This work was funded by the National Institutes of Health (R01 GM108756). We are grateful to Gary Karpen (University of California, Berkeley) for generously providing us with a GFP:CID S2 cell line stock from which our GFP:CID/mCherry:&#945;Tubulin line was generated10.

1. Preparation of S2 Cells for Treatment with RNAi
<ol>
	<li>From a stock culture of confluent pMT:GFP:CID and pMT:mCherry:&#945;-tubulin stably transfected S2 cells (~80% viable; grown at 24-28 &#730;C), seed cells in a 6 well sterile culture plate at a density of 1 x 106 cells/mL in fresh, warmed, 10% fetal bovine serum (FBS) supplemented Schneider&#8217;s insect media (SIM). 
	NOTE: Stably transfected S2 cells can be generated by following a protocol from the Drosophila RNAi Screening Center (DRSC) (https://dgrc.bio.indiana.edu/Protocols?tab=cells). Although the specific stable S2 line used herein utilizes GFP and mCherry, numerous alternatives exist both within these fluorescent spectra as well as others. A detailed discussion of these fluorescent proteins is beyond the scope of this protocol, but their properties and potential advantages/disadvantages have been expertly reviewed elsewhere 11.
	<ol>
		<li>Using a cell counter or hemocytometer, determine the density of confluent cell stock.</li>
		<li>Determine the volume of confluent cell suspension necessary to achieve 1 x 106 cells/mL in a total volume of 4 mL (e.g., ~4 x 106 total cells). Add this volume of cell stock to a volume of fresh SIM to make 4 mL/well total volume. Determine the density of the cell stock (cells/mL) by diluting 100 &#956;L of cells directly from the stock flask with 100 &#956;L of media and counting this 1:2 dilution either manually with a hemocytometer or with an automatic cell counter if available. 
		NOTE: If using S2 cells not stably transfected, a transient transfection assay with fluorescently tagged (GFP, mCherry, etc.) &#945;- or &#946;-tubulin, and a DNA marker (CENP-A, Histone H2B, etc.) can be performed. Further, stably transfected cell lines with various mitotic spindle and DNA markers are available from the Drosophila Genetics Resource Center that may better suit the exact experimental needs of the user (https://dgrc.bio.indiana.edu/cells/Catalog).</li>
		<li>Place freshly seeded cells into 24-28 &#730;C incubator for 36-48 h.</li>
	</ol>
	</li>
</ol>
2. Treatment of Seeded Cells with dsRNA Against Gene(s) of Interest
NOTE: The following example and Results section will use Shortstop (Shot), an actin-microtubule crosslinking agent as the gene of interest. Use a mock treatment with no dsRNA or with dsRNA directed against an irrelevant gene (e.g., beta-galactosidase, lacZ) as a negative control.
<ol>
	<li>Warm Schneider&#8217;s insect media not supplemented with FBS (Serum free media; SFM) in 24-28 &#730;C incubator.</li>
	<li>Transfer cells from the previously seeded well (from step 1.1.2) into a 15 mL sterile tube, noting the total volume of cells transferred.
	<ol>
		<li>Retain a small volume of cells (~100 &#181;L) for counting in a cell counter or hemocytometer.</li>
	</ol>
	</li>
	<li>Gently pellet cells by centrifuging at 1,000 x g for 3 min at room temperature.
	<ol>
		<li>While centrifuging cells, determine the concentration of cells per mL using a cell counter or hemocytometer. Multiply this number by the total volume being centrifuged (noted in 1.2.2.) to obtain the total number of cells.</li>
	</ol>
	</li>
	<li>Aspirate the supernatant from the pelleted cells.</li>
	<li>Re-suspend the cells in warmed SFM to obtain a concentration of 3 x 106 cells/mL.
	<ol>
		<li>Transfer 1 mL of the resuspended cells to a new well in a 6 well plate.</li>
	</ol>
	</li>
	<li>Add 10-50 &#181;g of dsRNA (diluted in 100 &#956;L of RNAase-free water) against Shot (or the target gene of interest) directly to the cells and swirl the plate to mix. Incubate the plate for 1 h in the 24-28 &#730;C incubator to allow for direct dsRNA uptake into cells.</li>
	<li>During the 1 h incubation, warm the 10% FBS supplemented SIM in the 24-28 &#730;C incubator.</li>
	<li>After incubating cells with dsRNA for 1 h, add 2 mL of 10 % FBS supplemented SIM directly to the well without removing the 1 mL of media.</li>
	<li>Place cells into 24-28 &#730;C incubator for 3-7 days. 
	NOTE: As with dsRNA concentration, total treatment time may need to be optimized for the desired target gene. To evaluate RNAi efficacy, perform a western blot on the whole cell lysates using an antibody against the target gene. If the initial dsRNA target sequence proves ineffective, designing alternative dsRNAs targeted to unique sequences within the target gene.</li>
</ol>
3. Induction of Fluorescent Protein Expression and Preparation of Cells for Imaging
<ol>
	<li>If stably transfected cells were generated using the pMT plasmid (containing an inducible metallothionein promotor) as described here, induce expression of fluorescent proteins by treating cells with copper sulfate (CuSO4) at a final concentration of 500 &#181;M for 24-36 h prior to imaging and 4 days after RNAi treatment. Use of constitutive expression plasmids such as pAct does not require copper induction.</li>
	<li>Prepare Cells for Imaging
	<ol>
		<li>Warm the 10% FBS supplemented SIM in the 24-28 &#730;C incubator for 1 h.</li>
		<li>Transfer cells into a 15 mL sterile tube, noting the total volume of cells transferred.
		<ol>
			<li>Retain a small volume of cells (~100 &#181;L) for counting in a cell counter or hemocytometer.</li>
		</ol>
		</li>
		<li>Gently pellet cells by centrifuging at 1,000 x g for 3 min at room temperature.
		<ol>
			<li>While centrifuging cells, determine the concentration (cells/mL) using a cell counter or hemocytometer. Multiply this number by the total volume being centrifuged (noted in 2.2.2.) to obtain the total number of cells.</li>
		</ol>
		</li>
		<li>Aspirate the supernatant from the pelleted cells.</li>
		<li>Resuspend the cells in fresh, warmed 10% FBS supplemented SIM to obtain a concentration of 2 x 106 cells/mL. Add an appropriate amount of CuSO4 to maintain 500 &#181;M concentration.</li>
		<li>Transfer 200-500 &#181;L of re-suspended cells to one well of a multi-well live cell chamber and place on the inverted fluorescent microscope. Allow the cells to settle in the chamber for 15-30 min prior to imaging experiments. 
		NOTE: Live cell chamber wells can be pre-coated with poly-L-lysine for additional adherence.</li>
	</ol>
	</li>
</ol>
4. Set Up a Live Cell Imaging Program
NOTE: Live Cell Imaging in this experiment was performed using an inverted imaging system and its associated software (e.g., Olympus IX83 with the cellSens Dimensions software package). Details will differ by microscope manufacturer and software package; thus, general guidelines and operations are listed below.
<ol>
	<li>Using the software that runs the inverted fluorescent microscope, prepare a program for imaging a cell (or cells) over time.
	<ol>
		<li>Open the software by double-clicking on the software desktop icon.</li>
		<li>Create a new experimental file by clicking File, then New experimental file.</li>
		<li>First, insert a time lapse loop over which images will be taken. Do this by clicking the Stopwatch icon (Time-lapse Loop icon) from the icon bar. Set the interval in s in the Experiment Manager tab, and then set the number of cycles in the same tab by dividing the desired overall experiment length (in s) by the interval. Allow the loop to repeat over the desired total time period. 
		NOTE: Typically, images are taken every 30-60 s over a period of 3-4 h.</li>
		<li>Within the time lapse loop layer, insert an infrared focus check to maintain the focus of the objective (if available). To do this, add a Z-drift Compensation (ZDC) step within the Time-lapse Loop layer by clicking on the Square with Two Arrows icon (Move XY icon) and selecting Z-Drift Compensation from the dropdown menu. 
		NOTE: The term infrared focus check refers to a system by which an infrared pulse is used to maintain a constant distance between the objective and the slide/imaging chamber. Many microscopes have such a system, each with their own proprietary nomenclature. Readers should consult their operation manual or representative for specific naming details.</li>
		<li>After the infrared focus check, add a step in the program to take a multichannel image (e.g., FITC for GFP and TRITC for mCherry) over a 3-5 z-stacks by first inserting a z stack step, then specifying the channel. Do this by adding a Multichannel Group layer within the Time-lapse loop layer by clicking on the Color Wheel icon (Multichannel Group icon). Next, add a Z-Stack Loop layer within the Multichannel Group layer by clicking on the 3-layered icon (Z-stack loop icon). Set the desired Step Size and Number of Slices in the Experiment Manager tab and set the exposure of each channel as low as possible to minimize photobleaching. 
		NOTE: The z-stack interval, exposure time, and percent transmittance for LEDs can vary. Typical in these experiments, use three z-stacks taken over a range of 3 &#181;m, giving an interval of 1 &#181;m between each stack. Defining the center of the cell rather than the top and bottom tends to lead to the best results. Images are then collected for each channel(s) at an exposure of 50 ms and percent transmittance of 50% with no neutral density (ND) filter engaged.</li>
	</ol>
	</li>
</ol>
5. Image Dividing Cells Using Live Cell Imaging Program
NOTE: S2 cells do not require CO2 and grow optimally at 23-27 &#176;C. All imaging was performed at ambient temperature in a well-controlled room.
<ol>
	<li>Using the oculars, find the top (or bottom) corner of the well and focus the objective (40-60x oil immersion) onto the cells using the mCherry (&#945;-tubulin) channel.</li>
	<li>Scan along the top (or bottom) of the well to move away from the vertical well divider. 
	NOTE: Imaging too close to the well dividers can interfere with infrared focus checks.</li>
	<li>Find a cell (or cells) that are in late G2 or early M-phase (prophase). 
	NOTE: These cells can be best identified by the existence of exactly two &#8216;star-like&#8217; microtubule structures (centrosomes) and an intact nucleus (indicated by a circular region of refracted light within the cell), both easily distinguished using the &#945;-tubulin (red) excitation filter. Selection of cells in earlier interphase, notable by one or zero easily visible centrosomes, can result in wasted time due to a lag in G2/M progression. Conversely, selection of cells after NEBD will prevent accurate calculation of mitotic timing and imaging of early spindle assembly and chromosome dynamics. Also, S2 cells often contain &#62;2 centrosomes. Although these typically clusters into a bipolar spindle12, it is advised that users avoiding these cells unless otherwise desired for the experimental design. Images and discussion of appropriate cells to select can be found in the Representative Results section.</li>
	<li>Click the Live view button to begin viewing cells in the software screen. Using the fine focus knob of the microscope, focus the cell (or cells) of interest. Set the infrared focus check by clicking the Set Focus button.</li>
	<li>Initiate the time lapse imaging program by clicking the Start button. Adjust the histograms as needed by selecting the desired channel and adjusting the mean pixel intensities to see the cell (or cells) of interest clearly.</li>
	<li>Allow the program to run, checking the cell after 15-20 min to ensure nuclear envelope breakdown (NEBD) has occurred, which is determined by the disappearance of the round dark, a refracted spot near the cell center.</li>
	<li>Continue allowing the program to run, checking intermittently (every 15-20 min) to determine once anaphase onset has occurred. Stop the program at this point if there is more time remaining in the total time period and save the file. Alternatively, if events during telophase and cytokinesis are of interest, allow the program to run a sufficient time in order to image these processes as well.</li>
	<li>Perform analysis of NEBD to anaphase onset timing by noting the time of NEBD (t&#173;NEBD) in min and the time of initial chromosome separation (tanaphase onset) in min and subtracting t&#173;NEBD from tanaphase onset to obtain the NEBD to anaphase onset time for a given cell.
	<ol>
		<li>Do this by clicking the frame up button and noting the frames where NEBD and anaphase onset occur, subtracting these to determine the total number of elapse frames, and multiplying this by the time interval between imaging frames.</li>
	</ol>
	</li>
	<li>Continue scanning for pre-dividing cells to obtain multiple n&#8217;s for a given condition. Cells can be imaged in the live cell chamber well for up to 12 h after their initial settling.</li>
</ol>

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The authors have nothing to declare.

Identifying an Appropriate Cell
Key to imaging dividing S2 cells is first locating the proper cell. Time can be wasted imaging cells that are mistakenly thought to be ready to divide but fail to do so in a reasonable timeframe. Cells must be identified that have two distinct and visible centrosomes and an intact nucleus. Centrosomes must have microtubule fibers emanating from them, giving them a &#8220;star-like&#8221; appearance. An intact nucleus refracts light when adjusting focus, and also makes the approximate center of the cell darker in appearance. The nucleus will also contain the GFP:CID &#8220;dots&#8221;, another distinguishing feature to help in its identification. Cells with &#62;2 centrosomes, tubulin puncta without tubulin fibers emanating from them, or cells with only one centrosome should be avoided. Additionally, if two centrosomes are visible, but a nucleus is not, NEBD has already occurred and the cell is too far advanced in its division to be imaged if a complete M-phase analysis is desired. Once an appropriate cell is found, speed in starting imaging is crucial as NEBD occurs quickly (typically within 3-5 min) and delaying the start of imaging can lead to missed opportunities. Focus the cell in the imaging software quickly such that both centrosomes are visible, or if centrosomes are out of plane with each other, set the focal point between the two, such that each can be captured when Z stacks are collected above and below the defined center point. Once focused, immediately set the offset between the stage and the live cell chamber surface using the infrared focus check (if available) and begin the imaging program. Although the protocol presented here is specifically tailored for experiments assessing NEBD-to-anaphase dynamics, simple modifications could suit readers studying alternative mitotic events. For NEBD-to-metaphase and anaphase-to-telophase imaging, we suggest 10 s image capture intervals as these processes are highly dynamic and occur quickly in S2 cells (5-10 min). Metaphase-to-anaphase transition (our typical experimental focus) is a longer process (20-30 min) in S2 cells, and we collect images at 30 s intervals. Lastly, telophase-through-cytokinesis occurs quite slowly in S2 cells, and we suggest 60 s intervals providing enough resolution for this approximate hour-long process.
Maintaining Focus Throughout the Imaging Program
If an infrared focus check device is not available, be sure to avoid bumping the microscope or table it sits upon, as this can result in the cell moving out of focus during the imaging program, leading to ambiguous, un-interpretable results. Pre-coating the live cell chamber wells with Poly-L-lysine can help in cell adherence, which can help avoid cell movement and is especially useful for setups without infrared focus checks. Additionally, setups including shock-absorbing platforms or air tables can further help avoid cells moving out of focus. Lastly, some software programs have pause functions allowing the user to refocus and resume the program.
Imaging Multiple Cells
Helpful to accumulation of data is that often multiple cells ready to divide can be placed within the same field of view for imaging. This can be especially useful for experiments in which cells have long M-phase durations or prolonged arrest (e.g., conditions that induce activation of the SAC). For these cells, we typically image until the cells are photobleached (approximately 2-3 h), but the full timing of arrested cells should be at the discretion of the researcher. Sometimes the multiple pre-dividing cells can be in different focal planes, in which case it may be useful to add z-stacks to accommodate all cells. Adding more z stacks can also help improve resolution. Caution should be taken in doing this, however, as it will expose cells to more radiation and lead to quicker photobleaching, as well as lead to large file sizes on hard disk storage devices.
Avoiding Photobleaching and Phototoxicity
Our method describes specific settings for exposure times, imaging intervals, z stacks, and percent transmittance for our LED light source that generally work for our experimental setup and desired outcomes. As microscopes and experimental goals may vary, what might work well for our system may lead to premature photobleaching or phototoxicity in others. Photodamage may present a lack of spindle movement, microtubule fragmentation, defective microtubule dynamics, and prolonged spindle checkpoint activation. One potential method to avoid such pitfalls is to limit the exposure time. Most modern cameras now possess wide dynamic ranges, and histograms can be adjusted to visualize structures even at very low exposures. Another technique is to increase the imaging interval (e.g., to several minutes between images) such that the number of times a cell is exposed to light over a total collection interval is reduced. This is advantageous particularly in imaging for longer periods of time (many hours), and can additionally help in decreasing the file size of such experiments. Limiting the number of z stacks taken can also help reduce light exposure, using 2 or even just 1 instead of 3. Further, adjusting the percent transmittance of light (for LED light sources), or utilizing neutral density (ND) filters (for both Halogen Lamp and LED light sources) will decrease the light intensity and coupled with longer exposure times can preserve visibility. By choosing cells with centrosomes already separated, overall exposure can be limited in that these cells usually enter mitosis quickly (within a minute or two) after beginning imaging. One can also limit the time that cells are allowed to be imaged before NEBD. Our lab typically sets this time at 10 min, but a more conservative limit can easily be imposed by the user. Additionally, a more &#8216;invasive&#8217; measure we recommend is treatment with dsRNA targeted against a component of the SAC (such as Rod or Mad2). If an arrest phenotype is due to non-specific cell damage (e.g., defective microtubule dynamics), such a treatment is less likely to suppress the arrest compared to a bona fide effect of the gene of interest in the original experiment.
Future Directions
The method described here can be utilized on a relatively simple epifluorescent microscope to rapidly image live cell divisions and can be easily adapted to suit a researcher&#8217;s particular experimental design and goals. Several excellent methods for culturing, RNAi knockdown approaches, transient transfection, and fluorescence microscopy have been published for S2 and other Drosophila cells9,14,15,16,17. Our protocol offers a couple of advantages. (1) The use of a double stable cell line (GFP:CID, mCherry:&#945;-Tubulin) simultaneously marks two key mitotic structures, avoids the hassle and potential complications of transient transfection, and ensures that nearly all cells will express fluorescent markers as potential candidates for imaging. (2) The adaptation to mitosis adds to published protocols examining cytoskeletal dynamics in motile, non-dividing cells and extends to examining mitotic events in real time. While we believe ours is a powerful technique that adds to the repertoire of others, there can always be improvements made. One particular area of cell division that we have found hard to image is centrosome division. This occurs prior to NEBD and it is difficult to determine when a single centrosome is ready to separate into two. Using fluorescent cell cycle markers (Cyclin A and Cyclin B) could help resolve this problem, but comes at the expense of channels that could be used to visualize cell division components. Our best strategy for attempting to visualize centrosome division is to add multipoint acquisition to the imaging program (defining different points in the XY plane to image), but this can result in large files, and can also require (depending on the number of points) increasing the interval between image collection, decreasing time resolution of the resulting movie. Another solution could be synchronization of cells at a desired cell cycle stage using drugs that target cell cycle regulators followed by subsequent washout prior to imaging, although these methods may not be reliable in S2 cells specifically.
The protocol presented here provides a foundation for future applications in live cell imaging as well. With improved imaging and software technology, truly high throughput adaptations of this protocol using higher capacity, multi-chambered plates could be possible. Such innovations would make large-scale RNAi screens, such as those already achieved in fixed preparations12,18, more tractable in a live cell format. Small molecule drug screens could also be envisioned as a means of identifying novel compounds targeting cell division processes. Improvement of fluorophores and optical filters could also lead to imaging of multiple mitotic components (not just the two we describe), enabling imaging of specific mitotic regulators and their interactions with DNA and/or the spindle. For example, generating cells with fluorescent reporters that express following the DNA damage or during apoptosis would be useful tools to identify novel genes involved in these processes. Similar approaches could be used to examine cell cycle dynamics using the expression of fluorescently tagged cyclins19.

Cell Division is a process critical to all multicellular organisms, both in their development and homeostasis1. Drosophila has long been used as a model for the study of cell division, with experiments in various tissue types and genetic conditions providing key insights into the process. While many of these insights come from fixed cell conditions, cell division is a dynamic procedure with many moving parts, making visualization of live cells integral to assessing RNAi or genetic knockout effects on numerous parts of cell division, including spindle formation, chromosome congressional and segregation, and cytokinesis.
Many protocols have been developed and utilized over the years to visualize Drosophila&#173; cell divisions in vivo. Various groups have cultivated techniques to image divisions in both larval imaginal discs and larval brains2,3,4,5,6,7,8. These techniques, while useful for imaging in specific tissues, are limited in throughput and often require generation and maintenance of genetic stocks to generate fluorescent components and alter the expression of genes of interest. Cultured S2 cells from Drosophila provide a higher throughput alternative to rapidly test the effects of various genes in cell division. Furthermore, with the ability to transfect various fluorescent proteins, S2 cells can be quickly modified to determine the effects of RNAi knockdown on numerous components of cell division. Fluorescently tagged genes of interest can also be observed in cell division, allowing for dynamic characterization of their function9.
Here we provide a detailed protocol for live imaging of mitotic S2 cells using methods we recently described10. Our method utilizes stably transfected cells with fluorescent markers for microtubules and centromeres whose expression are under control of a copper-inducible promotor (metallothionein; pMT). This methodology can be used to image spindle and chromosome dynamics in all phases of mitosis utilizing a relatively simple fluorescent microscope with basic imaging software. It can be further customized to fit individual research needs, with transient transfection and RNAi offering expanded possibilities for determining the role of candidate genes in mitosis. Because of the relative simplicity of the protocol, it can be used for small-scale loss-of-function screens to identify genes for which further study in vivo would be beneficial, allowing for more focused efforts and efficiency with subsequent genetic manipulation in flies.

<table>
	<tbody>
		<tr>
			<td>Bright-Line Hemacytometer</td>
			<td>Sigma-Aldrich</td>
			<td>Z359629-1EA</td>
			<td>for cell counting</td>
		</tr>
		<tr>
			<td>cellSens imaging software</td>
			<td>Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CELLSTAR Cell Culture Flask, 50ML, 25 CM2, PS, Red Filter Screw Cap, Clear, Sterile, 10PCS/BAG</td>
			<td>Greiner Bio-One</td>
			<td>690175</td>
			<td></td>
		</tr>
		<tr>
			<td>CELLSTAR Cell Culture Multiwell Plate, 6 well, PS, Clear, TC, Lid with condensation rings, sterile, single packed</td>
			<td>Greiner Bio-One</td>
			<td>657160</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge 5804 R</td>
			<td>eppendorf</td>
			<td>Cat. 022623508</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper(II) sulfate pentahydrate, minimum 98%</td>
			<td>Sigma-Aldrich</td>
			<td>C3036-250G</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Fetal Bovine Serum</td>
			<td>Fisher Scientific</td>
			<td>MT35015CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Effectene Transfection Reagent 1 mL</td>
			<td>Qiagen</td>
			<td>301425</td>
			<td>for transient transfection</td>
		</tr>
		<tr>
			<td>IX-83 Inverted Epifluorescent Microscope</td>
			<td>Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>LabTek Chambered Slide Insert</td>
			<td>Applied Scientific Instruments</td>
			<td>I-3016</td>
			<td></td>
		</tr>
		<tr>
			<td>MEGAscript T7 Transcription Kit</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM1334</td>
			<td>For dsRNA production</td>
		</tr>
		<tr>
			<td>MOXI Z Mini Automated Cell Counter Kit</td>
			<td>ORFLO</td>
			<td>MXZ001</td>
			<td>for cell counting</td>
		</tr>
		<tr>
			<td>MS-2000 XY Flat-Top Automated Stage and Controller</td>
			<td>Applied Scientific Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nunc Lab-Tek II Chambered Coverglass (no 1.5 borosilicate glass) 8-well</td>
			<td>Thermo Fisher Scientific</td>
			<td>155409</td>
			<td></td>
		</tr>
		<tr>
			<td>Orca-Flash 4.0 LT Camera</td>
			<td>Hammamatsu Photonics K.K.</td>
			<td>C11440-42U</td>
			<td></td>
		</tr>
		<tr>
			<td>pMT/V5-His A Drosophila Expression Vector</td>
			<td>Thermo Fisher Scientific</td>
			<td>V412020</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-Lysine</td>
			<td>Cultrex</td>
			<td>3438-100-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Purifier Logic+ Class II, Type A2 Biosafety Cabinets</td>
			<td>Labconco</td>
			<td>302310000</td>
			<td></td>
		</tr>
		<tr>
			<td>Schneider&#39;s insect medium</td>
			<td>Sigma-Aldrich</td>
			<td>S0146-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Spectra Tub Centrifuge Tubes</td>
			<td>VWR</td>
			<td>470224-998</td>
			<td></td>
		</tr>
		<tr>
			<td>Uner Counter BOD Incubator</td>
			<td>Sheldon manufacturing (VWR)</td>
			<td>89409-346</td>
			<td></td>
		</tr>
		<tr>
			<td>X-CITE 120 LED</td>
			<td>ExcelitasTechnologies</td>
			<td></td>
			<td>Led light source</td>
		</tr>
		<tr>
			<td>Z -Drift Compensator (ZDC)</td>
			<td>Olympus</td>
			<td></td>
			<td>infrared focus check</td>
		</tr>
	</tbody>
</table>

use of drosophila s2 cells for live imaging of cell division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a rapid and high-throughput manner. S2 cells have proven amenable to both fixed- and live-cell imaging applications. Notably, live-cell imaging can yield valuable information about how loss or knockdown of a gene can affect the kinetics and dynamics of key events during cell division, including mitotic spindle assembly, chromosome congression, and segregation, as well as overall cell cycle timing. Here we utilize S2 cells stably transfected with fluorescently tagged mCherry:&#945;-tubulin to mark the mitotic spindle and GFP:CENP-A (referred to as &#8216;CID&#8217; gene in Drosophila) to mark the centromere to analyze the effects of key mitotic genes on the timing of cell divisions, from prophase (specifically at Nuclear Envelope Breakdown; NEBD) to the onset of anaphase. This imaging protocol also allows for the visualization of the spindle microtubule and chromosome dynamics throughout mitosis. Herein, we aim to provide a simple yet comprehensive protocol that will allow readers to easily adapt S2 cells for live imaging experiments. Results obtained from such experiments should expand our understanding of genes involved in the cell division by defining their role in several simultaneous and dynamic events. Observations made in this cell culture system can be validated and further investigated in vivo using the impressive toolkit of genetic approaches in flies.

Methods we describe above will result in the identification and imaging of Drosophila S2 cells undergoing cell division. Cells that are about to divide (i.e., just prior to entering M-phase) can be targeted by the presence of two centrosomes and an intact nucleus indicated by refracted light and a darker spot within the cell when viewed in the &#945;-tubulin channel (Figure 1, left-most panels, red channel; arrowhead in Figure 1A). We estimate that approximately 2-5% of cells fall into this category, and between imaging experiments, we typically spend a maximum of 2-3 minutes scanning for another qualified cell. NEBD can be visualized through the disappearance of this dark spot, resulting in the uniform coloring of the cytoplasm (Figure 1, panels marked &#8220;00:00&#8221;, red). After NEBD, the time each cell takes to form the spindle, congress chromosomes, and segregate chromosomes can be measured simply by noting the time points of these events relative to NEBD.
We utilize these divisions to assess mitotic timing, specifically of NEBD to anaphase onset, noting the point at which chromosomes begin to separate. A majority (~90%) of copper-induced cells, expressed both mCherry:&#945;-tubulin and GFP:CID. A small percentage of cells (~10%) expressed only one marker or neither. These cells were avoided during cell selection. Typically, S2 cells display NEBD to anaphase onset timing of between 20-30 minutes (Figure 1A, Control). We next treated cells with dsRNA targeted against Shortstop (Shot), an actin-microtubule crosslinking protein we suspected may affect cell cycle dynamics. Indeed, following Shot knockdown cells exhibited a significant mitotic delay (Figure 1B, ShotRNAi delay), with many cells arresting at metaphase and never transitioning to anaphase during the entire 2-3-hour imaging experiment (Figure 1D, ShotRNAi arrest). We reasoned this delay/arrest phenotype could likely be due to activation of the spindle assembly checkpoint (i.e. the M-phase checkpoint). To directly test this hypothesis, we co-treated cells with dsRNA against Shot and Rough deal (Rod), an important component of this checkpoint13. This led to a suppression of the arrest phenotype, resulting in NEBD to anaphase times similar to Control (Figure 1C, ShotRNAi+RodRNAi). Thus, this live imaging protocol allowed us to conclude that Shot is required for timely M-phase progression and that its loss leads to checkpoint activation that delays or arrests mitotic cells.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60049/60049fig01.jpg" /> 
Figure 1: Live imaging reveals cell cycle defects due to mitotic checkpoint activation in S2 cells. 
In all conditions, live-cell imaging was conducted on S2 cells stably co-transfected with inducible GFP:CID and mCherry:&#945;Tubulin as described herein. (A) Cartoons illustrate important landmarks during mitosis, and corresponding images are shown from representative movies of indicated genotypes with time points relative to NEBD (t=00:00) indicated. Control cells typically progress to anaphase within 20-30 min. ShotRNAi-treated cells display NEBD-anaphase delay (B) and frequently undergo metaphase arrest, never entering anaphase within the imaging experiment (D). Co-treatment of cells with ShotRNAi and RodRNAi, a component of the spindle assembly checkpoint, suppresses the Shot phenotype leading to anaphase onset kinetics similar to Control cells (C). The arrows in (A) indicate the &#8216;star-like&#8217; centrosome structures, and the large arrowhead indicates the nucleus. Each scale bar represents 5 microns. This figure is adapted and republished with permission from10 (https://www.molbiolcell.org/info-for-authors). <a href="https://www.jove.com/files/ftp_upload/60049/60049fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Time-lapse movie of &#8216;Control&#8217; S2 cell division. Video shows a representative movie of S2 cell division in &#8216;Control&#8217; genotype. This movie corresponds to Figure 1A. This video is adapted and republished with permission from10 (https://www.molbiolcell.org/info-for-authors). <a href="https://cloudflare.jove.com/files/ftp_upload/60049/Dewey_2019_JoVE_Video_1_(Control).mov" target="_blank">Please click here to download this video.</a>
Video 2: Time-lapse movie of delayed &#8216;ShotRNAi&#8217; S2 cell division. Video shows a representative movie of S2 cell division in &#8216;ShotRNAi&#8217; genotype that leads to a phenotypic mitotic delay. This movie corresponds to Figure 1B. <a href="https://cloudflare.jove.com/files/ftp_upload/60049/Dewey_2019_JoVE_Video_2_(ShotDelay).mov" target="_blank">Please click here to download this video.</a>
Video 3: Time-lapse movie of &#8216;ShotRNAi+RodRNAi&#8217; S2 cell division. Video shows a representative movie of S2 cell division in &#8216;ShotRNAi+RodRNAi&#8217; genotype. This movie corresponds to Figure 1C. This video is adapted and republished with permission from10 (https://www.molbiolcell.org/info-for-authors). <a href="https://cloudflare.jove.com/files/ftp_upload/60049/Dewey_2019_JoVE_Video_3_(ShotRod).mov" target="_blank">Please click here to download this video.</a>
Video 4: Time-lapse movie of arrested &#8216;ShotRNAi&#8217; S2 cell division. Video shows a representative movie of S2 cell division in &#8216;ShotRNAi&#8217; genotype that leads to a phenotypic mitotic arrest. This movie corresponds to Figure 1D. <a href="https://cloudflare.jove.com/files/ftp_upload/60049/Dewey_2019_JoVE_Video_4_(ShotArrest).mov" target="_blank">Please click here to download this video.</a>

Watch this Scientific Journal Video about Use of Drosophila S2 Cells for Live Imaging of Cell Division at JoVE.com

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and quantified.

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Drosophila S2 cells are an important tool in studying mitosis in tissue culture, providing molecular insights into this fundamental cellular process in a ...

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8.1K Views.  University of New Mexico. Cell divisions can be visualized in real time using fluorescently tagged proteins and time-lapse microscopy. Using the protocol presented here, users can analyze cell division timing dynamics, mitotic spindle assembly, and chromosome congression and segregation. Defects in these events following RNA interference (RNAi)-mediated gene knockdown can be assessed and quantified.

Video: Use of Drosophila S2 Cells for Live Imaging of Cell Division

Immunostaining for DNA Modifications: Computational Analysis of Confocal Images

For several decades, 5-methylcytosine (5mC) has been thought to be the only DNA modification with a functional significance in metazoans. The discovery of enzymatic oxidation of 5mC to 5-hydroxymethylcytosine (5hmC), 5-formylcytosine (5fC) and 5-carboxylcytosine (5caC) as well as detection of N6-methyladenine (6mA) in the DNA of multicellular organisms provided additional degrees of complexity to the epigenetic research. According to a growing body of experimental evidence, these novel DNA modifications may play specific roles in different cellular and developmental processes. Importantly, as some of these marks (e. g. 5hmC, 5fC and 5caC) exhibit tissue- and developmental stage-specific occurrence in vertebrates, immunochemistry represents an important tool allowing assessment of spatial distribution of DNA modifications in different biological contexts. Here the methods for computational analysis of DNA modifications visualized by immunostaining followed by confocal microscopy are described. Specifically, the generation of 2.5 dimension (2.5D) signal intensity plots, signal intensity profiles, quantification of staining intensity in multiple cells and determination of signal colocalization coefficients are shown. Collectively, these techniques may be operational in evaluating the levels and localization of these DNA modifications in the nucleus, contributing to elucidating their biological roles in metazoans.

Newly discovered oxidized forms of 5-methylcytosine (oxi-mCs), 5-hydroxymethylcytosine (5hmC), 5-formylcytosine (5fC) and 5-carboxylcytosine (5caC) may represent distinct DNA modifications with unique functional roles. Here a semi-quantitative workflow for visualization of oxi-mCs' spatial distribution, signal intensity profiling and colocalization is described.

For several decades, 5-methylcytosine (5mC) has been thought to be the only DNA modification with a functional significance in metazoans. The discovery of ...

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

Purification of Low-abundant Cells in the Drosophila Visual System

Recent improvements in the sensitivity of next generation sequencing have facilitated the application of transcriptomic and genomic analyses to small numbers of cells. Utilizing this technology to study development in the Drosophila visual system, which boasts a wealth of cell type-specific genetic tools, provides a powerful approach for addressing the molecular basis of development with precise cellular resolution. For such an approach to be feasible, it is crucial to have the capacity to reliably and efficiently purify cells present at low abundance within the brain. Here, we present a method that allows efficient purification of single cell clones in genetic mosaic experiments. With this protocol, we consistently achieve a high cellular yield after purification using fluorescence activated cell sorting (FACS) (~25% of all labeled cells), and successfully performed transcriptomics analyses on single cell clones generated through mosaic analysis with a repressible cell marker (MARCM). This protocol is ideal for applying transcriptomic and genomic analyses to specific cell types in the visual system, across different stages of development and in the context of different genetic manipulations.

Here, we present a cell dissociation protocol for efficiently isolating cells present at low abundance within the Drosophila visual system through fluorescence activated cell sorting (FACS).

Recent improvements in the sensitivity of next generation sequencing have facilitated the application of transcriptomic and genomic analyses to small ...

Genetic Engineering of Dictyostelium discoideum Cells Based on Selection and Growth on Bacteria

Dictyostelium discoideum is an intriguing model organism for the study of cell differentiation processes during development, cell signaling, and other important cellular biology questions. The technologies available to genetically manipulate Dictyostelium cells are well-developed. Transfections can be performed using different selectable markers and marker re-cycling, including homologous recombination and insertional mutagenesis. This is supported by a well-annotated genome. However, these approaches are optimized for axenic cell lines growing in liquid cultures and are difficult to apply to non-axenic wild-type cells, which feed only on bacteria. The mutations that are present in axenic strains disturb Ras signaling, causing excessive macropinocytosis required for feeding, and impair cell migration, which confounds the interpretation of signal transduction and chemotaxis experiments in those strains. Earlier attempts to genetically manipulate non-axenic cells have lacked efficiency and required complex experimental procedures. We have developed a simple transfection protocol that, for the first time, overcomes these limitations. Those series of large improvements to Dictyostelium molecular genetics allow wild-type cells to be manipulated as easily as standard laboratory strains. In addition to the advantages for studying uncorrupted signaling and motility processes, mutants that disrupt macropinocytosis-based growth can now be readily isolated. Furthermore, the entire transfection workflow is greatly accelerated, with recombinant cells that can be generated in days rather than weeks. Another advantage is that molecular genetics can further be performed with freshly isolated wild-type Dictyostelium samples from the environment. This can help to extend the scope of approaches used in these research areas.

Genetic Engineering of Dictyostelium discoideum Cells Based on Selection and Growth on Bacteria

Dictyostelium discoideum is a popular model organism to study complex cellular processes such as cell migration, endocytosis, and development. The utility of the organism is dependent on the feasibility of genetic manipulation. Here, we present methods to transfect Dictyostelium discoideum cells that overcome existing limitations of culturing cells in liquid media.

Dictyostelium discoideum is an intriguing model organism for the study of cell differentiation processes during development, cell signaling, and other ...

The Use of Mouse Mammary Tumor Cells in an In Vitro Invasion Assay as a Measure of Oncogenic Cell Behavior

The in vitro invasion assay uses a protein-rich matrix in a Boyden chamber to measure the ability of cultured cells to pass through the matrix and a porous membrane in a process analogous to the initial steps of cancer cell metastasis. The tested cells can be altered for the gene expression or treated with inhibitors to test for changes in the invasion potential. This experiment tests the aggressive phenotype of the mouse mammary tumor cells to discover and characterize the potential oncogenes that promote cell invasion. This technique, however, can be versatile and adapted to many different applications. The experiment itself can be done in one day and the results are acquired by light microscopy in less than a day. The results include counts of the number of invading cells for comparison and analysis. The in vitro invasion assay is a rapid, inexpensive, and clear-cut method for determining cell behavior in a culture that can be used as an initial assessment before more involved in vivo assays.

The in vitro cell invasion assay is used to measure the potential of cancer metastasis by quantifying the cellular potential for invasion and migration using cell culture inserts containing protein matrix. Cells are challenged to migrate through the protein matrix and a porous membrane, towards a chemoattractant, and then quantified by light microscopy.

The in vitro invasion assay uses a protein-rich matrix in a Boyden chamber to measure the ability of cultured cells to pass through the matrix and a ...

Use of Frozen Tissue in the Comet Assay for the Evaluation of DNA Damage

The comet assay is gaining popularity as a means to assess DNA damage in cultured cells and tissues, particularly following exposure to chemicals or other environmental stressors. Use of the comet assay in regulatory testing for genotoxic potential in rodents has been driven by adoption of an Organisation for Economic Co-operation and Development (OECD) test guideline in 2014. Comet assay slides are typically prepared from fresh tissue at the time of necropsy; however, freezing tissue samples can avoid logistical challenges associated with simultaneous preparation of slides from multiple organs per animal and from many animals per study. Freezing also enables shipping samples from the exposure facility to a different laboratory for analysis, and storage of frozen tissue facilitates deferring a decision to generate DNA damage data for a given organ. The alkaline comet assay is useful for detecting exposure-related DNA double- and single-strand breaks, alkali-labile lesions, and strand breaks associated with incomplete DNA excision repair. However, DNA damage can also result from mechanical shearing or improper sample processing procedures, confounding the results of the assay. Reproducibility in collection and processing of tissue samples during necropsies may be difficult to control due to fluctuating laboratory personnel with varying levels of experience in harvesting tissues for the comet assay. Enhancing consistency through refresher training or deployment of mobile units staffed with experienced laboratory personnel is costly and may not always be feasible. To optimize consistent generation of high quality samples for comet assay analysis, a method for homogenizing flash frozen cubes of tissue using a customized tissue mincing device was evaluated. Samples prepared for the comet assay by this method compared favorably in quality to fresh and frozen tissue samples prepared by mincing during necropsy. Moreover, low baseline DNA damage was measured in cells from frozen cubes of tissue following prolonged storage.

This protocol describes several procedures for preparing high quality frozen tissue samples at the time of necropsy for use in the comet assay to assess DNA damage: 1) minced tissue, 2) scraped epithelial cells from the gastrointestinal tract, and 3) cubed tissue samples, requiring homogenization using a tissue mincing device.

The comet assay is gaining popularity as a means to assess DNA damage in cultured cells and tissues, particularly following exposure to chemicals or other ...

Use of the Pyrimidine Analog, 5-Iodo-2&prime;-Deoxyuridine (IdU) with Cell Cycle Markers to Establish Cell Cycle Phases in a Mass Cytometry Platform

The regulation of cell cycle phase is an important aspect of cellular proliferation and homeostasis. Disruption of the regulatory mechanisms governing the cell cycle is a feature of a number of diseases, including cancer. Study of the cell cycle necessitates the ability to define the number of cells in each portion of cell cycle progression as well as to clearly delineate between each cell cycle phase. The advent of mass cytometry (MCM) provides tremendous potential for high throughput single cell analysis through direct measurements of elemental isotopes, and the development of a method to measure the cell cycle state by MCM further extends the utility of MCM. Here we describe a method that directly measures 5-iodo-2&#8242;-deoxyuridine (IdU), similar to 5-bromo-2&#180;-deoxyuridine (BrdU), in an MCM system. Use of this IdU-based MCM provides several advantages. First, IdU is rapidly incorporated into DNA during its synthesis, allowing reliable measurement of cells in the S-phase with incubations as short as 10-15 minutes. Second, IdU is measured without the need for secondary antibodies or the need for DNA degradation. Third, IdU staining can be easily combined with measurement of cyclin B1, phosphorylated retinoblastoma protein (pRb), and phosphorylated histone H3 (pHH3), which collectively provides clear delineation of the five cell cycle phases. Combination of these cell cycle markers with the high number of parameters possible with MCM allow combination with numerous other metrics.

Use of the Pyrimidine Analog, 5-Iodo-2&#8242;-Deoxyuridine IdU with Cell Cycle Markers to Establish Cell Cycle Phases in a Mass Cytometry Platform

This protocol adapts cell cycle measurements for use in a mass cytometry platform. With the multi-parameter capabilities of mass cytometry, direct measurement of iodine incorporation allows identification of cells in S-phase while intracellular cycling markers enable characterization of each cell cycle state in a range of experimental conditions.

The regulation of cell cycle phase is an important aspect of cellular proliferation and homeostasis. Disruption of the regulatory mechanisms governing the ...

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. The CRISPR/Cas9 system allows researchers to modify the genome with unprecedented efficiency, fidelity, and simplicity. Harnessing this technology, researchers are seeking a rapid, efficient, and easy protocol for generating GM mice. Here we introduce an improved method for cryopreservation of one-cell embryos that leads to a higher developmental rate of the freeze-thawed embryos. By combining it with optimized electroporation conditions, this protocol allows for the generation of knockout and knock-in mice with high efficiency and low mosaic rates within a short time. Furthermore, we show a step-by-step explanation of our optimized protocol, covering CRISPR reagent preparation, in vitro fertilization, cryopreservation and thawing of one-cell embryos, electroporation of CRISPR reagents, mouse generation, and genotyping of the founders. Using this protocol, researchers should be able to prepare GM mice with unparalleled ease, speed, and efficiency.

Here, we present a modified method for cryopreservation of one-cell embryos as well as a protocol that couples the use of freeze-thawed embryos and electroporation for the efficient generation of genetically modified mice.

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. ...

Cryopreservation of PGCs for Long-Term Preservation of Drosophila Strains

Primordial Germ Cell Cryopreservation and Revival of Drosophila Strains

Drosophila strains must be maintained by the frequent transfer of adult flies to new vials. This carries a danger of mutational deterioration and phenotypic changes. Development of an alternative method for long-term preservation without such changes is therefore imperative. Despite previous successful attempts, cryopreservation of Drosophila embryos is still not of practical use because of low reproducibility. Here, we describe a protocol for primordial germ cell (PGC) cryopreservation and strain revival via transplantation of cryopreserved PGCs into agametic Drosophila melanogaster (D. melanogaster) host embryos. PGCs are highly permeable to cryoprotective agents (CPAs), and developmental and morphological variation among strains is less problematic than in embryo cryopreservation. In this method, PGCs are collected from approximately 30 donor embryos, loaded into a needle after CPA treatment, and then cryopreserved in liquid nitrogen. To produce donor-derived gametes, the cryopreserved PGCs in a needle are thawed and then deposited into approximately 15&#160;agametic host embryos. A frequency of at least 15% fertile flies was achieved with this protocol, and the number of progeny per fertile couple was always more than enough to revive the original strain (the average progeny number being 77.2 &#177; 7.1), indicating the ability of cryopreserved PGCs to become germline stem cells. The average number of fertile flies per needle was 1.1 &#177; 0.2, and 9 out of 26 needles produced two or more fertile progeny. It was found that 11 needles are enough to produce 6 or more progeny, in which at least one female and one male are likely included. The agametic host makes it possible to revive the strain quickly by simply crossing newly emerged female and male flies. In addition, PGCs have the potential to be used in genetic engineering applications, such as genome editing.

Author Spotlight: PGC Cryopreservation &#8211; An Alternative Approach Towards Long-Term Preservation of Drosophila Strains 

A long-term preservation method for Drosophila strains as an alternative to the frequent transfer of adult flies to fresh food vials is highly desirable. This protocol describes the cryopreservation of Drosophila primordial germ cells and strain revival via their transplantation to agametic host embryos.