This work was supported by an EMBO Long-term fellowship (ABW), a National Science Foundation International Postdoctoral Research Fellowship (ABW), the Max Planck Gesellschaft (ABW, DS, JS, ACO), a DIGS-BB fellowship (AO), and the European Research Council under the European Communities Seventh Framework Programme STG-207634 (DS, ACO). We thank Ravi Desai for helpful comments on the manuscript. We would also like to thank the MPI-CBG protein expression facility for production of the zebrafish Fibronectin1 fragment, the MPI-CBG fish facility staff for care and maintenance of our fish lines and the MPI-CBG light microscopy facility for imaging support.

1. Before Dissection
<ol>
	<li>On the day before dissection, obtain embryos from an incross of zebrafish pairs heterozygous for the transgenic allele.
	<ol>
		<li>Raise embryos in E3 medium without methylene blue at 28 oC until shield stage (6 hr post-fertilization).</li>
		<li>Transfer embryos in E3 medium without methylene blue to 20 oC O/N. At 20 oC, embryos will form 1 somite per hr once they reach tailbud stage. After 17-20 hr O/N at 20 oC, embryos should be at 5 to 8 somite stage the following morning at the beginning of the dissection protocol.</li>
	</ol>
	</li>
	<li>Assemble tools and reagents needed for dissection.
	<ol>
		<li>Fire-polished glass pipette for transfer of embryos and tissue</li>
		<li>Pair of fine forceps for removal of chorions from embryos</li>
		<li>Sylgard-coated 35-mm dish for dissection - Pour Sylgard polymer into a 35 mm dish and cure O/N in a 37 &#176;C. Make the dissection well using a needle tip to remove a small volume of cured polymer. The dish can be cleaned and subsequently reused.</li>
		<li>Sharpened, flattened tungsten wire tools for PSM tissue manipulation</li>
		<li>Micro-scalpel for cutting desired pieces of PSM to culture</li>
		<li>35-mm plastic dish for trypsin incubation</li>
		<li>L15 medium containing 10% fetal bovine serum</li>
		<li>0.25% Trypsin/EDTA solution</li>
		<li>Sigmacoted gel-loading tips for dispersal</li>
		<li>Plastic petri dishes containing E3 medium without methylene blue.</li>
	</ol>
	</li>
	<li>Cover glass-bottom imaging dish with Fibronectin1 substrate (10 &#956;g/ml in PBS). Leave dish on bench to coat during dissection.</li>
	<li>Use stereoscope with appropriate fluorescence filters to identify and sort positive transgenic embryos (5 to 8 somite stage).
	<ol>
		<li>Identify transgenic embryos by examining the presomitic mesoderm (PSM) for YFP expression under fluorescence channel (Figure 1A). Fluorescence should be visible in the region from the last formed somite to the tailbud. Select embryos with brightest signal; 25% of offspring should be homozygous embryos carrying 2 copies of the transgene. The number of identified embryos needed varies based on the experiment. One dissected tailbud piece will yield 1,000 cells, on average. Typically, a few additional positive embryos are useful in case errors are made during the dissection.</li>
		<li>Use transmitted light as a positional reference to distinguish any autofluorescence, particularly in the yolk cell, from signal.</li>
		<li>Transfer positive embryos to a separate plastic petri dish containing E3 medium without methylene blue.</li>
	</ol>
	</li>
</ol>
2. PSM Dissection and Dispersal
<ol>
	<li>Prepare embryos for dissection.
	<ol>
		<li>Under a dissection microscope, using fine forceps, carefully remove the chorion from each embryo in the E3 medium. Be sure not to damage the embryo or disrupt the yolk cell.</li>
		<li>Fill Sylgard-coated dissecting dish with L15 medium with serum. Using forceps or another flat tool, remove any air bubbles from the surface of the Sylgard.</li>
		<li>Transfer dechorionated embryos using the fire-polished glass pipette to the dissection dish.</li>
		<li>Using the wire tools, move all embryos to one side of the dissecting dish.</li>
	</ol>
	</li>
	<li>Dissect PSM from a single embryo.
	<ol>
		<li>Orient a single embryo on its lateral side in the small well made in the Sylgard layer within the dissecting dish (Figure 1B).</li>
		<li>Using the micro-scalpel, slice through the embryo and yolk cell just anterior to the hindbrain and through the ventral pole of the embryo (Figure 1B, dotted red line).</li>
		<li>Remove the anterior piece of the embryo from the well, moving it away to the side of the dish, and then use the wire tools to scrape away remaining yolk cell granules from the posterior section including the PSM.</li>
		<li>Once the yolk has been scraped away, flatten and orient the PSM with the anterior end pointing away from the experimenter and the posterior pointing towards the experimenter (Figure 1C).</li>
		<li>If the thin layer of ectoderm has not pulled off by this point, use the wire tools to peel it away from the top of the PSM tissue.</li>
		<li>Using the micro-scalpel cut the tailbud, the most posterior tip of the PSM past the end of the notochord, away from the rest of the tissue (Figure 1C, dotted red line). NOTE: Other pieces of tissue from the PSM, for example anterior PSM closer to the last formed somite, can also be taken into culture, depending on the experimental question.</li>
		<li>Move the tailbud piece to one corner of the dish away from the dissecting area. Use the wire tools to clear any debris and unwanted embryo tissue from the dissecting field.</li>
		<li>Repeat with the next embryo.</li>
	</ol>
	</li>
	<li>Pool tailbud pieces from multiple embryos, depending on how many cells will be required for the experiment. On average, a single piece of tailbud tissue yields 1,000 cells.</li>
	<li>Fill empty 35-mm plastic dish with a small volume of trypsin-EDTA.</li>
	<li>Using the fire-polished glass pipette, transfer tailbud pieces from dissecting dish into dish containing trypsin/EDTA. Incubate tailbud pieces in trypsin/EDTA for 20 min at RT.</li>
	<li>While the tissue is being incubated in trypsin, remove the Fibronectin1 solution from the glass-bottomed imaging dish.
	<ol>
		<li>Wash the solution from glass 3 times with MilliQ water. Use suction to remove each wash and ensure the dish is completely dry.</li>
	</ol>
	</li>
	<li>Disperse tailbud pieces in medium for imaging.
	<ol>
		<li>Pipette 100&#160;&#956;l of L15 medium with serum into imaging dish.</li>
		<li>Using a coated gel tip, remove the tailbud pieces from the trypsin/EDTA in as small a volume as possible.</li>
		<li>Pipette the tailbud pieces into the medium in the imaging dish. Pipette the pieces up and down multiple times to break them apart and suspend cells in the medium. Be careful not to introduce air bubbles. Check for cell clumps under the microscope and disperse more as needed.</li>
		<li>Allow dispersed cells in the suspension to settle onto the Fibronectin1-coated glass for 20 min at RT.</li>
	</ol>
	</li>
	<li>Add a small volume of additional L15 medium with serum to the cells before beginning imaging. Use care not to disrupt settled cells.</li>
	<li>Given the experimental question, add any supplemental or drug treatments to the culture.</li>
</ol>
3. Imaging of Dispersed PSM cells
<ol>
	<li>Set up imaging dish in temperature chamber on the time-lapse imaging microscope. Set Warner chamber to desired temperature for the experiment. Allow dish to equilibrate for at least 30 min before beginning image acquisition. Crosscheck temperature with external temperature probe placed into dish, if required. NOTE: Due to the relationship between temperature and somitogenesis rate 21 stable temperature control is essential for accurate measurements of period in single PSM cells.</li>
	<li>Acquire test images in the fluorescence channel to check that the exposure time and gain provide a large dynamic range of intensities without saturation to ensure good signal to noise levels. A typical fluorescence image acquired from dispersed cells generated from our lines using this protocol requires 400 msec and 40 msec for a transmitted light image with an EMCCD camera operating at an EM Gain of 85. In addition, pre-amp gain and readout speed from the camera are also essential to maximize signal over noise.</li>
	<li>Using the transmitted light channel, choose fields of cells for the time-lapse acquisition.</li>
	<li>Run time-lapse acquisition protocol for desired length of time.
	<ol>
		<li>Acquire one transmitted light and one fluorescence image per field. NOTE: Set an interval between acquisition rounds that will capture temporal dynamics without photo-bleaching over extended imaging or inducing toxicity in the cells. This protocol uses a 2 min interval.</li>
	</ol>
	</li>
	<li>Check time-lapse set-up occasionally during recording to ensure that cells remain in focus, no software or hardware errors, etc.</li>
</ol>
4. Image Processing of Acquired Time-lapse Movies
<ol>
	<li>Open movie file for an acquired field in an image processing software. NOTE: Fiji was used for all processing in this protocol.
	<ol>
		<li>Split transmitted light and fluorescence frames from one field to create 2 stacks of images.</li>
	</ol>
	</li>
	<li>Track a single cell in the transmitted light channel.
	<ol>
		<li>Place a circular ROI (region of interest) around the selected cell in the first frame in the transmitted light channel. NOTE: Only cells that 1. are healthy at the end of the recording, 2. do not move outside of the field, and 3. do not come into contact with other cells are tracked.</li>
		<li>Save an ROI every few frames to the ROI manager. Track cell until the last frame.</li>
	</ol>
	</li>
	<li>Measure intensity using the saved ROIs on the fluorescence channel.
	<ol>
		<li>Select the fluorescence stack and with the saved ROIs, use the custom circle interpolator plug-in and macro to measure the intensity of the tracked cell over time.</li>
		<li>Check the output trace from the macro. If the trace qualitatively captures features of the fluorescence time-lapse, export the values to an Excel worksheet.</li>
		<li>Save the ROI list for the cell.</li>
	</ol>
	</li>
	<li>Repeat tracking in transmitted channel and measuring in fluorescence channel for other cells in the field.</li>
	<li>Repeat all steps for additional fields from the experiment.</li>
</ol>

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Author Contributions: 
ABW developed and refined the dispersal, culture, imaging, and cell tracking protocols. DS generated the transgenic lines and oversaw the design of the time-lapse fluorescence microscope used in this protocol. AO pioneered initial proof-of-principle experiments to dissociate and image PSM cells in culture. JS wrote the circle interpolator plugin tool in Fiji used to measure fluorescence intensity from time-lapse movies. ABW and ACO wrote the manuscript.

To study a cellular process that takes place over the course of embryonic development, biologists typically utilize an approach in the context of the entire embryo. However, to fully understand how a single cell behaves within a developmental time frame, a method to examine and perturb individual cells in isolation is also highly beneficial. By generating dispersed PSM cell cultures using transgenic zebrafish embryos we now have a tool to directly study the cell autonomous nature of genetic oscillations in segmentation clock in a quantitative way. It is possible to measure dynamics of the oscillations in our fluorescent reporter in hundreds of cells under a variety of conditions.
The observed period of single cells in culture is longer than the somitogenesis period in the intact embryo. We observe that an intact PSM explant in culture also exhibits slower oscillations than the intact embryo (data not shown), suggesting that the longer period observed in isolated cells is not simply due to damage from dispersal. A variable period and amplitude is observed in most of our single-cell time series. The source of this variability is unknown, but should reveal important details about the segmentation clock's pace-making circuits.
With this method we can study the genetic components of segmentation at the cellular level, and address questions that are challenging to examine in the whole embryo. For example, by controlled addition of known signaling molecules found in the developing embryo to our cultures, we can examine their effects on the single PSM cellular oscillator in a robust and reproducible assay. Our PSM culture system opens the door to a rigorous evaluation of what factors, alone, or in combination promote oscillations in these cells, what factors inhibit these oscillations, and to test the interplay between such molecules. With these tools in hand, we aim to evaluate existing models of somitogenesis that are based on published tissue-level data, as well as use results in single cells to generate predictions that can be tested in the whole embryo. 
To our knowledge, this is the first zebrafish primary cell culture protocol for acute time-lapse recording of fluorescence in single cells; further development and refinement of this protocol is no doubt possible. Other protocols often use zebrafish embryos to generate stable cell lines that can be transfected with reporters and used for long-term imaging 27-29. While stable lines are useful for imaging a process that is not tied to development timing, such as the circadian clock, the question of embryonic segmentation necessitates immediate imaging while cells are still in their oscillatory, progenitor state. Once cells stop oscillating, they assume a differentiated cell fate, and when in the tissue, would be incorporated into a somite. It is possible that with the right factor or factors present in vitro we could generate PSM-like zebrafish cell culture lines that would remain oscillatory, enabling significantly longer-term observations, multiple sequential perturbations, or high-throughput screening.
We expect that this method of preparing primary cultures of dispersed cells for time-lapse imaging is well-suited for the study of any cellular process that occurs within a developmental time frame that is not approachable using stable zebrafish cell lines. The isolation of distinct cell-types at distinct developmental stages from the appropriate transgenic reporter lines, either via dissection or via FACS after embryonic dissociation, could provide the starting cells. Some optimization of culture conditions, guided by the embryonic origin of the cells, may be required. By combining this flexible and sensitive protocol with the rapidly growing collection of transgenic zebrafish lines, we hope to facilitate an in vitro developmental biology approach that is complementary to classical genetic and embryological methods.

The periodic formation of segments along the vertebrate body axis, or somitogenesis, is evidence of a spatial and temporal oscillator in the developing embryo. The favored mechanism controlling somitogenesis is conceptually described by a &#34;clock and wavefront&#34; model 1, wherein the &#34;clock&#34; consisting of cellular oscillators, now thought to be intracellularly driven by the rhythmic expression of a set of cyclic genes2, ticks off the formation of somites from the presomitic mesoderm (PSM). As the embryo develops, a maturation &#34;wavefront&#34; in the PSM moves in concert with the regressing tissue towards the posterior, slowing and arresting cellular oscillators as it passes 3. Together, this spatiotemporally dynamic system is termed the segmentation clock. Current approaches to study the segmentation clock span three increasing levels of organization from the genetic oscillator in single cells to local coupling that occurs between cells and finally to global regulation of positional information in the collective PSM tissue 4.
Previous studies suggest the cell-autonomous segmentation oscillator in zebrafish consists of genes and protein products from the her/hes transcription factorfamily, which are thought to form a negative feedback loop through transcriptional repression 5-7. The Delta/Notch signaling pathway synchronizes oscillations between neighboring cells and regulates the collective period of the population 8-10. Fgf signaling molecules produced in the tailbud appear to build a gradient across the zebrafish PSM, and are thereby hypothesized to contribute to slowing and arresting oscillating cells in the anterior 11. Until now, the functional roles of each of these molecules in somitogenesis have been investigated by genetic mutation, morpholino injection, heat-shock over-expression, and antagonist drug treatment of clock components and signaling between cells5,7,10,12 . Using these perturbations, segmentation clock function has been inferred from tissue-level descriptions of somite defects and the loss of uniform oscillations in expression of cyclic genes like her1, her7, and deltaC. However, nearly all of these data are from fixed embryos and fail to accurately capture changes that are inherent to the dynamic function of the segmentation clock. More recently, multiple-embryo time-lapse imaging has revealed the first mutants with altered oscillator period, but these observations were also made at the tissue level 7,13 . Thus, behavior of the hypothesized cell-autonomous oscillator during somitogenesis has not been observed.
Static snapshots of somitogenesis give an incomplete picture because, inherently, the process is driven by an oscillatory system. Earlier work in mouse and chick cells showed that levels of transcript and protein rise and fall, but sampling an approximately 2 hr oscillation every 30 or 45 min necessarily restricts the data collected and therefore the conclusions that can be drawn14,15 . Study of other biological oscillators, most notably, circadian clocks, has moved away from staged measurements of gene and protein expression to real-time monitoring using fluorescent and bioluminescent reporters 16,17. These tools are essential for demonstrating clock properties of single cells 18. A bioluminescent reporter of the Hes1 cyclic gene has been developed and briefly characterized in single mouse PSM cells 19. The average period and variance were calculated for a small number of cells, showing that oscillations persist for several cycles in vitro. However, these studies did not quantitatively address the stability and robustness of oscillator frequency and amplitude, whether cells can spontaneously enter or exit oscillations, and how the cells maintain their phase relations. Additionally, the effects of signaling molecules found in the embryo on the cell-autonomous clock have not been directly tested. Consequently, these fundamental properties of the single cell oscillator remain completely unknown.
We have recently developed transgenic fish lines using BAC recombineering 20 to drive Venus (YFP) fluorescence reporters of her1 expression30. Such lines exploit the regulatory cues of the intact chromosomal locus in which they are embedded and recapitulate the temporal dynamics and spatial pattern of her1. This breakthrough allows for real-time monitoring of gene expression in the developing zebrafish embryo in vivo. To study the fundamental properties of cell-autonomous oscillations and how such expression is regulated over time, we recently developed a reliable method to isolate and record from PSM cells in vitro. This protocol describes how we have utilized our transgenic reporter lines to generate dispersed cell cultures, from which we can characterize the oscillations of the zebrafish segmentation clock in single cells. We can thereby tackle outstanding questions in the field that were not accessible with static or tissue-level analysis, as well as directly manipulate the segmentation clock at the single cell level with signaling molecules and inhibitors.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Epifluorescence microscope</td>
 <td>Olympus</td>
 <td>Model: SZX16</td>
 <td ></td>
 </tr>
 <tr>
 <td>Epifluorescence microscope</td>
 <td>X-cite Illumination</td>
 <td>Series: 120Q</td>
 <td ></td>
 </tr>
 <tr>
 <td>Dissection microscope</td>
 <td>Olympus</td>
 <td>Model: SZX12</td>
 <td></td>
 </tr>
 <tr>
 <td>Fine forceps no. 55</td>
 <td>Fine Science Tools</td>
 <td>11295-51</td>
 <td></td>
 </tr>
 <tr>
 <td>Glass transfer pipettes</td>
 <td>Assistent</td>
 <td>567/2</td>
 <td></td>
 </tr>
 <tr>
 <td>35 mm plastic petri dishes</td>
 <td>Greiner</td>
 <td>627102</td>
 <td></td>
 </tr>
 <tr>
 <td>60 mm plastic petri dishes</td>
 <td>Greiner</td>
 <td>628102</td>
 <td></td>
 </tr>
 <tr>
 <td>Sylgard polymer</td>
 <td>SASCO</td>
 <td>266727</td>
 <td></td>
 </tr>
 <tr>
 <td>Manipulation tools</td>
 <td>Made in-house</td>
 <td></td>
 <td>For description of manipulation tools Ref. 22</td>
 </tr>
 <tr>
 <td>Microsurgical knife</td>
 <td>World Precision Instruments</td>
 <td>500249</td>
 <td></td>
 </tr>
 <tr>
 <td>L15 medium</td>
 <td>Invitrogen</td>
 <td>57322</td>
 <td></td>
 </tr>
 <tr>
 <td>Penicillin/streptomyocin</td>
 <td>PAA</td>
 <td>P11-010</td>
 <td></td>
 </tr>
 <tr>
 <td>Fetal bovine serum</td>
 <td>Invitrogen</td>
 <td>257322</td>
 <td></td>
 </tr>
 <tr>
 <td>0.05% trypsin / 0.02% EDTA</td>
 <td>PAA </td>
 <td>L11-004</td>
 <td></td>
 </tr>
 <tr>
 <td>Gel loading tips</td>
 <td>Fisher Scientific</td>
 <td>253188</td>
 <td></td>
 </tr>
 <tr>
 <td>Sigmacote</td>
 <td>Sigma Aldrich</td>
 <td>254589</td>
 <td></td>
 </tr>
 <tr>
 <td>Micropipette set</td>
 <td>Gilson International</td>
 <td>F167300</td>
 <td></td>
 </tr>
 <tr>
 <td>Zebrafish Fibronectin1 70 kD fragment</td>
 <td>MPI-CBG protein facility</td>
 <td></td>
 <td>Generated in-house from construct based on previously published work Ref. 23-25</td>
 </tr>
 <tr>
 <td>Glass bottom imaging dishes</td>
 <td>Mattek (single well)</td>
 <td>P35G-1.5-14-C</td>
 <td></td>
 </tr>
 <tr>
 <td>Glass bottom imaging dishes</td>
 <td>Greiner (CellView -multi-well)</td>
 <td>262502</td>
 <td></td>
 </tr>
 <tr>
 <td>E3 medium without methylene blue</td>
 <td>Made in-house</td>
 <td></td>
 <td>From The Zebrafish Book, 5th Ed. Ref. 26 </td>
 </tr>
 <tr>
 <td>Plastic transfer pipettes</td>
 <td>Ratiolab</td>
 <td>260011</td>
 <td></td>
 </tr>
 <tr>
 <td>Warner heating/cooling chamber</td>
 <td>Warner Instruments</td>
 <td>TC-324B/344B</td>
 <td></td>
 </tr>
 <tr>
 <td>EM-CCD camera</td>
 <td>Andor</td>
 <td>Model: iXOn 888</td>
 <td></td>
 </tr>
 <tr>
 <td>Wide-field fluorescence microscope with Venus filter set</td>
 <td>Zeiss</td>
 <td>Model: Axiovert 200M</td>
 <td></td>
 </tr>
 <tr>
 <td>Wide-field fluorescence microscope with Venus filter set</td>
 <td></td>
 <td>NeoFluor 40x, NA 0.75</td>
 <td></td>
 </tr>
 <tr>
 <td>Wide-field fluorescence microscope with Venus filter set</td>
 <td>Lumencor Light Engine</td>
 <td>Model: Spectra X</td>
 <td></td>
 </tr>
 <tr>
 <td>Wide-field fluorescence microscope with Venus filter set</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Wide-field fluorescence microscope with Venus filter set</td>
 <td>BrightLine HC 575/15</td>
 <td>F39-575</td>
 <td></td>
 </tr>
 </tbody>
</table>

generation of dispersed presomitic mesoderm cell cultures for imaging of the zebrafish segmentation clock in single cells

Segmentation is a periodic and sequential morphogenetic process in vertebrates. This rhythmic formation of blocks of tissue called somites along the body axis is evidence of a genetic oscillator patterning the developing embryo. In zebrafish, the intracellular clock driving segmentation is comprised of members of the Her/Hes transcription factor family organized into negative feedback loops. We have recently generated transgenic fluorescent reporter lines for the cyclic gene her1 that recapitulate the spatio-temporal pattern of oscillations in the presomitic mesoderm (PSM). Using these lines, we developed an in vitro culture system that allows real-time analysis of segmentation clock oscillations within single, isolated PSM cells. By removing PSM tissue from transgenic embryos and then dispersing cells from oscillating regions onto glass-bottom dishes, we generated cultures suitable for time-lapse imaging of fluorescence signal from individual clock cells. This approach provides an experimental and conceptual framework for direct manipulation of the segmentation clock with unprecedented single-cell resolution, allowing its cell-autonomous and tissue-level properties to be distinguished and dissected.

This protocol produces cultures of viable, dispersed, single PSM cells for time-lapse imaging of fluorescence signal (Figure 2). Our transgene generates a reporter whose cycle of production and degradation occurs with similar dynamics to the endogenous gene and protein in the embryo, on the order of half an hour. Due to its fast turnover, the YFP signal in single cells should be detected quickly to minimize bleaching and with a high temporal resolution to capture the features of each oscillatory cycle. Also, given the relative dimness of the signal, culture and acquisition conditions have been carefully tuned to ensure sensitive and robust results. We have found the following factors to be important in generating optimal PSM cell cultures for imaging: 1. An ECM substrate used for coating the glass-bottom culture dishes. 2. Addition of serum to the dissection and imaging medium. 3. Dissection of PSM from identified embryos taken after mating heterozygous pairs, whose offspring potentially carry 2 copies of the transgene. 4. Image acquisition within an optimal magnification range, using a higher NA objective to ensure efficient capture of the fluorescence signal. 5. Illumination with a solid-state light source to minimize intensity fluctuations that could contribute to background noise. 6. Signal detection with a highly sensitive EM-CCD camera to maximize signal readout.
Sub-optimal cultures will contain cells that do not remain rounded and healthy throughout the recording. We hypothesized that our ECM substrate, a fragment of zebrafish fibronectin1 keeps cells in an undifferentiated PSM-like state, in comparison to other commonly used substrates like poly-lysine or laminin. Other substrates we tested caused cells to flatten on the glass and the loss of oscillating fluorescent signal over the course of the recording. We also found that the addition of serum to the medium during dissection, dispersal, and recording was not only important to quench the trypsin used for dissociation, but also increased fluorescence intensity over background, likely due to improved cell viability. To ensure optimal signal capture from single cells we used a 40x objective specifically designed for fluorescence imaging (Zeiss Plan NeoFluor series) with a high NA. We also found that a solid-state light source provided more stable illumination than traditional mercury lamps, which is critical to minimize background fluctuations that contribute to noisy images. These modifications are important to ensure robust results.
Using this protocol we expect cultures in which the majority of cells in a given field are fluorescent at some point during the recording. We find that fluorescent cells typically remain rounded and are sometimes quite motile during recordings. A few cells, including fluorescent cells, may become apoptotic during the recording. These cells are excluded from any analysis. We also exclude cells that come into contact with other cells or move outside of the field of view. On average, we see a 12% reduction in cell number per field by the end of the 10 hr recording as measured by counting the number of healthy cells in the transmitted light channel. This loss includes both cell death and cells that have moved out of the field. Given these caveats we can, on average, track 5 fluorescent cells per field that remain viable and visible, and typically record 6 fields per condition. For example, an experiment with 4 conditions will have 24 total fields acquired and typically about 30 tracked cells per condition. Under standard imaging conditions with L15 medium containing 10% fetal bovine serum, we find that PSM cells (n=101 cells from 4 experimental replicates) can produce between 2 and 7 peaks, with the mean and standard deviation of 3&#177;1 peaks (2-3 cycles). The median peak number, as well as the 25% percentile, is 2 peaks and the 75% percentile is 3 peaks. Using our semi-automated tracking and analysis, we can quickly generate fluorescence intensity over time traces for individual PSM cells, with a representative cell trace shown in Figure 2C. These raw traces can then be used to make quantitative measurements of properties of the oscillating PSM cells, such as frequency, amplitude, number of cycles, and timing of peaks.
<img alt="Figure 1" fo:content-width="4in" fo:src="/files/ftp_upload/50307/50307fig1highres.jpg" src="/files/ftp_upload/50307/50307fig1.jpg" /> 
Figure 1. Identification of transgenic embryos and schematics of their dissection. (A) Lateral view of a transgenic zebrafish embryo expressing YFP fluorescence at ~ 5-somite stage. Image is an overlay of transmitted light and fluorescence channel. Arrows indicate areas of signal starting at the tip of the tailbud, throughout the PSM, towards the last formed somite. Inset shows a schematic of the reporter transgene (for more information on its development and in vivo behavior refer to Soroldoni et al.30. (B) Schematic of lateral view of embryo prior to first cut during dissection. Dotted red line indicates the first cut made across the hindbrain, though the yolk cell just behind the tailbud. (C) Schematic view of flattened PSM following removal of yolk granules and epidermal layer, oriented along its anterior-posterior axis. Dotted red line indicates the second cut to remove the tip of the tailbud for dispersal and culture. <a href="https://www.jove.com/files/ftp_upload/50307/50307fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="4in" fo:src="/files/ftp_upload/50307/50307fig2highres.jpg" src="/files/ftp_upload/50307/50307fig2.jpg" /> 
Figure 2. Representative images and traces from a single cell in a dispersed PSM culture. (A) Montage of transmitted light images of a single PSM cell in a 6-h time-lapse recording. Note that the cell remains rounded and healthy throughout the course of the recording. (B) Corresponding montage of fluorescence images from a single PSM cell. Peaks in intensity of the cell are numbered over the course of the recording. (C) Average intensity over time measured from an ROI placed on this cell. Values are taken from every frame from a movie with a frame rate of one per 2 min using the circle interpolator plug-in and custom macro written in Fiji. Peaks in intensity are again numbered throughout the trace. <a href="https://www.jove.com/files/ftp_upload/50307/50307fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Generation of Dispersed Presomitic Mesoderm Cell Cultures for Imaging of the Zebrafish Segmentation Clock in Single Cells at JoVE.com

Generation of Dispersed Presomitic Mesoderm Cell Cultures for Imaging of the Zebrafish Segmentation Clock in Single Cells

Somitogenesis is a rhythmic developmental process that spatially patterns the body axis of vertebrate embryos. Previously, we developed transgenic zebrafish lines that use fluorescent reporters to observe the cyclic genes that drive this process. Here, we culture dispersed cells from these lines and image their oscillations over time in vitro.

Segmentation is a  periodic and sequential morphogenetic process in vertebrates. This rhythmic  formation of blocks of tissue called somites along the ...

generation-dispersed-presomitic-mesoderm-cell-cultures-for-imaging

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13.7K Views.  Max Planck Institute of Molecular Cell Biology and Genetics. Somitogenesis is a rhythmic developmental process that spatially patterns the body axis of vertebrate embryos. Previously, we developed transgenic zebrafish lines that use fluorescent reporters to observe the cyclic genes that drive this process. Here, we culture dispersed cells from these lines and image their oscillations over time in vitro.

Video: Generation of Dispersed Presomitic Mesoderm Cell Cultures for Imaging of the Zebrafish Segmentation Clock in Single Cells

Live Imaging of Cell Extrusion from the Epidermis of Developing Zebrafish

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

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

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

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...

Generation of Mice Derived from Induced Pluripotent Stem Cells

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also produce a source of patient-matched cells for regenerative therapies. iPSCs may be generated using multiple protocols and derived from multiple cell sources. Once generated, iPSCs are tested using a variety of assays including immunostaining for pluripotency markers, generation of three germ layers in embryoid bodies and teratomas, comparisons of gene expression with embryonic stem cells (ESCs) and production of chimeric mice with or without germline contribution2. Importantly, iPSC lines that pass these tests still vary in their capacity to produce different differentiated cell types2. This has made it difficult to establish which iPSC derivation protocols, donor cell sources or selection methods are most useful for different applications.
The most stringent test of whether a stem cell line has sufficient developmental potential to generate all tissues required for survival of an organism (termed full pluripotency) is tetraploid embryo complementation (TEC)3-5. Technically, TEC involves electrofusion of two-cell embryos to generate tetraploid (4n) one-cell embryos that can be cultured in vitro to the blastocyst stage6. Diploid (2n) pluripotent stem cells (e.g. ESCs or iPSCs) are then injected into the blastocoel cavity of the tetraploid blastocyst and transferred to a recipient female for gestation (see Figure 1). The tetraploid component of the complemented embryo contributes almost exclusively to the extraembryonic tissues (placenta, yolk sac), whereas the diploid cells constitute the embryo proper, resulting in a fetus derived entirely from the injected stem cell line.
Recently, we reported the derivation of iPSC lines that reproducibly generate adult mice via TEC1. These iPSC lines give rise to viable pups with efficiencies of 5-13%, which is comparable to ESCs3,4,7 and higher than that reported for most other iPSC lines8-12. These reports show that direct reprogramming can produce fully pluripotent iPSCs that match ESCs in their developmental potential and efficiency of generating pups in TEC tests. At present, it is not clear what distinguishes between fully pluripotent iPSCs and less potent lines13-15. Nor is it clear which reprogramming methods will produce these lines with the highest efficiency. Here we describe one method that produces fully pluripotent iPSCs and "all- iPSC" mice, which may be helpful for investigators wishing to compare the pluripotency of iPSC lines or establish the equivalence of different reprogramming methods.

Generating induced pluripotent stem cell (iPSC) lines produces lines of differing developmental potential even when they pass standard tests for pluripotency. Here we describe a protocol to produce mice derived entirely from iPSCs, which defines the iPSC lines as possessing full pluripotency1.

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also ...

Monitoring Cell-autonomous Circadian Clock Rhythms of Gene Expression Using Luciferase Bioluminescence Reporters

In mammals, many aspects of behavior and physiology such as sleep-wake cycles and liver metabolism are regulated by endogenous circadian clocks (reviewed1,2). The circadian time-keeping system is a hierarchical multi-oscillator network, with the central clock located in the suprachiasmatic nucleus (SCN) synchronizing and coordinating extra-SCN and peripheral clocks elsewhere1,2. Individual cells are the functional units for generation and maintenance of circadian rhythms3,4, and these oscillators of different tissue types in the organism share a remarkably similar biochemical negative feedback mechanism. However, due to interactions at the neuronal network level in the SCN and through rhythmic, systemic cues at the organismal level, circadian rhythms at the organismal level are not necessarily cell-autonomous5-7. Compared to traditional studies of locomotor activity in vivo and SCN explants ex vivo, cell-based in vitro assays allow for discovery of cell-autonomous circadian defects5,8. Strategically, cell-based models are more experimentally tractable for phenotypic characterization and rapid discovery of basic clock mechanisms5,8-13.
Because circadian rhythms are dynamic, longitudinal measurements with high temporal resolution are needed to assess clock function. In recent years, real-time bioluminescence recording using firefly luciferase as a reporter has become a common technique for studying circadian rhythms in mammals14,15, as it allows for examination of the persistence and dynamics of molecular rhythms. To monitor cell-autonomous circadian rhythms of gene expression, luciferase reporters can be introduced into cells via transient transfection13,16,17 or stable transduction5,10,18,19. Here we describe a stable transduction protocol using lentivirus-mediated gene delivery. The lentiviral vector system is superior to traditional methods such as transient transfection and germline transmission because of its efficiency and versatility: it permits efficient delivery and stable integration into the host genome of both dividing and non-dividing cells20. Once a reporter cell line is established, the dynamics of clock function can be examined through bioluminescence recording. We first describe the generation of P(Per2)-dLuc reporter lines, and then present data from this and other circadian reporters. In these assays, 3T3 mouse fibroblasts and U2OS human osteosarcoma cells are used as cellular models. We also discuss various ways of using these clock models in circadian studies. Methods described here can be applied to a great variety of cell types to study the cellular and molecular basis of circadian clocks, and may prove useful in tackling problems in other biological systems.

Circadian clocks function within individual cells, i.e., they are cell-autonomous. Here, we describe methods for generating cell-autonomous clock models using non-invasive, luciferase-based real-time bioluminescence technology. Reporter cells provide tractable, functional model systems for studying circadian biology.

In mammals, many aspects of behavior and physiology such as sleep-wake cycles and liver metabolism are regulated by endogenous circadian clocks ...

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  ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Efficient Generation Human Induced Pluripotent Stem Cells from Human Somatic Cells with Sendai-virus

A few years ago, the establishment of human induced pluripotent stem cells (iPSCs) ushered in a new era in biomedicine. Potential uses of human iPSCs include modeling pathogenesis of human genetic diseases, autologous cell therapy after gene correction, and personalized drug screening by providing a source of patient-specific and symptom relevant cells. However, there are several hurdles to overcome, such as eliminating the remaining reprogramming factor transgene expression after human iPSCs production. More importantly, residual transgene expression in undifferentiated human iPSCs could hamper proper differentiations and misguide the interpretation of disease-relevant in vitro phenotypes. With this reason, integration-free and/or transgene-free human iPSCs have been developed using several methods, such as adenovirus, the piggyBac system, minicircle vector, episomal vectors, direct protein delivery and synthesized mRNA. However, efficiency of reprogramming using integration-free methods is quite low in most cases.
Here, we present a method to isolate human iPSCs by using Sendai-virus (RNA virus) based reprogramming system. This reprogramming method shows consistent results and high efficiency in cost-effective manner.

Here, we present our established method to reprogram human somatic cells into transgene-free human iPSCs with Sendai virus, which shows consistent outcome and enhanced efficiency.

A few years ago, the establishment of human induced pluripotent stem cells (iPSCs) ushered in a new era in biomedicine. Potential uses of human iPSCs ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...