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, 50 mL, 25 CM2, PS, Red Filter Screw Cap, Clear, Sterile, 10 PCS/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&#039;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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8.2K 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

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Developmental Biology

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Neuroscience

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

Biology

Imaging Live Brain Explants from Drosophila melanogaster Third Instar Larvae

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a differentiating ganglion mother cell (GMC), which will undergo one additional division to give rise to two neurons or glia. Studies in NBs have uncovered the molecular mechanisms underlying cell polarity, spindle orientation, neural stem cell self-renewal, and differentiation. These asymmetric cell divisions are readily observable via live-cell imaging, making larval NBs ideally suited for investigating the spatiotemporal dynamics of asymmetric cell division in living tissue. When properly dissected and imaged in nutrient-supplemented medium, NBs in explant brains robustly divide for 12-20 h. Previously described methods are technically difficult and may be challenging to those new to the field. Here, a protocol is described for the preparation, dissection, mounting, and imaging of live third-instar larval brain explants using fat body supplements. Potential problems are also discussed, and examples are provided for how this technique can be used.

Author Spotlight: Investigating Asymmetric Cell Division Dynamics: A Protocol for Live-Imaging of Drosophila Larval Brain Explants 

Here, we discuss a workflow to prepare, dissect, mount, and image live explant brains from Drosophila melanogaster third instar larvae to observe the cellular and subcellular dynamics under physiological conditions.

Drosophila neural stem cells (neuroblasts, NBs hereafter) undergo asymmetric divisions, regenerating the self-renewing neuroblast, while also forming a ...

Real-Time Imaging of Germline Stem Cells in Drosophila Ex Vivo Germaria Culture

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to explore the dynamics of a myriad of developmental processes, such as cell division, stemness, differentiation, migration, apoptosis, autophagy, cellular adhesion, etc., over time. Recently, we have implemented an extended ex vivo culture and live imaging of the female Drosophila GSC niche. Using a Drosophila line harboring a GFP::Par-1 transgene as an example, this method allows the visualization of the GSCs&#39; asymmetric division within their niche and the description of the changes in the spectrosome morphology along the cell cycle. Here, we present a detailed protocol for the ex vivo culture of Drosophila germaria, enabling prolonged visualization of the female GSC niche. Importantly, this protocol is broadly applicable to live imaging GSCs with multiple fluorescently tagged proteins of interest that are available in stock centers and/or in the Drosophila research community.

Here, we provide a detailed protocol to perform live imaging of the asymmetric division of germline stem cells (GSCs) in the Drosophila ovarian niche. We use a transgenic line that ubiquitously expresses a green fluorescent protein (GFP) fusion of the spectrosome protein Par-1.

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized microtubules. Organelles move along these processes by microtubule motors. Easy maintenance, high sensitivity to RNAi-mediated protein knock-down and efficient procedure for creating stable cell lines make Drosophila S2 cells an ideal model system to study cargo transport by live imaging. The results obtained with S2 cells can be further applied to a more physiologically relevant system: axonal transport in primary neurons cultured from dissociated Drosophila embryos. Cultured neurons grow long neurites filled with bundled microtubules, very similar to S2 processes. Like in S2 cells, organelles in cultured neurons can be visualized by either organelle-specific fluorescent dyes or by using fluorescent organelle markers encoded by DNA injected into early embryos or expressed in transgenic flies. Therefore, organelle transport can be easily recorded in neurons cultured on glass coverslips using living imaging. Here we describe procedures for culturing and visualizing cargo transport in Drosophila S2 cells and primary neurons. We believe that these protocols make both systems accessible for labs studying cargo transport.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Summary

Explore More Videos

Live-imaging of the Drosophila Pupal Eye

Live Imaging of Drosophila Larval Neuroblasts

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Preparation of Drosophila S2 cells for Light Microscopy

Live-Cell Imaging of Drosophila melanogaster Third Instar Larval Brains

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Imaging Through the Pupal Case of Drosophila melanogaster