DLM is supported by an NSF GRFP. ALM is supported by funding from the Smith Family Award for Excellence in Biomedical Research.

1. Generation of hTERT-RPE-1 cells stably expressing RFP-Histone 2B (RFP-H2B) and &#945;-tubulin-EGFP (tub-EGFP)
NOTE: All steps follow aseptic techniques and take place in a biosafety level II+ (BSL2+) safety cabinet.
<ol>
	<li>Generate retrovirus carrying the genes of interest (&#945;-tubulin-EGFP and RFP-H2B) by the transfection of 293T cells with the appropriate lentiviral plasmids according to the manufacturer&#39;s instructions of the lipid-based transfection delivery system.
	<ol>
		<li>Day 1: Use a disposable glass Pasteur pipette to aspirate the cell culture medium from a plate of sub-confluent 293T cells and wash off the residual medium with phosphate buffered saline (PBS) by adding 5 mL of PBS. Swirl to distribute PBS over the plate bottom, then aspirate the PBS with a sterile disposable glass Pasteur pipette.</li>
		<li>Add 2 mL of 0.05% trypsin that has been pre-warmed to 37 &#176;C and return the plate to a humidified incubator at 37 &#176;C with 5% CO2 for 2 to 5 min to allow adherent cells to be released from the cell culture dish surface.</li>
		<li>Add 8 mL of fresh Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin to the plate containing trypsin.</li>
		<li>Resuspend the cells by gently pipetting and transfer the suspension to a sterile 15 mL conical tube. Place the conical tube in a balanced centrifuge and spin at 161 x g for 5 min at room temperature to gently pellet the cells.</li>
		<li>Aspirate the medium/trypsin solution from pelleted cells and resuspend cells in 10 mL of DMEM supplemented with 10% FBS and 1% penicillin/streptomycin. Use a hemocytometer to count the 293T cell suspension and plate 2 x 106 293T cells per well of a 6 well plate.</li>
		<li>Culture cells in a total volume of 2 mL of Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin and maintain&#160;in a humidified incubator at 37 &#176;C with 5% CO2.</li>
		<li>Day 2: Pipette 7 &#181;L of lipid-based transfection delivery reagent and 100 &#181;L of reduced serum medium into a 1.5 mL microcentrifuge tube and allow it to incubate at room temperature for 5 min (tube #1).</li>
		<li>In a separate 1.5 mL microcentrifuge tube, pipette 1 &#181;g of RFP-H2B expression vector (or 1 &#181;g of &#945;-tubulin-EGFP expression vector), together with 5 &#181;L enhancer reagent (as required per manufacturer&#39;s guidelines), 0.5 &#181;g pMD2.G, and 1 &#181;g psPAX2 and 100 &#181;L of reduced serum medium (tube #2).</li>
		<li>Combine the contents of step 1.1.7 and step 1.1.8 by carefully pipetting tube #2 into tube #1 and incubate for 20 min at room temperature. 
		NOTE: The reaction can be scaled as needed for transfection of cells in additional wells.</li>
		<li>Pipette the transfection reaction dropwise to the desired well containing 293T cells in 2 mL of medium and return the dish to the humidified incubator at 37 &#176;C with 5% CO2. 
		NOTE: To generate cells expressing both RFP-H2B and &#945;-tubulin-EGFP, generate separate virus for each expression construct (steps 1.1.7- 1.1.9).</li>
		<li>Day 3: Aspirate and replace the medium with 2 mL of fresh DMEM supplemented with 10% FBS and 1% penicillin/streptomycin. Return the dish to the humidified incubator at 37 &#176;C with 5% CO2.</li>
	</ol>
	</li>
	<li>Day 4: Collect the medium containing expressed virus particles, being cautious not to disrupt or remove 293T cells. Filter the medium containing virus particles by passing it through a 0.45 &#181;M filter attached to a 5 mL syringe. Aliquot virus for short term storage at 4 &#176;C, or long term storage at -80 &#176;C. 
	NOTE: Viral particles obtained in this manner will be present in a range of ~1 x 107 to 1 x 108 Transducing Units/mL. As the viral-producing cells and cells to be infected are both cultured in the same medium, viral concentration to replace the medium is not required and filtered viral particles can be used directly to infect cells.</li>
	<li>Seed 2 x 105 hTERT-RPE-1 cells per well of a 6-well dish in preparation for the viral infection. Culture cells in 2 mL of DMEM supplemented with 10% FBS and 1% penicillin/streptomycin and maintain in a humidified incubator at 37 &#176;C with 5% CO2.</li>
	<li>Day 5: Add hexadimethrine bromide (e.g., polybrene)to the hTERT-RPE-1 cells to a final concentration of 8 &#181;g/mL. Infect cells with a mixture of 500 &#181;L of virus diluted in 500 &#181;L of cell culture medium. 
	NOTE: Hexadimethrine bromide stock solution is prepared in sterile distilled water, then filtered through a 0.45 &#956;m filter.</li>
	<li>Day 6: Using a disposable glass Pasteur pipette, aspirate the medium from wells containing the virus infected cells. Replace the cell culture medium with 2 mL of DMEM containing 10% FBS, 1% penicillin/streptomycin, and appropriate concentrations of antibiotic to select for the plasmid integration and expression. 
	NOTE: The &#945;-tubulin-EGFP expression plasmid used in the described experiments carries a puromycin-resistance gene and the RFP-H2B expression plasmid carries a blasticidin resistance gene. Therefore, 10 &#181;g/mL of puromycin and 2 &#181;g/mL of blasticidin are used for the selection of &#945;-tubulin-EGFP, RFP-H2B expressing hTERT RPE-1 cells. Concentrations of antibiotic used for the selection may differ for various cell lines used.</li>
	<li>Maintain cells under antibiotic selection for 5-7 days, replacing the medium every 3 days with fresh medium containing appropriate selection reagents. 
	NOTE: Cells should be maintained at sub-confluence during the selection and should be expanded as needed, as described in steps 1.1.1 to 1.1.4.</li>
	<li>Use immunofluorescence imaging to confirm the expression of tagged constructs. 
	NOTE: If desired, single cell clones can be derived to obtain uniform expression levels within the cell population. Once stable clones are confirmed, cells can be maintained under the standard culture conditions in the absence of antibiotic selection.</li>
</ol>
2. Preparation of cells for live cell imaging following mitotic spindle perturbations
NOTE: Use aseptic techniques and perform the steps in a BSL2 safety cabinet.
<ol>
	<li>Using a sterile disposable glass Pasteur pipette, aspirate the medium from the culture plate containing the cell line that carries the expression construct(s) (from step 1.7). Briefly wash the cells with 10 mL of sterile PBS. Swirl to distribute PBS over the plate bottom, then aspirate PBS with a sterile disposable glass Pasteur pipette.</li>
	<li>Add 2 mL of 0.05% trypsin to the 10 cm plate. Incubate the plate at 37 &#176;C for 2-5 min or until cells have detached from the plate surface.</li>
	<li>Add 8 mL of fresh medium to the plate containing trypsin. Resuspend the cells by gently pipetting and transfer the suspension to a sterile 15 mL conical tube. Place the conical tube in a balanced centrifuge and spin at 161 x g for 5 min at room temperature to gently pellet the cells.</li>
	<li>Carefully aspirate the supernatant and resuspend in 10 mL of PBS by pipetting gently with a 10 mL serological pipette. Place the conical tube in a balanced centrifuge and spin at 161 x g for 5 min at room temperature to gently pellet the cells.</li>
	<li>Carefully aspirate the supernatant and resuspend in 10 mL of the fresh medium by pipetting gently with a 10 mL serological pipette.</li>
	<li>Count cells, then calculate the cell number using a hemacytometer and dilute to a concentration of 1-2 x 105 cells/mL in the cell culture medium. Seed 500 &#181;L of the cell suspension to each well of a sterile 12-well imaging bottom plate. Place the plate in the cell culture incubator and allow cells to adhere to the plate surface. 
	NOTE: Live cell imaging requires cells to be well-adhered so that they remain associated with the imaging plane during image acquisition. The duration of the time needed for cells to adhere following plating can differ from one cell line to the next and should be optimized for the cell line under study.</li>
	<li>Up to 30 min prior to initiating time-lapse imaging, add a relevant concentration of a mitotic drug to one or more of the wells seeded with cells. To account for the potential impact of the organic solvent on cellular behavior, add an equal volume of the inhibitor&#39;s diluent to cells as controls. For example, ensure that the addition of 100 nM of the specific inhibitor of the mitotic kinase Aurora A (e.g., alisertib) is paralleled with an equal volume of DMSO, the diluent for this drug, in a control well of cells. 
	NOTE: Comparative analysis of dynamic changes in the mitotic progression require wells of cells to be prepared in the absence of perturbations to enable imaging of normal mitoses in parallel to experimental conditions.</li>
</ol>
3. Microscope set up for time-lapse imaging of RFP-H2B, &#945;-tubulin-GFP expressing cells (Figure 1, Figure 2)
<ol>
	<li>Place the cell culture plate containing RFP-H2B, &#945;-tubulin-GFP expressing hTERT RPE-1 cells to be imaged into an appropriate stage insert on an inverted epifluorescence microscope that is equipped with a high resolution camera (pixel size of 0.67 &#956;m at 20x), an environmental chamber preheated to 37 &#176;C, and a delivery system for humidified 5% CO2. 
	NOTE: Either enclosed environmental chambers or temperature controlled stage inserts may be appropriate provided humidified 5% CO2 can be delivered and stable temperature control obtained.</li>
	<li>Use a 20x air objective with a numerical aperture of 0.5 and equipped for the high contrast fluorescence and phase contrast or brightfield imaging. View cells with the phase contrast or brightfield and adjust the course and fine focus on the microscope to bring cells into focus.</li>
	<li>Identify and set the optimal exposure times for brightfield, GFP, and RFP image acquisition by selecting the respective filter cube with appropriate excitation and emission for the fluorophores that will be imaged. Alternatively, click the auto exposure button to input&#160;a predetermined exposure time. If the signal is not sufficiently intense, select pixel binning to enable shorter exposure times by clicking on the binning tab and selecting 2 x 2 pixel binning from the drop down menu. 
	NOTE: Phototoxicity can impair mitotic progression and cell viability. Failure of mitotic cells in control populations to complete normal mitoses may be an indication that exposure times and/or imaging duration needs to be further optimized.</li>
	<li>Use an acquisition and analysis software that enables multi-coordinate, multi-well imaging to be acquired concurrently and define the parameters for the image acquisition.
	<ol>
		<li>Select and calibrate the microscope stage to the multi-well dish, according to the manufacturer&#39;s instructions. Use the image acquisition software to highlight or otherwise select the wells that will be imaged by clicking on the relevant wells in the diagram of the plate format that is being used.</li>
		<li>Under the GeneratedPoints control panel (Figure 2), define the coordinates within each well that will be imaged by clicking on the Point Placement tab and selecting predefined or random coordinate placement from the drop down menu. Select the Working Area tab and select restricted from the drop down menu to restrict the coordinate selection area to exclude the boundaries of the well. Click the Count and Distribution tabs to select the number and distribution of points to be captured per well, respectively. 
		NOTE: The number of coordinates that can be imaged per well within the 5 min timepoint increment will be limited by the number of wells to be imaged, as well as the exposure time for each channel. Typically, 5-8 coordinates per well are sufficient to observe at least 50 cells in each condition progress through mitosis within 4 h.</li>
		<li>Select and input the time interval and duration to collect images by clicking on and inputting the values in the TimeSequence control panel. 
		NOTE: Acquisition of images every 1 to 5 minutes is appropriate to monitor the dynamics of mitotic progression. The overall&#160;duration of imaging should reflect the desired endpoint and proliferation rate of the cell line. For example, 4 hours is sufficient to see many cells progress through mitosis, 16 hours is sufficient to see most RPE-1 cells in an asynchronous population progress through mitosis.</li>
	</ol>
	</li>
</ol>
4. Time-lapse image analysis to determine metaphase timing and mitotic cell fate following mitotic spindle perturbations
NOTE: Perform the image analysis using an image acquisition software (Table of Materials), ImageJ, or comparable image analysis software.
<ol>
	<li>Visualize RFP-H2B labeled chromatin by selecting the images captured with the RFP filter cube in place. Identify a cell entering mitosis as indicated by initial chromatin compaction (Figure 2C, blue arrowhead) and nuclear envelope break down (when &#945;-tubulin-EGFP is no longer excluded by the nuclear boundary).</li>
	<li>Determine the mitotic timing of metaphase alignment and anaphase onset in individual cells: Track the cell through consecutive timepoints in the acquired movie to determine the number of timepoints/minutes from mitotic entry until RFP-H2B-labelled chromatin completes alignment at the cell equator during metaphase (Figure 2C, yellow arrowhead).</li>
	<li>To monitor mitotic timing, mitotic fidelity, and cell fate, continue to track the cell through consecutive timepoints to identify the time coordinate at which anaphase chromosome segregation is apparent (Figure 2C, white arrowhead) and/or where chromatin decompaction and nuclear envelope reformation (as indicated by tubulin exclusion from the nucleus) has occurred. 
	NOTE: Perturbations in spindle assembly or mitotic progression can be assessed as a function of the time required to obtain a bipolar mitotic spindle and achieve complete chromosome alignment, or to complete anaphase chromosome segregation.</li>
	<li>Visualize RFP-H2B to identify cells in each population that exhibit mitotic defects including lagging chromosomes and chromatin bridges during anaphase chromosome segregation. 
	NOTE: Compromised mitotic fidelity may also result in multinucleated or micronucleated daughter cells and can be visualized in cells post mitotic exit, as in Figure 3.</li>
</ol>

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The authors have nothing to disclose.

The temporal resolution provided by the time-lapse imaging allows for the visualization and assessment of sequential cellular events within single cells. Approaches that make use of cellular synchronization followed by the collection and fixation of cells at sequential time points are limited in that comparisons are ultimately made between populations of cells. In contexts where the cellular response to perturbations may be non-uniform, or where the process being visualized is dynamic, live cell time-lapse imaging is bet...

Image-based analysis of fixed cells is commonly used to assess the cell population level changes in response to various perturbations. When combined with cell synchronization, followed by the collection and imaging of serial time points, such approaches can be used to suggest a cellular sequence of events. Nevertheless, fixed cell imaging is limited in that temporal relationships are implied for a population and not demonstrated at the level of individual cells. In this way, while fixed cell imaging and analysis is sufficient to observe robust phenotypes and steady-state changes, the ability to detect transient changes over time and changes that impact only a subpopulation of the cells is imperfect. In contrast, live cell imaging is an eloquent tool that can be used to observe cellular and subcellular processes within a single cell, or cellular population, over time and without the aid of synchronization approaches that may themselves impact cellular behavior1,2,3,4,5,6.
The formation of a bipolar mitotic spindle is essential for the proper chromosome segregation during cell division, resulting in two genetically identical daughter cells. Defects in mitotic spindle structure that corrupt mitotic progression and compromise the fidelity of chromosome segregation can result in catastrophic cell divisions and reduced cell viability. For this reason, mitotic poisons that alter spindle formation are promising therapeutics to limit the rapid proliferation of cancer cells7,8,9. Nevertheless, fixed cell analysis of spindle structure following the addition of mitotic poisons is limited in its ability to assess the dynamic process of spindle formation and may not indicate whether observed changes in spindle structure are permanent or are instead transient and may be overcome to permit successful cell division.
In this protocol, we describe an approach to assess the dynamics of mitosis following spindle perturbations by live cell imaging. Using the hTERT immortalized RPE-1 cell line engineered to express an RFP-tagged Histone 2B to visualize chromatin, together with an EGFP-tagged &#945;-tubulin to visualize microtubules, the timing of metaphase chromosome alignment, anaphase onset, and ultimately mitotic cell fate are assessed using visual cues of chromosome movement, compaction, and nuclear morphology.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.05% Trypsin</td>
			<td>Gibo-Life sciences</td>
			<td>25-510</td>
			<td>A serine protease used to release adherent cells from culture dishes</td>
		</tr>
		<tr>
			<td>15ml centrifuge tubes</td>
			<td>Olympus Plastics</td>
			<td>28-101</td>
			<td></td>
		</tr>
		<tr>
			<td>20x CFI Plan Fluor objective&#160;</td>
			<td>Nikon</td>
			<td></td>
			<td>For use in Live-cell imaging to visualize both bright field and fluorescence</td>
		</tr>
		<tr>
			<td>293T Cells</td>
			<td>ATCC</td>
			<td>CRL-3216</td>
			<td>For use in retroviral transfection; used in step 1.1</td>
		</tr>
		<tr>
			<td>a-tubulin-EGFP&#160;</td>
			<td>Addgene</td>
			<td>various numbers</td>
			<td>Expression vector for alpha tubulin fused to a green fluorescent protein tag; for use in the visualization of tubulin in live-cell imaging: commercially available through addgene and other vendors</td>
		</tr>
		<tr>
			<td>Alisertib</td>
			<td>Selleckchem</td>
			<td>S1133</td>
			<td>Small molecule inhibitor of the mitotic kinase Aurora A. Stock concentration is prepared at 10mM in DMSO, and used at a final concentration of 100nM.</td>
		</tr>
		<tr>
			<td>Blasticidin</td>
			<td>Invitrogen</td>
			<td>A11139-03</td>
			<td>Antibiotic selection agent; used to select for a-tub-EGFP expressing cells</td>
		</tr>
		<tr>
			<td>C02</td>
			<td>Airgas</td>
			<td></td>
			<td>For use in cell culture and live cell imaging</td>
		</tr>
		<tr>
			<td>Chroma ET-DS Red (TRITC/Cy3)</td>
			<td>Chroma</td>
			<td>49005</td>
			<td>Single band filter set; excitation wavelength 545nm with 25nm bandwidth and emission at 605nm wavelength with 70nm bandwidth; for visualization of H2B-RFP</td>
		</tr>
		<tr>
			<td>Chroma ET-EGFP (FITC/Cy2)</td>
			<td>Chroma</td>
			<td>49002</td>
			<td>Single band filter set; excitation wavelength 470nm with 40nm bandwidth and emission at 525nm wavelength with 50nm bandwidth; for visualization of GFP-tubulin</td>
		</tr>
		<tr>
			<td>disposable glass Pastuer pipets, sterilized&#160;</td>
			<td>Fisher Scientific</td>
			<td>13-678-6A</td>
			<td>For use in aspirating cells&#160;</td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s Modified Eagle Medium (DMEM)</td>
			<td>Gibo-Life sciences</td>
			<td>11965-084</td>
			<td>Cell culture medium for growth of RPE-1 and 293T cells</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>Gibo-Life sciences</td>
			<td>10438-026</td>
			<td>Cell culture medium supplement</td>
		</tr>
		<tr>
			<td>Lipofectamine 3000 and p3000</td>
			<td>Invitrogen</td>
			<td>L3000-015</td>
			<td>Lipid based transfection reagent for transfection of plasmids; used in 1.1.4</td>
		</tr>
		<tr>
			<td>Multi well Tissue Culture dishes</td>
			<td>Corning</td>
			<td>various</td>
			<td>for use in cell culture, transfection/infection, and live cell imaging</td>
		</tr>
		<tr>
			<td>Nikon Ti-E microscope</td>
			<td>Nikon</td>
			<td></td>
			<td>Inverted epifluorescence microscope for use in live-cell imaging</td>
		</tr>
		<tr>
			<td>NIS Elements HC&#160;</td>
			<td>Nikon</td>
			<td>Version 4.51</td>
			<td>Image acquisition and analysis software; used in sections 3 &#38; 4</td>
		</tr>
		<tr>
			<td>OPTI-MEM</td>
			<td>Gibo-Life sciences</td>
			<td>31985-070</td>
			<td>Reduced serum medium for cell transfection; used in step 1.1.3</td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin&#160;</td>
			<td>Gibo-Life sciences</td>
			<td>15140-122</td>
			<td>antibiotic used in cell culture medium</td>
		</tr>
		<tr>
			<td>phosphate bufferred saline (PBS)</td>
			<td>Caisson labs</td>
			<td>PBP06-10X1LT</td>
			<td>sterile saline solution for use with cell culture</td>
		</tr>
		<tr>
			<td>pMD2.G</td>
			<td>Addgene</td>
			<td>12259</td>
			<td>Lentiviral VSV-G envelope expression construct; used in step 1.1.4</td>
		</tr>
		<tr>
			<td>Polybrene</td>
			<td>Sigma-Aldrich</td>
			<td>H9268</td>
			<td>Cationic polymer used to enhane viral infection efficiency; used in step 1.1.10</td>
		</tr>
		<tr>
			<td>psPAX</td>
			<td>Addgene</td>
			<td>12260</td>
			<td>2nd generation lentiviral packaging plasmid; used in step 1.1.4</td>
		</tr>
		<tr>
			<td>Puromycin</td>
			<td>Invitrogen</td>
			<td>ant-pr-1</td>
			<td>Antibiotic selection agent; used to select for RFP-H2B expressing cells</td>
		</tr>
		<tr>
			<td>RFP- Histone 2B (H2B)</td>
			<td>Addgene</td>
			<td>various numbers</td>
			<td>Expression vector for red fluorescent protein-tagged histone 2B; for use in the visualization of chromatin in live-cell imaging: commercially available through Addgene and other vendors</td>
		</tr>
		<tr>
			<td>RNAi Max</td>
			<td>Invitrogen</td>
			<td>13778-150</td>
			<td>Lipid based transfection reagent for transfection of siRNA constructs</td>
		</tr>
		<tr>
			<td>RPE-1 cells</td>
			<td>ATCC</td>
			<td>CRL-4000</td>
			<td>Human retinal pigment epithelial cell line</td>
		</tr>
		<tr>
			<td>Tissue culture dish 100x20mm</td>
			<td>Corning&#160;</td>
			<td>353003</td>
			<td>for use in culturing adherent cells</td>
		</tr>
		<tr>
			<td>Zyla sCMOS camera&#160;</td>
			<td>Nikon</td>
			<td></td>
			<td>Camera attached to the micrscope, used for capturing images of cells</td>
		</tr>
	</tbody>
</table>

live cell imaging to assess the dynamics of metaphase timing and cell fate following mitotic spindle perturbations

Live cell time-lapse imaging is an important tool in cell biology that provides insight into cellular processes that might otherwise be overlooked, misunderstood, or misinterpreted by the fixed-cell analysis. While the fixed cell imaging and analysis is robust and sufficient to observe cellular steady-state, it can be limited in defining a temporal order of events at the cellular level and is ill-equipped to assess the transient nature of dynamic processes including mitotic progression. In contrast, live cell imaging is an eloquent tool that can be used to observe cellular processes at the single-cell level over time and has the capacity to capture the dynamics of processes that would otherwise be poorly represented in fixed cell imaging. Here we describe an approach to generate cells carrying fluorescently labeled markers of chromatin and microtubules and their use in live cell imaging approaches to monitor metaphase chromosome alignment and mitotic exit. We describe imaging-based techniques to assess the dynamics of spindle formation and mitotic progression, including the identification of cells at various stages in mitosis, identification&#160;and tracking of mitotic defects, and analysis of spindle dynamics and mitotic cell fate following the treatment with mitotic inhibitors.

Assessment of mitotic progression in the presence of spindle perturbations 
The regulation of spindle pole focusing is an essential step in proper bipolar spindle formation. Disruption in this process through protein depletions, drug inhibition, or alterations in centrosome number corrupt spindle structure and delay or halt mitotic progression10,11,12,<sup class="...

Watch this Scientific Journal Video about Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations at JoVE.com

Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations

Here we present a protocol to assess the dynamics of spindle formation and mitotic progression. Our application of time-lapse imaging enables the user to identify cells at various stages of mitosis, track and identify mitotic defects, and analyze spindle dynamics and mitotic cell fate upon exposure to anti-mitotic drugs.

Live cell time-lapse imaging is an important tool in cell biology that provides insight into cellular processes that might otherwise be overlooked, ...

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Unit 2

Cell Proliferation

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8.1K Views.  Worcester Polytechnic Institute. Here we present a protocol to assess the dynamics of spindle formation and mitotic progression. Our application of time-lapse imaging enables the user to identify cells at various stages of mitosis, track and identify mitotic defects, and analyze spindle dynamics and mitotic cell fate upon exposure to anti-mitotic drugs.

Video: Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Assembly, Loading, and Alignment of an Analytical Ultracentrifuge Sample Cell

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed centrifugation. When properly planned and executed, an AUC sedimentation velocity or sedimentation equilibrium experiment can reveal a great deal about a protein in regards to size and shape, sample purity, sedimentation coefficient, oligomerization states and protein-protein interactions.
This technique, however, requires a rigorous level of technical attention. Sample cells hold a sectored center piece sandwiched between two window assemblies. They are sealed with a torque pressure of around 120-140 in/lbs. Reference buffer and sample are loaded into the centerpiece sectors and then after sealing, the cells are precisely aligned into a titanium rotor so that the optical detection systems scan both sample and reference buffer in the same radial path midline through each centerpiece sector while rotating at speeds of up to 60, 000 rpm and under very high vacuum
Not only is proper sample cell assembly critical, sample cell components are very expensive and must be properly cared for to ensure they are in optimum working condition in order to avoid leaks and breakage during experiments. Handle windows carefully, for even the slightest crack or scratch can lead to breakage in the centrifuge. The contact between centerpiece and windows must be as tight as possible; i.e. no Newton s rings should be visible after torque pressure is applied. Dust, lint, scratches and oils on either the windows or the centerpiece all compromise this contact and can very easily lead to leaking of solutions from one sector to another or leaking out of the centerpiece all together. Not only are precious samples lost, leaking of solutions during an experiment will cause an imbalance of pressure in the cell that often leads to broken windows and centerpieces. In addition, plug gaskets and housing plugs must be securely in place to avoid solutions being pulled out of the centerpiece sector through the loading holes by the high vacuum in the centrifuge chamber. Window liners and gaskets must be free of breaks and cracks that could cause movement resulting in broken windows.
This video will demonstrate our procedures of sample cell assembly, torque, loading and rotor alignment to help minimize component damage, solution leaking and breakage during the perfect AUC experiment.

The analytical ultracentrifuge (AUC) sample cell holds sample and reference buffer and during experiments and is exposed to high vacuum and rotor speeds up to 60,000 rpm. This video will demonstrate the rigorous attention to detail necessary for assembly, loading and alignment of this very important component of an AUC experiment.

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed ...

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function. Our studies have found that dying cells send signals to their live neighbors to form and contract a ring of actin and myosin that ejects it out from the epithelial sheet while closing any gaps that might have resulted from its exit, a process termed cell extrusion1. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe a method to induce and image extrusion in the larval zebrafish epidermis. To visualize extrusion, we inject a red fluorescent protein labeled probe for F-actin into one-cell stage transgenic zebrafish embryos expressing green fluorescent protein in the epidermis and induce apoptosis by addition of G418 to larvae. We then use time-lapse imaging on a spinning disc confocal microscope to observe actin dynamics and epithelial cell behaviors during the process of apoptotic cell extrusion. This approach allows us to investigate the extrusion process in live epithelia and will provide an avenue to study disease states caused by the failure to eliminate apoptotic cells.

Dying cells are extruded from epithelial tissues by concerted contraction of neighboring cells without disrupting barrier function. The optical clarity of developing zebrafish provides an excellent system to visualize extrusion in living epithelia. Here we describe methods to induce and image extrusion in the larval zebrafish epidermis at cellular resolution.

Homeostatic maintenance of epithelial tissues requires the continual removal of damaged cells without disrupting barrier function.  Our studies have found ...

Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal which tissues descend from each cell of the embryo. Although fate maps are very useful for identifying the precursors of an organ and for elucidating the developmental path by which the descendant cells populate that organ in the normal embryo, they do not illustrate the full developmental potential of a precursor cell or identify the mechanisms by which its fate is determined. To test for cell fate commitment, one compares a cell's normal repertoire of descendants in the intact embryo (the fate map) with those expressed after an experimental manipulation. Is the cell's fate fixed (committed) regardless of the surrounding cellular environment, or is it influenced by external factors provided by its neighbors? Using the comprehensive fate maps of the Xenopus embryo, we describe how to identify, isolate and culture single cleavage stage precursors, called blastomeres. This approach allows one to assess whether these early cells are committed to the fate they acquire in their normal environment in the intact embryo, require interactions with their neighboring cells, or can be influenced to express alternate fates if exposed to other types of signals.

The fate of an individual embryonic cell can be influenced by inherited molecules and/or by signals from neighboring cells. Utilizing fate maps of the cleavage stage Xenopus embryo, single blastomeres can be identified for culture in isolation to assess the contributions of inherited molecules versus cell-cell interactions.

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal  which tissues descend from each cell of the embryo.  Although fate maps ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single procedure. Due to advances in microscope technology it is relatively easy to capture live cell images for the purpose of investigating cellular events in real time with minimal concern regarding phototoxicity to the cells. This protocol describes how to take live cell timelapse images of primary rat neonatal cardiomyocytes using a confocal spinning disk microscope following lentiviral and adenoviral transduction to modulate properties of the cell. The application of two different types of viruses makes it easier to achieve an appropriate transduction rate and expression levels for two different genes. Well focused live cell images can be obtained using the microscope&#8217;s autofocus system, which maintains stable focus for long time periods. Applying this method, the functions of exogenously engineered proteins expressed in cultured primary cells can be analyzed. Additionally, this system can be used to examine the functions of genes through the use of siRNAs as well as of chemical modulators.

This protocol describes a method of live cell imaging using primary rat neonatal cardiomyocytes following lentiviral and adenoviral transduction using confocal spinning disk microscopy. This enables detailed observations of cellular processes in living cardiomyocytes.

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single ...

Identification of MyoD Interactome Using Tandem Affinity Purification Coupled to Mass Spectrometry

Skeletal muscle terminal differentiation starts with the commitment of pluripotent mesodermal precursor cells to myoblasts. These cells have still the ability to proliferate or they can differentiate and fuse into multinucleated myotubes, which maturate further to form myofibers. Skeletal muscle terminal differentiation is orchestrated by the coordinated action of various transcription factors, in particular the members of the Muscle Regulatory Factors or MRFs (MyoD, Myogenin, Myf5, and MRF4), also called the myogenic bHLH transcription factors family. These factors cooperate with chromatin-remodeling complexes within elaborate transcriptional regulatory network to achieve skeletal myogenesis. In this, MyoD is considered the master myogenic transcription factor in triggering muscle terminal differentiation. This notion is strengthened by the ability of MyoD to convert non-muscle cells into skeletal muscle cells. Here we describe an approach used to identify MyoD protein partners in an exhaustive manner in order to elucidate the different factors involved in skeletal muscle terminal differentiation. The long-term aim is to understand the epigenetic mechanisms involved in the regulation of skeletal muscle genes, i.e., MyoD targets. MyoD partners are identified by using Tandem Affinity Purification (TAP-Tag) from a heterologous system coupled to mass spectrometry (MS) characterization, followed by validation of the role of relevant partners during skeletal muscle terminal differentiation. Aberrant forms of myogenic factors, or their aberrant regulation, are associated with a number of muscle disorders: congenital myasthenia, myotonic dystrophy, rhabdomyosarcoma and defects in muscle regeneration. As such, myogenic factors provide a pool of potential therapeutic targets in muscle disorders, both with regard to mechanisms that cause disease itself and regenerative mechanisms that can improve disease treatment. Thus, the detailed understanding of the intermolecular interactions and the genetic programs controlled by the myogenic factors is essential for the rational design of efficient therapies.

MyoD is a myogenic transcription factor with a strong capacity to induce myogenic transdifferentiation of many fully differentiated non-muscle cell lines. The epigenetic mechanisms involved in this transdifferentiation are largely unknown. Here we describe a double-affinity purification method followed by mass spectrometry to exhaustively characterize MyoD partners.

Skeletal muscle terminal differentiation starts with the commitment of pluripotent mesodermal precursor cells to myoblasts. These cells have still the ...

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we showed that D. discoideum is remarkably resilient to protein aggregation and can be used to gain insights into the cellular protein quality control system. However, the use of D. discoideum as a model system poses several challenges to microscopy-based experimental approaches, such as the high motility of the cells and their susceptibility to photo-toxicity. The latter proves to be especially challenging when studying protein homeostasis, as the phototoxic effects can induce a cellular stress response and thus alter to behavior of the protein quality control system. 
Temperature increase is a commonly used way to induce cellular stress. Here, we describe a temperature-controllable imaging protocol, which allows observing temperature-induced perturbations in D. discoideum. Moreover, when applied at normal growth temperature, this imaging protocol can also noticeably reduce photo-toxicity, thus allowing imaging with higher intensities. This can be particularly useful when imaging proteins with very low expression levels. Moreover, the high mobility of the cells often requires the acquisition of multiple fields of view to follow individual cells, and the number of fields needs to be balanced against the desired time interval and exposure time.

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging

The social amoebae Dictyostelium discoideum has recently been established as a system to study protein misfolding and proteostasis. Here, we describe a new imaging-based methodology to study temperature-induced protein aggregation and the cellular stress response in D. discoideum.

The complex lifestyle of the social amoebae Dictyostelium discoideum makes it a valuable model for the study of various biological processes. Recently, we ...

Live Cell Imaging to Assess the Dynamics of Metaphase Timing and Cell Fate Following Mitotic Spindle Perturbations

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Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

Assembly, Loading, and Alignment of an Analytical Ultracentrifuge Sample Cell

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development

4D Imaging of Protein Aggregation in Live Cells

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy

Identification of MyoD Interactome Using Tandem Affinity Purification Coupled to Mass Spectrometry

Studying the Protein Quality Control System of D. discoideum Using Temperature-controlled Live Cell Imaging