The authors have nothing to disclose.

1. Preparation of C. elegans Embryos for Time-lapse Microscopy using Microbeads
<ol>
	<li>Transfer gravid adult worms by using a worm pick (platinum wire) into a drop of M9 buffer and clean off bacteria by first vigorously agitating and then transferring each worm to an adjacent drop of M9 using an eye lash mounted onto a tooth pick/pipette tip or use a pointed glass capillary.</li>
	<li>Dilute the dispersion containing the 20 &#181;m diameter polystyrene beads 10-fold in M9 buffer (1 L M9 buffer contains 3 g KH2PO4, 6 g Na2HPO4, 5 g NaCl, and, after autoclaving (20 min, 120 &#176;C), add an additional 1 ml of a 1 M MgSO4 solution).</li>
	<li>Make a mouth pipette from any type of thin glass capillary (e.g., melting point capillaries or capillaries used to pull injection needles), a roughly 0.5 m long piece of rubber, silicone or similar material tubing (ideally transparent), a 200 &#181;l pipette tip, the lid of a 200 &#181;l PCR tube, a hydrophobic filter for small syringes (e.g., a 0.2-0.45 &#181;m pore size and 15-50 mm diameter Nylon, PTFE or similar material filter), and a 1000 &#181;l pipette tip. This type of mouth pipette ensures that no biological material can get in contact with the experimenter.</li>
	<li>Assemble the pipette by inserting the 200 &#181;l pipette tip with the tip first into the tubing and plug it with the lid of a PCR tube which has a central perforation that matches the diameter of the capillary used. A very pointed awl or preparation needle can be used to generate the perforation.</li>
	<li>Draw a thin capillary and insert it into the perforation. Insert the filter into the other end of the tubing and insert a 1000 &#181;l pipette tip as mouth piece (Figure 1).</li>
	<li>Dissect gravid adults by cutting worms transversely at the vulva with a clean razor blade or scalpel. 
	NOTE: Usually, many embryos will be expulsed after cutting. However, if early embryos are needed, they sometimes have to be released from the carcass using an eye lash or pointed needle.</li>
	<li>Pipette one drop of the diluted polystyrene bead dispersion (~4 &#181;l) onto a 24x60 mm cover glass and transfer embryos into this drop using the mouth pipette.</li>
	<li>Carefully place a small (e.g.,&#160;18x18 or 20x20 mm) cover glass on top of the drop and make sure the drop has fully spread out. 
	NOTE: The drop should also be small enough not to touch the edges of the small cover glass.</li>
	<li>Seal the mount with liquefied vaseline. Add a drop of immersion oil on the 24x60 mm cover slip side and proceed to the &#39;Time-lapse Microscopy&#39; section.</li>
</ol>
2. Preparation of Adult C. elegans Animals for Time-lapse Microscopy using Nanobeads
NOTE: In general, the protocol for mounting adult animals is an adaptation of the protocol from ref. 18. With this protocol, it is possible to perform long-term time-lapse confocal microscopy of adult animals.
<ol>
	<li>Prepare a 1.5% (w/v) agarose solution in M9 buffer, boil (1 min, 100 &#176;C in water bath or microwave). The use of higher concentrations of agarose is not recommended since this can lead to extrusion of the intestine and/or parts of the gonad.</li>
	<li>Stick down 2-3 layers of tape onto two microscope slides each. Place a slide without tape between the two slides with tape.</li>
	<li>Use a pipette tip and place one drop of the hot agarose solution in the center of the middle slide. Quickly place another slide onto the drop so that it is supported by the tape layers on each side. This will create a round and flat agar pad of around 10 mm diameter.</li>
	<li>Put a drop of 100 nm bead solution (undiluted, which is 2.6% (w/v)) onto the pad. Transfer 1-10 cleaned adult worms into the drop with a worm pick. Seal with 18x18 mm cover glass and vaseline as explained in Protocol 1.</li>
</ol>
3. Time-lapse microscopy
<ol>
	<li>Put a drop of immersion oil onto the microscope slide or the cover glass sandwich. Use brightfield at low magnification (5x or 10x lenses) to locate the sample. 
	NOTE: Limit the exposure to blue light as much as possible (in particular for C. elegans embryos), e.g., do not use epi-fluorescence microscopy to find/focus the specimen but use brightfield instead.</li>
	<li>Change to higher magnification (40x or 60x objectives), bring the sample into focus (either central plane or top/bottom planes of the sample, depending on the acquisition software).</li>
	<li>Set the sampling intervals to 30 planes at 1 &#181;m spacing recorded every 1min. In case an automated xy- or xyz-stage is available, multiple spots can be recorded in one session. Switch to fluorescence imaging and check the focus again. 
	NOTE: Try to avoid using high laser powers, rather use slightly longer exposure times at very low intensities. Measure acquired images right away, either in the image acquisition program or in ImageJ to avoid saturation, which crops the dynamics range and also means excessive radiation exposure.</li>
	<li>Start recording the time-lapse series. 
	NOTE: To exclude analyzing artifacts, start with long (3-10 min) time intervals between sample scanning and gradually increase temporal sampling to the desired intervals. Developmental timing as well as end points in both cases should be similar when performing a statistical analysis. Moreover, it is often necessary to assess phototoxicity and survival rates. Therefore, recover specimen from the mount after long-term time-lapse microscopy and check whether they continue to develop/live.</li>
</ol>
4. Reconstructing a 3D Cell Shape model with IMOD
NOTE: IMOD software is available from the IMOD web page: http://bio3d.colorado.edu/imod/. Installation of the software is described there. When using Windows OS, a Unix toolkit has to be installed, which can also be found as a package on the IMOD webpage. Moreover, there is an excellently compiled 3dmod introduction available on the IMOD webpage (http://bio3d.colorado.edu/imod/doc/3dmodguide.html).
<ol>
	<li>Open the Unix shell and type in &#34;imod&#34;. The IMOD window will open. Select the file with the raw data (ideally a stacked tif or similar) from which to generate a 3D model.</li>
	<li>Locate the bottom and top of the objects in the ZaP window by using the information window (up&#38;down scrolling tool).</li>
	<li>Start tracing the cell outlines. Importantly, ensure that each cell is one object. Begin with the top plane of each cell and create the first contour there using the information window. Scroll down in z and create a new contour for each z-plane.</li>
	<li>Do the same for as many cells as needed to model in 3D.&#160; Do not forget to create a new object when starting with a new cell. 
	NOTE: There are many different ways for contour tracing and several helpful plugins. Please visit the IMOD webpage and online tutorials for details. In the case shown here, the default setting has been used.</li>
	<li>Open the &#34;Model View&#34; window to inspect the objects and their contours.</li>
	<li>In the &#34;Model View&#34; toolbar choose &#34;Edit&#34; -&#34;Objects&#34;. The respective editing window will open.</li>
	<li>To model cells as filled and closed objects, choose &#34;Mesh&#34; as &#34;Draw Data Type&#34; and &#34;Fill&#34; as &#34;Drawing Style&#34;. Click on &#34;Meshing&#34; in the scrollable selection at the lower left of the window. Check the option &#34;Cap&#34; in the selection at the lower right. Check as many objects as necessary and click &#34;Mesh All&#34;. The program will generate solid objects from the stacks of contours. The z-sampling factor can be adjusted by changing &#34;Z-scale&#34; in the &#34;View&#34; window.</li>
	<li>Move/rotate the 3D object (&#34;Edit&#34; - &#34;View&#34;) and save snapshots or movies (&#34;File&#34; -&#34;Snap Tiff As ...&#34; or &#34;Movie/Montage&#34;).</li>
</ol>
5. Tracking Fluorescently Labeled Structures with Endrov
NOTE: In order to track cell division remnants, use a strain with a nuclear or plasma membrane marker for cell identification (e.g., H2B, PH-PLC-d1) and a cell division remnant marker (e.g., NMY-2, ZEN-4). The nuclear or plasma membrane marker is important to determine which cell inherits the cell division remnant.
<ol>
	<li>For fast loading and ease of use, the raw data has to be converted to a tiff file (e.g., in ImageJ). Next, open Endrov and load the file to be analysed by choosing the &#34;Load file&#34; option from the file menu. Click on the &#34;2D viewer&#34; icon on the toolbar.</li>
	<li>Adjust the histogram by clicking on the &#34;Fit range&#34; icon (the blue icon at the bottom right corner of the Endrov screen). Set the XYZ-resolution by going to &#34;Data&#34; -&#34;filename&#34;-&#34;Imageset&#34; -&#34;Channel&#34;-&#34;Set XYZ-resolution&#34;. Type in values for X, Y, and Z, e.g., 5, 5, 1 if X and Y resolution is 0.2 &#181;m and sample every 1 &#181;m in Z.</li>
	<li>For annotating the nuclei, move to the plane of interest and click on &#34;2D viewer&#34;. Select &#34;Lineage&#34; from the dropdown menu and choose &#34;New lineage&#34;.</li>
	<li>By clicking and dragging on the nucleus, it can be marked. To name the nucleus, right click and choose &#34;Rename particle&#34; from the dropdown menu. To increase or decrease the diameter of the circle, drag the arrow icon that appears on the left of the marked circle. To mark the central plane of the nucleus, click on the right icon. When using a plasma membrane marker, a large sphere can be drawn that will cover most of the cell.</li>
	<li>To proceed in time and plane, choose the &#34;Frame&#34; and &#34;Z&#34; options at the bottom toolbar. After going to the next time point, adjust the position or size of the circle that marks the nucleus accordingly. Repeat this until the tracked cell divides.</li>
	<li>When there is a cell division, mark the daughter nuclei and switch to &#34;Lineage viewer&#34;. This can be selected by clicking on the &#34;Windows&#34; icon on the top toolbar. Mark all three nuclei simultaneously, go to the &#34;Lineage&#34; option in the top toolbar and select &#34;Associate parent&#34;.</li>
	<li>When cell tracking is finished, switch to the channel marking the cell division remnant for tracking. To switch channels, click on the file name on the bottom left corner of the toolbar. The cell division remnant (or any other structure) can be tracked as described above for the nucleus.</li>
	<li>When returning to the software after closing, it is necessary to go to &#34;2D Viewer&#34; -&#34;Lineage&#34;-&#34;Edit&#34; and to click on the lineage number that will be edited.</li>
</ol>
6. Analyzing Cortical Actomyosin Flow with PIVlab
NOTE: PIVlab is a freely available software from: &#160;http://pivlab.blogspot.de/; it can be invoked from within the Matlab command line environment by typing PIVlab_GUI. When working with PIVlab, it is recommended to save the image series in a separate folder.
<ol>
	<li>Start a new session from &#34;File menu&#34;. Load the image files by clicking the &#34;Load images&#34; button.</li>
	<li>Select the sequencing style 1-2, 2-3, ... and navigate to the directory that contains the sequence of desired images. Select the images and click on the &#34;Add&#34; button and import.</li>
	<li>Go to &#34;Analyses settings&#34; and select &#34;Exclusions (ROI,Mask)&#34;. Alternatively, press &#34;Ctrl + E&#34;. Use the button &#34;Draw mask(s) for current frame&#34; to inactivate the undesired part of the image/s (i.e., area outside embryo).</li>
	<li>Select &#34;PIV settings&#34; from the &#34;Analyses settings&#34; menu. Alternatively, press &#34;Ctrl + S&#34;. Choose &#34;FFT window deformation&#34; as the desired PIV algorithm. 
	NOTE: Fast Fourier transformation (FFT) reduces the computational cost but is only recommended the signal is well above noise and if the displacement of the particles studied does not exceed half the pass area analyzed (see 6.5).</li>
	<li>Enter the following values for the three passes, which are the area of interrogation for intensity cross correlation between subsequent images: Pass 1: Interrogation area of 64 pixels with a step of 32. Pass 2: Interrogation area of 32 pixels with a step of 16. Select &#34;linear&#34; from the Window deformation drop down menu. Choose Gaussian 2*3-point method for sub-pixel displacement estimation.</li>
	<li>Select &#34;Analyze!&#34; from Analysis menu and use the button &#34;Analyze all frames&#34; to begin an analysis.</li>
	<li>For post processing select &#34;Vector validation&#34; from &#34;Post processing&#34; menu and set the standard deviation (Stdev) filter threshold to 7 and apply to all frames.</li>
	<li>Choose &#34;Derive parameters/ modify data&#34; from &#34;Plot&#34; menu. Select &#34;Vectors [px/frame]&#34; from dropdown menu. Adjust the smoothness of vectors and high pass filter and apply to all frames.</li>
	<li>To calculate mean vectors of all frames, set the &#34;Used frames&#34; to 1:end and click on &#34;Calculate mean vectors&#34; button.</li>
</ol>

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Research in the lab of CP is funded by the Deutsche Forschungsgemeinschaft (EXC 115; FOR 1756) and the LOEWE Research Cluster Ubiquitin Networks. CP is further supported by the European Union Framework Program 7 (Marie Curie Actions Project 326632).

Through object tracking in development, in particular nuclear tracking, it has been possible to elucidate central patterning mechanisms of C. elegans embryogenesis1,23,24. Expanding this strategy to higher throughput, it has been recently possible to uncover additional patterning rules and to propose a method how to deduce patterning rules de novo10. For many mutants, however, the precise patterning defects are still unknown. The methods described here are tools that can be instrum...

With the steady improvements of fluorescent proteins, genome engineering, light microscopy, and computer soft- and hardware, it is now possible to record development of many model organisms at unprecedented spatio-temporal resolution. This allows researchers to ask questions that could not be addressed previously or to revisit known developmental processes in order to search for overlooked aspects. This progress has sparked the field of quantitative developmental biology, which aims at transforming qualitative, informal models into quantitative models by thorough measurements and statistical analyses.
Tracking cells and subcellular structures has made it possible to derive quantitative models of embryonic development, nervous system activity, or cell division1-12. By tracking cell division remnants during early development of the C. elegans embryo, we could recently reveal that they follow a stereotyped path and constitute important polarizing factors13,14.
Here, protocols are presented that make quantitative developmental biology approaches accessible for non-experts. The focus lies on three straight-forward, freely available tools that are implementable in any lab that has access to standard confocal microscopy and computers. These include a protocol to generate 3D cell shapes, a protocol to track cell division remnants, and a protocol to quantitatively describe cortical actomyosin dynamics. The nematode C. elegans is used as an exemplary case, however, the methods and tools discussed here are suited for a variety of questions in other biological models, e.g., cultured cells, tissue explants, organoids or spheroids, other embryos, etc.
Generally, some of the analyses shown here can also be performed with the popular open source tool ImageJ (http://imagej.nih.gov/ij/docs/index.html; or FIJI, the &#8216;batteries included&#8217; version of ImageJ, http://fiji.sc/Fiji) for which many plugins for different quantitative analyses are available. However, the programs discussed here are designed to tackle specific problems.
Firstly, IMOD, an image processing, modeling and display suite can be used for 3D reconstructions of serial sections from electron or light microscopy15. IMOD also contains tools for viewing the 3D data from any orientation. Secondly, Endrov, a Java program designed to perform image analysis, data processing, and annotation of networks or tracks (among others) on the basis of an extended plugin architecture, with ImageJ plugin compatibility16. It contains over 140 image-processing operations and an extensible user interface in which model and raw data are displayed separately. Its source code can be found at https://github.com/mahogny/Endrov. Thirdly, PIVlab, a MATLAB tool for digital particle image velocimetry that allows the user to quantitatively and qualitatively analyze particle flow fields17. The use of this programs requires a MATLAB license that includes the Image Processing Toolbox (http://mathworks.com). PIVlab is a program designed to quantitatively describe flow. It calculates the velocity distribution, magnitude, vorticity, divergence, or shear within image pairs or series. For this, it cross-correlates small areas of images (called &#8216;passes&#8217; in the protocol section) of an image pair to derive the most probable particle displacement. This cross-correlation yields a correlation matrix that can be analyzed in the space or frequency domain using either direct cross-correlation or a fast Fourier transformation (FFT), respectively.
The equipment used here is an inverted microscope equipped with a Nipkow (&#39;spinning&#39;) disc, an EMCCD, 488 and 561 nm standard solid-state lasers, and 20x air or 40x or 60x plan/apochromat oil- or water-immersion objectives. However, it is also possible to perform time-lapse imaging with other imaging modalities, e.g., point-, line- or sheet-scanning laser-based microscopy, multi-photon microscopy, as well as epi-fluorescence microscopy combined with deconvolution or structured illumination. The advantage of using a Nipkow disc system is the extremely fast image acquisition, especially if a streaming mode (continuous movement and scanning of the object in the z dimension) is available. In addition, to improve resolution, a 1.5-fold magnifying extender in front of the EMCCD can be used.

<table border="0" cellpadding="0" cellspacing="0" width="573">
	<tbody>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">Stereo microscope</td>
			<td style="width:71px;">Motic/VWR</td>
			<td style="width:119px;">OT4005S</td>
			<td style="width:167px;">Stereo microscope for dissection and mounting</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Polybead Polystyrene Microspheres,&#160;</td>
			<td rowspan="2" style="width:71px;">Polysciences</td>
			<td rowspan="2" style="width:119px;">18329</td>
			<td rowspan="2" style="width:167px;">Embryo mounting</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">20 &#181;m</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Polybead Polystyrene Microspheres,&#160;</td>
			<td rowspan="2" style="width:71px;">Polysciences</td>
			<td rowspan="2" style="width:119px;">876</td>
			<td rowspan="2" style="width:167px;">Adult animal mounting</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">0.1 &#181;m</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Microscope slides</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">631-0902</td>
			<td style="width:167px;">Adult animal mounting</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Cover glass 18x18 mm</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">631-1331</td>
			<td style="width:167px;">Embryo/adult mounting</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Cover glass 24x60 mm</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">631-1339</td>
			<td style="width:167px;">Embryo mounting</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Scalpel</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">233-5455</td>
			<td style="width:167px;">Embryo dissection</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Silicone tubing</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">228-1501</td>
			<td style="width:167px;">Tubing for mouth pipette</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">30 mm PTFE membrane filter</td>
			<td style="width:71px;">NeoLab</td>
			<td style="width:119px;">Jul-01</td>
			<td style="width:167px;">Filter for mouth pipette</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Capillary tubes</td>
			<td style="width:71px;">VWR</td>
			<td style="width:119px;">621-0003</td>
			<td style="width:167px;">Pipette tip for mouth pipette</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Vaseline</td>
			<td style="width:71px;">Roth</td>
			<td style="width:119px;">E746.1</td>
			<td style="width:167px;">Embryo/adult mounting</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Agar</td>
			<td style="width:71px;">Roth</td>
			<td style="width:119px;">5210.5</td>
			<td style="width:167px;">Adult animal mounting</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Potassium-di-hydrogenphosphate</td>
			<td style="width:71px;">Roth</td>
			<td style="width:119px;">P018.2</td>
			<td style="width:167px;">M9 buffer</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Di-sodium- hydrogenphosphate</td>
			<td style="width:71px;">Roth</td>
			<td style="width:119px;">P030.2</td>
			<td style="width:167px;">M9 buffer</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">Sodium chloride</td>
			<td style="width:71px;">Roth</td>
			<td style="width:119px;">3957.1</td>
			<td style="width:167px;">M9 buffer</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">VisiScope Spinning Disc Confocal System</td>
			<td style="width:71px;">Visitron Systems</td>
			<td style="width:119px;">n/a</td>
			<td style="width:167px;">Confocal microscopy</td>
		</tr>
	</tbody>
</table>

tracking and quantifying developmental processes in c. elegans using open-source tools

Quantitatively capturing developmental processes is crucial to derive mechanistic models and key to identify and describe mutant phenotypes. Here protocols are presented for preparing embryos and adult C. elegans animals for short- and long-term time-lapse microscopy and methods for tracking and quantification of developmental processes. The methods presented are all based on C. elegans strains available from the Caenorhabditis Genetics Center and on open-source software that can be easily implemented in any laboratory independently of the microscopy system used. A reconstruction of a 3D cell-shape model using the modelling software IMOD, manual tracking of fluorescently-labeled subcellular structures using the multi-purpose image analysis program Endrov, and an analysis of cortical contractile flow using PIVlab (Time-Resolved Digital Particle Image Velocimetry Tool for MATLAB) are shown. It is discussed how these methods can also be deployed to quantitatively capture other developmental processes in different models, e.g., cell tracking and lineage tracing, tracking of vesicle flow.

By using protocols 2, 3, and 4, time-lapse imaging of gonads in wild type C. elegans adults is performed (strain OD58 (unc-119(ed3) III; ltIs38[pAA1; pie-1::GFP::PH(PLC1delta1) + unc-119(+)]), expressing a membrane marker from a germline promoter). Focusing on the turn of the gonad, a 3D model of the germ cells is generated from the microscopy data (Figure 2). This model allows to analyze changes in cell size while the cells transit form the distal to the proximal arm, reveals the organization o...

Watch this Scientific Journal Video about Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools at JoVE.com

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Here it is shown how to track and quantify developmental processes in C. elegans. The methods presented are based on open-source tools that can be easily implemented. It is demonstrated how to reconstruct 3D cell-shape models, how to manually track subcellular structures, and how to analyze cortical contractile flow.

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Quantitatively capturing developmental processes is crucial to derive mechanistic models and key to identify and describe mutant phenotypes. Here ...

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8.7K Views.  Goethe University. Here it is shown how to track and quantify developmental processes in C. elegans. The methods presented are based on open-source tools that can be easily implemented. It is demonstrated how to reconstruct 3D cell-shape models, how to manually track subcellular structures, and how to analyze cortical contractile flow.

Video: Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Using Micro-Electro-Mechanical Systems (MEMS) to Develop Diagnostic Tools

Using Micro-Electro-Mechanical Systems MEMS to Develop Diagnostic Tools

Generation of Stable Transgenic C. elegans Using Microinjection

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.

This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1.  The plasmid DNA rearranges to ...

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using fluorescent-labeled reporter proteins or differential interference contrast (DIC) microscopy, allows for the examination of the temporal progression of these dynamic events which is otherwise inferred from analysis of fixed samples1,2. Moreover, the study of the developmental regulations of cellular processes necessitates conducting time-lapse experiments on an intact organism during development. The Caenorhabiditis elegans embryo is light-transparent and has a rapid, invariant developmental program with a known cell lineage3, thus providing an ideal experiment model for studying questions in cell biology4,5and development6-9. C. elegans is amendable to genetic manipulation by forward genetics (based on random mutagenesis10,11) and reverse genetics to target specific genes (based on RNAi-mediated interference and targeted mutagenesis12-15). In addition, transgenic animals can be readily created to express fluorescently tagged proteins or reporters16,17. These traits combine to make it easy to identify the genetic pathways regulating fundamental cellular and developmental processes in vivo18-21. In this protocol we present methods for live imaging of C. elegans embryos using DIC optics or GFP fluorescence on a compound epifluorescent microscope. We demonstrate the ease with which readily available microscopes, typically used for fixed sample imaging, can also be applied for time-lapse analysis using open-source software to automate the imaging process.

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

The C. elegans embryo is a powerful system for studying cell biology and development. We present a protocol for live imaging of C. elegans embryos utilizing DIC optics or fluorescence using readily available epifluorescent microscopes and open-source software.

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using ...

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1. Apoptosis can be triggered when cells encounter a wide range of cytotoxic stresses. These insults initiate signaling cascades that ultimately cause the release of cytochrome c from the mitochondrial intermembrane space to the cytoplasm2. The release of cytochrome c from mitochondria is a key event that triggers the rapid activation of caspases, the key cellular proteases which ultimately execute cell death3-4.
The pathway of apoptosis is regulated at points upstream and downstream of cytochrome c release from mitochondria5. In order to study the post-mitochondrial regulation of caspase activation, many investigators have turned to direct cytoplasmic microinjection of holocytochrome c (heme-attached) protein into cells6-9. Cytochrome c is normally localized to the mitochondria where attachment of a heme group is necessary to enable it to activate apoptosis10-11. Therefore, to directly activate caspases, it is necessary to inject the holocytochrome c protein instead of its cDNA, because while the expression of cytochrome c from cDNA constructs will result in mitochondrial targeting and heme attachment, it will be sequestered from cytosolic caspases. Thus, the direct cytosolic microinjection of purified heme-attached cytochrome c protein is a useful tool to mimic mitochondrial cytochrome c release and apoptosis without the use of toxic insults which cause cellular and mitochondrial damage.
In this article, we describe a method for the microinjection of cytochrome c protein into cells, using mouse embryonic fibroblasts (MEFs) and primary sympathetic neurons as examples. While this protocol focuses on the injection of cytochrome c for investigations of apoptosis, the techniques shown here can also be easily adapted for microinjection of other proteins of interest.

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

In this protocol, we describe the direct cytoplasmic microinjection of cytochrome c protein into fibroblasts and primary sympathetic neurons. This technique allows for the introduction of cytochrome c protein into the cytoplasm of cells and mimics the release of cytochrome c from mitochondria, which occurs during apoptosis.

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1.  Apoptosis can be triggered when cells encounter a ...

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

To stain C. elegans with antibodies, the relatively impermeable cuticle must be bypassed by chemical or mechanical methods. "Freeze-cracking" is one method used to physically pull the cuticle from nematodes by compressing nematodes between two adherent slides, freezing them, and pulling the slides apart. Freeze-cracking provides a simple and rapid way to gain access to the tissues without chemical treatment and can be used with a variety of fixatives. However, it leads to the loss of many of the specimens and the required compression mechanically distorts the sample. Practice is required to maximize recovery of samples with good morphology. Freeze-cracking can be optimized for specific fixation conditions, recovery of samples, or low non-specific staining, but not for all parameters at once. For antibodies that require very hard fixation conditions and tolerate the chemical treatments needed to chemically permeabilize the cuticle, treatment of intact nematodes in solution may be preferred. If the antibody requires a lighter fix or if the optimum fixation conditions are unknown, freeze-cracking provides a very useful way to rapidly assay the antibody and can yield specific subcellular and cellular localization information for the antigen of interest.

Antibody Staining in C. Elegans Using &quot;Freeze-Cracking&quot;

"Freeze-cracking," a method for exposing the inner tissues of the nematode C. elegans to antibodies for protein localization, is demonstrated.

To stain C.  elegans with antibodies, the relatively impermeable cuticle must be  bypassed by chemical or mechanical methods.  "Freeze-cracking" is one ...

Methods for Skin Wounding and Assays for Wound Responses in C. elegans

The C. elegans epidermis and cuticle form a simple yet sophisticated skin layer that can repair localized damage resulting from wounding. Studies of wound responses and repair in this model have illuminated our understanding of the cytoskeletal and genomic responses to tissue damage. The two most commonly used methods to wound the C. elegans adult skin are pricks with microinjection needles, and local laser irradiation. Needle wounding locally disrupts the cuticle, epidermis, and associated extracellular matrix, and may also damage internal tissues. Laser irradiation results in more localized damage. Wounding triggers a succession of readily assayed responses including elevated epidermal Ca2+ (seconds-minutes), formation and closure of an actin-containing ring at the wound site (1-2 hr), elevated transcription of antimicrobial peptide genes (2-24 hr), and scar formation. Essentially all wild type adult animals survive wounding, whereas mutants defective in wound repair or other responses show decreased survival. Detailed protocols for needle and laser wounding, and assays for quantitation and visualization of wound responses and repair processes (Ca dynamics, actin dynamics, antimicrobial peptide induction, and survival) are presented.

Methods for Skin Wounding and Assays for Wound Responses in C. elegans

The adult C. elegans skin is a tractable model for studies of epithelial wound responses, including wound closure, scar formation, and innate immunity.

The C. elegans epidermis and cuticle form a simple yet sophisticated skin layer that can repair localized damage resulting from wounding. Studies of wound ...

Open Source High Content Analysis Utilizing Automated Fluorescence Lifetime Imaging Microscopy

We present an open source high content analysis instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions using F&#246;rster resonance energy transfer (FRET) based readouts of fixed or live cells in multiwell plates. This provides a means to screen for cell signaling processes read out using intramolecular FRET biosensors or intermolecular FRET of protein interactions such as oligomerization or heterodimerization, which can be used to identify binding partners. We describe here the functionality of this automated multiwell plate FLIM instrumentation and present exemplar data from our studies of HIV Gag protein oligomerization and a time course of a FRET biosensor in live cells. A detailed description of the practical implementation is then provided with reference to a list of hardware components and a description of the open source data acquisition software written in &#181;Manager. The application of FLIMfit, an open source MATLAB-based client for the OMERO platform, to analyze arrays of multiwell plate FLIM data is also presented. The protocols for imaging fixed and live cells are outlined and a demonstration of an automated multiwell plate FLIM experiment using cells expressing fluorescent protein-based FRET constructs is presented. This is complemented by a walk-through of the data analysis for this specific FLIM FRET data set.

We present an open source high content analysis (HCA) instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions using F&#246;rster resonance energy transfer (FRET) based readouts. Data acquisition for this openFLIM-HCA instrument is controlled by software written in &#181;Manager and data analysis is undertaken in FLIMfit.

We present an open source high content analysis instrument utilizing automated fluorescence lifetime imaging (FLIM) for assaying protein interactions ...

Open-source Single-particle Analysis for Super-resolution Microscopy with VirusMapper

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm allows for the routine imaging of nanoscale biological complexes and processes. This increase in resolution also means that methods popular in electron microscopy, such as single-particle analysis, can readily be applied to super-resolution fluorescence microscopy. By combining this analytical approach with super-resolution optical imaging, it becomes possible to take advantage of the molecule-specific labeling capacity of fluorescence microscopy to generate structural maps of molecular elements within a metastable structure. To this end, we have developed a novel algorithm &#8212; VirusMapper &#8212; packaged as an easy-to-use, high-performance, and high-throughput ImageJ plugin. This article presents an in-depth guide to this software, showcasing its ability to uncover novel structural features in biological molecular complexes. Here, we present how to assemble compatible data and provide a step-by-step protocol on how to use this algorithm to apply single-particle analysis to super-resolution images.

This manuscript uses the Fiji-based open-source software package VirusMapper to apply single-particle analysis to super-resolution microscopy images in order to generate precise models of nanoscale structure.

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm ...

Measuring RAN Peptide Toxicity in C. elegans

C. elegans is commonly used to model age-related neurodegenerative diseases caused by repeat expansion mutations, such as Amyotrophic Lateral Sclerosis (ALS) and Huntington&#8217;s disease. Recently, repeat expansion-containing RNA was shown to be the substrate for a novel type of protein translation called repeat-associated non-AUG-dependent (RAN) translation. Unlike canonical translation, RAN translation does not require a start codon and only occurs when repeats exceed a threshold length. Because there is no start codon to determine the reading frame, RAN translation occurs in all reading frames from both sense and antisense RNA templates that contain a repeat expansion sequence. Therefore, RAN translation expands the number of possible disease-associated toxic peptides from one to six. Thus far, RAN translation has been documented in eight different repeat expansion-based neurodegenerative and neuromuscular diseases. In each case, deciphering which RAN products are toxic, as well as their mechanisms of toxicity, is a critical step towards understanding how these peptides contribute to disease pathophysiology. In this paper, we present strategies to measure the toxicity of RAN peptides in the model system C. elegans. First, we describe procedures for measuring RAN peptide toxicity on the growth and motility of developing C. elegans. Second, we detail an assay for measuring postdevelopmental, age-dependent effects of RAN peptides on motility. Finally, we describe a neurotoxicity assay for evaluating the effects of RAN peptides on neuron morphology. These assays provide a broad assessment of RAN peptide toxicity and may be useful for performing large-scale genetic or small molecule screens to identify disease mechanisms or therapies.

Measuring RAN Peptide Toxicity in C. elegans

Repeat-associated non-ATG-dependent translational products are emerging pathogenic features of several repeat expansion-based diseases. The goal of the protocol described is to evaluate toxicity caused by these peptides using behavioral and cellular assays in the model system C. elegans.

C. elegans is commonly used to model age-related neurodegenerative diseases caused by repeat expansion mutations, such as Amyotrophic Lateral Sclerosis ...

Worm-align and Worm_CP, Two Open-Source Pipelines for Straightening and Quantification of Fluorescence Image Data Obtained from Caenorhabditis elegans

An issue often encountered when acquiring image data from fixed or anesthetized C. elegans is that worms cross and cluster with their neighbors. This problem is aggravated with increasing density of worms and creates challenges for imaging and quantification. We developed a FIJI-based workflow, Worm-align, that can be used to generate single- or multi-channel montages of user-selected, straightened and aligned worms from raw image data of C. elegans. Worm-align is a simple and user-friendly workflow that does not require prior training of either the user or the analysis algorithm. Montages generated with Worm-align can aid the visual inspection of worms, their classification and representation. In addition, the output of Worm-align can be used for subsequent quantification of fluorescence intensity in single worms, either in FIJI directly, or in other image analysis software platforms. We demonstrate this by importing the Worm-align output into Worm_CP, a pipeline that uses the open-source CellProfiler software. CellProfiler&#8217;s flexibility enables the incorporation of additional modules for high-content screening.&#160;As a practical example, we have used the pipeline on two datasets: the first dataset are images of heat shock reporter worms that express green fluorescent protein (GFP) under the control of the promoter of a heat shock inducible gene hsp-70, and the second dataset are images obtained from fixed worms, stained for fat-stores with a fluorescent dye.

Worm-align and Worm_CP, Two Open-Source Pipelines for Straightening and Quantification of Fluorescence Image Data Obtained from Caenorhabditis elegans

Worm-align/Worm_CP is a simple FIJI/CellProfiler workflow that can be used to straighten and align Caenorhabditis elegans samples and to score whole-worm image-based assays without the need for prior training steps. We have applied Worm-align/Worm_CP to the quantification of&#160;heat-shock induced expression in live animals or lipid droplets&#160;in fixed samples.

An issue often encountered when acquiring image data from fixed or anesthetized C. elegans is that worms cross and cluster with their neighbors. This ...