This work was supported by funding from the Swiss National Science Foundation (project grant 31003A_173188; www.snf.ch) and the University of Bern (www.unibe.ch) to BS. The funders had no role in study design, data collection, and analysis, decision to publish, or preparation of the manuscript.

1. Preparation of reagents and stocks for Click-it EdU assay
NOTE: Refer to the Table of Materials and Table 1 for details about the kit and reagents supplied with the kit.
<ol>
	<li>Bring the vials to room temperature before preparing the solutions.</li>
	<li>Prepare 10 mM EdU (component A) stock solution by adding 2 mL of dimethylsulfoxide (DMSO, component C). Mix well and store at -20 &#176;C.</li>
	<li>Prepare a working solution of Alexa Fluor 647-azide (component B) by adding 70&#956;L of DMSO (component C). Mix well and store at -20 &#176;C.</li>
	<li>Prepare a 1x solution of Click-iT EdU reaction buffer (component D) by mixing 4 mL of this solution with 36 mL of deionized water. Store the remaining solution at 2-6 &#176;C.</li>
	<li>Make a 10x stock (200 mg/mL) of the Click-iT EdU buffer additive (component F) by adding 2 mL of deionized water. Mix well and store at -20 &#176;C.</li>
	<li>Store Hoechst 33342 (component G) at 2-6 &#176;C. Store DMSO (component C) in a desiccator at -20 &#176;C. 
	&#8203;NOTE: Gloves should be used to handle DMSO and Hoechst.</li>
</ol>
2. Dissection of third instar larval brains and EdU incorporation
NOTE: The protocol for brain dissections has been described previously12. Before starting dissections, make sure sufficient amounts of the EdU and PFA solutions are prepared and thawed as described in 2.6 and 3.1.
<ol>
	<li>Prepare 10x Phosphate-buffered saline (PBS) by adding 25.6 g Na2HPO4.7H20, 80 g NaCl, 2 g KCl, and 2 g KH2PO4 to 1 L of deionized water. Adjust the pH to 7.4, and subsequently prepare a 1x solution in deionized water.</li>
	<li>Add Schneiders dissection medium (SM) to two consecutive depressions of a glass (3 or 9) depression glass spot plate. Add 1x PBS to the next consecutive depression.</li>
	<li>Using a pair of forceps, pick wandering third instar larvae and place on a tissue wetted with PBS to clean off fly food residue. Place the larva in the depression with PBS and then in the consecutive depression with SM.</li>
	<li>Using a pair of fine forceps, remove 3/4th of the lower body of the larva.
	<ol>
		<li>Gently grab the larval mouth hooks with one pair of forceps, and hold the cuticle at the other end with the other pair of forceps.</li>
		<li>Turn the larval head inside out by pushing inward the mouth hooks and simultaneously peeling off the tissue at the other end.</li>
	</ol>
	</li>
	<li>Observe the larval brain with attached imaginal discs. Remove other tissues attached to the brain, and transfer the brain to the next consecutive depression with SM.</li>
	<li>Thaw the 10 mM EdU stock solution. Prepare a 100 &#956;M solution by diluting this 10 mM stock in SM.</li>
	<li>Add 100 &#956;L of this 100 &#956;M EdU+SM solution in another depression on the Pyrex plate. Incubate ~5-10 dissected brains in 100 &#956;L of 100 &#956;M EdU+SM solution for 2 h at 25 &#176;C. 
	&#8203;NOTE: As NBs divide once in ~2 h, this protocol aims to analyze one full cell cycle. Only use intact brains for further steps. Discard brains that are damaged during dissection.</li>
</ol>
3. Fixation and immunostaining
<ol>
	<li>Prepare 4% paraformaldehyde (PFA) solution in a fume hood.
	<ol>
		<li>Add 4 g of paraformaldehyde powder to 80 mL of 1x PBS in a beaker placed on a magnetic stirrer and heat to 60 &#176;C. To dissolve PFA, add a few drops of 1 N NaOH. Check the pH, and adjust to 7.0 with a few drops of 1 M HCl if required.</li>
		<li>Adjust the volume to 100 mL with 1x PBS. Aliquot the PFA solution and store at 2-6 &#176;C for up to 1 month. Add 0.3% non-ionic detergent (see the Table of Materials) to the PFA solution for efficient fixation of large tissues such as the larval brain.</li>
	</ol>
	</li>
	<li>After EdU incorporation, transfer the brains to 0.6 mL microcentrifuge tubes containing 4% PFA. Incubate at room temperature on a nutator for 15 min. Tap the tube on the laboratory bench so that the brains settle down at the bottom of the tube.</li>
	<li>Remove the PFA and wash 3 times with PBS + 0.3% non-ionic detergent (PBST) at room temperature. Ensure that each wash lasts for at least 10 min on a nutator. After the last wash, remove PBST, add blocking solution (PBST + 5% bovine serum albumin), and incubate for 30 min at room temperature.</li>
	<li>Dilute anti-Miranda antibody (1:250) and anti pH3 antibody (1:200) in PBST (both antibodies in the same tube). Remove the blocking buffer, and add the primary antibody solution. Incubate overnight at 2-6 &#176;C on a nutator. 
	NOTE: Miranda is a membrane protein present on CB NBs. It serves to identify these large round cells in the CB region13.</li>
	<li>Remove the primary antibody solution and wash 3 times with PBST. Ensure that each wash lasts for 10 min on a nutator at room temperature.</li>
	<li>Prepare secondary antibody solution by adding 1:500 diluted Alexa Fluor 488 anti-Rabbit and Alexa Fluor 568 anti-Rat in PBST. Remove the PBST, and add the secondary antibody solution to the dissected brains. Incubate overnight at 2-6 &#176;C in the dark, on a nutator, and wash 3 times with PBST (step 3.5).</li>
</ol>
4. EdU detection, DNA staining, and mounting
<ol>
	<li>&#160;To prepare the EdU detection cocktail, mix the components (see Table 2) in a 1.5 mL tube.</li>
	<li>After the last wash (step 3.6), remove PBST, and add the EdU detection cocktail to the dissected brains. Incubate at room temperature in the dark for 30 min on a nutator. Wash 2 times with PBST for 10 min each, in the dark.</li>
	<li>For DNA staining, dilute Hoechst 33342 (component G) 1:2,000 in PBST to prepare a 5 &#956;g/mL solution.</li>
	<li>Remove PBST after the second wash (step 4.2), and add 500 &#956;L of 5 &#956;g/mL Hoechst solution to the dissected brains. Incubate in the dark for 10 min. Remove Hoechst, and wash once with PBST for 10 min.</li>
	<li>Tap the tube onto the lab bench, and let the brains settle down. Remove the PBST from the tube, but leave behind only 50-100 &#956;L.</li>
	<li>Cut the end of a 200 &#956;L micropipette tip, and carefully transfer the brains onto a clean glass slide. Remove excess PBST from the slide by blotting with filter paper strips. Be careful not to let the filter paper touch the brains.</li>
	<li>Put a drop of water-soluble, non-fluorescing mounting medium onto the brains, and orient the brains such that the ventral nerve cord faces the slide, and the lobes face upwards. Arrange the brains in a single file so that it is easier to image the brains serially.</li>
	<li>Gently place a coverslip on top of the brains. Place the slide at 2-6 &#176;C overnight. 
	&#8203;NOTE: The mounting medium hardens upon overnight storage so that there is no need to seal the coverslip with nail polish.</li>
</ol>
5. Imaging
NOTE: See the Table of Materials for details on the laser scanning microscope and oil-immersion objective used in this protocol.
<ol>
	<li>From the Acquisition software, select the 63x objective.</li>
	<li>Put a drop of immersion oil on the coverslip just above the mounted brains to make it easier to locate the tissue through the eyepiece.</li>
	<li>Using the DAPI/Hoechst 33342 channel, find the brain through the eyepiece, and then switch to the acquisition mode in the software.</li>
	<li>Set up 4 channels to image Hoechst 33342 (DNA), Alexa Fluor 488 (pH3), Alexa Fluor 568 (Miranda), and Alexa Fluor 647 (EdU). Use the dye-assistant tool, which automatically sets up the excitation lasers and emission filters for the selected dyes.</li>
	<li>Set the field of view such that it encompasses the entire brain lobe. Image the entire volume of the brain lobe by acquiring z-stacks spaced 0.8 &#956;m apart. Store all images from an imaging session in a *.lif library format.</li>
</ol>
6. Image analysis
NOTE: The following steps describe the analysis of acquired images and how to sort cells into G0/G1 phase, S phase, S&#62;G2/M (progression from S to G2/M), and M phase using the ImageJ software.
<ol>
	<li>Download Fiji (Fiji is ImageJ) from the following URL: https://fiji.sc/. Open Fiji, then drag and drop the .lif files into Fiji. 
	NOTE: Fiji is a version of ImageJ that comes pre-installed with several plugins14. Transferring .lif files into Fiji will open the Bio-formats plugin, which is needed to process the .lif files. Bio-formats plugin is also required to open image files generated from some other microscope brands, e.g., nd2 files generated from a Nikon microscope. This plugin comes pre-installed with Fiji.</li>
	<li>Select Data browser from the stack viewing tab, and use virtual stack from the memory management tab in the bio-formats plugin. 
	NOTE: Lif files with z-stacks from 8-10 brains are often 300-400 megabytes in size. On computers with low RAM, opening a few such files can rapidly deplete the available RAM on ImageJ and prevent any further image processing. Virtual stack is &#39;read-only&#39;, does not need high RAM for processing, and is an ideal option to load large datasets onto Fiji.</li>
	<li>Observe the multichannel image displayed in ImageJ. Change the color of the channels from the menu bar using Image | color | channels tool. Observe the Miranda-labeled NBs as large round cells in the CB region.</li>
	<li>Draw a region of interest (ROI) using the ellipse tool over each NB to avoid counting the NB twice.
	<ol>
		<li>From the ImageJ menu bar, select analyze | tools | ROI manager. Mark all NBs in the current z-section, and press t after marking each cell.</li>
	</ol>
	</li>
	<li>Once all NBs in the current z-section are marked, change the channels to pH3 and EdU, and manually count the number of EdU-positive NBs, pH3-positive NBs, NBs staining positively for both EdU and pH3, and NBs not staining for both markers.</li>
	<li>Search for NBs in subsequent z-sections, delete old ROIs, and add new ROIs to count NBs in different stacks.</li>
	<li>Prepare a spreadsheet with the following columns: 1. EdU- pH3-: this section represents the double-negative cells that are in the G0/G1 phase of the cell cycle; 2. EdU+: the cells in this category have incorporated EdU and are undergoing DNA replication (S-phase); 3. EdU+ pH3+: these dual-positive cells have completed the S phase and have progressed to G2 or M phase; 4. pH3+: these NBs are undergoing mitosis.</li>
	<li>Calculate percentages of NBs present in all of the above 4 categories for each lobe. Prepare a bar graph using the spreadsheet software showing the pooled data for percentages of NBs in each category.</li>
</ol>

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The authors have declared that no competing interests exist.

EdU incorporation and its subsequent &#39;click&#39; reaction with cell-permeable azide presents practical advantages of this technique over the BrdU method used earlier7. However, this reaction is catalyzed by Cu(I) ions, and several dyes may be unstable in the presence of this copper catalyst, as is clearly advised by the Click-it EdU kit manufacturer. When immunostaining experiments had been performed after executing the EdU detection step, the expected signal intensity was not observed with th...

The cell division cycle comprises a G1 phase (first gap phase), a replicative/S-phase, a G2 (second gap phase), and an M (mitotic) phase. Passing through these phases, the cell undergoes dramatic changes in cellular transcription, translation, and re-organization of cytoskeletal machinery1,2. In response to developmental and environmental cues, cells may temporarily cease to divide and become quiescent (G0) or differentiate and permanently cease to divide3. Other scenarios, such as DNA damage, may cause premature differentiation or apoptosis3,4. Response to such cues is mediated by cell cycle checkpoints, which act as a surveillance system to ensure the integrity of essential cellular processes before the cell commits to the next phase of the division cycle5. Therefore, studies on genes regulating DNA replication, checkpoints, and mitotic machinery need to analyze possible cell cycle progression defects that may occur in mutant cells or upon siRNA knockdown of these genes. Additionally, such analyses may be employed to test overall cell health as well as cellular responses to drug treatment.
5-bromo-2&#39;-deoxyuridine (BrdU) is a thymidine analog that is incorporated into DNA during replication6. This method was used extensively to identify cells in S-phase. However, the cells are then subjected to harsh DNA denaturation procedures to allow detection of BrdU through the use of anti-BrdU antibodies6. This harsh treatment may damage cellular epitopes and prevent further characterization of the sample through immunostaining. EdU incorporation and subsequent detection by a copper-catalyzed, &#39;click reaction&#39; with small, cell-permeable, fluorescently tagged azide dyes eliminates the need for harsh denaturation procedures7. This method, therefore, emerged as a more practical alternative to BrdU incorporation.
Further, pH3 has been described as a reliable marker for mitotic/M phase cells8. Histone H3 is a DNA-associated core histone protein that becomes phosphorylated in around the late G2 phase to early M phase and is de-phosphorylated toward the end of anaphase8. Several commercial antibodies can be used to detect pH3 using standard immunostaining protocols. Dual-staining of EdU and pH3 would therefore enable the detection of cells in S-phase as well as M-phase. Additionally, cells in the G1 and early G2 phase would not stain positively for either of the markers.
Drosophila neural stem cells or neuroblasts (NBs) offer a well-characterized stem cell model wherein cells divide asymmetrically to produce one identical self-renewing NB and a ganglion mother cell (GMC), which is fated for differentiation9. Additionally, several genetic tools and NB-specific antibodies make this system suitable for genetic manipulation and live-cell imaging. Consequently, several studies have utilized NBs to study genes regulating asymmetric divisions and cell fate determination9. Distinct populations of NBs exist in the central brain (CB) and the optic lobe (OL) of the larval brain9; CB NBs were used for the current study. These third instar larval CB NBs are large cells that are also suitable for studying factors regulating mitotic spindle assembly. A protocol to analyze cell cycle progression defects would be a vital tool in such studies.
Protocols published earlier employed commercial kits, such as Click-iT EdU Alexa Fluor Cell Proliferation Kit, which provide several reaction components and azide dyes tagged with a variety of Alexa Fluor dyes for EdU incorporation and detection10. However, the reagents supplied with such kits are not compatible with some fluorescent tags often used with secondary antibodies. This EdU detection kit (Click-iT EdU Alexa Fluor Cell Proliferation Kit supplied with Alexa Fluor 647-conjugated azide dye) was tested in Drosophila third instar larval NBs, and co-staining was attempted with antibodies against pH3 and Miranda, a marker for NBs. Further, Alexa Fluor 568- or Cy3-tagged secondary antibodies were used for detection of Miranda labeling on the plasma membrane of NBs11. However, the expected signal intensity and staining pattern (unpublished results) were not observed with these secondary antibodies when immunostaining was performed after EdU detection.
For EdU incorporation, the protocol described by Daul and colleagues required feeding of the larvae with Kankel-White medium mixed with EdU and bromophenol blue (BPB)10. The larvae fed on the EdU and BPB-spiked food, which could be seen by its blue color upon ingestion in the larval gut. Although this method was used for EdU incorporation in Mms19 loss-of-function (Mms19P) third instar larvae, the Mms19P larvae apparently did not feed as hardly any blue color was detected in the larval gut (unpublished results). The Mms19P larvae show drastic developmental deformities and eventually arrest in the third instar stage. This may somehow affect the feeding behavior of the third instar larvae and render the EdU-feeding protocol unsuitable for such cases.
After studying the available literature and working extensively on the standardization of essential steps, an alternative approach was proposed for EdU/pH3 dual-labeling in Drosophila NBs, which does not require feeding EdU to larvae. A previous study employed dual EdU/pH3 staining to analyze the cell cycle in NBs, but did not present a detailed protocol4. This presents an unnecessary hurdle for labs trying to implement this method. Furthermore, evaluating the compatibility of various reagents with the EdU kit and performing further optimization can be a time-consuming process. This paper presents a step-by-step protocol that covers EdU incorporation in dissected larval brains and immunostaining with anti-pH3 antibodies, followed by confocal microscopy and image analysis to allocate NBs to four distinct categories: G0/G1 phase, S phase, S&#62;G2/M (progression from S to G2/M), and M phase. The steps that need optimization are outlined and tips provided for image analysis of large datasets. Additionally, the EdU/pH3 readout in wild-type NBs is analyzed and compared with Mms19P&#160;NBs, which were recently reported to show a cell cycle delay11.

<table><tbody><tr><td>&lt;strong&gt;fly stocks&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>P{&lt;em&gt;EPgy2&lt;/em&gt;}&lt;em&gt;Mms19EY00797&lt;/em&gt;/TM3, &lt;em&gt;Sb1 Ser1&lt;/em&gt; &lt;em&gt;(Mms19&lt;sup&gt;p&lt;/sup&gt;)&lt;/em&gt;</td><td>Bloomington stock center</td><td>#15477</td><td>P-element insertion in the third exon of Mms19</td></tr><tr><td>&amp;nbsp;+; Mms19::eGFP, Mms19p</td><td>Generated in house</td><td></td><td>eGFP-tagged Mms19 protein expressed in Mms19p background</td></tr><tr><td>w&lt;sup&gt;1118&lt;/sup&gt;</td><td>Bloomington stock center</td><td>#3605</td><td>wild-type stock (w;+;+)</td></tr><tr><td>Primary antibodies</td><td></td><td></td><td>Dilution</td></tr><tr><td>Rat anti-Miranda</td><td>Abcam</td><td>Ab197788</td><td>1/250</td></tr><tr><td>Rabbit anti-pH3</td><td>Cell Signaling</td><td>9701</td><td>1/200</td></tr><tr><td>&lt;strong&gt;Secondary antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Goat anti-Rat Cy3</td><td>Jackson Immuno</td><td>112-165-167</td><td>1/150</td></tr><tr><td>Goat anti-Rat Alexa Fluor 568</td><td>Invitrogen</td><td>A11077</td><td>1/500</td></tr><tr><td>Goat anti-Rabbit Alexa Fluor 488</td><td>Invitrogen</td><td>A27034</td><td>1/500</td></tr><tr><td>&lt;strong&gt;Reagent/Kit&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Aqua Poly/Mount mounting medium</td><td>Polysciences Inc</td><td>18606-20</td><td></td></tr><tr><td>Click-it EdU incorporation kit, Alexa Flour 647</td><td>Thermo Fischer Scientific</td><td>C10340</td><td></td></tr><tr><td>Schneider&amp;rsquo;s Drosophila medium</td><td>Thermo Fischer Scientific</td><td>21720-024</td><td></td></tr><tr><td>Bovine serum albumin (BSA) fraction V</td><td>Merck</td><td>10735078001</td><td></td></tr><tr><td>Triton X-100</td><td>Fischer Scientific</td><td>9002-93-1</td><td>non-ionic detergent</td></tr><tr><td>&lt;strong&gt;Software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Fiji (Imagej)</td><td>https://imagej.net/Fiji</td><td></td><td></td></tr><tr><td>Leica Application Suite (LAS X)</td><td>Leica microsystems</td><td></td><td></td></tr><tr><td>PRISM</td><td>Graph pad software</td><td>Version 5</td><td></td></tr><tr><td>Microsoft Excel</td><td>Microsoft office</td><td>2016</td><td></td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Leica TCS SP8 laser scanning confocal microscope with 63x oil-immersion, 1.4 NA Plan-apochromat objective</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Materials&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Aqua Poly/Mount mounting medium</td><td></td><td></td><td>water-soluble, non-fluorescing mounting medium</td></tr><tr><td>Pyrex 3 or 9 depression glass spot plate</td><td></td><td></td><td></td></tr><tr><td>Whatman filter paper</td><td></td><td></td><td></td></tr></tbody></table>

5-ethynyl-2'-deoxyuridine/phospho-histone h3 dual-labeling protocol for cell cycle progression analysis in drosophila neural stem cells

In vivo cell cycle progression analysis is routinely performed in studies on genes regulating mitosis and DNA replication. 5-Ethynyl-2'-deoxyuridine (EdU) has been utilized to investigate replicative/S-phase progression, whereas antibodies against phospho-histone H3 have been utilized to mark mitotic nuclei and cells. A combination of both labels would enable the classification of G0/G1 (Gap phase), S (replicative), and M (mitotic) phases and serve as an important tool to evaluate the effects of mitotic gene knockdowns or null mutants on cell cycle progression. However, the reagents used to mark EdU-labelled cells are incompatible with several secondary antibody-fluorescent tags. This complicates immunostaining, where primary and tagged secondary antibodies are used to mark pH3-positive mitotic cells. This paper describes a step-by-step protocol for the dual-labeling of EdU and pH3 in Drosophila larval neural stem cells, a system utilized extensively to study mitotic factors. Additionally, a protocol is provided for image analysis and quantification to allocate labeled cells in 3 distinct categories, G0/G1, S, S&gt;G2/M (progression from S to G2/M), and M phases.

The bi-lobed Drosophila third instar larval brain has been utilized as a model system to study fundamental cellular and developmental processes9. The focus of the current study was to present a protocol for analysis of cell cycle progression in EdU- and pH3-labeled NBs of the CB region (Figure 1). The CB NBs are sub-divided into type I and type II, and they display the characteristic asymmetric division pattern9. Each type I NB divisio...

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5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

Cell cycle analysis with 5-ethynyl-2'-deoxyuridine (EdU) and phospho-histone H3 (pH3) labeling is a multi-step procedure that may require extensive optimization. Here, we present a detailed protocol that describes all steps for this procedure including image analysis and quantification to distinguish cells in different cell cycle phases.

5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

In vivo cell cycle progression analysis is routinely performed in studies on genes regulating mitosis and DNA replication. 5-Ethynyl-2'-deoxyuridine (EdU) ...

5-ethynyl-2-deoxyuridinephospho-histone-h3-dual-labeling-protocol-for

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Biology

3.7K Views.  University of Bern. Cell cycle analysis with 5-ethynyl-2'-deoxyuridine (EdU) and phospho-histone H3 (pH3) labeling is a multi-step procedure that may require extensive optimization. Here, we present a detailed protocol that describes all steps for this procedure including image analysis and quantification to distinguish cells in different cell cycle phases. 

Video: 5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

Studying Cell Cycle-regulated Gene Expression by Two Complementary Cell Synchronization Protocols

The gene expression program of the cell cycle represents a critical step for understanding cell cycle-dependent processes and their role in diseases such as cancer. Cell cycle-regulated gene expression analysis depends on cell synchronization into specific phases. Here we describe a method utilizing two complementary synchronization protocols that is commonly used for studying periodic variation of gene expression during the cell cycle. Both procedures are based on transiently blocking the cell cycle in one defined point. The synchronization protocol by hydroxyurea (HU) treatment leads to cellular arrest in late G1/early S phase, and release from HU-mediated arrest provides a cellular population uniformly progressing through S and G2/M. The synchronization protocol by thymidine and nocodazole (Thy-Noc) treatment blocks cells in early mitosis, and release from Thy-Noc mediated arrest provides a synchronized cellular population suitable for G1 phase and S phase-entry studies. Application of both procedures requires monitoring of the cell cycle distribution profiles, which is typically performed after propidium iodide (PI) staining of the cells and flow cytometry-mediated analysis of DNA content. We show that the combined use of two synchronization protocols is a robust approach to clearly determine the transcriptional profiles of genes that are differentially regulated in the cell cycle (i.e. E2F1 and E2F7), and consequently to have a better understanding of their role in cell cycle processes. Furthermore, we show that this approach is useful for the study of mechanisms underlying drug-based therapies (i.e. mitomycin C, an anticancer agent), because it allows to discriminate genes that are responsive to the genotoxic agent from those solely affected by cell cycle perturbations imposed by the agent.

We report two cell synchronization protocols that provide a context for studying events related to specific phases of the cell cycle. We show that this approach is useful for analyzing the regulation of specific genes in an unperturbed cell cycle or upon exposure to agents affecting the cell cycle.

The gene expression program of the cell cycle represents a critical step for understanding cell cycle-dependent processes and their role in diseases such ...

Genetics

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.

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.

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

Assessing Cell Cycle Progression of Neural Stem and Progenitor Cells in the Mouse Developing Brain after Genotoxic Stress

Neurons of the cerebral cortex are generated during brain development from different types of neural stem and progenitor cells (NSPC), which form a pseudostratified epithelium lining the lateral ventricles of the embryonic brain. Genotoxic stresses, such as ionizing radiation, have highly deleterious effects on the developing brain related to the high sensitivity of NSPC. Elucidation of the cellular and molecular mechanisms involved depends on the characterization of the DNA damage response of these particular types of cells, which requires an accurate method to determine NSPC progression through the cell cycle in the damaged tissue. Here is shown a method based on successive intraperitoneal injections of EdU and BrdU in pregnant mice and further detection of these two thymidine analogues in coronal sections of the embryonic brain. EdU and BrdU are both incorporated in DNA of replicating cells during S phase and are detected by two different techniques (azide or a specific antibody, respectively), which facilitate their simultaneous detection. EdU and BrdU staining are then determined for each NSPC nucleus in function of its distance from the ventricular margin in a standard region of the dorsal telencephalon. Thus this dual labeling technique allows distinguishing cells that progressed through the cell cycle from those that have activated a cell cycle checkpoint leading to cell cycle arrest in response to DNA damage.
An example of experiment is presented, in which EdU was injected before irradiation and BrdU immediately after and analyzes performed within the 4 hr&#160;following irradiation. This protocol provides an accurate analysis of the acute DNA damage response of NSPC in function of the phase of the cell cycle at which they have been irradiated. This method is easily transposable to many other systems in order to determine the impact of a particular treatment on cell cycle progression in living tissues.

Administration of two analogs of thymidine, EdU and BrdU, in pregnant mice allows the analysis of cell cycle progression in neural and progenitor cells in the embryonic mouse brain. This method is useful to determine the effects of genotoxic stress, including ionizing radiation, during brain development.

Neurons of the cerebral cortex are generated during brain development from different types of neural stem and progenitor cells (NSPC), which form a ...

Neuroscience

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 ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

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.

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.

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

Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization

This protocol describes a method to permit the tracking of cells through the cell cycle without requiring the cells to be synchronized. Achieving cell synchronization can be difficult for many cell systems. Standard practice is to block cell cycle progression at a specific stage and then release the accumulated cells producing a wave of cells progressing through the cycle in unison. However, some cell types find this block toxic resulting in abnormal cell cycling, or even mass death. Bromodeoxyuridine (BrdU) uptake can be used to track the cell cycle stage of individual cells. Cells incorporate this synthetic thymidine analog, while synthesizing new DNA during S phase. By providing BrdU for a brief period it is possible to mark a pool of cells that were in S phase while the BrdU was present. These cells can then be tracked through the remainder of the cell cycle and into the next round of replication, permitting the duration of the cell cycle phases to be determined without the need to induce a potentially toxic cell cycle block. It is also possible to determine and correlate the expression of both internal and external proteins during subsequent stages of the cell cycle. These can be used to further refine the assignment of cell cycle stage or assess effects on other cellular functions such as checkpoint activation or cell death.

This protocol describes the use of bromodeoxyuridine (BrdU) uptake to permit the temporal tracking of cells that were in S phase at a specific point in time. Addition of DNA dyes and antibody labeling facilitates detailed analysis of the fate of the S phase cells at later times.

This protocol describes a method to permit the tracking of cells through the cell cycle without requiring the cells to be synchronized. Achieving cell ...

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

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

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

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

Assessment of Global DNA Double-Strand End Resection using BrdU-DNA Labeling coupled with Cell Cycle Discrimination Imaging

The study of the DNA damage response (DDR) is a complex and essential field, which has only become more important due to the use of DDR-targeting drugs for cancer treatment. These targets are poly(ADP-ribose) polymerases (PARPs), which initiate various forms of DNA repair. Inhibiting these enzymes using PARP inhibitors (PARPi) achieves synthetic lethality by conferring a therapeutic vulnerability in homologous recombination (HR)-deficient cells due to mutations in breast cancer type 1 (BRCA1), BRCA2, or partner and localizer of BRCA2 (PALB2). 
Cells treated with PARPi accumulate DNA double-strand breaks (DSBs). These breaks are processed by the DNA end resection machinery, leading to the formation of single-stranded (ss) DNA and subsequent DNA repair. In a BRCA1-deficient context, reinvigorating DNA resection through mutations in DNA resection inhibitors, such as 53BP1 and DYNLL1, causes PARPi resistance. Therefore, being able to monitor DNA resection in cellulo is critical for a clearer understanding of the DNA repair pathways and the development of new strategies to overcome PARPi resistance. Immunofluorescence (IF)-based techniques allow for monitoring of global DNA resection after DNA damage. This strategy requires long-pulse genomic DNA labeling with 5-bromo-2&#8242;-deoxyuridine (BrdU). Following DNA damage and DNA end resection, the resulting single-stranded DNA is specifically detected by an anti-BrdU antibody under native conditions. Moreover, DNA resection can also be studied using cell cycle markers to differentiate between various phases of the cell cycle. Cells in the S/G2 phase allow the study of end resection within HR, whereas G1 cells can be used to study non-homologous end joining (NHEJ). A detailed protocol for this IF method coupled to cell cycle discrimination is described in this paper.

In the present protocol, we demonstrate how to visualize DNA double-strand end resection during S/G2 phase of the cell cycle using an immunofluorescence-based method.

The study of the DNA damage response (DDR) is a complex and essential field, which has only become more important due to the use of DDR-targeting drugs ...

Cell Cycle Analysis

Cell cycle refers to the set of events through which a cell grows, replicates its genome, and ultimately divides into two daughter cells through the process of mitosis. Because the amount of DNA in a cell shows characteristic changes throughout the cycle, techniques known as cell cycle analysis can be used to separate a population of cells according to the different phases of cell cycle they are in, based on their varying DNA content.This video will cover the principles behind cell cycle analysis via DNA-staining. We will review a generalized protocol for performing this staining using bromodeoxyuridine (BrdU, a thymidine analog that is incorporated into newly synthesized DNA strands) and propidium iodide (PI, a DNA dye that stains all DNA), followed by analysis of the stained cells with flow cytometry. During flow cytometry, a single cell suspension of fluorescently labeled cells is passed through an instrument with a laser beam and the fluorescence of each cell is read. We will then discuss how to interpret data from flow cytometric scatter plots, and finally, look at a few applications of this technique.

Cell Cycle Analysis: Principle, BrdU and Flow Cytometry

Cell cycle refers to the set of events through which a cell grows, replicates its genome, and ultimately divides into two daughter cells through the ...

5-Ethynyl-2'-Deoxyuridine/Phospho-Histone H3 Dual-Labeling Protocol for Cell Cycle Progression Analysis in Drosophila Neural Stem Cells

Summary

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Chapters in this video

Studying Cell Cycle-regulated Gene Expression by Two Complementary Cell Synchronization Protocols

Use of Drosophila S2 Cells for Live Imaging of Cell Division

Assessing Cell Cycle Progression of Neural Stem and Progenitor Cells in the Mouse Developing Brain after Genotoxic Stress

Analysis of Cell Cycle Position in Mammalian Cells

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

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

Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization

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

Assessment of Global DNA Double-Strand End Resection using BrdU-DNA Labeling coupled with Cell Cycle Discrimination Imaging

Cell Cycle Analysis