We kindly thank Mr. Chao Fang for cell culture. This work was supported by grants from the Natural Science Foundation of China (Grant No. 81570403, 81371284); Anhui Provincial Natural Science Foundation (Grant No. 1408085MH158); Outstanding Young Investigator of Anhui Medical University; Supporting Program for Excellent Young Talents in Universities of Anhui Province.

1. Cell Culture and Transfection
<ol>
	<li>Cell culture 
	Note: HN4, an HNSCC cell line, was derived from patients with HNSCC14.
	<ol>
		<li>Culture HN4 cells in Dulbecco&#39;s modified Eagle medium (DMEM, 4.5 g/L glucose) supplemented with 10% fetal bovine serum and antibiotics (100 U/mL penicillin and 100 &#181;g/mL streptomycin). Incubate cells at 37 &#176;C with 5% CO2.</li>
	</ol>
	</li>
	<li>Cell transfection
	<ol>
		<li>One day before transfection, plate 5 x 105 cells in 2 mL/well of culture medium without antibiotics in 6-well plates.</li>
		<li>Dilute 100 pmol CLIC4 siRNA or scrambled SiRNA in 250 &#181;L of reduced serum media, gently. Dilute 5 &#181;L liposome reagent in 250 &#181;L of reduced serum media and incubate for 5 min. Combine the diluted siRNA with the diluted liposome reagent, mix gently, and incubate for 20 min at room temperature.</li>
		<li>Add the mixture to each well containing cells and medium. Mix gently by rocking the plate back-and-forth. Incubate the cells at 37 &#176;C in a CO2 incubator for 24 h before proceeding with the following Rh123 labeling procedure. Replace the medium with normal growth medium 4 h after transfection.</li>
	</ol>
	</li>
</ol>
2. Rh123 Labeling
Note: Rh123 was utilized to measure the MMP in HN4 cells1.
<ol>
	<li>After CLIC4 siRNA treatment (Section 1), use HN4 cells in 6-well plates for the following treatment. The amount of HN4 cells in each well is enough to repeat the experiment 10 times on coverslips.</li>
	<li>Use tweezers to put circular coverslips on the bottom of new 12-well plates. The number of plates is set according to the experimental design (Figure 1A).</li>
	<li>Put 150 &#181;L of 100 &#181;g/mL polylysine on the middle of every coverslip inside the 12-well plates and wait 2 min. Pay attention to not put polylysine outside the coverslips. Then remove the polylysine by pipettor thoroughly (Figure 1B).</li>
	<li>Remove the culture medium of the HN4 cells inside the dish and add 1 mL 0.25% trypsin to detach the cells. After 6 min, add 2 mL culture medium to neutralize the trypsin. Gently mix the HN4 cells by pipettor for 5 times and seed 2 - 6 &#1093; 104 cells on the circular coverslips. Place in an incubator at 37 &#176;C with 5% CO2 (Figure 1C).</li>
	<li>After 8 h incubation, incubate the HN4 cells with 2 &#181;mol/L Rh123 for 40 min at 37 &#176;C. Gently mix the medium up-and-down several times to ensure even distribution of Rh123 in the medium (Figure 1D).</li>
	<li>Prepare enough normal physiological saline solution (NPSS); to make NPSS (in mmol/L): 140 NaCl, 5 KCl, 1 MgCl2, 1 CaCl2, 10 glucose, and 5 HEPES, pH 7.4). Preheat to 37 &#176;C in a water bath.</li>
	<li>Use tweezers to remove the coverslips and wash the remaining liquid via briefly placing the coverslips into the preheated NPBS buffer (Figure 1E).</li>
	<li>Place the coverslips on the bottom of the 500 &#181;L stainless steel chamber (see Table of Materials) with a replaceable 25 mm glass coverslip bottom sealed in place by an o-ring. Fix it by tightening the lid of the chamber (Figure 1E).</li>
	<li>Add 450 &#181;L preheated NPSS buffer to the assembled chamber and put the bath under the visual field of the fluorescence microscope or confocal laser scanning fluorescence microscope (Figure 1E).</li>
</ol>
3. Detection of the MMP
<ol>
	<li>Common fluorescence microscope
	<ol>
		<li>Turn on the fluorescence microscope following the manufacturer&#39;s instructions. 
		NOTE: Common fluorescence microscope was accomplished with a mercury lamp fiberoptic light source, a FITC filter set for Rh123 (480/40 excitation filter, 505 LP dichroic mirror and 535/40 emission filter) at room temperature. 20X&#160;objective was used to perform live-cell imaging.</li>
		<li>Open the imaging system software and select the &#34;New&#34; button from the command bar to begin a new experiment (Figure 2A).</li>
		<li>Adjust the visual field and focus to find the location of the cells in white light.</li>
		<li>Turn off the light and push the button to close white light. Click the &#34;Focus&#34; button from the Command bar and select &#34;Start Focusing&#34; to switch on the fluorescence. Find the location of the cells again via microscope with a 507 nm excitation and 529 nm long pass emission wavelength (Set before the experiment). Adjust the visual field and focus again if needed (Figure 2B).</li>
		<li>Select a visual field containing only 20 to 30 separated HN4 cells. The change of intracellular fluorescence signal for each cell will be recorded accurately.</li>
		<li>Once a clear visual field is achieved, click &#34;Close Focusing&#34; from the Focus bar. Click &#34;Acq One&#34; from the Command Bar to acquire one set of images. Click &#34;Regions&#34; from the Command Bar and click the &#34;OK&#34; button to edit measurement regions. 
		NOTE: From the Edit Region bar, different region shapes can be used depending on the study.</li>
		<li>To fit for different shapes of individual cells, select the &#34;Trace Region&#34; tool and circle the region of one single cell and double click when finished. Repeat the procedure until all cells are circled (Figure 2C). Click &#34;Done&#34; and select &#34;Save Images&#34; to find the saved location.</li>
		<li>Click &#34;Zeroclock &#34;and quickly push F4 to start monitoring the change of fluorescence intensity. Record the real-time change of fluorescence intensity once every 5 s for 5 min (Figure 2D).</li>
		<li>When the baseline becomes stable, add 50 &#181;L NPSS mixed with 0.5 &#181;L of 100 mmol/L ATP to the chamber via pipettor. Remember to not touch the chamber to avoid removing the visual field (Figure 2E). 
		NOTE: The fluorescence image will be recorded for 20 min and then the experiment can end via manual operation. The time duration is routinely settled for 20 min in order to observe the entire fluorescence change. However, manual operation to stop capture is applied when unexpected factors influence the stability of the curve or when the entire process takes less than 20 min.</li>
	</ol>
	</li>
	<li>Confocal laser scanning fluorescence microscope
	<ol>
		<li>Turn on the confocal laser scanning fluorescence microscope following the manufacturer&#39;s instructions, and open the bundled software. 
		NOTE: Here, Rh123 was excited by an Argon laser set at 488 nm and the emitted signal was recorded over the range of 545 - 700 nm via application of the TD488/543/633 filter.</li>
		<li>Click &#34;Objective&#34; under the &#34;Acquire&#34; menu to select 63X/1.4 N.A. oil objective lens. Observe the specimen and find a clear visual field under the light field.</li>
		<li>Push the &#34;TL/IL&#34; button to switch to fluorescence mode and select the correct fluorescence filters to find the location of the cell under darkness via microscope. Push &#34;SHUTTER&#34; to protect the specimen when observation is finished.</li>
		<li>Under the &#34;Acquire&#34; menu, adjust suitable PMT channel and click &#34;Achieve&#34; to activate the settled optical path (Figure 3A). Click on the &#34;Configuration&#34; menu and choose &#34;Laser&#34; to determine the needed laser type. &#34;Argon&#34; is recommended for this protocol (Figure 3B).</li>
		<li>Click &#34;Live&#34; to get real-time image and make sure the visual field contains only 20 to 30 separated HN4 cells. The change of intracellular fluorescence signal for each cell will be recorded accurately.</li>
		<li>Select &#34;xyt&#34; in Acquisition Mode in the &#34;Acquisition&#34; submenu under the &#34;Acquire&#34; menu (Figure 3C). Set 5 s and 20 min for time interval and duration, respectively. Click&#34; Apply&#34; when all parameters are settled.</li>
		<li>Under the &#34;Quantify&#34; menu, use the &#34;Draw Polyline&#34; tool to circle the recording area of each cell. Select &#34;Line Profile&#34; and choose &#34;Start&#34; to conduct observation. Record the real-time change of fluorescence intensity for 5 min (Figure 3D - E).</li>
		<li>When the baseline becomes stable, add 50 &#181;L NPSS mixed with 0.5 &#181;L of 100 mmol/L ATP to the chamber via pipettor. Remember to not touch the chamber to avoid removing the visual field. 
		NOTE: The fluorescence image will be recorded for 20 min and then the experiment is completed.</li>
	</ol>
	</li>
</ol>
4. Statistical Analysis
<ol>
	<li>Common fluorescence microscope
	<ol>
		<li>Turn on the fluorescence microscope routinely following the operator&#39;s manual. Open the imaging system software and select the &#34;Open&#34; button from the command bar to open a saved experiment. Click &#34;Regions&#34; from the &#39;Command Bar&#39; and click &#34;OK&#34; button to edit measurement regions.</li>
		<li>Select the &#34;Trace region&#34; tool and circle the region of one single cell and double click when done. Repeat the procedure until all cells have been circled. Click &#34;Done&#34; and select the &#34;F4: forward&#34; button to obtain a trace showing fluorescence intensity.</li>
		<li>Once finished, click the down arrow button located on the lower right corner of the graph and click &#34;Show Graph Data&#34; (Figure 2F). Click &#34;OK&#34; twice and choose the &#34;Log Data&#34; button to open the interface of data processing software. Click &#34;Log Data&#34; again and find the records of real time fluctuation of fluorescence intensity appear on the spreadsheet. 
		NOTE: For each cell, changes in MMP were displayed as the ratio of fluorescence relative to the intensity before the application of ATP or TG (F1/F0). Each fluorescence intensity data were the average of 20 - 30 cells. Data for Rh123 are presented as mean &#177; SE. Results from 2 groups were tested by the two-tailed independent t test and P value &#60;0.05 was considered as statistical significance. Statistical analysis software was applied to conduct the statistical analyses in this experiment.</li>
	</ol>
	</li>
	<li>Confocal laser scanning fluorescence microscope
	<ol>
		<li>Turn on the confocal laser microscope and log into the bundled software following the manufacturer&#39;s instructions.</li>
		<li>Select &#34;Fire&#34; to open a recorded experiment.</li>
		<li>Select the &#34;Stack profile&#34; tool under the &#34;Quantify&#34; menu, choose &#34;All in one&#34; from &#34;Sort charts by Channels and ROIs.&#34;</li>
		<li>Use the &#34;Draw polygon&#34; tool to circle cell regions for observation.</li>
		<li>Select the &#34;Graph&#34; menu and click the right mouse button, and choose &#34;Export to excel&#34; to add the values of fluorescent intensity to a spreadsheet. The following analysis is consistent with normal fluorescence microscopes (Figure 3F).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

It is well documented that Cl&#8722; channels are essential in maintaining the hemostasis of the internal environment and play important roles in cell proliferation and apoptosis15,16. Therefore, understanding the relationship between ion channel-targeted intervention and apoptosis is of great need and significance to find a better therapeutic approach for various cancers17. Mitochondria maintain the normal biological state of a cell and its function highly relates to its membrane permeability and transmembrane potential. Apart from cancer, accumulating neuropathological investigations have documented that mitochondrial abnormalities contribute to myopathy, cardiovascular disease, and neurological symptoms including sensorineural deafness, cerebellar ataxia, dementia, and epilepsy18,19. All these have spurred the research in mitochondrial-targeted intervention with which to ameliorate clinical treatment. Rh123, a mitochondrion-targeted fluorescence dye, can penetrate the cell membrane, and serves as a universal fluorescence probe to detect MMP. In the early state of cell apoptosis, opening of MPTP elevates the mitochondrial membrane permeability allowing Rh123 to be released outside the mitochondria and finally stronger green fluorescence signal may be detected. Under this situation, bilateral ions distribute freely and lead to the rapid drop of MMP causing decoupling of the electron transport chain and decreased ATP production, which severely affects the normal energy supply of cells. Additionally, opening of MPTP triggers the outflow of pro-apoptotic bioactive compounds including Cyt C, apoptosis inducing factor, apoptotic protease activating factor-1, and ROS, and finally leads to irreversible apoptosis20,21.
ATP may induce mitochondrial membrane depolarization as demonstrated by elevated fluorescence of Rh123. By comparing ATP-induced mitochondrial membrane depolarization of cells under different treatments, it is possible to decipher the biological function of the change in mitochondria and the degree and state of apoptosis22. Both common fluorescence microscope and confocal laser scanning fluorescence microscope can achieve real-time monitoring of MMP via recording the fluctuation of fluorescence intensity of Rh123, but advantages and shortcomings of both methods need to be considered. Compared with the common fluorescence microscope, the images captured by confocal laser scanning fluorescence microscope have higher resolution and quality. Therefore, more subcellular details can be visualized. Additionally, it diminishes the impacts of the light cross-talk when multiple fluorescence dyes are used. Hence, the confocal microscope can precisely record the real-time fluorescence fluctuation of Rh123 and reflect the real cellular bioactivity. However, our methods to monitor MMP need continuously recording for a relative long time and a series of cytotoxins, including single oxygen and ROS, may be produced under the laser radiation leading to cell damage23. Moreover, high laser intensity usually causes fading of fluorescence dyes during the continuous scanning24. By contrast, the common fluorescence microscope cannot take images with high quality like the confocal microscope, but it does not have the aforementioned adverse effects and thus makes long time recording possible. By comparing our data from both common fluorescence microscope and confocal microscope, no obvious difference could be found, which indicates that the common fluorescence microscope is competent enough for monitoring the fluctuation of MMP, as well as cost-effective and more convenient data analysis. HN4 cells were seeded on coverslips before the experiment. Although polylysine is applied to enhance the cellular attachment, a small fraction of cells still detached from the coverslips and rough operation leads to the impairment of cell viability. Besides, adding Rh123 or ATP into the chamber, care should be taken to not touch the chamber and coverslips.
Besides Rh123, DioC6, JC-1, and tetramethyl rhodamine methyl ester (TMRM) are fluorescence dyes that may be used to detect the MMP. Instead of real-time recording via microscope, flow cytometry also can accomplish this task. However, recording the real-time change of fluorescence intensity with the microscope is more suitable to discover the cellular bioactivity and generate intuitionistic images, giving more reliable results.

Rh123 is a cationic fluorescence dye, which serves as an indicator for transmembrane potential. Rh123 is capable of penetrating the cell membrane and entering the mitochondrial matrix depending on the potential difference of inside and outside the membrane1. Apoptosis leads to damage of mitochondrial membrane integrity. The mitochondrion permeability transition pore (MPTP) will open and lead to collapse of the MMP, which in turn results in the release of Rh123 to the outside of the mitochondria. Finally, stronger green fluorescence signal will be detected under fluorescence microscope. It is well documented that depletion of the MMP and elevated membrane permeability are early signs of cell apoptosis2. Therefore, Rh123 may be applied to the detection of MMP change and the occurrence of cell apoptosis.
As the 6th most common carcinoma in the world, head and neck cancer severely deteriorates a person&#39;s health3. Although many approaches were developed in recent years, clinical outcome of treatment for patients suffering from head and neck squamous cell carcinoma (HNSCC) is still not ideal4. Exploring new therapeutic methods can improve the treatment for HNSCC5. Ion channels involving numerous biological processes display an important role in the development of different cancers6. Partial or total participation of Cl- channels are highly involved in various properties of neoplastic transformation including active migration, high rate of proliferation and invasiveness. In light of this, the CLIC, a novel protein family, has been listed as a promising class of therapeutic targets for cancer treatment6,7. Recent studies have revealed that members of the CLIC family including CLIC1, CLIC4, and CLIC5, localize to cardiac mitochondrial and the reactive oxygen species (ROS) level is upregulated by CLIC5, indicating the functional role of mitochondrial-located Cl- channels in the apoptotic response8. CLIC4, one members of the CLIC family (also known as mtCLIC, P64H1, and RS43), has been most extensively studied for its apoptotic regulation properties in cancer cells and subcellular location including Golgi, endoplasmic reticulum, and mitochondrion in human keratinocytes7,9,10. The expression profile of CLIC4 was regulated by tumor necrosis factor-&#945; (TNF-&#945;), P53, and external stimulus. Overexpression and downregulation of CLIC4 trigger an apoptotic response mainly through the mitochondrial pathway accompanied with the imbalance of Bcl-2 family members, activation of caspase cascade, and release of cytochrome C11,12,13. Therefore, MMP measurement is crucial to explore CLIC4-related apoptosis, and Rh123 serves as an ideal fluorescence indicator.
The present study describes a detailed protocol for the detection of MMP to study CLIC4 knockdown-induced apoptosis in HN4 cells. Rh123 is used as a fluorescence probe to observe the change of the MMP. Under common fluorescence microscope and confocal laser scanning fluorescence microscope, the real-time fluctuation of the MMP may be resolved. We discuss the advantages and limitations of the application of confocal laser scanning fluorescence microscope in detail, and also compare it with other methods. This protocol also can be applied to other apoptosis-related studies.

<table><tbody><tr><td>HNSCC cells</td><td>ATCC</td><td>CRL-3241</td><td></td></tr><tr><td>Polylysine</td><td>Thermo Fisher Scientific</td><td>P4981</td><td></td></tr><tr><td>Specific siRNA for human CLIC4</td><td>Biomics</td><td>NM_013943</td><td>(accession numbers, NM_013943; corresponding to the cDNA sequence&lt;br/&gt; 5-GCTGAAGGAGGAGGACAAAGA-3) and scrambled siRNA (5 ACGCGUAACGCGGGAAUUU-3) were designed and obtained from Biomics Company</td></tr><tr><td>Lipofectamine 2000 Transfection Reagent&amp;nbsp;</td><td>Thermo Fisher Scientific</td><td>L3000-015</td><td></td></tr><tr><td>Opti-MEM I Reduced Serum Medium, GlutaMAX</td><td>Thermo Fisher Scientific</td><td>51985-042</td><td></td></tr><tr><td>Rhodamine 123, FluoroPure grede</td><td>Thermo Fisher Scientific</td><td>R22420</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;smodified Eagle medium (DMEM, 4.5 g/L glucose)</td><td>Gibco</td><td>11965-084</td><td></td></tr><tr><td>Fetal Bovine Serum, Qualified, Australia Origin</td><td>Gibco</td><td>10099141</td><td></td></tr><tr><td>Trypsin-EDTA Solution</td><td>Beyotime</td><td>C0201</td><td></td></tr><tr><td>Antibiotic-Antimycotic, 100X</td><td>Gibco</td><td>15240062</td><td></td></tr><tr><td>Laser Scanning Confocal Microscopy</td><td>Leica Microsystems GmbH</td><td>LEICA.SP5-DMI6000-DIC</td><td></td></tr><tr><td>Nikon Eclipse TE300 Inverted Microscope</td><td>Nikon</td><td>N/A</td><td></td></tr><tr><td>Metaflour, V7.5.0.0</td><td>Universal Imaging Corporation</td><td>N/A</td><td></td></tr><tr><td>Leica application suite, v2.6.0.7266</td><td>Leica Microsystems GmbH</td><td>N/A</td><td></td></tr><tr><td>Microsoft office Excel 2007</td><td>Microsoft</td><td>N/A</td><td></td></tr><tr><td>Sigma Plot 12.5</td><td>Systat Software</td><td>N/A</td><td></td></tr><tr><td>Attofluor Cell Chamber</td><td>Thermo Fisher Scientific</td><td>A7816</td><td></td></tr></tbody></table>

detection of mitochondria membrane potential to study clic4 knockdown-induced hn4 cell apoptosis in vitro

Depletion of the mitochondrial membrane potential (MMP, &#916;&#936;m) is considered the earliest event in the apoptotic cascade. It even occurs ahead of nuclear apoptotic characteristics, including chromatin condensation and DNA breakage. Once the MMP collapses, cell apoptosis will initiate irreversibly. A series of lipophilic cationic dyes can pass through the cell membrane and aggregate inside the matrix of mitochondrion, and serve as fluorescence marker to evaluate MMP change. As one of the six members of the Cl- intracellular channel (CLIC) family, CLIC4 participates in the cell apoptotic process mainly through the mitochondrial pathway. Here we describe a detailed protocol to measure MMP via monitoring the fluorescence fluctuation of Rhodamine 123 (Rh123), through which we study apoptosis induced by CLIC4 knockdown. We discuss the advantages and limitations of the application of confocal laser scanning and normal fluorescence microscope in detail, and also compare it with other methods.

In the present study, Rh123 was applied to detect the MMP. Initially, HN4 cells were cultured for the following fluorescence staining experiments. Tweezers were used to put circular coverslips on the bottom of 6-well plates (Figure 1A). The coverslips were coated with polylysine for 5 min and then the polylysine removed via pipettor (Figure 1B). Then HN4 cells were trypsinized and seeded on the coverslips placed in the bottom of 6-well plates. Next, the appropriate amount of Rh123 was added and mixed well (Figure 1C-D). After washing with the preheated NPSS, the coverslip was assembled into the viewing chamber. The appropriate amount of NPSS was added for the following experiment (Figure 1E).
After the procedure of Rh123 staining, both the common fluorescence microscope and confocal microscope were used to record the real-time Rh123 fluorescence with different steps and software. For the common fluorescence microscope, the imaging software was turned on to create a new experiment. The shutter was opened to show the green fluorescence light and the focus and location were adjusted to obtain an appropriate visual field. After taking a cell image, the shape trace tools were used to draw the region of every single cell and then record the fluorescence change. Once the baseline became stable, ATP was added into the chamber to trigger mitochondrial membrane depolarization. In the experiment of confocal laser scanning fluorescence microscope, the bundled software was opened and a suitable optical path was set. After the argon light source was chosen, we adjusted the focus and position of the chamber to achieve a clear visual of the cell shapes on screen. The &#34;xyt&#34; scan mode was selected and the &#34;Draw polygon&#34; tool was applied to circle the cell region to record the fluorescence change within every single cell. To analyze the raw data, F0 was the value of fluorescence intensity before ATP application and F1 was the maximum value of fluorescence intensity after ATP application. F1/F0 was used to assess the mitochondrial membrane depolarization (Figure 2 and Figure 3). From Figure 4B and Figure 5B, the levels of fluorescent intensity were elevated following the treatment of ATP and decreased back to the baseline after about 10 min. Figure 4C and Figure 5C clearly demonstrate that the peak of fluorescence intensity of Rh123 was higher in the group under the CLIC4 siRNA treatment than that in control group. Figure 4A and Figure 5A show representative images of the fluorescent change in each stage. Compared with the common fluorescence microscope, the confocal microscope obtained a higher quality of image, showing cell nucleus and mitochondria. However, when comparing the data from the two methods, no significant differences were found. Results from fluorescence and confocal microscope demonstrated that knockdown of CLIC4 enhanced ATP-induced mitochondrial membrane depolarization in HN4 cells (Figure 4 and Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56317/56317fig1.jpg" /> 
Figure 1. Rh123 labeling. (A) Tweezers were used to put circular coverslips on the bottom of 6-well plates. (B) Coating coverslips with polylysine via pipettor for 5 min and finally suction back into pipettor to remove the polylysine. (C) HN4 cells were seeded on the coverslips on the bottom of 6-well plates. (D) Adding a suitable amount of Rh123 and mixing well. (E) Washing the coverslips with preheated NPPS and assembling the chamber with a suitable amount of NPSS for the following experiment. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56317/56317fig1large.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/56317/56317fig2.jpg" /> 
Figure 2. Steps to monitor real-time fluctuation of Rh123 fluorescence by common fluorescence microscope. (A) Open a new experiment. (B) Start focusing and choose a suitable visual field under fluorescence. (C) Using the &#34;Regions&#34; tool from the Command Bar to edit observation regions. (D) Click &#34;Zero clock&#34; and quickly push F4 to start monitoring the change of fluorescence intensity. (E) Adding ATP (100 &#181;mol/L) to trigger mitochondrial membrane depolarization after the baseline becomes stable. (F) Once the experiment is finished, click the down arrow located on the lower right corner of the graph to export data. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56317/56317fig2large.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/56317/56317fig3.jpg" /> 
Figure 3. Steps to monitor real-time fluctuation of Rh123 fluorescence by confocal laser scanning fluorescence microscope. (A) Select the PMT channel and set a suitable wavelength range under the &#34;Acquire&#34; menu. (B) Select the correct laser type from &#34;Current available laser&#34; and set its power. (C) Choose the &#34;xyt&#34; observing mode and set a series of recording parameters. (D) Choose the tools &#34;Stack Profile&#34; and &#34;All in One&#34; from &#34;Sort charts by Channels and ROIs.&#34;&#160;(E) Use the tool &#34;Draw polygon&#34; to circle the cell region for the real-time data. (F) Export the values of fluorescent intensity to data processing software. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56317/56317fig3large.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/56317/56317fig4.jpg" /> 
Figure 4. Effect of CLIC4 knockdown on ATP-induced mitochondrial membrane depolarization resolved by common fluorescence microscope in HN4 cells. (A) Representative images of fluorescence fluctuation for Rh123. Representative traces (B) and summarized data (C) showing ATP (100 &#181;mol/L)-induced changes in the HN4 cell MMP. The change in the membrane potential was indicated by the ratio of the fluorescence intensity before and after the application of ATP. The ratio increase represents the membrane potential depolarization. The HN4 cells were transfected with CLIC4siRNA (CLIC4 siRNA) or scrambled siRNA (Con siRNA). Values are shown as the mean &#177; SEM. n = 5. *P &#60;0.05 vs. control (Con siRNA) group. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56317/56317fig4large.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/56317/56317fig5.jpg" /> 
Figure 5. Effect of CLIC4 knockdown on ATP-induced mitochondrial membrane depolarization resolved by confocal laser scanning fluorescence microscope in HN4 cells. (A) Representative images of fluorescence fluctuation for Rh123. Representative traces (B) and summarized data (C) showing ATP (100 &#181;mol/L)-induced changes in the HN4 cell MMP. The change in the membrane potential was indicated by the ratio of the fluorescence density before and after the application of ATP. The ratio increase represents the membrane potential depolarization. The HN4 cells were transfected with CLIC4 siRNA (CLIC4 siRNA) or scrambled siRNA (Con siRNA). Values are shown as the mean &#177; SEM. n = 5. *P &#60;0.05 vs. control (Con siRNA) group.&#160;<a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56317/56317fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Detection of Mitochondria Membrane Potential to Study CLIC4 Knockdown-induced HN4 Cell Apoptosis In Vitro at JoVE.com

Detection of Mitochondria Membrane Potential to Study CLIC4 Knockdown-induced HN4 Cell Apoptosis In Vitro

Here we present a detailed protocol for the application of rhodamine 123 to identify the mitochondrial membrane potential (MMP) and study CLIC4 knockdown-induced HN4 cell apoptosis in vitro. Under common fluorescence microscope and confocal laser scanning fluorescence microscope, the real-time change of the MMP was recorded.

Detection of Mitochondria Membrane Potential to Study CLIC4 Knockdown-induced HN4 Cell Apoptosis In Vitro

Depletion of the mitochondrial membrane potential (MMP, &#916;&#936;m) is considered the earliest event in the apoptotic cascade. It even occurs ahead of ...

detection-mitochondria-membrane-potential-to-study-clic4-knockdown

Nanyang Technological University

Research

JoVE Journal

Cancer Research

10.3K Views.  Anhui Medical University. Here we present a detailed protocol for the application of rhodamine 123 to identify the mitochondrial membrane potential (MMP) and study CLIC4 knockdown-induced HN4 cell apoptosis in vitro. Under common fluorescence microscope and confocal laser scanning fluorescence microscope, the real-time change of the MMP was recorded.

Video: Detection of Mitochondria Membrane Potential to Study CLIC4 Knockdown-induced HN4 Cell Apoptosis In Vitro

Experimental Protocol for Detecting Mitochondrial Function in Hepatocytes Exposed to Organochlorine Pesticides

This paper presents detailed methods on detecting hepatic mitochondrial function for a better understanding the cause of metabolic disorders caused by environmental organochlorine pesticides (OCPs) in hepatocytes. HepG2 cells were exposed to &#946;-hexachlorocyclohexane (&#946;-HCH) for 24 h at an equivalent dose of internal exposure in general population. Ultrastructure in hepatocytes was examined by transmission electron microscopy (TEM) to show the damage of mitochondria. Mitochondrial function was further evaluated by mitochondrial fluorescence intensity, adenosine 5'-triphosphate (ATP) levels, oxygen consumption rate (OCR) and mitochondrial membrane potential (MMP) in HepG2 cells incubated with &#946;-HCH. The mitochondria fluorescence intensity after stained by mitochondrial green fluorescent probe was observed with a fluorescence microscopy. The luciferin-luciferase reaction was used to determine ATP levels. The MMP was detected by the cationic dye JC-1 and analyzed under flow cytometry. OCR was measured with an extracellular flux analyzer. In summary, these protocols were used in detecting mitochondrial function in hepatocytes with to investigate mitochondria damages.

Understanding the influence of environmental organochlorine pesticides (OCPs) on mitochondrial function in hepatocytes is important in exploring the mechanism of OCPs causing metabolic disorders. This paper presents detailed methods on detecting hepatic mitochondrial function.

This paper presents detailed methods on detecting hepatic mitochondrial function for a better understanding the cause of metabolic disorders caused by ...

Environment

Monitoring Cleaved Caspase-3 Activity and Apoptosis of Immortalized Oligodendroglial Cells using Live-cell Imaging and Cleaveable Fluorogenic-dye Substrates Following Potassium-induced Membrane Depolarization

The central nervous system can experience a number of stresses and neurological insults, which can have numerous adverse effects that ultimately lead to a reduction in neuronal population and function. Damaged axons can release excitatory molecules including potassium or glutamate into the extracellular matrix, which in turn, can produce further insult and injury to the supporting glial cells including astrocytes and oligodendrocytes 8, 16. If the insult persists, cells will undergo programmed cell death (apoptosis), which is regulated and activated by a number of well-established signal transduction cascades 14. Apoptosis and tissue necrosis can occur after traumatic brain injury, cerebral ischemia, and seizures. A classical example of apoptotic regulation is the family of cysteine-dependent aspartate-directed proteases, or caspases. Activated proteases including caspases have also been implicated in cell death in response to chronic neurodegenerative diseases including Alzheimer's, Huntington's, and Multiple Sclerosis 4, 14, 3, 11, 7.
In this protocol we describe the use of the NucView 488 caspase-3 substrate to measure the rate of caspase-3 mediated apoptosis in immortalized N19-oligodendrocyte (OLG) cell cultures 15, 5, following exposure to different extracellular stresses such as high concentrations of potassium or glutamate. The conditionally-immortalized N19-OLG cell line (representing the O2A progenitor) was obtained from Dr. Anthony Campagnoni (UCLA Semel Institute for Neuroscience) 15, 5, and has been previously used to study molecular mechanisms of myelin gene expression and signal transduction leading to OLG differentiation (e.g.6, 10). We have found this cell line to be robust with respect to transfection with exogenous myelin basic protein (MBP) constructs fused to either RFP or GFP (red or green fluorescent protein) 13, 12. Here, the N19-OLG cell cultures were treated with either 80 mM potassium chloride or 100 mM sodium glutamate to mimic axonal leakage into the extracellular matrix to induce apoptosis 9. We used a bi-functional caspase-3 substrate containing a DEVD (Asp-Glu-Val-Asp) caspase-3 recognition subunit and a DNA-binding dye 2. The substrate quickly enters the cytoplasm where it is cleaved by intracellular caspase-3. The dye, NucView 488 is released and enters the cell nucleus where it binds DNA and fluoresces green at 488 nm, signaling apoptosis. Use of the NucView 488 caspase-3 substrate allows for live-cell imaging in real-time 1, 10. In this video, we also describe the culturing and transfection of immortalized N19-OLG cells, as well as live-cell imaging techniques.

Monitoring Cleaved Caspase-3 Activity and Apoptosis of Immortalized Oligodendroglial Cells using Live-cell Imaging and Cleaveable Fluorogenic-dye Substrates Following Potassium-induced Membrane Depola

Live-cell imaging of caspase-3 mediated apoptosis in immortalized N19-oligodendrocyte cell cultures using the NucView 488 caspase-3 substrate. This technique is applicable for programmed cell death assays in real-time in a variety of cell types and tissues.

The central nervous system can experience a number of stresses and neurological insults, which can have numerous adverse effects that ultimately lead to a ...

Neuroscience

Use of a Caspase Multiplexing Assay to Determine Apoptosis in a Hypothalamic Cell Model

The ability to multiplex assays in studies of complex cellular mechanisms eliminates the need for repetitive experiments, provides internal controls, and decreases waste in costs and reagents. Here we describe optimization of a multiplex assay to assess apoptosis following a palmitic acid (PA) challenge in an in vitro hypothalamic model, using both fluorescent and luminescent based assays to measure viable cell counts and caspase-3/7 activity in a 96-well microtiter plate format. Following PA challenge, viable cells were determined by a resazurin-based fluorescent assay. Caspase-3/7 activity was then determined using a luminogenic substrate, DEVD, and normalized to cell number. This multiplexing assay is a useful technique for determining change in caspase activity following an apoptotic stimulus, such as saturated fatty acid challenge. The saturated fatty acid PA can increase hypothalamic oxidative stress and apoptosis, indicating the potential importance of assays such as that described here in studying the relationship between saturated fatty acids and neuronal function.

Multiplex assays can provide beneficial information for basic cellular mechanisms&#160;and eliminate waste of reagents and unnecessary repetitive experiments. We describe here a multiplex caspase-3/7 activity assay, using fluorescent- and luminescent-based methods, to determine cell viability in an in vitro hypothalamic model following oxidative challenge with palmitic acid.

The ability to multiplex assays in studies of complex cellular mechanisms eliminates the need for repetitive experiments, provides internal controls, and ...

Validation of FAM83A as a Target Gene in Cervical Cancer

Knockdown of FAM83A to Verify Its Role in Cervical Cancer Cell Growth and Cisplatin Sensitivity

The exploration of tumor target genes holds paramount importance for the prevention and treatment of cervical cancer. In this study, we outline the steps involved in the identification of a tumor target gene FAM83A in cervical cancer. First, the Cancer Genome Atlas dataset was employed to validate the expression and prognostic significance of FAM83A in women. A small interfering RNA (siRNA) was used for knockdown of the FAM83A gene in HeLa and C33a cells. Next, 5-ethynyl-2&#39;-deoxyuridine (EdU) staining was conducted to determine the effects on the proliferation capabilities of the tumor cells. Wound healing and porous membrane insert assays were performed to evaluate tumor cell migration and invasion abilities.
Western blotting was used to quantify apoptosis-related protein levels. JC-1 staining was employed to evaluate mitochondrial function alterations. Furthermore, cisplatin (diaminedichloroplatinum, DDP) intervention was used to assess the therapeutic potential of the target gene. Flow cytometry and colony formation assays were conducted to further validate the anticancer characteristics of the gene. As a result, FAM83A knockdown was shown to inhibit the proliferation, migration, and invasion of cervical cancer cells and sensitize these cells to cisplatin. These comprehensive methodologies collectively validate FAM83A as a tumor-associated target gene, holding promise as a potential therapeutic target in the prevention and treatment of cervical cancer.

Author Spotlight: Exploring the Role of FAM83A in Cervical Cancer 

Here, we show the procedures for FAM83A knockdown; the assays to detect its effects on proliferation, migration, and invasion of cervical cancer cells; and the sensitization of these cells to cisplatin. This study provides a promising target gene for cervical cancer and a reference for further drug research.

The exploration of tumor target genes holds paramount importance for the prevention and treatment of cervical cancer. In this study, we outline the steps ...

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, ...

Biology

Assessment of Mitochondrial Functions and Cell Viability in Renal Cells Overexpressing Protein Kinase C Isozymes

The protein kinase C (PKC) family of isozymes is involved in numerous physiological and pathological processes. Our recent data demonstrate that PKC regulates mitochondrial function and cellular energy status. Numerous reports demonstrated that the activation of PKC-a and PKC-&epsilon; improves mitochondrial function in the ischemic heart and mediates cardioprotection. In contrast, we have demonstrated that PKC-&alpha; and PKC-&epsilon; are involved in nephrotoxicant-induced mitochondrial dysfunction and cell death in kidney cells. Therefore, the goal of this study was to develop an in vitro model of renal cells maintaining active mitochondrial functions in which PKC isozymes could be selectively activated or inhibited to determine their role in regulation of oxidative phosphorylation and cell survival. Primary cultures of renal proximal tubular cells (RPTC) were cultured in improved conditions resulting in mitochondrial respiration and activity of mitochondrial enzymes similar to those in RPTC in vivo. Because traditional transfection techniques (Lipofectamine, electroporation) are inefficient in primary cultures and have adverse effects on mitochondrial function, PKC-&epsilon; mutant cDNAs were delivered to RPTC through adenoviral vectors. This approach results in transfection of over 90% cultured RPTC.
Here, we present methods for assessing the role of PKC-&epsilon; in: 1. regulation of mitochondrial morphology and functions associated with ATP synthesis, and 2. survival of RPTC in primary culture. PKC-&epsilon; is activated by overexpressing the constitutively active PKC-&epsilon; mutant. PKC-&epsilon; is inhibited by overexpressing the inactive mutant of PKC-&epsilon;. Mitochondrial function is assessed by examining respiration, integrity of the respiratory chain, activities of respiratory complexes and F0F1-ATPase, ATP production rate, and ATP content. Respiration is assessed in digitonin-permeabilized RPTC as state 3 (maximum respiration in the presence of excess substrates and ADP) and uncoupled respirations. Integrity of the respiratory chain is assessed by measuring activities of all four complexes of the respiratory chain in isolated mitochondria. Capacity of oxidative phosphorylation is evaluated by measuring the mitochondrial membrane potential, ATP production rate, and activity of F0F1-ATPase. Energy status of RPTC is assessed by determining the intracellular ATP content. Mitochondrial morphology in live cells is visualized using MitoTracker Red 580, a fluorescent dye that specifically accumulates in mitochondria, and live monolayers are examined under a fluorescent microscope. RPTC viability is assessed using annexin V/propidium iodide staining followed by flow cytometry to determine apoptosis and oncosis.
These methods allow for a selective activation/inhibition of individual PKC isozymes to assess their role in cellular functions in a variety of physiological and pathological conditions that can be reproduced in in vitro.

The effects of activation of protein kinase C (PKC) isozymes on mitochondrial functions associated with respiration and oxidative phosphorylation and on cell viability are described. The approach adapts adenoviral technique to selectively overexpress PKC isozymes in primary cell culture and a variety of assays to determine mitochondrial functions and energy status of the cell.

The protein kinase C (PKC) family of isozymes is involved in numerous physiological and pathological processes. Our recent data demonstrate that PKC ...

Preparation of Cell-lines for Conditional Knockdown of Gene Expression and Measurement of the Knockdown Effects on E4orf4-Induced Cell Death

Functional inactivation of gene expression in mammalian cells is crucial for the study of the contribution of a protein of interest to various pathways1,2. However, conditional knockdown of gene expression is required in cases when constitutive knockdown is not tolerated by cells for a long period of time3-5. Here we describe a protocol for preparation of cell lines allowing conditional knockdown of subunits of the ACF chromatin remodeling factor. These cell lines facilitate the determination of the contribution of ACF to induction of cell death by the adenovirus E4orf4 protein6. Sequences encoding short hairpin RNAs for the Acf1 and SNF2h subunits of the ACF chromatin remodeling factor were cloned next to a doxycycline-inducible promoter in a plasmid also containing a gene for the neomycin resistance gene. Neomycin-resistant cell clones were selected in the presence of G418 and isolated. The resulting cell lines were induced by doxycycline treatment, and once Acf1 or SNF2h expression levels were reduced, the cells were transfected with a plasmid encoding E4orf4 or an empty vector. To confirm the specific effect of the shRNA constructs, Acf1 or SNF2h protein levels were restored to WT levels by cotransfection with a plasmid expressing Acf1 or SNF2h which were rendered resistant to the shRNA by introduction of silent mutations. The ability of E4orf4 to induce cell death in the various samples was determined by a DAPI assay, in which the frequency of appearance of nuclei with apoptotic morphologies in the transfected cell population was measured7-9.
The protocol described here can be utilized for determination of the functional contribution of various proteins to induction of cell death by their protein partners in cases when constitutive knockdown may be cell lethal.

Contribution of the ACF chromatin remodeling factor to E4orf4-induced cell death was measured. The protocol includes selection of cell clones in which doxycycline treatment induces conditional knockdown of the ACF subunits Acf1 and SNF2h, and use of the DAPI assay to measure E4orf4-induced cell death in the inducible cell lines.

Functional inactivation of gene expression in mammalian cells is crucial for the study of the contribution of a protein of interest to various ...

A Flow Cytometry-based Assay for Measuring Mitochondrial Membrane Potential in Cardiac Myocytes After Hypoxia/Reoxygenation

Timely and efficient reperfusion of the occluded coronary artery is the best strategy for decreasing myocardial infarct size in patients with a ST-segment elevated myocardial infarction. However, reperfusion per se can result in further cardiomyocyte death, a phenomenon known as reperfusion injury. The opening of the mitochondrial permeability transition pore (mPTP), with the decrease of the mitochondrial membrane potential (MMP), or mitochondrial depolarization, is universally recognized as the final step of reperfusion injury and is responsible for mitochondrial and cardiomyocyte death. JC-1 is a lipophilic cationic dye that accumulates in mitochondria depending on the value of MMP. The higher the MMP is, the more JC-1 accumulates in the mitochondria. The increasing amounts of JC-1 in mitochondria can be reflected by a fluorescence emission shift from green (~530 nm) to red (~590 nm). Therefore, the reduction of the red/green fluorescence intensity ratio can indicate the depolarization of mitochondria. Here, we take advantage of JC-1 to measure MMP, or the opening of mPTP in human cardiac myocytes after hypoxia/reoxygenation, detected by flow cytometry.

Here, we present a protocol that uses JC-1 dye to assess the mitochondrial membrane potential of cells after being exposed to hypoxia/reoxygenation with or without a protective agent.

Timely and efficient reperfusion of the occluded coronary artery is the best strategy for decreasing myocardial infarct size in patients with a ST-segment ...

Medicine

Strategies for Tracking Anastasis, A Cell Survival Phenomenon that Reverses Apoptosis

Anastasis (Greek for &#8220;rising to life&#8221;) refers to the recovery of dying cells. Before these cells recover, they have passed through important checkpoints of apoptosis, including mitochondrial fragmentation, release of mitochondrial cytochrome c into the cytosol, activation of caspases, chromatin condensation, DNA damage, nuclear fragmentation, plasma membrane blebbing, cell shrinkage, cell surface exposure of phosphatidylserine, and formation of apoptotic bodies. Anastasis can occur when apoptotic stimuli are removed prior to death, thereby allowing dying cells to reverse apoptosis and potentially other death mechanisms. Therefore, anastasis appears to involve physiological healing processes that could also sustain damaged cells inappropriately. The functions and mechanisms of anastasis are still unclear, hampered in part by the limited tools for detecting past events after the recovery of apparently healthy cells. Strategies to detect anastasis will enable studies of the physiological mechanisms, the hazards of undead cells in disease pathology, and potential therapeutics to modulate anastasis. Here, we describe effective strategies using live cell microscopy and a mammalian caspase biosensor&#160;for identifying and tracking anastasis in mammalian cells.

The term anastasis refers to the phenomenon in which dying cells reverse a cell suicide process at a late stage, repair themselves, and ultimately survive. Here we demonstrate protocols for detecting and tracking cells that undergo anastasis.

Anastasis (Greek for &#8220;rising to life&#8221;) refers to the recovery of dying cells. Before these cells recover, they have passed through important ...

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

Detection of Mitochondria Membrane Potential to Study CLIC4 Knockdown-induced HN4 Cell Apoptosis In Vitro

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Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue