Name: a faster, high resolution, mtpa-gfp-based mitochondrial fusion assay acquiring kinetic data of multiple cells in parallel using confocal microscopy
Uploaded: 2012-07-20T12:00:00
Duration: 10 min 45 s
Description: Watch this Scientific Journal Video about A Faster, High Resolution, mtPA-GFP-based Mitochondrial Fusion Assay Acquiring Kinetic Data of Multiple Cells in Parallel Using Confocal Microscopy at JoVE.co

We thank the Tufts Center for Neuroscience Research, P30 NS047243 (Jackson), the Mitochondria Affinity Research Collaborative (mtARC) supported by the Evans Center for Interdisciplinary Biomedical Research at Boston University Medical Campus, Link Medicine Corporation, and Zeiss for supporting this work.

1. Image Plate Preparation
<ol>
 <li>Culture INS1 cells in RPMI media containing 10% standard fetal bovine serum, 1% penicillin-streptomycin, 2 mM L-glutamine, 50 &mu;M 2-mercaptoethanol, 5 mM NaHCO3, 2 mM HEPES, 2 mM pyruvic acid, and 11 mM glucose to 80% confluence.</li>
 <li>Trypsinize the INS1 cell cultures with 0.05% trypsin and plate them onto poly-D-lysine coated coverslip-bottomed imaging plates (30-40% confluence).</li>
 <li>Allow for plates to reach 60-80% confluence (~2 days) and add mitochondrial matrix targeted (COXVIII) adenoviral PA-GFP for 24 hr (MOI=5). Exchange media and allow cells to grow for another 2 days before imaging.</li>
 <li>On the day of imaging, add 7-15 nM TMRE to the imaging plates, and equilibrate for at least 45 min.</li>
 <li>During this time turn on the incubator (stage top incubator has been used here) and allow the microscope to equilibrate to 37 &deg;C for about 1 hour. Turn on the 5% CO2.</li>
</ol>
2. Imaging Parameters for the Zeiss LSM 710 Confocal Microscope
<ol>
 <li>After the microscope has equilibrated, under the Acquisition tab, click on "show manual tools", open the imaging set up panel, and choose "channel mode" and switch track every "frame".</li>
 <li>In the light path panel, choose LSM and channel mode. Working from the specimen icon upward, select "rear", MBS 690+, and MBS 488. At the bottom of the panel, choose "T-PMT" to enable visualization of the bright field or DIC channel.</li>
 <li>Finish adjusting the light path parameters by setting the range for the GFP dye to 490-540 nm and to 580-700 nm for the TMRE dye.</li>
 <li>In the acquisition mode, use a 512 &times; 512 pixel scan field, average 4x, and choose a zoom factor (which you may want to keep consistent for calibration purposes).</li>
</ol>
3. Optimizing the Imaging Parameters
<ol>
 <li>Using the plan apochromat 100x (1.4 NA) objective lens, focus on your cells with the halogen light, not fluorescent light, to protect mitochondria from phototoxicity.</li>
 <li>Screen for PA-GFP expressing cells using a maximally open pinhole and scanning to find the cells that are brighter green.</li>
 <li>Find the lowest power of the 2-photon laser needed to activate the PA-GFP and optimize the imaging parameters ensuring that the signal does not saturate the detector. Check that the PA-GFP signal colocalizes with the TMRE signal for a few z-series. Loss of the TMRE signal indicates mitochondrial depolarization or phototoxicity, and cells exhibiting these characteristics should not be used for analysis.</li>
 <li>Find a PA-GFP expressing cell and specify a zoom factor.</li>
 <li>Set the z-series range to collect 6 slices (this range will need to satisfy all 10 cells unless a specific z-focus is set at each position).</li>
 <li>Save the imaging method for each cell (cell 1, cell 2 etc.).</li>
</ol>
4. Adjust the Automated Portion of the Program and Designate the 10 Cells You Will Follow for the 1 Hour Mitochondrial Fusion Assay
<ol>
 <li>On the left panel of the multitime window, select the "saving" panel, and specify the location where files will be saved.</li>
 <li>In the "Acquisition" panel, load the acquisition method saved for cell 1 in the scan configuration and check the z stack box. Also choose "marked Z- middle of z stack".</li>
 <li>In the "blocks" panel, select "single block at each location". Click on "add blocks" for each interval to be measured.</li>
 <li>In the "timing" panel, select "wait interval" and type "0" in the "wait interval before block at first location only". Blocks 2-4 will have "15 min" in this section.</li>
 <li>In the location panel, choose "move focus to load position between locations" and "load scan config when 'move to loc' or 'next loc' clicked. Under the "edit locations list" choose "clear all" and then select "multiple locations motorized stage".</li>
 <li>Under the "Bleach" panel, click the "bleach" box, and designate the configuration file in the "config" pull down menu, as designated in the window of the main software. Then, in the "bleaching" window, save the appropriate photoactivation method. In the regions panel, choose the ROI for this particular cell, and add this ROI to the multitime by choosing "add current region to ROI list" window. Be sure to select the same location in the "ROI" pull down menu.</li>
</ol>
For the timing to work properly and for each cell to have a 15 minute interval, two methods need to be saved. In the list of blocks, the first one will have the "real" photoactivation configuration, and the rest will have a "mock" configuration that does not use the two-photon laser. The first block will also be the only block that does not have a delay (BKIntv=0). Therefore, the method begins with one baseline scan, followed by a photoactivation scan. The rest of the blocks have a 900 sec BkIntv, and at each time point there are two scans just as at time 0 sec, to maintain the timing consistency.
<ol start="7">
 <li>For each of the 10 cells to be followed over time, perform this sequence: 
 Find a PAGFP expressing cell-save its imaging method 
 Location panel- mark the stage position, specify imaging method 
 Main ROI panel-choose a region of interest to be photoactivated Bleach panel-save specific ROI and also load it in the ROI box: 
 Erase the ROI and reset the scan zoom to 1 
 Find the next cell and set the zoom 
 Repeat the process for the next 9 cells</li>
</ol>
Check that each stage location and all blocks have the appropriate imaging method (scan configuration) associated. Also make sure that the first block at each location corresponds to the appropriate photoactivation method while the rest contain a "mock" method. Finally, select "run".
5. Analyze the PA-GFP Signal Intensity
<ol>
 <li>For the cells in which the photoactivatable-GFP signal colocalizes with the red TMRE signal, subtract the background in the PAGFP images (here we use metamorph).</li>
 <li>Then export the data to excel and calculate the average intensity for z-stacks at each time point. Discard optical sections that have no signal.</li>
 <li>Measure the signal dilution in each z-stack by calculating the percentage of the original photoactivated signal.</li>
</ol>
Each data point is scanned and recorded twice, resulting in duplicate z-stack information for each time point. Check that the z-stack values agree. If not; this is indicative of a problem (e.g. focus, cell movement).
6. Representative Results
When a fusion event occurs between an activated and non-activated PA-GFP mitochondrion, the PAGFP within the mitochondrial matrix mixes with the non-labeled matrix and becomes diluted over a larger area, decreasing the signal intensity (Figure 1A). In the INS1 cell, a significant decrease in signal intensity occurs every 15 minutes, until equilibration of mitochondrial fusion has been reached (about 1 hour). Note that the cell in Figure 1B exhibits nearly complete colocalization of PA-GFP and TMRE signals. In these assays a very low concentration of TMRE (15 nM) is used to help target the photoactivation of PAGFP, and also to monitor cell health. Cells with an abundance of depolarized mitochondria will have incomplete colocalization of PAGFP and TMRE and should not be analyzed because this indicates either phototoxicity, or cells in a dying state.
The equilibration time for mitochondrial fusion is usually 1 hr in INS1 cells, when ~15% of the mitochondrial volume is activated. Sometimes, even if a small area is illuminated, most of mitochondria become photoactivated due to being highly networked, in which case further fusion is difficult to detect. Other cell types may exhibit different equilibration times and should be tested at shorter intervals and over a longer period to characterize mitochondrial dynamics. To inhibit mitochondrial fusion, cells can be placed within a lipotoxic environment. It has previously been demonstrated that 0.4 mM palmitate fragments mitochondria, and inhibits mitochondrial fusion9. This effect can be seen in Figure 2, where mitochondria are fragmented, but the signal intensity of the mtPAGFP does not change as much as under normal conditions (Figure 1). Therefore, the dilution of the PA-GFP signal is likely to be due to mitochondrial fusion rather than fission. In other cells types, we suggest using other known ways to induce mitochondrial fragmentation, such as silencing OPA1 which is necessary for mitochondrial fusion14.
<img alt="Figure 1" src="/files/ftp_upload/3991/3991fig1.jpg" /> 
Figure 1. Typical dilution of the PA-GFP signal after photoactivation during a mitochondrial fusion assay. A. PA-GFP becomes progressively dimmer due to mitochondrial fusion events leading to the dilution of the protein over an increasing area as can be seen in these projected images of 6 optical sections quantified every 15 minutes. B. TMRE with PA-GFP shows that the mitochondria are active and not depolarized. <a href="https://www.jove.com/files/ftp_upload/3991/3991fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 2" src="/files/ftp_upload/3991/3991fig2.jpg" /> 
Figure 2. Mitochondrial fusion inhibited with 0.4 mM palmitate decreases the dilution of PA-GFP. A. Projections of 6 optical sections showing short mitochondria with an unchanging signal intensity over time. B. Colocalization of PA-GFP with TMRE shows that mitochondria are not depolarized. <a href="https://www.jove.com/files/ftp_upload/3991/3991fig2large.jpg" target="_blank"> Click here to view larger figure</a>.

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Production and Free Access to this article is sponsored by Zeiss, Inc.

This method allows for the imaging of around 10 cells at a time, if acquisition occurs every 15 minutes after photoactivation. The exact number of cells will depend on how quickly one is able to locate and mark the mtPAGFP expressing cells within the culture dish, and how quickly one can set up all the software parameters. To make the automation run smoothly an even layer of cells should be used because the designated z-stack margins will apply for all cells. 
The size of this initial photoactivation area will govern the equilibration time. To be able to measure mitochondrial fusion, it is important to photoactivate only 10-20% of the network, such that the spread of the signal to the rest of the network can be monitored over time. If too much of the network is photoactivated, it is possible that complete fusion will occur too quickly, and the event will not be captured. 
Extreme care must be taken to adjust the laser power of the two-photon laser as well as the TMRE concentration to avoid phototoxicity which leads to mitochondrial depolarization. Ensuring that the mtPAGFP signal colocalizes with the TMRE signal can help in assessing phototoxicity and general cell health 8,15. Illumination with epifluoerescent light should be avoided. While searching for cells expressing mtPAGFP, the pinhole should be maximally open while scanning with a low power 488nm excitation. Adjusting the two-photon laser power to photoactivate enough PA-GFP to measure the signal over 1 hour but not to oversaturate any of the cells can be tricky 8. However, time should be spent in this optimization step because once automated program is started, it is tedious to stop it, to choose more cells, and resume.
For quality control, acquisition of a differential interference contrast (DIC) image (or transmitted light) to monitor the focus on the cells can be very helpful and also a good way to detect bubbles formed in the immersion oil during scanning; this sometimes happens from the movements of the stage. 
Although using this mtPAGFP method gathers data on the unidirectional movement of mitochondrial matrix proteins from photoactivated mitochondria to those that are not labeled, it is conceivable to utilize this technique to study other processes. For example, specific fluorochromes can be attached to membrane proteins to observe their specific movement during fusion events, as has been shown for ABC-me, the fusion of which occurs on a different time scale from the mixing of soluble matrix proteins 15.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>COXIII-adenoviral PA-GFP</td>
 <td>Dr. Lippincott-Schwartz</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>TMRE</td>
 <td>Invitrogen</td>
 <td>T669</td>
 </tr>
 <tr>
 <td>Zeiss LSM 710 confocal</td>
 <td>Zeiss</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

a faster, high resolution, mtpa-gfp-based mitochondrial fusion assay acquiring kinetic data of multiple cells in parallel using confocal microscopy

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

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

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

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

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Biology

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Analysis of SNARE-mediated Membrane Fusion Using an Enzymatic Cell Fusion Assay

The interactions of SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) proteins on vesicles (v-SNAREs) and on target membranes (t-SNAREs) catalyze intracellular vesicle fusion1-4. Reconstitution assays are essential for dissecting the mechanism and regulation of SNARE-mediated membrane fusion5. In a cell fusion assay6,7, SNARE proteins are expressed ectopically at the cell surface. These "flipped" SNARE proteins drive cell-cell fusion, demonstrating that SNAREs are sufficient to fuse cellular membranes. Because the cell fusion assay is based on microscopic analysis, it is less efficient when used to analyze multiple v- and t-SNARE interactions quantitatively.
Here we describe a new assay8 that quantifies SNARE-mediated cell fusion events by activated expression of &beta;-galactosidase. Two components of the Tet-Off gene expression system9 are used as a readout system: the tetracycline-controlled transactivator (tTA) and a reporter plasmid that encodes the LacZ gene under control of the tetracycline-response element (TRE-LacZ). We transfect tTA into COS-7 cells that express flipped v-SNARE proteins at the cell surface (v-cells) and transfect TRE-LacZ into COS-7 cells that express flipped t-SNARE proteins at the cell surface (t-cells). SNARE-dependent fusion of the v- and t-cells results in the binding of tTA to TRE, the transcriptional activation of LacZ and expression of &beta;-galactosidase. The activity of &beta;-galactosidase is quantified using a colorimetric method by absorbance at 420 nm.
The vesicle-associated membrane proteins (VAMPs) are v-SNAREs that reside in various post-Golgi vesicular compartments10-15. By expressing VAMPs 1, 3, 4, 5, 7 and 8 at the same level, we compare their membrane fusion activities using the enzymatic cell fusion assay. Based on spectrometric measurement, this assay offers a quantitative approach for analyzing SNARE-mediated membrane fusion and for high-throughput studies.

We have developed a cell fusion assay that quantifies SNARE-mediated membrane fusion events by activated expression of &beta;-galactosidase.

The interactions of SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor)  proteins on vesicles (v-SNAREs) and on target membranes ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

High-resolution Respirometry to Assess Mitochondrial Function in Permeabilized and Intact Cells

A high-resolution oxygraph is a device for measuring cellular oxygen consumption in a closed-chamber system with very high resolution and sensitivity in biological samples (intact and permeabilized cells, tissues or isolated mitochondria). The high-resolution oxygraph device is equipped with two chambers and uses polarographic oxygen sensors to measure oxygen concentration and calculate oxygen consumption within each chamber. Oxygen consumption rates are calculated using software and expressed as picomoles per second per number of cells. Each high-resolution oxygraph chamber contains a stopper with injection ports, which makes it ideal for substrate-uncoupler-inhibitor titrations or detergent titration protocols for determining effective and optimum concentrations for plasma membrane permeabilization. The technique can be applied to measure respiration in a wide range of cell types and also provides information on mitochondrial quality and integrity, and maximal mitochondrial respiratory electron transport system capacity.

High-resolution respirometry is used to determine mitochondrial oxygen consumption. This is a straightforward technique to determine mitochondrial respiratory chain complexes&#39; (I-IV) respiratory rates, maximal mitochondrial electron transport system capacity, and mitochondrial outer membrane integrity.

A high-resolution oxygraph is a device for measuring cellular oxygen consumption in a closed-chamber system with very high resolution and sensitivity in ...

Combining Mitotic Cell Synchronization and High Resolution Confocal Microscopy to Study the Role of Multifunctional Cell Cycle Proteins During Mitosis

Study of the various regulatory events of the cell cycle in a phase-dependent manner provides a clear understanding about cell growth and division. The synchronization of cell populations at specific stages of the cell cycle has been found to be very useful in such experimental endeavors. Synchronization of cells by treatment with chemicals that are relatively less toxic can be advantageous over the use of pharmacological inhibitory drugs for the study of consequent cell cycle events and to obtain specific enrichment of selected mitotic stages. Here, we describe the protocol for synchronizing human cells at different stages of the cell cycle, including both in S phase and M phase with a double thymidine block and release procedure for studying the functionality of mitotic proteins in chromosome alignment and segregation. This protocol has been extremely useful for studying the mitotic roles of multifunctional proteins which possess established interphase functions. In our case, the mitotic role of Cdt1, a protein critical for replication origin licensing in G1 phase, can be studied effectively only when G2/M-specific Cdt1 can be depleted. We describe the detailed protocol for depletion of G2/M-specific Cdt1 using double thymidine synchronization. We also explain the protocol of cell fixation, and live cell imaging using high resolution confocal microscopy after thymidine release. The method is also useful for analyzing the function of mitotic proteins under both physiological and perturbed conditions such as for Hec1, a component of the Ndc80 complex, as it enables one to obtain large sample sizes of mitotic cells for fixed and live cell analysis as we show here.&#160;&#160;

We present a protocol for double thymidine synchronization of HeLa cells followed by analysis using high resolution confocal microscopy. This method is key to obtaining large number of cells that proceed synchronously from S phase to mitosis, enabling studies on mitotic roles of multifunctional proteins which also possess interphase functions.

Study of the various regulatory events of the cell cycle in a phase-dependent manner provides a clear understanding about cell growth and division. The ...

High-resolution Respirometry to Measure Mitochondrial Function of Intact Beta Cells in the Presence of Natural Compounds

High-resolution respirometry allows for the measurement of oxygen consumption of isolated mitochondria, cells and tissues. Beta cells play a critical role in the body by controlling blood glucose levels through insulin secretion in response to elevated glucose concentrations. Insulin secretion is controlled by glucose metabolism and mitochondrial respiration. Therefore, measuring intact beta cell respiration is essential to be able to improve beta cell function as a treatment for diabetes. Using intact 832/13 INS-1 derived beta cells we can measure the effect of increasing glucose concentration on cellular respiration. This protocol allows us to measure beta cell respiration in the presence or absence of various compounds, allowing one to determine the effect of given compounds on intact cell respiration. Here we demonstrate the effect of two naturally occurring compounds, monomeric epicatechin and curcumin, on beta cell respiration under the presence of low (2.5 mM) or high glucose (16.7 mM) conditions. This technique can be used to determine the effect of various compounds on intact beta cell respiration in the presence of differing glucose concentrations.

The goal of this protocol is to measure the effect of glucose-mediated changes to mitochondrial respiration in the presence of natural compounds on intact 832/13 beta cells using high-resolution respirometry.

High-resolution respirometry allows for the measurement of oxygen consumption of isolated mitochondria, cells and tissues. Beta cells play a critical role ...

Parallel High Throughput Single Molecule Kinetic Assay for Site-Specific DNA Cleavage

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable of measuring SSDC in many single DNA molecules simultaneously. Bead-tethered substrate DNAs, each containing a single copy of the target sequence, are prepared in a microfluidic flow channel. An external magnet applies a weak force to the paramagnetic beads. The integrity of up to 1,000 individual DNAs can be monitored by visualizing the microbeads under darkfield imaging using a wide-field, low magnification objective. Injecting of a restriction endonuclease, NdeI, initiates the cleavage reaction. Video microscopy is used to record the exact moment of each DNA cleavage by observing the frame in which the associated bead moves up and out of the focal plane of the objective. Frame-by-frame bead counting quantifies the reaction, and an exponential fit determines the reaction rate. This method allows collection of quantitative and statistically significant data on single molecule SSDC reactions in a single experiment.

A highly parallel method for measuring the site-specific cleavage of DNA at the single molecule level is described. This protocol demonstrates the technique using the restriction endonuclease NdeI. The method can easily be modified to study any process that results in site-specific DNA cleavage.

Site-specific DNA cleavage (SSDC) is a key step in many cellular processes, and it is crucial to gene editing. This work describes a kinetic assay capable ...

Real-Time Analysis of Bioenergetics in Primary Human Retinal Pigment Epithelial Cells Using High-Resolution Respirometry

Metabolic dysfunction of retinal pigment epithelial cells (RPE) is a key pathogenic driver of retinal diseases such as age-related macular degeneration (AMD) and proliferative vitreoretinopathy (PVR). Since RPE are highly metabolically-active cells, alterations in their metabolic status reflect changes in their health and function. High-resolution respirometry allows for real-time kinetic analysis of the two major bioenergetic pathways, glycolysis and mitochondrial oxidative phosphorylation (OXPHOS), through quantification of the extracellular acidification rate (ECAR) and oxygen consumption rate (OCR), respectively. The following is an optimized protocol for conducting high-resolution respirometry on primary human retinal pigment epithelial cells (H-RPE). This protocol provides a detailed description of the steps involved in producing bioenergetic profiles of RPE to define their basal and maximal OXPHOS and glycolytic capacities. Exposing H-RPE to different drug injections targeting the mitochondrial and glycolytic machinery results in defined bioenergetic profiles, from which key metabolic parameters can be calculated. This protocol highlights the enhanced response of BAM15 as an uncoupling agent compared to carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) to induce the maximal respiration capacity in RPE. This protocol can be utilized to study the bioenergetic status of RPE under different disease conditions and test the efficacy of novel drugs in restoring the basal metabolic status of RPE.

The metabolic status of human retinal pigment epithelial cells (H-RPE) reflects their health and function. Presented here is an optimized protocol for examining the real-time metabolic flux of H-RPE using high-resolution respirometry.

Metabolic dysfunction of retinal pigment epithelial cells (RPE) is a key pathogenic driver of retinal diseases such as age-related macular degeneration ...

Quantification of Mitochondrial Membrane Potential and Superoxide Levels

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

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

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

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

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