We thank Aida de la Cruz for developing and teaching the original protocol on which this version is based10. Work in the Buttitta Lab is supported by NIH grant GM086517.

1. Fly Husbandry
<ol>
 <li>Cross flies of desired genotypes in narrow plastic vials with 10 ml Yeast-Glucose Medium3 or other protein-rich media of your choice. The extensive Drosophila transgenic tools available for tissue specific expression and lineage tracing with in vivo fluorescent protein expression are described in detail elsewhere4,5. Transfer parents to fresh vials daily to obtain series of 24 hr progeny collections, or for more precise staging, collect embryos on agar plates and transfer newly hatched larva to vials as described10. To ensure uniform conditions across experiments and avoid overcrowding, the total number of F1 progeny in a single vial should be no more than 100. Depending on the genetic design of the experiment, keep vials in an appropriate incubator until the desired developmental stage.</li>
 <li>Collect experimental animals: you will need mature larvae at the 3rd larval instar (L3) at about 110-120 hr of development for larval dissections or white prepupa (WPP) for accurate pupal staging. Dramatic cell cycle changes, including a switch to a non-proliferative state occur at distinct timepoints during metamorphosis12,13. Therefore correct staging of pupa (within 1 hr of development) is essential for reproducible cell cycle data during metamorphosis. WPP corresponds to 0 hr after pupa formation (0 hr APF) as shown in Figure 1, panel I. Collect WPP in a 35 mm Petri dish on a folded Kimwipe wet with 2.5 ml of water to maintain some humidity. Age pupa until desired stage (hours APF) at appropriate temperature. Note that development proceeds 1.2 times faster at 29 &deg;C and 2.2 times more slowly at 18 &deg;C than at the standard 25 &deg;C14.</li>
 <li>Heat-shock animals to activate heat-shock flipase (hs-flp)-induced recombination for lineage-tracing clones if needed9,11. For the heat-shock of larva, immerse vials in a water bath (set at 37 &deg;C) and ensure larvae are fully submerged. For the heat-shock of pupa, seal the Petri dish with Parafilm and fully submerge it into the water-bath. Be sure to remove Parafilm after the heat-shock, to allow sufficient oxygen exchange for animal survival. Duration of the heat-shock depends on activity of the flipase transgene used and number of clones desired. We routinely use 7-8 min for inducing "flipout" lineage tracing clones15 in larvae with the hs-flp transgene on the first chromosome and 2-4 min for heat shock of pupae in 35 mm dishes, while the hs-flp transgene on the second chromosome requires a 20 min. heat shock to obtain similar numbers of clones.</li>
</ol>
The rate of cell division and timing of dissection determines clone size in each tissue. Under normal conditions, we find that clones induced in the larval wing with a 20 min heat shock (using the hs-flp transgene on the second chromosome) and dissected 48 hr later, contain approximately 20-30 well separated clones with sizes ranging from 10 cells per clone to two cells per clone, with an average around four cells per clone. In contrast, a 7 min heat-shock with the same hs-flp transgene of pupae in a Petri dish at 0 hr APF dissected 36 hr later, yields well separated clones (approximately 15-25) ranging from one to four cells with an average of 2.2 cells per clone. Such data can be used to determine the average cell doubling time for the tissue under study.
2. Dissection
<ol>
 <li>Gently transfer the larvae/pupae to a clear glass dissection dish containing 1X PBS.</li>
 <li>Using a dissecting microscope and ample light, gently grasp a larva by the abdomen with a sharp Inox #5 forcep, cut the anterior third with micro-scissors, and invert the anterior third by pushing the mouth area inwards toward the posterior while holding the larval cuticle steady (Figure 1A). Carefully remove opaque fat body and any remaining gut to increase visibility of desired tissues, such as wing, eye or brain. Dissect the desired tissues; cleanly removing wings from trachea and cuticle with forceps, eyes from mouth-hooks, or brain from eyes (Figure 1G).</li>
 <li>For dissecting pupa; use a forcep to gently grasp a pupa by the anterior tip of the pupal case where there is an air bubble, to avoid damaging the pupa inside (Figure 1C). Forceps or micro-scissors in the opposing hand can be used to help position the floating pupa for proper grasping. Cleanly cut through the posterior tip of the pupa with micro-scissors. This cross-sectional cut should be made at a slight angle toward the dorsal part of the animal, to release the internal pressure without forcing the histolyzed fat into the desired tissues in the thorax and head. Cut carefully along the dorsal median, avoiding the wings and the anterior eye-brain complex. Carefully remove the pupa from the pupal case by grasping the posterior edge of the pupal epidermis and pulling it out of the pupal case (Figure 1D). Poke a small hole in the cuticle anterior to the eye-brain complex with forceps. Using a glass Pasteur pipette with a rubber bulb, gently wash PBS dissection solution through the partially dissected pupa to remove histolyzed fat and any remaining gut tissue. This also effectively lubricates the glass pipette with lipids and proteins, which reduces the incidence of dissected tissues from sticking to the interior of the glass pipette later.</li>
 <li>To obtain pupal wings, the cuticle enveloping the wing and the wing itself must be pried from the pupa (Figure 1E). This can be done by holding the pupa down by the head with one forcep while using another forcep to either grasp the most posterior tip of the wing cuticle and pulling out or maneuvering the forcep under the wing near the hinge before dislodging it from the side of the carcass (Figure 1E). The wing can then be cut or torn at the hinge to remove it from the body and put in a pile away from the carcasses and other material.</li>
 <li>To obtain pupal eyes or brain, wash the pupa until the eye-brain complex becomes dislodged and free from the pupa (Figure 1F, 1H). Using a forcep, gently pull the pupal eye away from the brain and save the desired tissue.</li>
 <li>Do not take longer than 45 min to dissect each sample, as results becomes less accurate the longer cells are left in PBS after opening the animals.</li>
</ol>
3. Tissue Dissociation and DNA Staining
<ol>
 <li>Using a lubricated Pasteur pipette, carefully transfer the dissected tissues into 0.5 ml of Live DNA Stain Solution in a 5 ml polystyrene tube (Figure 1G). Transfer as little PBS as possible (less than 300 &mu;l) into the DNA Stain Solution, as dilution of trypsin will inhibit tissue dissociation.</li>
 <li>Incubate tubes with shaking at 23 &deg;C at 500 rpm (we use an Eppendorf Thermomixer).</li>
 <li>Incubate for 70 min, then vortex gently for 5 sec at a medium speed (setting of 5). Excessive vortexing will cause cell disruption and form debris, while under-vortexing will lead to cell clumps. Return the samples to shaking at 23 &deg;C at 500 rpm for an additional 15 to 20 min before vortexing them once more for 5 sec at speed 5.</li>
 <li>For brains and pupal eyes older than 36hAPF, a modified protocol is required. Dissociate tissues in DNA Stain Solution at 23 &deg;C at 500 rpm for 45-60 min in 1.7 ml microcentrifuge tubes. Then one-by-one manually dissociate large chunks of tissue by transferring to dissecting dish along with about 50 &mu;l DNA Stain Solution and rapid pipetting up and down with a manually drawn Pasteur pipette 16 (of approximately 100-150 &mu;m opening). Transfer each dissociated specimen with the rest of the 0.5 ml of DNA Stain Solution into a 5 ml tube. Incubate an additional 45-30 min (up to 90 min of total incubation time) at 23 &deg;C with shaking at 500 rpm until clumps are no longer visible.</li>
 <li>Fully dissociated samples should have very few large visible chunks of tissue, although note that the acellular transparent pupal wing cuticle will not dissociate.</li>
</ol>
4. Flow Cytometry
<ol>
 <li>Turn on the Attune cytometer and open the Attune software. The cytometer lasers should be turned on via the startup menu, and a daily performance test should be performed with Performance Tracking Beads with adequate scores before each day of data acquisition. Fluid levels should be at least half full and waste emptied before the performance test. Details for startup, proper calibration and operation of the Attune can be found in the Attune User Guide (<a href="http://tools.invitrogen.com/content/sfs/manuals/cms_082577.pdf" target="_blank"> http://tools.invitrogen.com/content/sfs/manuals/cms_082577.pdf</a>)</li>
 <li>Design a new template by creating 4 dot plots and 2 histograms in a blank workspace with the parameters indicated in 4.3, or select a previously designed template suitable for the experiment, and indicate the number of samples to be collected. We have provided Attune templates (files in .get format) for live Drosophila cell cycle analysis with Vybrant DyeCycle Violet and GFP or RFP expression in Appendix I.</li>
 <li>The templates provided include four dot plots and two histograms with the parameters indicated in Figure 2. Channels for detection include: FSC and SSC - forward scatter and side scatter indicating cell size and membrane complexity, VL1 height (H), width (W) and area (A)- indicating Violet laser excitation of DyeCycle Violet staining the DNA, and BL1-A - Blue laser Channel 1 Area, indicating GFP fluorescence. The provided template for RFP analysis is identical to the template for GFP analysis, except that it uses the Blue laser channel 3 (BL3-A), which is optimized for RFP detection.</li>
 <li>The dot plots in the templates have gates to restrict analysis to the desired populations and to limit debris and cell clumps from analysis (Figure 2). Gate 1 excludes debris and clumps based on SSC vs. FSC (Figure 2A). Gate 2 excludes unstained cells, sub G1 apoptotic cells and cell clumps based upon identification of singlets (VL1Area vs. VL1Width) (Figure 2B). Gate 3 is a derived gate, encompassing cells within gates 1 and 2, and identifies GFP (or RFP) negative cells vs. DNA content based on VL1Height (Figure 2C). Gate 4 is a derived gate, encompassing cells within gates 1 and 2, and identifies GFP (or RFP) positive cells vs. DNA content (Figure 2C). Dot plots of cell size as measured by GFP (or RFP) vs. forward scatter (FSC) are used to generate Gates 5 and 6 (Figure 2D). The two histograms plot DNA content on the X-axis and cell count on the Y-axis for the populations defined by Gate 3 and Gate 4. (Figure 2E and 2F) Histograms can also be included for cell size if desired (not shown). For cell size histograms, populations should be set to Gates 5 and 6, plotting FSC on the X-axis and counts on the y-axis.</li>
 <li>Set recording to stop at 10,000 GFP or RFP positive events (this will be Gate 4, the derived Gate of Gate 1, Gate 2 and the GFP or RFP positive gate). Ranges from 6,000 - 10,000 GFP positive events have traditionally been the goal for publication quality profiles9,11. We suggest using a minimum of 15 wings or eyes, that are approximately 50% GFP positive to obtain sufficient Gate 4 events10 for high quality profiles. However, we have successfully obtained clear data for preliminary cell cycle analysis from as few as 6 wings (Figure 3A). Set the acquisition volume for 300 &mu;l, this will use about 460 &mu;l of the total 0.5 ml sample. Table 2 shows the appropriate threshold and voltage settings provided in the template for the example in Figure 2.</li>
 <li>Run samples at standard sensitivity or high sensitivity 100 &mu;l/min and record. We find little to no difference in our analysis with the different sensitivity levels. Due to the acoustic focusing of cells, this cytometer can accurately measure samples at very high flow rates. Unlike traditional cytometers, we find no data compromise by acquiring at speeds up to 1,500 events per second. The recorded data files generated by the Attune software are in .fcs format, which is compatible with most cell cycle modeling software.</li>
</ol>
5. Data Analysis
<ol>
 <li>Histograms of cell size and DNA content for GFP (or RFP) positive and negative populations can be compared. They can be manually layered by copying and pasting using Adobe Photoshop software. If the y-axis is set to automatic, the Y-axis will scale with the highest count for each sample. Overlaying two graphs with the Y-axis set to autoscale creates an overlay of histograms with the axes set for the global maximum for each population. This allows easy visual comparison for relative changes in cell cycle phasing, even when total cell numbers of GFP positive and GFP negative populations are not equivalent (example in Figure 3B). Alternatively, a custom y-axis (y-axis manual option) can be set to a fixed value for the histograms. In this case the relative cell numbers for two populations can be compared. As the Attune uses a defined volume of sample per run, absolute quantification of cells for each run, in each gate with percentages of the total population, can be obtained from the statistics table automatically generated during the run in the workspace (example in Figure 3F). Dot plots and histograms can also be easily layered in the Attune software, by simply dragging and dropping different samples from the sample file menu at right onto the desired graph of an open sample for comparison. However to overlay subpopulations from within a single sample (i.e. GFP positive and GFP negative within the same sample), we use Photoshop.</li>
 <li>Two methods can be used to find relative percentages in different phases of the cell cycle for a population. The approximation method relies upon user defined regions to delineate boundaries of G1, S, G2 and &gt;G2 peaks. This method allows quick and easy estimations of percentages within the user-defined regions (Figure 3E). We set our regions based upon a direct test with Drosophila Kc cells, labeled for S-phase using Ethynyl-deoxyUridine (EdU) incorporation. The second method uses third party modeling software for estimation of cell cycle distribution. We used ModFitLT (Verity Software House) software for example, in Figure 3F.</li>
</ol>

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The authors have nothing to disclose.

The protocol described here allows for analysis of cell cycle, relative cell size and relative cell number in live Drosophila tissues at various developmental stages. When this analysis is coupled with cell-type specific fluorescent protein expression or lineage tracing, detailed information can be obtained about cellular responses to discreet cell cycle or growth perturbations. As proof of principle, we disrupted quiescence in the pupal fly brain by expressing G1-S cell cycle regulators in GFP labeled cel...

Flow cytometry can be used for measurements of cell viability, relative cell size, DNA content and fluorescent protein expression in live cell populations. Due to the replication of nuclear DNA during S-phase, information about DNA content in a population of cells can be used to infer relative percentages in different cell cycle phases 1-3. This method has become a cornerstone of cell cycle analysis in model systems from yeast to mammals.
The fruit fly Drosophila melanogaster has become an excellent model system for genetic in vivo analyses of cell cycle regulation. The extensive genetic tools available in flies allow for elegant tissue specific and temporally regulated manipulations of cell cycle regulators along with in vivo fluorescent protein-based lineage tracing 4-6. Flow cytometry has been used to study DNA content in a number of Drosophila cell types, including endoreplicating cells and cultured mitotic cells 7,8. An important advance for in vivo cell cycle studies was made by de la Cruz and Edgar, with the development of a protocol for flow cytometric analysis of live diploid Drosophila imaginal discs 9,10, a protocol which has been used and adapted by many labs. This technique, when coupled with genetic in vivo lineage tracing via inducible fluorescent protein expression and tissue specific labeling, allows one to obtain information about gene manipulation effects on overall cell doubling time, cell size and to determine precise timing of cell cycle phases in vivo 9,11. However this method has thus far relied on the use of the cell permeable Hoechst 33342 DNA-intercalating dye to stain and quantify DNA in live cells, which has limited users to flow cytometers with a UV laser capable of exciting the Hoechst dye. These are generally found only in sorters (i.e. BD FACS Vantage, BD FACSAria) or expensive multicolor benchtop systems (i.e. BD LSR), usually requiring support by institutional flow core facilities.
We have modified the Hoechst-based protocol to use a new live-cell DNA dye from Invitrogen, Vybrant DyeCycle Violet. This dye is compatible with a violet 405 nm laser, more common in smaller benchtop analyzers and available in the small self-contained benchtop analyzer, the Attune Acoustic Focusing Cytometer. Here we present a detailed protocol for cell cycle analysis that can be coupled with cell type, cell size, cell number and lineage analysis in a variety of Drosophila tissues during various stages of development using DyeCycle Violet and the Attune. This protocol expands the number of cytometers suitable for such analysis with Drosophila tissues and provides examples of how this type of live cell cycle analysis can be modified for additional tissue types and developmental stages.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>12x75 mm Polystyrene Round-Bottom 5 ml Test Tube </td>
 <td>BD Falcon </td>
 <td>352058 </td>
 <td>5 ml tubes</td>
 </tr>
 <tr>
 <td>Attune Acoustic Focusing Cytometer</td>
 <td>Life Technologies/ Applied Biosystems</td>
 <td>4445315</td>
 <td>Blue / Violet configuration</td>
 </tr>
 <tr>
 <td>Attune Cytometer Software (version 1.2.5)</td>
 <td>Life Technologies/ Applied Biosystems</td>
 <td>Free </td>
 <td>PC only</td>
 </tr>
 <tr>
 <td>Attune Performance Tracking Beads (5 x 106 beads/ ml) </td>
 <td>Life Technologies/ Applied Biosystems</td>
 <td>4449754 </td>
 <td>For daily performance test</td>
 </tr>
 <tr>
 <td>Dumont #5 Inox forceps </td>
 <td>Fine Science Tools</td>
 <td>11251-20 </td>
 <td></td>
 </tr>
 <tr>
 <td>Embryo dishes 
 30 mm x 12mm </td>
 <td>Electron Microscopy Sciences</td>
 <td>70543-30 </td>
 <td>Glass dissection dishes </td>
 </tr>
 <tr>
 <td>Eppendorf Thermomixer</td>
 <td>Eppendorf</td>
 <td>022670051 </td>
 <td></td>
 </tr>
 <tr>
 <td>Trypsin-EDTA Solution (10x)</td>
 <td>Sigma</td>
 <td>T4174 </td>
 <td></td>
 </tr>
 <tr>
 <td>Vannas-T&uuml;bingen Spring Scissors </td>
 <td>Fine Science Tools</td>
 <td>15003-08 </td>
 <td>Straight 5mm Cutting Edge </td>
 </tr>
 <tr>
 <td>Vybrant DyeCycle Violet Stain</td>
 <td>Life Technologies/ Invitrogen</td>
 <td>V35003 </td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Table 1. Required reagents and instruments.</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Live DNA Stain Solution (10 ml):
 1 ml 10X Ca2+ Mg2+ free PBS (pH7.2) 
 9 ml 10X Trypsin-EDTA (Sigma) 
 5 &mu;l Invitrogen Vybrant DyeCycle Violet (note that this is 0.25X the recommended concentration for mammalian cells. We find that higher concentrations are toxic to Drosophila cells.)
 10X Ca2+ Mg2+ free PBS (pH7.2): 1.37M NaCl, 27 mM KCl, 100mM Na2HPO4 (dibasic), 20mM KH2PO4 (monobasic) adjusted to pH 7.2
 </td>
</tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td><table border="1">
 <tbody>
 <tr>
 <td></td>
 <td>FSC</td>
 <td>SSC</td>
 <td>BL1</td>
 <td>VL1</td>
 </tr>
 <tr>
 <td>Threshold (x1000)</td>
 <td>100</td>
 <td>10</td>
 <td>10</td>
 <td>10</td>
 </tr>
 <tr>
 <td>Voltage (mV)</td>
 <td>2950</td>
 <td>4250</td>
 <td>1800</td>
 <td>1150</td>
 </tr>
 </tbody>
 </table>
Table 2. Threshold and voltage setting for the analysis in Figure 2.</td>
 </tr>
 </tbody>
 </table>

live cell cycle analysis of drosophila tissues using the attune acoustic focusing cytometer and vybrant dyecycle violet dna stain

Flow cytometry has been widely used to obtain information about DNA content in a population of cells, to infer relative percentages in different cell cycle phases. This technique has been successfully extended to the mitotic tissues of the model organism Drosophila melanogaster for genetic studies of cell cycle regulation in vivo. When coupled with cell-type specific fluorescent protein expression and genetic manipulations, one can obtain detailed information about effects on cell number, cell size and cell cycle phasing in vivo. However this live-cell method has relied on the use of the cell permeable Hoechst 33342 DNA-intercalating dye, limiting users to flow cytometers equipped with a UV laser. We have modified this protocol to use a newer live-cell DNA dye, Vybrant DyeCycle Violet, compatible with the more common violet 405nm laser. The protocol presented here allows for efficient cell cycle analysis coupled with cell type, relative cell size and cell number information, in a variety of Drosophila tissues. This protocol extends the useful cell cycle analysis technique for live Drosophila tissues to a small benchtop analyzer, the Attune Acoustic Focusing Cytometer, which can be run and maintained on a single-lab scale.

Figure 2 shows representative results for a larval wing sample, expressing GFP in the posterior half of the tissue, using the provided GFP template. Similar results are obtained with the same tissue type and expression pattern for RFP using the provided RFP template (Figure 3A). The provided templates and voltages (Table 2) are suitable for analysis of larval eyes (Figure 3B), brains and wings, as well as pupal eyes, brains (Figure 3D) and wings. H...

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Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

A protocol for cell cycle analysis of live Drosophila tissues using the Attune Acoustic Focusing Cytometer is described. This protocol simultaneously provides information about relative cell size, cell number, DNA content and cell type via lineage tracing or tissue specific expression of fluorescent proteins in vivo.

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

Flow cytometry has  been widely used to obtain information about DNA content in a population of  cells, to infer relative percentages in different cell ...

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13.5K Views.  University of Michigan. A protocol for cell cycle analysis of live Drosophila tissues using the Attune Acoustic Focusing Cytometer is described. This protocol simultaneously provides information about relative cell size, cell number, DNA content and cell type via lineage tracing or tissue specific expression of fluorescent proteins in vivo.

Video: Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Analysis of Cell Cycle Position in Mammalian Cells

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of cell proliferation underlies the pathology of diseases like cancer. As such there is great need to be able to investigate cell proliferation and quantitate the proportion of cells in each phase of the cell cycle. It is also of vital importance to indistinguishably identify cells that are replicating their DNA within a larger population. Since a cell&prime;s decision to proliferate is made in the G1 phase immediately before initiating DNA synthesis and progressing through the rest of the cell cycle, detection of DNA synthesis at this stage allows for an unambiguous determination of the status of growth regulation in cell culture experiments.
DNA content in cells can be readily quantitated by flow cytometry of cells stained with propidium iodide, a fluorescent DNA intercalating dye. Similarly, active DNA synthesis can be quantitated by culturing cells in the presence of radioactive thymidine, harvesting the cells, and measuring the incorporation of radioactivity into an acid insoluble fraction. We have considerable expertise with cell cycle analysis and recommend a different approach. We Investigate cell proliferation using bromodeoxyuridine/fluorodeoxyuridine (abbreviated simply as BrdU) staining that detects the incorporation of these thymine analogs into recently synthesized DNA. Labeling and staining cells with BrdU, combined with total DNA staining by propidium iodide and analysis by flow cytometry1 offers the most accurate measure of cells in the various stages of the cell cycle. It is our preferred method because it combines the detection of active DNA synthesis, through antibody based staining of BrdU, with total DNA content from propidium iodide. This allows for the clear separation of cells in G1 from early S phase, or late S phase from G2/M. Furthermore, this approach can be utilized to investigate the effects of many different cell stimuli and pharmacologic agents on the regulation of progression through these different cell cycle phases.
In this report we describe methods for labeling and staining cultured cells, as well as their analysis by flow cytometry. We also include experimental examples of how this method can be used to measure the effects of growth inhibiting signals from cytokines such as TGF-&beta;1, and proliferative inhibitors such as the cyclin dependent kinase inhibitor, p27KIP1. We also include an alternate protocol that allows for the analysis of cell cycle position in a sub-population of cells within a larger culture5. In this case, we demonstrate how to detect a cell cycle arrest in cells transfected with the retinoblastoma gene even when greatly outnumbered by untransfected cells in the same culture. These examples illustrate the many ways that DNA staining and flow cytometry can be utilized and adapted to investigate fundamental questions of mammalian cell cycle control.

Determining the cell cycle position of a population of cells, or understanding how signals affect proliferation, can be readily measured by flow cytometry using this protocol. We report a simple experimental approach to staining cells and quantifying their position in the cell cycle.

The regulation of cell proliferation is central to tissue morphogenesis during the development of multicellular organisms. Furthermore, loss of control of ...

Assessment of GFP Expression and Viability Using the Tali Image-Based Cytometer

Single-cell and population information are commonly obtained either by flow cytometry or fluorescence microscopy. However, these two methods provide different information. Flow cytometry gives quantitative multi-parametric information about physical characteristics and staining or expression, but doesn't allow for visualization. Stand-alone fluorescence microscopy provides visual data, but doesn't allow for straightforward quantitative measurements1.
Image-based cytometry bridges the gap between these two methods, enabling the quick visualization and simultaneous quantitative analysis of thousands of cells in heterogeneous populations2. Here, we present a method for performing cell viability and green fluorescent protein (GFP) expression assays using the Tali Image-Based Cytometer3. The Tali instrument is a 3-channel (bright field, green fluorescence, red fluorescence) benchtop assay platform that offers several advantages over flow cytometry and fluorescence microscopy. The Tali cytometer is less expensive, takes up less bench space, requires less maintenance, and the work flow has been simplified so that the operation and analysis is much simpler and quicker. The Tali cytometer is capable of performing a range of suspension cell-based assays, including GFP and red fluorescent protein (RFP) expression, apoptosis4-6 and cell viability analysis with propidium iodide (PI)7-11.
Here, we demonstrate the use of the Tali instrument in performing a cell viability assay in cells expressing GFP. GFP-transduced cells are stained using the Tali Viability Kit - Dead Cell Red. The cells are then pipetted into a Tali Cellular Analysis Slide and loaded into the cytometer. Bright field, red fluorescence and green fluorescence images are captured and analyzed using assay specific algorithms. Histograms are then generated to display cell size, PI fluorescence intensity, and GFP fluorescence intensity. These parameters can then be thresholded to home in on a specific cell population.
A side-by side comparison of the Tali Image-Based Cytometer and traditional flow cytometry demonstrates that the two methods provide comparable data regarding cell viability and protein expression. However, the Tali instrument provides additional visual information about the cell population that cannot be obtained using a flow cytometer.

This protocol describes how to perform cell viability and fluorescence expression assays using the Tali Image-Based Cytometer.

Single-cell and population information are commonly obtained either by flow cytometry or fluorescence microscopy. However, these two methods provide ...

Performing Vaginal Lavage, Crystal Violet Staining, and Vaginal Cytological Evaluation for Mouse Estrous Cycle Staging Identification

A rapid means of assessing reproductive status in rodents is useful not only in the study of reproductive dysfunction but is also required for the production of new mouse models of disease and investigations into the hormonal regulation of tissue degeneration (or regeneration) following pathological challenge. The murine reproductive (or estrous) cycle is divided into 4 stages: proestrus, estrus, metestrus, and diestrus. Defined fluctuations in circulating levels of the ovarian steroids 17-&beta;-estradiol and progesterone, the gonadotropins luteinizing and follicle stimulating hormones, and the luteotropic hormone prolactin signal transition through these reproductive stages. Changes in cell typology within the murine vaginal canal reflect these underlying endocrine events. Daily assessment of the relative ratio of nucleated epithelial cells, cornified squamous epithelial cells, and leukocytes present in vaginal smears can be used to identify murine estrous stages. The degree of invasiveness, however, employed in collecting these samples can alter reproductive status and elicit an inflammatory response that can confound cytological assessment of smears. Here, we describe a simple, non-invasive protocol that can be used to determine the stage of the estrous cycle of a female mouse without altering her reproductive cycle. We detail how to differentiate between the four stages of the estrous cycle by collection and analysis of predominant cell typology in vaginal smears and we show how these changes can be interpreted with respect to endocrine status.

Here, we describe how to identify the stage of the murine reproductive (proestrus, estrus, metestrus, or diestrus) by simple, non-invasive collection and cytological assessment of vaginal smear samples. We further describe how vaginal cytology reflects circulating hormonal levels underlying transition through the murine reproductive cycle.

A rapid means of assessing reproductive status in rodents is useful not only in the study of reproductive dysfunction but is also required for the ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

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

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

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

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

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Drosophila eye is widely used as a model for studies of development and neuronal degeneration. With the powerful mitotic recombination technique, elegant genetic screens based on clonal analysis have led to the identification of signaling pathways involved in eye development and photoreceptor (PR) differentiation at larval stages. We describe here the Tomato/GFP-FLP/FRT method, which can be used for rapid clonal analysis in the eye of living adult Drosophila. Fluorescent photoreceptor cells are imaged with the cornea neutralization technique, on retinas with mosaic clones generated by flipase-mediated recombination. This method has several major advantages over classical histological sectioning of the retina: it can be used for high-throughput screening and has proved an effective method for identifying the factors regulating PR survival and function. It can be used for kinetic analyses of PR degeneration in the same living animal over several weeks, to demonstrate the requirement for specific genes for PR survival or function in the adult fly. This method is also useful for addressing cell autonomy issues in developmental mutants, such as those in which the establishment of planar cell polarity is affected.

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Tomato/GFP-FLP/FRT method involves visualizing mosaic photoreceptor cells in living Drosophila. It can be used to follow individual photoreceptor cell fates in the retina for days or weeks. This method is ideal for studies of retinal degeneration and neurodegenerative diseases or photoreceptor cell development.

The Drosophila eye is widely used as a model  for studies of development and neuronal degeneration. With the powerful mitotic  recombination technique, ...

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Reduced-gravity Environment Hardware Demonstrations of a Prototype Miniaturized Flow Cytometer and Companion Microfluidic Mixing Technology

Until recently, astronaut blood samples were collected in-flight, transported to earth on the Space Shuttle, and analyzed in terrestrial laboratories. If humans are to travel beyond low Earth orbit, a transition towards space-ready, point-of-care (POC) testing is required. Such testing needs to be comprehensive, easy to perform in a reduced-gravity environment, and unaffected by the stresses of launch and spaceflight. Countless POC devices have been developed to mimic laboratory scale counterparts, but most have narrow applications and few have demonstrable use in an in-flight, reduced-gravity environment. In fact, demonstrations of biomedical diagnostics in reduced gravity are limited altogether, making component choice and certain logistical challenges difficult to approach when seeking to test new technology. To help fill the void, we are presenting a modular method for the construction and operation of a prototype blood diagnostic device and its associated parabolic flight test rig that meet the standards for flight-testing onboard a parabolic flight, reduced-gravity aircraft. The method first focuses on rig assembly for in-flight, reduced-gravity testing of a flow cytometer and a companion microfluidic mixing chip. Components are adaptable to other designs and some custom components, such as a microvolume sample loader and the micromixer may be of particular interest. The method then shifts focus to flight preparation, by offering guidelines and suggestions to prepare for a successful flight test with regard to user training, development of a standard operating procedure (SOP), and other issues. Finally, in-flight experimental procedures specific to our demonstrations are described.

Spaceflight blood diagnostics need innovation. Few demonstrations have been published illustrating in-flight, reduced-gravity health diagnostic technology. Here we present a method for construction and operation of a parabolic flight test rig for a prototype point-of-care flow-cytometry design, with components and preparation strategies adaptable to other setups.

Until recently, astronaut blood samples were collected in-flight, transported to earth on the Space Shuttle, and analyzed in terrestrial laboratories. If ...