren (Ca 2+), waardoor ze vorm te geven cytosolische Ca 2+ het signaleren binnen de cel. We beschrijven een werkwijze voor het gelijktijdig meten van mitochondriën Ca2 + opname en ΔΨ m in levende cellen met fluorescerende kleurstoffen en confocale microscopie. 

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

Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ability to accurately measure the influx and efflux of Ca2+ from mitochondria is important for determining the role of mitochondrial Ca2+ handling in these processes. In this report, we present two methods for the measurement of mitochondrial Ca2+ handling in both isolated mitochondria and cultured cells. We first detail a plate reader-based platform for measuring mitochondrial Ca2+ uptake using the Ca2+ sensitive dye calcium green-5N. The plate reader-based format circumvents the need for specialized equipment, and the calcium green-5N dye is ideally suited for measuring Ca2+ from isolated tissue mitochondria. For our application, we describe the measurement of mitochondrial Ca2+ uptake in mitochondria isolated from mouse heart tissue; however, this procedure can be applied to measure mitochondrial Ca2+ uptake in mitochondria isolated from other tissues such as liver, skeletal muscle, and brain. Secondly, we describe a confocal microscopy-based assay for measurement of mitochondrial Ca2+ in permeabilized cells using the Ca2+ sensitive dye Rhod-2/AM and imaging using 2-dimensional laser-scanning microscopy. This permeabilization protocol eliminates cytosolic dye contamination, allowing for specific recording of changes in mitochondrial Ca2+. Moreover, laser-scanning microscopy allows for high frame rates to capture rapid changes in mitochondrial Ca2+ in response to various drugs or reagents applied in the external solution. This protocol can be applied to measure mitochondrial Ca2+ uptake in many cell types including primary cells such as cardiac myocytes and neurons, and immortalized cell lines.

Here, we present two protocols for the measurement of mitochondrial Ca2+ influx in isolated mitochondria and cultured cells. For isolated mitochondria, we detail a plate reader-based Ca2+ import assay using the Ca2+ sensitive dye calcium green-5N. For cultured cells, we describe a confocal microscopy method using the Ca2+ dye Rhod-2/AM.

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ...

Biochemistry

Monitoring Dynamic Changes In Mitochondrial Calcium Levels During Apoptosis Using A Genetically Encoded Calcium Sensor

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, differentiation, and apoptosis1. During apoptosis, calcium accumulation in mitochondria promotes the release of pro-apoptotic factors from the mitochondria into the cytosol2. It is therefore of interest to directly measure mitochondrial calcium in living cells in situ during apoptosis. High-resolution fluorescent imaging of cells loaded with dual-excitation ratiometric and non-ratiometric synthetic calcium indicator dyes has been proven to be a reliable and versatile tool to study various aspects of intracellular calcium signaling. Measuring cytosolic calcium fluxes using these techniques is relatively straightforward. However, measuring intramitochondrial calcium levels in intact cells using synthetic calcium indicators such as rhod-2 and rhod-FF is more challenging. Synthetic indicators targeted to mitochondria have blunted responses to repetitive increases in mitochondrial calcium, and disrupt mitochondrial morphology3. Additionally, synthetic indicators tend to leak out of mitochondria over several hours which makes them unsuitable for long-term experiments. Thus, genetically encoded calcium indicators based upon green fluorescent protein (GFP)4 or aequorin5 targeted to mitochondria have greatly facilitated measurement of mitochondrial calcium dynamics. Here, we describe a simple method for real-time measurement of mitochondrial calcium fluxes in response to different stimuli. The method is based on fluorescence microscopy of 'ratiometric-pericam' which is selectively targeted to mitochondria. Ratiometric pericam is a calcium indicator based on a fusion of circularly permuted yellow fluorescent protein and calmodulin4. Binding of calcium to ratiometric pericam causes a shift of its excitation peak from 415 nm to 494 nm, while the emission spectrum, which peaks around 515 nm, remains unchanged. Ratiometric pericam binds a single calcium ion with a dissociation constant in vitro of ~1.7 &mu;M4. These properties of ratiometric pericam allow the quantification of rapid and long-term changes in mitochondrial calcium concentration. Furthermore, we describe adaptation of this methodology to a standard wide-field calcium imaging microscope with commonly available filter sets. Using two distinct agonists, the purinergic agonist ATP and apoptosis-inducing drug staurosporine, we demonstrate that this method is appropriate for monitoring changes in mitochondrial calcium concentration with a temporal resolution of seconds to hours. Furthermore, we also demonstrate that ratiometric pericam is also useful for measuring mitochondrial fission/fragmentation during apoptosis. Thus, ratiometric pericam is particularly well suited for continuous long-term measurement of mitochondrial calcium dynamics during apoptosis.

This protocol describes a method for real-time measurement of mitochondrial calcium fluxes by fluorescent imaging. The method takes advantage of a circularly permutated YFP-based dual-excitation ratiometric calcium sensor (ratiometric pericam-mt) selectively expressed in mitochondria.

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, ...

Measuring Fast Calcium Fluxes in Cardiomyocytes

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to extracellular agents is predominantly regulated by the phospholipase C&beta;- G&alpha;q pathway localized on the plasma membrane which is stimulated by agents such as acetylcholine 3,4. We have recently found that plasma membrane protein domains called caveolae5,6 can entrap activated G&alpha;q 7. This entrapment has the effect of stabilizing the activated state of G&alpha;q and resulting in prolonged Ca2+ signals in cardiomyocytes and other cell types8. We uncovered this surprising result by measuring dynamic calcium responses on a fast scale in living cardiomyocytes. Briefly, cells are loaded with a fluorescent Ca2+ indicator. In our studies, we used Ca2+ Green (Invitrogen, Inc.) which exhibits an increase in fluorescence emission intensity upon binding of calcium ions. The fluorescence intensity is then recorded for using a line-scan mode of a laser scanning confocal microscope. This method allows rapid acquisition of the time course of fluorescence intensity in pixels along a selected line, producing several hundreds of time traces on the microsecond time scale. These very fast traces are transferred into excel and then into Sigmaplot for analysis, and are compared to traces obtained for electronic noise, free dye, and other controls. To dissect Ca2+ responses of different flux rates, we performed a histogram analysis that binned pixel intensities with time. Binning allows us to group over 500 traces of scans and visualize the compiled results spatially and temporally on a single plot. Thus, the slow Ca2+ waves that are difficult to discern when the scans are overlaid due to different peak placement and noise, can be readily seen in the binned histograms. Very fast fluxes in the time scale of the measurement show a narrow distribution of intensities in the very short time bins whereas longer Ca2+ waves show binned data with a broad distribution over longer time bins. These different time distributions allow us to dissect the timing of Ca2+fluxes in the cells, and to determine their impact on various cellular events.

We present a method to isolate rapid (microsecond) calcium events from slower fluxes in living cells using laser scanning confocal microscopy. The method measures fluorescence intensity fluctuations of calcium indicators by recording line scans of several hundred pixels in a cell. Histogram analysis allows us to isolate the time scales of different calcium fluxes.

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to ...

Mitochondrial Ca2+ Retention Capacity Assay and Ca2+-triggered Mitochondrial Swelling Assay

The production of ATP by oxidative phosphorylation is the primary function of mitochondria. Mitochondria in higher eukaryotes also participate in cytosolic Ca2+ buffering, and the ATP production in mitochondrial can be mediated by intramitochondrial free Ca2+ concentration. Ca2+ retention capacity can be regarded as the capability of mitochondria to retain calcium in the mitochondrial matrix. Accumulated intracellular Ca2+ leads to the permeability of the inner mitochondrial membrane, termed the opening of mitochondrial permeability transition pore (mPTP), which leads to the leakage of molecules with a molecular weight less than 1.5 kDa. Ca2+-triggered mitochondria swelling is used to indicate the mPTP opening. Here, we describe two assays to examine the Ca2+ retention capacity and Ca2+-triggered mitochondrial swelling in isolated mitochondria. After certain amounts of Ca2+ are added, all steps can be completed in one day and recorded by a microplate reader. Thus, these two simple and effective assays can be adopted to assess the Ca2+-related mitochondrial functions.

Mitochondrial Ca2+ Retention Capacity Assay and Ca2+-triggered Mitochondrial Swelling Assay

This protocol aims to describe a method to examine the Ca2+ retention capacity and Ca2+- triggered mitochondrial swelling of isolated mitochondria of SH-SY5Y cells step-by-step.

The production of ATP by oxidative phosphorylation is the primary function of mitochondria. Mitochondria in higher eukaryotes also participate in ...

Neuroscience

A Faster, High Resolution, mtPA-GFP-based Mitochondrial Fusion Assay Acquiring Kinetic Data of Multiple Cells in Parallel Using Confocal Microscopy

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in living cells by photo-conversion of matrix targeted photoactivatable GFP (mtPAGFP) in a subset of mitochondria. The rate at which the photoconverted molecules equilibrate across the entire mitochondrial population is used as a measure of fusion activity. Thus far measurements were performed using a single cell time lapse approach, quantifying the equilibration in one cell over an hour. Here, we scale up and automate a previously published live cell method based on using mtPAGFP and a low concentration of TMRE (15 nm). This method involves photoactivating a small portion of the mitochondrial network, collecting highly resolved stacks of confocal sections every 15 min for 1 hour, and quantifying the change in signal intensity. Depending on several factors such as ease of finding PAGFP expressing cells, and the signal of the photoactivated regions, it is possible to collect around 10 cells within the 15 min intervals. This provides a significant improvement in the time efficiency of this assay while maintaining the highly resolved subcellular quantification as well as the kinetic parameters necessary to capture the detail of mitochondrial behavior in its native cytoarchitectural environment.
Mitochondrial dynamics play a role in many cellular processes including respiration, calcium regulation, and apoptosis1,2,3,13. The structure of the mitochondrial network affects the function of mitochondria, and the way they interact with the rest of the cell. Undergoing constant division and fusion, mitochondrial networks attain various shapes ranging from highly fused networks, to being more fragmented. Interestingly, Alzheimer's disease, Parkinson's disease, Charcot Marie Tooth 2A, and dominant optic atrophy have been correlated with altered mitochondrial morphology, namely fragmented networks4,10,13. Often times, upon fragmentation, mitochondria become depolarized, and upon accumulation this leads to impaired cell function18. Mitochondrial fission has been shown to signal a cell to progress toward apoptosis. It can also provide a mechanism by which to separate depolarized and inactive mitochondria to keep the bulk of the network robust14. Fusion of mitochondria, on the other hand, leads to sharing of matrix proteins, solutes, mtDNA and the electrochemical gradient, and also seems to prevent progression to apoptosis9. How fission and fusion of mitochondria affects cell homeostasis and ultimately the functioning of the organism needs further understanding, and therefore the continuous development and optimization of how to gather information on these phenomena is necessary.
Existing mitochondrial fusion assays have revealed various insights into mitochondrial physiology, each having its own advantages. The hybrid PEG fusion assay7, mixes two populations of differently labeled cells (mtRFP and mtYFP), and analyzes the amount of mixing and colocalization of fluorophores in fused, multinucleated, cells. Although this method has yielded valuable information, not all cell types can fuse, and the conditions under which fusion is stimulated involves the use of toxic drugs that likely affect the normal fusion process. More recently, a cell free technique has been devised, using isolated mitochondria to observe fusion events based on a luciferase assay1,5. Two human cell lines are targeted with either the amino or a carboxy terminal part of Renilla luciferase along with a leucine zipper to ensure dimerization upon mixing. Mitochondria are isolated from each cell line, and fused. The fusion reaction can occur without the cytosol under physiological conditions in the presence of energy, appropriate temperature and inner mitochondrial membrane potential. Interestingly, the cytosol was found to modulate the extent of fusion, demonstrating that cell signaling regulates the fusion process 4,5. This assay will be very useful for high throughput screening to identify components of the fusion machinery and also pharmacological compounds that may affect mitochondrial dynamics. However, more detailed whole cell mitochondrial assays will be needed to complement this in vitro assay to observe these events within a cellular environment.
A technique for monitoring whole-cell mitochondrial dynamics has been in use for some time and is based on a mitochondrially-targeted photoactivatable GFP (mtPAGFP)6,11. Upon expression of the mtPAGFP, a small portion of the mitochondrial network is photoactivated (10-20%), and the spread of the signal to the rest of the mitochondrial network is recorded every 15 minutes for 1 hour using time lapse confocal imaging. Each fusion event leads to a dilution of signal intensity, enabling quantification of the fusion rate. Although fusion and fission are continuously occurring in cells, this technique only monitors fusion as fission does not lead to a dilution of the PAGFP signal6. Co-labeling with low levels of TMRE (7-15 nM in INS1 cells) allows quantification of the membrane potential of mitochondria. When mitochondria are hyperpolarized they uptake more TMRE, and when they depolarize they lose the TMRE dye. Mitochondria that depolarize no longer have a sufficient membrane potential and tend not to fuse as efficiently if at all. Therefore, active fusing mitochondria can be tracked with these low levels of TMRE9,15. Accumulation of depolarized mitochondria that lack a TMRE signal may be a sign of phototoxicity or cell death. Higher concentrations of TMRE render mitochondria very sensitive to laser light, and therefore great care must be taken to avoid overlabeling with TMRE. If the effect of depolarization of mitochondria is the topic of interest, a technique using slightly higher levels of TMRE and more intense laser light can be used to depolarize mitochondria in a controlled fashion (Mitra and Lippincott-Schwartz, 2010). To ensure that toxicity due to TMRE is not an issue, we suggest exposing loaded cells (3-15 nM TMRE) to the imaging parameters that will be used in the assay (perhaps 7 stacks of 6 optical sections in a row), and assessing cell health after 2 hours. If the mitochondria appear too fragmented and cells are dying, other mitochondrial markers, such as dsRED or Mitotracker red could be used instead of TMRE.
The mtPAGFP method has revealed details about mitochondrial network behavior that could not be visualized using other methods. For example, we now know that mitochondrial fusion can be full or transient, where matrix content can mix without changing the overall network morphology. Additionally, we know that the probability of fusion is independent of contact duration and organelle dimension, is influenced by organelle motility, membrane potential and history of previous fusion activity8,15,16,17.
In this manuscript, we describe a methodology for scaling up the previously published protocol using mtPAGFP and 15nM TMRE8 in order to examine multiple cells at a time and improve the time efficiency of data collection without sacrificing the subcellular resolution. This has been made possible by the use of an automated microscope stage, and programmable image acquisition software. Zen software from Zeiss allows the user to mark and track several designated cells expressing mtPAGFP. Each of these cells can be photoactivated in a particular region of interest, and stacks of confocal slices can be monitored for mtPAGFP signal as well as TMRE at specified intervals. Other confocal systems could be used to perform this protocol provided there is an automated stage that is programmable, an incubator with CO2, and a means by which to photoactivate the PAGFP; either a multiphoton laser, or a 405 nm diode laser.

Mitochondrial fusion was measured by tracking the equilibration of photoconverted matrix-targeted GFP across the mitochondrial network over time. Thus far, only one cell could be subjected to an hour long kinetic analysis at a time. We present a method that simultaneously measures multiple cells, thereby speeding up the data collection process.

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in ...

Simultaneous Measurements of Intracellular Calcium and Membrane Potential in Freshly Isolated and Intact Mouse Cerebral Endothelium

Cerebral arteries and their respective microcirculation deliver oxygen and nutrients to the brain via blood flow regulation. Endothelial cells line the lumen of blood vessels and command changes in vascular diameter as needed to meet the metabolic demand of neurons. Primary endothelial-dependent signaling pathways of hyperpolarization of membrane potential (Vm) and nitric oxide typically operate in parallel to mediate vasodilation and thereby increase blood flow. Although integral to coordinating vasodilation over several millimeters of vascular length, components of endothelium-derived hyperpolarization (EDH) have been historically difficult to measure. These components of EDH entail intracellular Ca2+ [Ca2+]i increases and subsequent activation of small- and intermediate conductance Ca2+-activated K+ (SKCa/IKCa) channels.
Here, we present a simplified illustration of the isolation of fresh endothelium from mouse cerebral arteries; simultaneous measurements of endothelial [Ca2+]i and Vm using Fura-2 photometry and intracellular sharp electrodes, respectively; and a continuous superfusion of salt solutions and pharmacological agents under physiological conditions (pH 7.4, 37 &#176;C). Posterior cerebral arteries from the Circle of Willis are removed free of the posterior communicating and the basilar arteries. Enzymatic digestion of cleaned posterior cerebral arterial segments and subsequent trituration facilitates removal of adventitia, perivascular nerves, and smooth muscle cells. Resulting posterior cerebral arterial endothelial &#34;tubes&#34; are then secured under a microscope and examined using a camera, photomultiplier tube, and one to two electrometers while under continuous superfusion. Collectively, this method can simultaneously measure changes in endothelial [Ca2+]i and Vm in discrete cellular locations, in addition to the spreading of EDH through gap junctions up to millimeter distances along the intact endothelium. This method is expected to yield a high-throughput analysis of the cerebral endothelial functions underlying mechanisms of blood flow regulation in the normal and diseased brain.

Demonstrated here are protocols for (1) freshly isolating intact cerebral endothelial "tubes" and (2) simultaneous measurements of endothelial calcium and membrane potential during endothelium-derived hyperpolarization. Further, these methods allow for pharmacological tuning of endothelial cell calcium and electrical signaling as individual or interactive experimental variables.

Cerebral arteries and their respective microcirculation deliver oxygen and nutrients to the brain via blood flow regulation. Endothelial cells line the ...

Measurement of Mitochondrial Membrane Potential In Vivo using a Genetically Encoded Voltage Indicator

Mitochondrial membrane potential (MMP, &#916;&#936;m) is critical for mitochondrial functions, including ATP synthesis, ion transport, reactive oxygen species (ROS) generation, and the import of proteins encoded by the nucleus. Existing methods for measuring &#916;&#936;m typically use lipophilic cation dyes, such as Rhodamine 800 and tetramethylrhodamine methyl ester (TMRM), but these are limited by low specificity and are not well-suited for in vivo applications. To address these limitations, we have developed a novel protocol utilizing genetically encoded voltage indicators (GEVIs). Genetically encoded voltage indicators (GEVIs), which generate fluorescent signals in response to membrane potential changes, have demonstrated significant potential for monitoring plasma membrane and neuronal potentials. However, their application to mitochondrial membranes remains unexplored. Here, we developed protein-based mitochondrial-targeted GEVIs capable of detecting &#916;&#936;m fluctuations in cells and the motor cortex of living animals. The mitochondrial potential indicator (MPI)offers a non-invasive approach to study &#916;&#936;m dynamics in real-time, providing a method to investigate mitochondrial function under both normal and pathological conditions.

This protocol describes the application of mitochondria-targeted genetically encoded voltage indicators (GEVIs). These GEVIs offer a significant advantage over traditional mitochondrial membrane potential dyes by enabling specific, in vivo, and real-time monitoring of mitochondrial membrane potential.

Mitochondrial membrane potential (MMP, &#916;&#936;m) is critical for mitochondrial functions, including ATP synthesis, ion transport, reactive oxygen ...

Quantification of Mitochondrial Membrane Potential and Superoxide Levels

Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, various mitochondrial quality control mechanisms cooperate to maintain a healthy mitochondrial network. One such pathway is mitophagy, where PTEN-induced kinase 1 (PINK1) and Parkin phospho-ubiquitination of damaged mitochondria facilitate autophagosome sequestration and subsequent removal from the cell via lysosome fusion. Mitophagy is important for cellular homeostasis, and mutations in Parkin are linked to Parkinson's disease (PD). Due to these findings, there has been a significant emphasis on investigating mitochondrial damage and turnover to understand the molecular mechanisms and dynamics of mitochondrial quality control. Here, live-cell imaging was used to visualize the mitochondrial network of HeLa cells, to quantify the mitochondrial membrane potential and superoxide levels following treatment with carbonyl cyanide m-chlorophenyl hydrazone (CCCP), a mitochondrial uncoupling agent. In addition, a PD-linked mutation of Parkin (ParkinT240R) that inhibits Parkin-dependent mitophagy was expressed to determine how mutant expression impacts the mitochondrial network compared to cells expressing wild-type Parkin. The protocol outlined here describes a simple workflow using fluorescence-based approaches to quantify mitochondrial membrane potential and superoxide levels effectively.

Author Spotlight: Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells  

This technique describes an effective workflow to visualize and quantitatively measure mitochondrial membrane potential and superoxide levels within HeLa cells using fluorescence-based live imaging.

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, ...

Simultaneous Measurement of Intracellular Calcium and Membrane Potential in a Mouse Cerebral Endothelium

Source: Hakim, M. A. et al. <a href="https://app.jove.com/v/58832">Simultaneous Measurements of Intracellular Calcium and Membrane Potential in Freshly Isolated and Intact Mouse Cerebral Endothelium.</a> J. Vis. Exp. (2019)
The video demonstrates the simultaneous measurement of intracellular calcium levels and changes in membrane potential in endothelial cells of mouse cerebral artery segments. The isolated endothelial tube is loaded with a fluorescent calcium indicator and treated with a drug that induces intracellular calcium release, resulting in a change in membrane potential. The indicator&#39;s fluorescence enables the measurement of calcium levels, and a recording electrode is inserted into a cell to measure changes in membrane potential.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Dissection and Isolation of ...

JoVE Encyclopedia of Experiments

Neurophysiology

Imaging and Optical Techniques

Calcium Influx Assay to Measure Mitochondrial Calcium Uptake in Cultured Cells

Source: Maxwell, J.T., et al., <a href="https://app.jove.com/v/57225">Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells.</a> J. Vis. Exp. (2018).
This video describes a confocal microscopy imaging-based assay to measure mitochondrial calcium uptake in cultured cells. The assay uses the calcium-sensitive fluorescent dye Rhod-2/AM to measure the mitochondrial calcium in permeabilized cultured cells.

Source: Maxwell, J.T., et al., Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells. J. Vis. Exp. (2018).
This video ...

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

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