This work was supported in part by NIH grants (1R01CA137494, R01CA132115, R01CA086072 to R.G.P.), the Kimmel Cancer Center NIH Cancer Center Core grant P30CA056036 (R.G.P.), a grant from the Breast Cancer Research Foundation, generous grants from the Dr. Ralph and Marian C. Falk Medical Research Trust (R.G.P.) and a grant from the Pennsylvania Department of Health (R.G.P.). In part this work was supported by an American Italian Cancer Foundation postdoctoral fellowship (G.D.) and Bioimaging Shared Resource of the Sidney Kimmel Cancer Center (NCI 5 P30 CA-56036).The Department specifically disclaims responsibility for an analysis, interpretations or conclusions. There are no conflicts of interest associated with this manuscript.

1. Electroporation of Fibroblast with Both Expression Vectors EYFP-Parkin and pDsRed2-Mito
<ol>
	<li>Grow the immortalized mouse embryonic fibroblast cells on 10 centimeter tissue-culture plate using DMEM (Dulbecco&#39;s modified Eagle&#39;s medium) medium supplemented with 10% fetal bovine serum, 2 mmol/l L-glutamine, 100 U/ml&#160;penicillin, and 100 mg/ml streptomycin in humidified atmosphere containing 5% CO2 at 37 &#176;C.
	<ol>
		<li>At 80% cell confluency, discard the complete DMEM medium by sterile suction and add 10 ml of sterile 1x phosphate buffer solution (PBS) (80 g of NaCl, 2.0 g of KCl, 14.4 g of Na2HPO4, 2.4 g of KH2PO4 and 1 L distilled H2O, PH: 7.4).</li>
		<li>Discard the PBS by sterile suction and add 1 ml of Trypsin-EDTA 0.25%. Incubate the plate at 37 &#176;C until the cells are detached (2 - 3 min). Add 4 ml of complete DMEM medium, resuspend the cells and take out 10 &#956;l&#160;of the cell suspension for the hemocytometer counting. 
		NOTE: The hemocytometer is designed so that the number of cells in one set of 16 corner squares is equivalent to the number of cells x 104/ml.</li>
	</ol>
	</li>
	<li>Seed 1 x 106 cells onto 10 cm tissue culture dishes 24 hours prior to the electroporation process.</li>
	<li>Discard the complete DMEM medium by sterile suction and add 10 ml of sterile PBS. Discard the PBS by sterile suction and add 1 ml of Trypsin-EDTA 0.25%. Incubate the plate at 37 &#176;C until the cells are detached (2 - 3 min). Add 4 ml of complete DMEM medium and resuspend the cells in a 15 ml tube.</li>
	<li>Spin the cells down at 250 x g for 5 min&#160;using a refrigerated-centrifuge (4 &#176;C). Discard the supernatant by sterile suction and resuspend the pellet in 1 ml of sterile PBS. Spin the cells down at 250 x g for 5 min&#160;at 4 &#176;C.</li>
	<li>Discard the supernatant by sterile suction, and add 100 &#181;l of solution mix for electroporation (82 &#181;l of electroporation Solution V + 18 &#181;l of Supplemental solution 1) to the cell pellet and gently resuspend the pellet by pipetting (For more details refer to user&#39;s guide). Add 2 &#181;g of EYFP-Parkin (excitation/emission 514/527) and 1 &#181;g pDsRed2-Mito (excitation/emission 565/620) at the cell suspension.</li>
	<li>Transfer the solution to the sterile cuvette using a disposable pasteur (both tools are provided with the kit) and electroporate using the pre-set program NIH/3T3 U-030 (Single pulse, voltage 200 V, capacitance 960 &#181;F, pulse time 20 milliseconds, pulse number: 1).</li>
	<li>Immediately after the electroporation, add 500 &#181;l of fresh pre-warmed complete DMEM medium and seed the cell on a 6 cm tissue-culture plate live-imaging-grade and incubate them in a humidified atmosphere containing 5% CO2 at 37 &#176;C for 24 hours.</li>
</ol>
2. Time-Lapse Video Microscopy
<ol>
	<li>Set the temperature of the microscope&#39;s chamber at 37 &#176;C prior to use.</li>
	<li>At this point, prepare a specific live imaging medium to proceed with the experimental protocol. Prepare the live imaging medium as follows; mix DMEM phenol-free supplemented with 10% fetal bovine serum, 2 mmol/l L-glutamine, 100 U/ml penicillin, and 100 mg/ml streptomycin. Pre-warm the live imaging medium at 37 &#176;C in a water bath.</li>
	<li>Discard the medium of the electroporated cells by sterile suction and add 1 ml of pre-warmed live imaging medium, using a P1000 pipet and incubate the plate for 30 min&#160;at 37 &#176;C.</li>
	<li>During the plate incubation period, influx 5% CO2 in the microscope&#39;s chamber already at a stable temperature of 37 &#176;C.</li>
	<li>Place the plate with the electroporated cells into the microscope&#39;s chamber avoiding major movements or oscillations.</li>
	<li>Using the software interface, set the microscope in order to detect both fluorescence signals from the fusion proteins encoded by the co-transfected vectors, EYFP-Parkin (Excitation range 495 to 510 nm and emission range 520 to 550 nm, green) and the pDsRed2-Mito (Excitation maximum 558 nm and emission maximum 583 nm, red).
	<ol>
		<li>Open the time-lapse video microscopy software. In the upper menu select the FITC for EYFP-Parkin and Rhodamine for pDsRed2-Mito. In the upper menu select the magnification (20X).</li>
	</ol>
	</li>
	<li>Using the software interface, look for the single cell expressing both co-transfected vectors and register the position. Repeat this step until a minimum of 10 cells is collected for every experimental condition (Experimental group).
	<ol>
		<li>Select the menu &#34;Apps&#34; and click on Multi Dimensional Acquisition. In the Multi Dimensional Acquisition windows select the parameters needed, such as the number of acquisitions, the interval of time between each acquisition and the position of the recorded cell. Click on &#34;Acquire&#34; to start the acquisition process.</li>
	</ol>
	</li>
	<li>Start the basal acquisition of both EYFP-Parkin and DsRed2-Mito fluorescent signal collecting images every 5 min for a total interval of 15 min.</li>
	<li>Prepare pre-warmed live imaging medium with a concentration of FCCP twice higher than the final working concentration (0.1 - 10 &#181;M, cell type-dependent).</li>
	<li>Interrupt the acquisition process and add gently 1 ml of pre-warmed live imaging medium with FCCP into the plate inside the microscope&#39;s chamber using a P1000 pipet.</li>
	<li>Restart the acquisition process as previously described, for a total period of 3 hours, using the recorded position. Save all the acquired images in order to analyze them and create a video of the mitophagic process when the acquisition is over.</li>
</ol>
3. Analysis
<ol>
	<li>Collect all the acquired images in &#34;.tiff &#34; format on a personal computer</li>
	<li>Open the images with a graphic software and define the temporal interval occurred from the mitochondrial induced-depolarization with FCCP to the first image showing the recruitment of Parkin into the mitochondrial membrane (Images are acquired every 5 min). Repeat this analysis for each cell in the experimental group.</li>
	<li>Label the first cell of the column on a spreadsheet with the experimental group name. For each experimental group, measure the temporal intervals for Parkin recruitment of a minimum of 10 cells. Make an average of the calculated temporal intervals for Parkin recruitment and define the standard deviation for each experimental group (Mean &#177; S.D., N = 10).
	<ol>
		<li>Organize into columns the single calculated temporal intervals. Every column contains the n = 10 measurements of the experimental group.</li>
		<li>Select the function &#34;average&#34; from the Formula Builder menu. Select the data in the column. Select the function &#34;Standard Deviation&#34; from the Formula Builder menu. Select the data in the column.</li>
	</ol>
	</li>
	<li>Using a statistical approach, compare the experimental groups applying the appropriate statistic in order to define a significant difference in the dynamic of mitophagy between the experimental groups. (For instance, two groups = Student&#39;s t-Test, three or more groups = ANOVA plus a post-hoc test)</li>
</ol>

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The authors have nothing to disclose.

Time-lapse microscopy can be defined as the technique that extends live cell imaging from a single observation in time to the observation of cellular dynamics over long periods of time. This methodology is distinguished from a simple confocal or live cell microscopy because it allows the observer to identify in real time a single fluorescent-tagged protein and follow its dynamic inside a single live cell. In fact, the confocal microscopy can easily identifies immuno-labeled proteins using fluorescent antibody, but it does not allow observing the cell and the molecular event in its live environment.
The time-lapse video microscopy is primarily used in research monitoring molecular processes in different cell types such as HEK293 16, primary neurons 17 and Hela cells 18 but has also clinical application such as pregnancy and predicting aneuploidy 19,20. Time-lapse microscopy provides embryologists with additional information about embryo development for a clinical pregnancy, and it is possible that this information can be used to improve our ability to select viable embryos for transfer. Additional advantages of the use of Time-lapse microscopy include less handling of embryos and their dishes for intermediate observations, which would reduce the risk of loss or contamination within the laboratory 19.
Cells are able to maintain a healthy mitochondrial homeostasis by degrading damaged and dysfunctional mitochondria through mitophagy. Defective mitophagy is linked to Parkinson&#39;s disease 21, apoptosis 22, aging 23 and cancer 24. Mitophagy is driven by the ubiquitin ligase Parkin and PINK121. Due to the importance of this process to maintain a healthy mitochondrial status, we found it important to study the dynamic of this event using a time-lapse video microscopy. The proposed protocol will provide evidence on how mitochondria homeostasis is affected under different cellular conditions, including changes in gene expression and protein functions.
In this study, we transiently transfected fibroblasts by electroporation. The electroporation method applies an electrical pulse at an optimized voltage lasting a few microseconds to the cell suspension in order to increase the permeability of the cell membrane by disturbing the phospholipid bilayer of the membrane. This action results in the formation of temporary pores. The membrane permeabilization allows the DNA to be introduced into the cell 25. We chose this process to transiently transfect fibroblasts in order to have higher transfection efficiency of both plasmids EYFP-Parkin and pDsRed2-Mito. In fact, the electroporation process is approximately ten times more effective than chemical transfection 26. The Images acquisition process can be compromised by both external factors; temperature fluctuations, vibrations, humidity and internal factors; pH, cell motility. The electroporation and fluorescent tracking are two steps considered to be the most critical for the success of this experimental protocol. Even if the transfection efficiency is higher than other methods, the electroporation kills ~ 20% of the electroporated cells. Another major issue is the acquisition quality of the images during the monitoring step. Since the cellular environment is dynamic, the focus plane needs continuous adjustments in order to maintain the focus throughout the acquisition process.
Mitochondrial damage can be caused by membrane mitochondrial depolarization and oxidative stress. These factors induce a mitophagic response. Evidence that mitochondria depolarization can induce selective mitophagy comes from photodamage experiments which lead to ROS production and mitochondrial injury 27-30. In order to induce mitochondrial membrane depolarization and mitophagic response, FCCP has been used as protonophore 10. FCCP is an ionophore that allows protons to cross lipid bilayers inducing membrane depolarization. Using this approach we induced mitophagy and followed the dynamic of the molecular marker.
The concentration of FCCP is crucial, as it can be cytotoxic and promote cellular death. Since this approach relies on the observation of living cells, the final concentration of FCCP has to be adjusted in a cell type dependent manner in order to induce mitophagy without trigger the mechanisms that lead to cellular death.
Overall, we think that following the dynamic of mitophagy, using EYFP-Parkin as a fluorescent marker, will provide new insights into mitochondrial function. The selective removal of mitochondria via mitophagy is a crucial process in maintaining mitochondrial quality, cell viability and homeostasis. This live imaging approach will help to understand the mitophagic process under healthy and stressed conditions.
Modern approaches are further extending time-lapse microscopy observations beyond making movies of cellular dynamics. Increasingly this boundary is blurred as cytometric techniques are being integrated with imaging techniques for monitoring and measuring dynamic activities of cells and subcellular structures 31.

Macro autophagy is an intracellular process that involves the catabolic degradation of both damaged and dysfunctional cellular components, such as organelles and proteins for the purpose of either recycling or energy production. To initiate this metabolic process, the cell engulfs the damaged cellular components into a double-membrane structure, known as an autophagosome, which fuses with a lysosome and its content is degraded and recycled 1,2. There are two major types of autophagy, the non-selective and selective. The non-selective autophagy process occurs when the cell is under nutrient deprivation conditions and needs to scavenge for both essential nutrients and energy. However, selective autophagy occurs to mediate the removal of both dysfunctional/damaged organelles and proteins that otherwise could be toxic. One of the most studied selective autophagy process is the removal of mitochondria, termed mitophagy 1,3-5.
Mitochondria are the central organelles for cell metabolism and the primary source of adenosine triphosphate (ATP) via oxidative phosphorylation through the electron transport chain, fatty acid oxidation, and tricarboxylic acid (TCA) cycle. Moreover, mitochondria regulate reactive oxygen species (ROS) production and release proteins that participate in cell death pathways 6-8.
PTEN-induced putative kinase 1 (PINK1) and Parkin RBR E3 ubiquitin ligase (Parkin) are the key proteins implicated in the mitophagy process. Parkin can protect against cell death by keeping the cell healthy through mitochondrial quality control9. Upon the loss of mitochondrial membrane potential, cytosolic Parkin is recruited to the mitochondria by PINK1. This recruitment triggers the sequential events of mitophagy 10. There is a broad range of evidence that mitophagy is a fundamental mitochondria quality control process and abnormalities in this process drive disease 7. For instance, autosomal recessive Parkinson's disease has been associated with mutations in the genes that encode for Parkin and PINK1 (PARK2 and PINK1, respectively) 11. The quality control of mitochondrial health is essential for the removal of mitochondria that contribute to the accumulation of ROS12. Excessive presence of intracellular ROS can lead to damage of both nuclear and mitochondrial DNA (DNA and mt DNA, respectively).
Herein, we show a time-lapse video microscopy approach to follow the aggregation of Parkin after the induction of Parkin-mediated mitophagy in immortalized mouse embryonic fibroblasts via in vitro administration of carbonyl cyanide 4-(trifluoromethoxy)-phenylhydrazone (FCCP), an uncoupling agent. FCCP disrupts ATP synthesis by short circuiting protons across the outer mitochondria membrane and hence uncoupling oxidative phosphorylation from the electron transport chain 13. Triggering the depolarization of the mitochondrial membrane leads to the disruption of mitochondria and selective Parkin-dependent removal. Therefore, transfecting the cells of interest with an expression vector encoding Parkin fused with a fluorescent marker (enhanced yellow fluorescent protein, EYFP) can be used as a fluorescent tag to follow the recruitment of Parkin during the mitophagic process. In order to visualize the mitochondria, we co-transfected pDsRed2-Mito, which encodes red fluorescent protein (DsRed2) that contains a mitochondrial targeting sequence of cytochrome c oxidase subunit VIII (Mito). pDsRed2-Mito is designed for fluorescent labeling of mitochondria14. The time required for Parkin translocation into the mitochondrial membrane can be measured and gives an indirect measure of cellular health. For example, we can say that if a cell line knocked-out for a particular gene of interest shows either a faster or slower recruitment of Parkin after the induction of mitophagy by FCCP, that gene product would be a key player in order to keep the metabolic rates of the cell at the physiological status and prevent the development of diseases. Therefore, the time-lapse video microscopy provides a very powerful tool for both basic and clinical research applications in following the dynamic of labeled proteins during their molecular processes and understanding how these processes are affected during a pathological condition.

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	<tbody>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">DMEM</td>
			<td style="width:357px;">Corning Life Science</td>
			<td style="width:276px;">10-013-CV</td>
			<td style="width:369px;">Pre-warm at +37 C before use</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">PHENOL-FREE DMEM</td>
			<td style="width:357px;">Corning Life Science</td>
			<td style="width:276px;">17-205-CV</td>
			<td>Pre-warm at +37 C before use</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">FETAL BOVINE SERUM</td>
			<td style="width:357px;">Sigma-Aldrich</td>
			<td style="width:276px;">F2442</td>
			<td>Pre-warm at +37 C before use</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">L-GLUTAMINE</td>
			<td style="width:357px;">Gibco</td>
			<td style="width:276px;">25030</td>
			<td>Pre-warm at +37 C before use</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">PENICILLIN/ STREPTOMYCIN</td>
			<td style="width:357px;">Corning Life Science</td>
			<td style="width:276px;">30-002-CI</td>
			<td>Pre-warm at +37 C before use</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">EYFP-PARKIN EXPRESSION VECTOR</td>
			<td style="width:357px;">Addgene</td>
			<td style="width:276px;">23955</td>
			<td></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">pDsRed2-Mito EXPRESSION VECTOR</td>
			<td style="width:357px;">Clontech</td>
			<td style="width:276px;">632421</td>
			<td></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">NUCLEOFECTOR 2B DEVICE</td>
			<td style="width:357px;">LONZA</td>
			<td style="width:276px;">AAD-1001S</td>
			<td></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;">NUCLEOFECTOR FOR KIT R NIH/3T3</td>
			<td style="width:357px;">LONZA</td>
			<td style="width:276px;">VCA-1001</td>
			<td></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;">ZEISS AXIOVERT 200M INVERTED MICROSCOPE</td>
			<td style="width:357px;">CARL ZEISS</td>
			<td style="width:276px;"></td>
			<td></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;">Carbonyl Cyanide 4-(trifluoromethoxy)-Phenylhydrazone (FCCP)</td>
			<td style="width:357px;">Sigma-Aldrich</td>
			<td style="width:276px;">C2920</td>
			<td></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:735px;">MetaMorph</td>
			<td style="width:357px;">Molecular Devices</td>
			<td style="width:276px;">Experimental Builder</td>
			<td></td>
		</tr>
		<tr height="36">
			<td height="36" style="height:36px;width:735px;">ImageJ</td>
			<td style="width:357px;">National Institute of Health</td>
			<td style="width:276px;">Experimental Builder</td>
			<td></td>
		</tr>
	</tbody>
</table>

time-lapse video microscopy for assessment of eyfp-parkin aggregation as a marker for cellular mitophagy

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

Herein, we show how time-lapse video microscopy is a powerful technique that can be used to follow molecular events of fluorescently-tagged proteins in a single cell. The representative results also show how this technique allows for the acquisition of high quality images. When images of the molecular process, are obtained, we have the opportunity to analyze them in different ways. Here, we analyze the interval of time between initiation of the induced mitophagy process, which is the crucial molecular process that maintain the cellular homeostasis selectively removing the damaged mitochondria.
This technique founds a broad range of applications. It can be used either for monitoring macro cellular process such as cell migration or micro molecular events through the labeling of the key molecules involved in the process that has to be studied.
Parkin-mediated Mitophagy
The selective Parkin-mediated removal of damaged and dysfunctional mitochondria process is summarized in Figure 1, where a membrane depolarization induced by the uncoupling agent FCCP triggers selective Parkin-mediated mitophagy, recruiting PINK1 to the outer mitochondrial membrane. Following, Parkin is recruited to the damaged mitochondria and interacts with PINK1. This complex Parkin-PINK1 leads to the ubiquitination of the damaged organelle and its incorporation into an autophagosome, which fuses with a lysosome. This process known as mitophagy is considered to be a crucial metabolic process in order to keep the cellular environment healthy and allow the regeneration of functional mitochondria 8,15.
Experimental Protocol
Figure 2 shows a schematic representation of the experimental protocol and the microscope set-up used for following the recruitment of EYFP-Parkin before and after FCCP-induced mitophagy process.
Parkin Recruitment to Depolarized Mitochondrial Membrane
In Figure 3A, we show Parkin distribution prior and post FCCP administration. During the basal time points (BT5, BT10 and BT15 min) EYFP-Parkin (Green) is homogeneously diffused throughout the cell and the mitochondria network (Red) appears to be well interconnected. Following FCCP administration (FT = 0 min) (Figure 3B), we observe mitochondria fractionation (FT = 5 min). However, EYFP-Parkin is still homogeneously diffused though out the cytoplasm. Mitophagy is stimulated at 55 min&#160;post FCCP administration, EYFP-Parkin is recruited at the damaged mitochondrial membranes to trigger the mitophagy process (FT = 55).
<img alt="Figure 1" src="/files/ftp_upload/53657/53657fig1.jpg" /> 
Figure 1.&#160;Schematic Representation of Parkin-mediated Mitophagy. A decreasing of membrane potential (&#936;m) occurs in damaged or dysfunctional mitochondria and can be induced by the uncoupling agent FCCP. Loss of membrane potential triggers selective Parkin-mediated mitophagy, recruiting PINK1 (Red) to the outer mitochondrial membrane. Parkin (Blue) is recruited to the damaged mitochondria by directly interacting with PINK1. The presence of Parkin leads to the ubiquitination (Yellow) of the damaged organelle and its incorporation into an autophagosome, which fuses with a lysosome. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53657/53657fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53657/53657fig2.jpg" /> 
Figure 2.&#160;Microscope Set-up and Representation of the Experimental Protocol.&#160;(A) Overview of the experimental events described in the protocol section. The reported experimental events allow the operator to transiently co-transfect the cells with EYFP-Parkin and pDsRed2-Mito by electroporation in order to follow the FCCP-induced mitophagy process using a (B) set-up for time-lapse video microscopy approach. BT = basal time point; FT = FCCP time point. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53657/53657fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53657/53657fig3.jpg" /> 
Figure 3. Fibroblast Cell Showing Parkin Recruitment after Mitochondrial Membrane Depolarization Induced by FCCP. EYFP-Parkin (green) translocation into the mitochondria (red) was monitored by time-lapse video microscopy during the basal time point (BT) and post FCCP administration (FT). From BT = 0, EYFP-Parkin and mitochondrial fluorescence were detected every 5 min. (A) EYFP-Parkin is homogenously diffused across the whole basal period and the mitochondrial network is intact (white arrows). (B) Mitochondrial fractionation due to membrane depolarization by FCCP is recorded at FT = 5 min&#160;(white arrows) and EYFP-Parkin recruitment to damaged mitochondrial membranes started at FT = 55 min&#160;(white arrows). The white boxes represent the magnified area of the panels (5X). For more details concerning the application of this experimental protocol refer to reference8. (Scale bar: 10 &#181;m). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53657/53657fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy at JoVE.com

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

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

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

time-lapse-video-microscopy-for-assessment-eyfp-parkin-aggregation-as

Nanyang Technological University

Research

JoVE Journal

Biology

7.1K Views.  Thomas Jefferson University, SKCC. Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

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

Monitoring Actin Disassembly with Time-lapse Microscopy

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

Time-lapse Imaging of Mitosis After siRNA Transfection

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

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

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

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

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

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These age-associated disorders are characterized by the appearance of protein aggregates with fibrillary structure in the brains of these patients. Exactly why normally soluble proteins undergo an aggregation process remains poorly understood. The discovery that protein aggregation is not limited to disease processes and instead part of the normal aging process enables the study of the molecular and cellular mechanisms that regulate protein aggregation, without using ectopically expressed human disease-associated proteins. Here we describe methodologies to examine inherent protein aggregation in Caenorhabditis elegans through complementary approaches. First, we examine how to grow large numbers of age-synchronized C. elegans to obtain aged animals and we present the biochemical procedures to isolate highly-insoluble-large aggregates. In combination with a targeted genetic knockdown, it is possible to dissect the role of a gene of interest in promoting or preventing age-dependent protein aggregation by using either a comprehensive analysis with quantitative mass spectrometry or a candidate-based analysis with antibodies. These findings are then confirmed by in vivo analysis with transgenic animals expressing fluorescent-tagged aggregation-prone proteins. These methods should help clarify why certain proteins are prone to aggregate with age and ultimately how to keep these proteins fully functional.

Methods to Study Changes in Inherent Protein Aggregation with Age in Caenorhabditis elegans

The goal of the method presented here is to explore protein aggregation during normal aging in the model organism C. elegans. The protocol represents a powerful tool to study the highly insoluble large aggregates that form with age and to determine how changes in proteostasis impact protein aggregation.

In the last decades, the prevalence of neurodegenerative disorders, such as Alzheimer's disease (AD) and Parkinson's disease (PD), has grown. These ...

Confocal Microscopy Reveals Cell Surface Receptor Aggregation Through Image Correlation Spectroscopy

Confocal microscopy provides an accessible methodology to capture sub-cellular interactions critical for the characterization and further development of pre-clinical agents labeled with fluorescent probes. With recent advancements in antibody based cytotoxic drug delivery systems, understanding the alterations induced by these agents within the realm of receptor aggregation and internalization is of critical importance. This protocol leverages the well-established methodology of fluorescent immunocytochemistry and the open source FIJI distribution of ImageJ, with its inbuilt autocorrelation and image mathematical functions, to perform spatial image correlation spectroscopy (ICS). This protocol quantitates the fluorescent intensity of labeled receptors as a function of the beam area of the confocal microscope. This provides a quantitative measure of the state of target molecule aggregation on the cell surface. This methodology is focused on the characterization of static cells with potential to expand into temporal investigations of receptor aggregation. This protocol presents an accessible methodology to provide quantification of clustering events occurring at the cell surface, utilizing well established techniques and non-specialized imaging apparatus.

Antibodies that bind to target receptors on the cell surface can confer conformation and clustering alterations. These dynamic changes have implications for characterizing drug development in target cells. This protocol utilizes confocal microscopy and image correlation spectroscopy through ImageJ/FIJI to quantify the extent of receptor clustering on the cell surface.

Confocal microscopy provides an accessible methodology to capture sub-cellular interactions critical for the characterization and further development of ...

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

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

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

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

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