The authors would like to thank the Yogev and Hammarlund labs for technical assistance, feedback, and discussions. We would like to especially thank Grace Swaim for guidance in live cell imaging and Grace and Brian Swaim for initially establishing the manual kymograph analysis in the lab. OG is supported by a Walter-Benjamin Scholarship funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) -Project# 465611822. SY is funded by the NIH grant R35-GM131744.

For a detailed protocol on how to maintain and prepare nematodes for live-cell imaging, refer to the work of S.Niwa 7.
1. Worm strain generation
In addition to generating nematode strains, the Caenorhabditis Genetics Center (CGC)8 contains a growing collection of nematode strains with endogenously fluorescently tagged proteins that can be directly obtained from their webpage.
<ol>
	<li>Choice of the fluorescence labeling strategy
	<ol>
		<li>Use the nematode strain MTS1161, which contains the rab-3(ox699) allele6, to visualize endogenous GFP labeled RAB-3 in the DA9 motor neuron (Strain will be deposited at the CGC). To visualize other axonal proteins, generate a nematode strain with an endogenous tagged protein by using the CRISPR Cas9 editing protocol from the Mello Lab9. 
		NOTE: MTS1161 utilizes a recombination approach to cell specifically label RAB-3 endogenously with a GFP tag6. A common alternative approach is based on the reconstitution of a split-fluorescent protein, which is recommended for especially low expressed proteins as it enhances the fluorescent copy number per endogenous protein by using multiple split fluorescent copies10. The copy number can initially be estimated based on transcriptome datasets from the CenGen webpage (https://cengen.shinyapps.io/CengenApp/)&#160;&#8203;11.</li>
	</ol>
	</li>
	<li>Choice of neuron to visualize axonal transport
	<ol>
		<li>In the strain MTS1161, visualize RAB-3 in the motor neuron DA9 (see Figure 1). For other axonal cargos, especially low expressed proteins, identify a neuron in which the expression levels of the analyzed protein are highest using the CenGen webpage. 
		NOTE: The CenGen project (cengen.org) provides a great online resource to estimate expression levels for a protein of interest in many neurons of the nematode based on a large transcriptomic dataset covering all 302 neurons of the C. elegans nervous system12,13.</li>
	</ol>
	</li>
</ol>
2. Worm handling and preparation for imaging
<ol>
	<li>Maintain nematodes on a lawn of bacteria (strain: OP50) seeded on nematode growth medium plates at 20 &#176;C and grown to an age of 1 day into adulthood for imaging. 
	NOTE: Measuring axonal transport at a defined age stage is important as transport rates declines with ageing consistently across different axonal cargos, neuron classes and organisms14,15,16,17,18-Nematodes at larval stage L4 or 1 day into adulthood have been used in most papers and thus offer the largest datasets for comparisons.</li>
	<li>Prepare nematodes for live cell imaging: Perform all subsequent steps with a stereo microscope to visualize and handle the worms.
	<ol>
		<li>Transfer nematodes to a droplet of 0.5 mM Levamisole using a platinum wire pick and incubate the nematodes for 20 min in the droplet.</li>
		<li>Carefully mount 10-20 nematodes from the Levamisole droplet into a 12 &#181;L droplet of M9 medium on a 10% agarose (dissolved in M9 medium) patch on a glass slide by using a hair pick. For a detailed procedure on how to make a 10% agarose patch, refer to the protocol of S.Niwa7. 
		NOTE: Mounting a low number of nematodes is sufficient since only a few animals will be imaged per slide before the animals start to suffer from hypoxia.</li>
		<li>Place a cover slip on top of the droplet (22 mm x 22 mm or 18 mm x 18 mm). For imaging transport in the DA9 axon, position the axon close to the cover slip. To do so, rotate the cover slip by 45&#176; after placing it on top of the agar patch, to roll the animals onto their back.</li>
		<li>Seal the space between the cover slip and the glass slide with a viscous gel like petrolatum. This will prevent the agarose patch from drying, which can cause the nematodes to drift out of the imaging field. 
		NOTE: It is crucial to treat the animals as gently as possible. Too high concentrations of levamisole or physical damage to the worm can impair axonal transport. Mild twitches of the tail during the imaging session are a good sign that the animal remains in a healthy state. Animals remain healthy and can even recover from the agarose patch after the imaging session.</li>
	</ol>
	</li>
</ol>
3. Live-cell Microscopy
NOTE: Exact acquisition parameter values can differ between microscopes. However, trends for each acquisition parameter should be independent of the microscope used. A spinning-disc confocal microscope that was equipped with a separate laser line for bleaching was used in this protocol (see Table of Materials for details on the microscope). Green fluorescence was excited by a 488 nm laser and emission was filtered by an ET525/36 emission filter. Bleaching was performed using a 488 nm laser line.
<ol>
	<li>Ensure that the temperature of the room, or if a temperature-controlled stage is available, is set to a constant temperature. For nematodes, set the temperature to 20 &#176;C.</li>
	<li>Mount the slide and use objective lens with 4x-20x magnification to locate the nematodes on the slide and memorize positions. Switch to 63x magnification lens for image acquisition.</li>
	<li>Take a single image to check initial acquisition parameters. Track the signal intensities in the field of view with the intensity histogram of the acquisition software. Intensity values should cover the lowest third of the maximum signal that the camera can detect. Avoid pixel saturation. Enhance the intensity of the excitation laser if the signal intensity is too low. 
	NOTE: Do not use too high excitation laser intensities as it bleaches the fluorescent protein during time-lapse.</li>
	<li>Alter the acquisition parameter and steps here to optimize fluorescence signal detection.
	<ol>
		<li>Implementation of a bleaching step: Bleach the region of the axon that is going to be analyzed using a 488 nm laser (power:15 mW) line for the bleaching step. Aim for a bleaching efficiency of at least 90%. Determine the bleaching efficiency by comparing the signal intensity of the region before and after the bleaching step. 
		NOTE: Bleaching enhances the signal of moving particles by lowering the signal that derives from stationary particles as well as from the signal that derives from the cytoplasmic fraction of the protein in the background. Membranous cargo, such as RAB-3 on synaptic vesicle precursors have a low cytoplasmic fraction so that the bleaching step only mildly improves the signal of the transport package over the cytoplasmic background (Figure 2A,D). However, bleaching improves tracing individual transport events over longer distances and allows quantifying pausing times (Figure 2A).</li>
		<li>Binning: Use 2 x 2 binning to enhance the signal intensity of individual transport events. Binning combines pixel arrays into one larger pixel. This will reduce the spatial resolution but enhance the signal intensity of individual transport events and is especially helpful for dim particles (Figure 2B,D).</li>
		<li>Exposure time: Enhance the exposure time to enhance the signal intensity. Enhancing the exposure time enhances the signal intensity of the sample to the cost of getting a lower temporal resolution for the transport events. For the experiment here, use a temporal resolution of 300 ms to 700 ms between consecutive timepoints (100- 500 ms exposure time) to follow individual transport events of RAB-3. (Figure 2C,D)</li>
	</ol>
	</li>
	<li>Time-lapse recording: Take a time lapse of at least 1-3 min depending on the signal intensity of the protein as well as the frequency of individual transport events. Once a time lapse has been recorded move to the next animal. Animals on a slide should not be imaged for more than 30 min to ensure that the recorded animals are in a healthy state. 
	NOTE: Mild twitches of the animals&#39; tail during the recording are an indicator that the animal is still alive.</li>
</ol>
4. Analyzing axonal transport data
NOTE: Use ImageJ/Fiji19 for the subsequent image analysis steps. Fiji is able to read data by all common microscopy software packages.
<ol>
	<li>Kymograph generation
	<ol>
		<li>Import data: Import the time-lapse data file into Fiji. In case it cannot import the data into Fiji, export the time-recording as a tiff file from the software package and load it subsequently into Fiji.</li>
		<li>Drift correction: Check whether the animal/axon segment moved or slightly drifted during the image acquisition. Correct for animal movement or drifting by running the plugin&#160;StackReg20. When running the plugin, choose the Rigid Body transformation.</li>
		<li>Manual generation of a kymograph: A detailed step by step procedure is provided in Figure 3. Utilize the drift corrected movie to generate the kymograph. Use the Segmented Line tool and adjust the line width to match the axon diameter by double clicking with the left mouse button on the segmented line icon to draw a line along the axon segment that will be analyzed. Run the ImageJ/Fiji plugin Kymoreslicewide to generate the kymograph and utilize the maximum intensity value across the width of the line in its parameter settings. 
		NOTE: Keeping consistency in the line drawing direction (e.g., always proximal to distal axon or reverse) simplifies tracing back movement direction in the kymographs.</li>
	</ol>
	</li>
	<li>Analysis of transport parameter
	<ol>
		<li>Calculate the transport parameter: event number, velocity, run length and pause duration from the kymograph by following the subsequent steps.
		<ol>
			<li>Trace transport events in the kymograph: Use the Straight Line&#160;tool to trace individual transport events on the kymograph. Save each line to the ROI manager and trace all transport events in the kymograph. Choose the following measurement parameters in ImageJ/Fiji: Area, Bounding rectangle, Mean gray value, Feret&#180;s diameter.</li>
			<li>Calculate transport parameter: Paste the results table into a spreadsheet and use the following columns of the result sections to calculate: 
			Run length: Multiply width (in pixel) by the resolution of the camera (e.g., x &#181;m/pxl) to determine the run length in &#181;m. 
			Velocity: Run length/Run duration. To determine the duration of a movement event in seconds, multiply height (in pixel) by the acquisition time between consecutive timepoints (e.g., x s/pixel). 
			Pausing time: Run duration in between two consecutive movements at which velocity is 0. 
			Event number: Total events can be normalized to the total length of the kymograph to determine event number per min and axon length segment. Events can further be categorized into anterograde and retrograde transport events. To determine the directionality of each movement event, utilize the Feret Angle tool. In kymographs that were initially generated by drawing the segmented line from proximal to distal and the anterograde transport events in the kymograph are pointing from right to left, a Feret Angle &#60; 90&#176; will indicate an anterograde and &#62;90&#176; a retrograde event, otherwise it will be the inverse.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

Enhancing Axonal Transport Analysis in Living C. elegans Using GFP-Tagged RAB-3

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The authors declare no competing or financial interests that could have appeared to influence the work reported in this paper.

Limitations of the method and alternative methods 
In this protocol, we optimized acquisition parameters to visualize the axonal transport of endogenously tagged RAB-3, which is associated with synaptic vesicle precursors. To visualize RAB-3, we made use of a recently published FLIP-on::GFP::RAB-3 strain6 and expressed the recombinase Flippase under a cell specific promoter (glr-4p)25. This strategy allows us to label RAB-3 with a single GFP fluorophor...

Throughout life, neurons rely on axonal transport to deliver proteins, lipids, and other molecules from the cell body to their final destination in the axon. Consequently, impairment of axonal transport is associated with a loss of neuronal function and is often involved in the pathology of neurodegenerative disorders1,2. Hence, understanding the mechanisms that underly axonal transport is of great interest. 
Several decades of research on axonal transport revealed many important insights into the molecular machinery that mediates this transport, their composition as well as regulatory mechanisms. Long-range axonal transport occurs on the microtubule cytoskeleton, which consists of partially overlapping microtubule polymers that are typically oriented with their plus end out in axons3. Consequently, anterograde transport is mediated by motor proteins that walk to the plus end of microtubules, kinesins, whereas retrograde transport depends on the minus end directed dynein motor. Although many aspects of transport have been revealed, for many axonal proteins it still remains unclear, how they are loaded into the transport machinery, how individual transport packages are organized, and how this transport is regulated3. 
Axonal transport was initially studied in radio-labeling experiments, in which radiolabeled amino acids were injected into the somatic compartment, where they were incorporated into nascent endogenous proteins and could be traced over time in the axonal compartment by autoradiography4. Although radiolabeling experiments allowed the study of axonal transport of endogenous proteins in vivo, it does not allow for the direct follow-up of the behavior of individual cargo to get mechanistic insights4. This limitation was overcome with the use of fluorescence microscopy. However, axonal transport is often not visualized on endogenous proteins but instead by expression of a fluorescent labeled copy. Especially for low expressed proteins, overexpression provides higher signal intensities which make visualization, preferably with single neuron resolution, possible. Moreover, ectopic expression of the fluorescent tagged protein circumvents the need and challenges of genome editing. Conversely, it has been argued that the behavior of ectopically expressed cargo may differ from the behavior of the endogenous cargo5. 
Recent improvements in genome editing made endogenous labeling strategies easier accessible. Hence, a lower signal intensity has become the major limitation to study axonal transport of a cargo by ectopic expression instead of endogenous labeling. Careful considerations in the endogenous labeling strategy paired with an optimization of the acquisition conditions can overcome this challenge. 
Nematodes provide an excellent research model to study axonal transport in vivo due to their transparency and ease in genetic manipulations. In this protocol, we describe a research strategy to visualize axonal transport of endogenous proteins with single neuron resolution in living Caenorhabditis elegans. We visualize the axonal transport of synaptic vesicle precursors by using a strain generated by the Jorgensen Lab6, in which the vesicle associated RAB GTPase, RAB3, is endogenously labeled with GFP, in the motor neuron DA9. By asking how small adaptations in different acquisition parameters and photobleaching can improve the visualization of individual transport events, the protocol provides ideas on how to optimize imaging conditions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agarose</td>
			<td>Sigma-Aldrich</td>
			<td>A9539</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover slips (22 mm x 22 mm, No1); Gold Seal Cover Glass</td>
			<td>Thomas Scientific</td>
			<td>6672A14</td>
			<td></td>
		</tr>
		<tr>
			<td>Levamisole</td>
			<td>ChemCruz</td>
			<td>sc-205730</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope: Nikon Ti2 inverted microscope, Yokogawa CSU-W1 SoRa Scanhead, Hamatsu Orca-Fusion BT sCMOS camera, Nikon CFI Plan Apo lambda 60x 1.4 NA oil immersion objective, Nikon photostimulation scanner at 488nm with an ET525/36 emission filter</td>
			<td>Nikon</td>
			<td></td>
			<td>Spinning Disc Confocal Microscope</td>
		</tr>
		<tr>
			<td>&#160;NIS-elements AR</td>
			<td>Nikon</td>
			<td></td>
			<td>Software for the Nikon Ti2&#160;</td>
		</tr>
		<tr>
			<td>Plain precleaned microscopy slides</td>
			<td>Thermo Scientific</td>
			<td>420-004T</td>
			<td></td>
		</tr>
		<tr>
			<td>Nematode strain</td>
			<td>Identifier</td>
			<td>Source</td>
			<td></td>
		</tr>
		<tr>
			<td>rab-3(ox699[GFP::flip-on::rab-3]) (II); shyIs43(glr-4p::FLP-NLSx2; odr-1p::RFP) (II)</td>
			<td>Park et al. (DOI: 10.1016/j.cub.2023.07.052)</td>
			<td>MTS1161&#160;</td>
			<td>Will be deposited at CGC (https://cgc.umn.edu/)</td>
		</tr>
	</tbody>
</table>

optimizing visualization of axonal transport of endogenous cargo by fluorescence microscopy in living caenorhabditis elegans

Axonal transport is a prerequisite to deliver axonal proteins from their site of synthesis in the neuronal cell body to their destination in the axon. Consequently, loss of axonal transport impairs neuronal growth and function. Studying axonal transport therefore improves our understanding of neuronal cell biology. With recent improvements in CRISPR Cas9 genome editing, endogenous labeling of axonal cargos has become accessible, enabling to move beyond ectopic expression-based visualization of transport. However, endogenous labeling often comes at the cost of low signal intensity and necessitates optimization strategies to obtain robust data. Here, we describe a protocol to optimize the visualization of axonal transport by discussing acquisition parameters and a bleaching approach to improve the signal of endogenous labeled cargo over diffuse cytoplasmic background. We apply our protocol to optimize the visualization of synaptic vesicle precursors (SVPs) labeled by green fluorescent protein (GFP)-tagged RAB-3 to highlight how fine-tuning acquisition parameters can improve the analysis of endogenously labeled axonal cargo in Caenorhabditis elegans (C. elegans).

Overview of the model system and measurement procedure 
To visualize axonal transport of synaptic vesicle precursors, we traced endogenously GFP labeled RAB-3. Here we make use of a recently generated GFP::Flip-on::RAB-3 strain6, in which expression of the recombinase Flippase under a cell specific promoter (glr-4p) labels endogenous RAB-3 in the DA9 motor neuron. DA9 is a bipolar motor neuron, with its cell body located in the posterior of the animal on the ventral ...

Watch this Scientific Journal Video about Optimizing Visualization of Axonal Transport of Endogenous Cargo by Fluorescence Microscopy in Living Caenorhabditis elegans at JoVE.com

Author Spotlight: Advanced Techniques for Visualizing Endogenous Axonal Transport Dynamics

The paper describes the optimization of fluorescence microscopy acquisition parameters to visualize the axonal transport of endogenous labeled cargos at single-neuron resolution in a living nematode.

Optimizing Visualization of Axonal Transport of Endogenous Cargo by Fluorescence Microscopy in Living Caenorhabditis elegans

Axonal transport is a prerequisite to deliver axonal proteins from their site of synthesis in the neuronal cell body to their destination in the axon. ...

Classical start is commonly used for axonal transport proteins for axonal transport visualization, but endogenous cargo behavior differs. Endogenous protein levels are low, making visualization challenging. We offer ideas to optimize the acquisition conditions for better visualization of endogenous GPR3, which labels synaptic vascular precursors.The abundance of cytosolic proteins in the axonal transport field, makes it difficult to visualize their dynamics, using conventional fluorescence microscopy. This protocol suggests using ablation step, along with temporally controlled labeling approaches to overcome these challenges. We have recently been able to visualize the axonal transport of the endogenous spectry.By optimizing the acquisition parameters described in this protocol, paired our temporal control labeling approach. This allowed us to identify the first components of the machinery that mediate the transport.

optimizing-visualization-axonal-transport-endogenous-cargo

Research

JoVE Journal

Biology

644 Views.  Yale School of Medicine. Classical start is commonly used for axonal transport proteins for axonal transport visualization, but endogenous cargo behavior differs. Endogenous protein levels are low, making visualization challenging. We offer ideas to optimize the acquisition conditions for better visualization of endogenous GPR3, which labels synaptic vascular precursors.The abundance of cytosolic proteins in the axonal transport field, makes it difficult to visualize their dynamics, using conventional fluoresc...

Video: Author Spotlight: Advanced Techniques for Visualizing Endogenous Axonal Transport Dynamics

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

The cuticle of C. elegans is a highly resistant structure that surrounds the exterior of the animal1-4. The cuticle not only protects the animal from the environment, but also determines body shape and plays a role in motility4-6. Several layers secreted by epidermal cells comprise the cuticle, including an outermost lipid layer7.
Circumferential ridges in the cuticle called annuli pattern the length of the animal and are present during all stages of development8. Alae are longitudinal ridges that are present during specific stages of development, including L1, dauer, and adult stages2,9. Mutations in genes that affect cuticular collagen organization can alter cuticular structure and animal body morphology5,6,10,11. While cuticular imaging using compound microscopy with DIC optics is possible, current methods that highlight cuticular structures include fluorescent transgene expression12, antibody staining13, and electron microscopy1. Labeled wheat germ agglutinin (WGA) has also been used to visualize cuticular glycoproteins, but is limited in resolving finer cuticular structures14. Staining of cuticular surface using fluorescent dye has been observed, but never characterized in detail15. We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. This optimized protocol for DiI staining is a simple, robust method for high resolution fluorescent visualization of annuli, alae, vulva, male tail, and hermaphrodite tail spike in C. elegans.

Visualization of Caenorhabditis elegans Cuticular Structures Using the Lipophilic Vital Dye DiI

We present a method to visualize cuticle in live C. elegans using the red fluorescent lipophilic dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate), which is commonly used in C. elegans to visualize environmentally exposed neurons. With this optimized protocol, alae and annular cuticular structures are stained by DiI and observed using compound microscopy.

The cuticle of C. elegans is a highly resistant structure that surrounds the exterior of the animal1-4. The cuticle not only protects the animal from the ...

Basic Caenorhabditis elegans Methods: Synchronization and Observation

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has since been used extensively as a model organism 1. C. elegans possesses key attributes such as simplicity, transparency and short life cycle that have made it a suitable experimental system for fundamental biological studies for many years 2. Discoveries in this nematode have broad implications because many cellular and molecular processes that control animal development are evolutionary conserved 3.
C. elegans life cycle goes through an embryonic stage and four larval stages before animals reach adulthood. Development can take 2 to 4 days depending on the temperature. In each of the stages several characteristic traits can be observed. The knowledge of its complete cell lineage 4,5 together with the deep annotation of its genome turn this nematode into a great model in fields as diverse as the neurobiology 6, aging 7,8, stem cell biology 9 and germ line biology 10. 
An additional feature that makes C. elegans an attractive model to work with is the possibility of obtaining populations of worms synchronized at a specific stage through a relatively easy protocol. The ease of maintaining and propagating this nematode added to the possibility of synchronization provide a powerful tool to obtain large amounts of worms, which can be used for a wide variety of small or high-throughput experiments such as RNAi screens, microarrays, massive sequencing, immunoblot or in situ hybridization, among others. 
Because of its transparency, C. elegans structures can be distinguished under the microscope using Differential Interference Contrast microscopy, also known as Nomarski microscopy. The use of a fluorescent DNA binder, DAPI (4',6-diamidino-2-phenylindole), for instance, can lead to the specific identification and localization of individual cells, as well as subcellular structures/defects associated to them.

Basic Caenorhabditis elegans Methods: Synchronization and Observation

The easiness of maintaining and propagating the nematode C. elegans make it a nice model organism to work with. The possibility of synchronizing worms allows the work with a significant amount of subjects at the same developmental stage, what facilitates the study of one particular process in many animals.

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein subcellular localization changes; in particular, a nuclear translocation under a heat stress condition. Proteins react correspondingly to external and internal signals. A common mechanism is to change its subcellular localization. This article describes a protocol to track protein localization that does not require an antibody, radioactive labeling, or a confocal microscope. In this article, GFP is used to tag the target protein EXL-1 in C. elegans, a member of the chloride intracellular channel proteins (CLICs) family, including mammalian CLIC4. An integrated translational exl-1::gfp transgenic line (with a promoter and a full gene sequence) was created by transformation and &#947;-radiation, and stably expresses the gene and gfp. Recent research showed that upon heat stress, not oxidative stress, EXL-1::GFP accumulates in the nucleus. Overlapping the GFP signal with both the nuclei structure and the DAPI signals confirms the EXL-1 subcellular localization changes under stress. This protocol presents two different fixation methods for DAPI staining: ethanol fixation and acetone fixation. The DAPI staining protocol presented in this article is fast and efficient and preserves both the GFP signal and the protein subcellular localization changes. This method only requires a fluorescence microscope with Nomarski, a FITC filter, and a DAPI filter. It is suitable for a small laboratory setting, undergraduate student research, high school student research, and biotechnology classrooms.

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4&#039;,6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

This protocol demonstrates how to track a protein nuclear translocation under heat stress by using a green fluorescence protein (GFP) fusion protein as a marker and 4',6-diamidino-2-phenylindole (DAPI) staining. The DAPI staining protocol is fast and preserves the GFP and protein subcellular localization signals.

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein ...

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...

Oligomerization Dynamics of Cell Surface Receptors in Living Cells by Total Internal Reflection Fluorescence Microscopy Combined with Number and Brightness Analysis

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of clustering. Here, we describe an imaging approach to determine the average oligomeric state of mEGFP-tagged-receptor homocomplexes in the membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis. N&#38;B is a method similar to fluorescence-correlation spectroscopy (FCS) and photon counting histogram (PCH), which are based on the statistical analysis of the fluctuations of the fluorescence intensity of fluorophores diffusing in and out of an illumination volume during an observation time. In particular, N&#38;B is a simplification of PCH to obtain information on the average number of proteins in oligomeric mixtures. The intensity fluctuation amplitudes are described by the molecular brightness of the fluorophore and the average number of fluorophores within the illumination volume. Thus, N&#38;B considers only the first and second moments of the amplitude distribution, namely, the mean intensity and the variance. This is, at the same time, the strength and the weakness of the method. Because only two moments are considered, N&#38;B cannot determine the molar fraction of unknown oligomers in a mixture, but it only estimates the average oligomerization state of the mixture. Nevertheless, it can be applied to relatively small time series (compared to other moment methods) of images of live cells on a pixel-by-pixel basis, simply by monitoring the time fluctuations of the fluorescence intensity. It reduces the effective time-per-pixel to a few microseconds, allowing acquisition in the time range of seconds to milliseconds, which is necessary for fast oligomerization kinetics. Finally, large cell areas as well as sub-cellular compartments can be explored.

We describe an imaging approach for the determination of the average oligomeric state of mEGFP-tagged-receptor oligomers induced by ligand binding in the plasma membrane of living cells. The protocol is based on Total Internal Reflection Fluorescence (TIRF) microscopy combined with Number and Brightness (N&#38;B) analysis.

Despite the importance and ubiquity of receptor oligomerization, few methods are applicable for detecting clustering events and measuring the degree of ...

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits obesity and many associated diseases including cardiovascular disorders, type II diabetes, and cancer. To advance the current understanding of lipid metabolic regulation, quantitative methods to precisely measure in vivo lipid storage levels in time and space have become increasingly important and useful. Traditional approaches to analyze lipid storage are semi-quantitative for microscopic assessment or lacking spatio-temporal information for biochemical measurement. Stimulated Raman scattering (SRS) microscopy is a label-free chemical imaging technology that enables rapid and quantitative detection of lipids in live cells with a subcellular resolution. As the contrast is exploited from intrinsic molecular vibrations, SRS microscopy also permits four-dimensional tracking of lipids in live animals. In the last decade, SRS microscopy has been widely used for small molecule imaging in biomedical research and overcome the major limitations of conventional fluorescent staining and lipid extraction methods. In the laboratory, we have combined SRS microscopy with the genetic and biochemical tools available to the powerful model organism, Caenorhabditis elegans, to investigate the distribution and heterogeneity of lipid droplets across different cells and tissues and ultimately to discover novel conserved signaling pathways that modulate lipid metabolism. Here, we present the working principles and the detailed setup of the SRS microscope and provide methods for its use in quantifying lipid storage at distinct developmental timepoints of wild-type and insulin signaling deficient mutant C. elegans.

Label-Free Imaging of Lipid Storage Dynamics in Caenorhabditis elegans using Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy allows selective, label-free imaging of specific chemical moieties and it has been effectively employed to image lipid molecules in vivo. Here, we provide a brief introduction to the principle of SRS microscopy and describe methods for its use in imaging lipid storage in Caenorhabditis elegans.

Lipid metabolism is a fundamental physiological process necessary for cellular and organism health. Dysregulation of lipid metabolism often elicits ...

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures the interaction of homologous chromosomes is the assembly of the synaptonemal complex (SC) in meiotic prophase I. Even though there is little sequence homology between protein components within the SC among different species, the general structure of the SC has been highly conserved during evolution. In electron micrographs, the SC appears as a tripartite, ladder-like structure composed of lateral elements or axes, transverse filaments, and a central element. 
However, precisely identifying the localization of individual components within the complex by electron microscopy to determine the molecular structure of the SC remains challenging. By contrast, fluorescence microscopy allows for the identification of individual protein components within the complex. However, since the SC is only ~100 nm wide, its substructure cannot be resolved by diffraction-limited conventional fluorescence microscopy. Thus, determining the molecular architecture of the SC requires super-resolution light microscopy techniques such as structured illumination microscopy (SIM), stimulated-emission depletion (STED) microscopy, or single-molecule localization microscopy (SMLM). 
To maintain the structure and interactions of individual components within the SC, it is important to observe the complex in an environment that is close to its native environment in the germ cells. Therefore, we demonstrate an immunohistochemistry and imaging protocol that enables the study of the substructure of the SC in intact, extruded Caenorhabditis elegans germline tissue with SMLM and STED microscopy. Directly fixing the tissue to the coverslip reduces the movement of the samples during imaging and minimizes aberrations in the sample to achieve the high resolution necessary to visualize the substructure of the SC in its biological context.

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

Super-resolution microscopy can provide a detailed insight into the organization of components within the synaptonemal complex in meiosis. Here, we demonstrate a protocol to resolve individual proteins of the Caenorhabditis elegans synaptonemal complex.

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures ...