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><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>&amp;nbsp;NIS-elements AR</td><td>Nikon</td><td></td><td>Software for the Nikon Ti2&amp;nbsp;</td></tr><tr><td>Plain precleaned microscopy slides</td><td>Thermo Scientific</td><td>420-004T</td><td></td></tr><tr><td>&lt;strong&gt;Nematode strain&lt;/strong&gt;</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&amp;nbsp;</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 ...

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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. ...

optimizing-visualization-axonal-transport-endogenous-cargo

Research

JoVE Journal

Biology

696 Views.  Yale School of Medicine. Klassieke start wordt vaak gebruikt voor axonale transporteiwitten voor axonale transportvisualisatie, maar het endogene ladingsgedrag verschilt. Het endogene eiwitgehalte is laag, waardoor visualisatie een uitdaging is. We bieden ideeën om de acquisitiecondities te optimaliseren voor een betere visualisatie van endogene GPR3, dat synaptische vasculaire voorlopers labelt.De overvloed aan cytosolische eiwitten in het axonale transportveld maakt het moe...

Video: Auteur in de schijnwerpers: geavanceerde technieken voor het visualiseren van endogene axonale transportdynamiek

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

Neuroscience

Automated Quantification of Synaptic Fluorescence in C. elegans

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall neural circuit function. Synapse strength critically depends on the abundance of neurotransmitter receptors clustered at synaptic sites on the postsynaptic membrane. Receptor levels are established developmentally, and can be altered by receptor trafficking between surface-localized, subsynaptic, and intracellular pools, representing important mechanisms of synaptic plasticity and neuromodulation. Rigorous methods to quantify synaptically-localized neurotransmitter receptor abundance are essential to study synaptic development and plasticity. Fluorescence microscopy is an optimal approach because it preserves spatial information, distinguishing synaptic from non-synaptic pools, and discriminating among receptor populations localized to different types of synapses. The genetic model organism Caenorhabditis elegans is particularly well suited for these studies due to the small size and relative simplicity of its nervous system, its transparency, and the availability of powerful genetic techniques, allowing examination of native synapses in intact animals.
Here we present a method for quantifying fluorescently-labeled synaptic neurotransmitter receptors in C. elegans. Its key feature is the automated identification and analysis of individual synapses in three dimensions in multi-plane confocal microscope output files, tabulating position, volume, fluorescence intensity, and total fluorescence for each synapse. This approach has two principal advantages over manual analysis of z-plane projections of confocal data. First, because every plane of the confocal data set is included, no data are lost through z-plane projection, typically based on pixel intensity averages or maxima. Second, identification of synapses is automated, but can be inspected by the experimenter as the data analysis proceeds, allowing fast and accurate extraction of data from large numbers of synapses. Hundreds to thousands of synapses per sample can easily be obtained, producing large data sets to maximize statistical power. Considerations for preparing C. elegans for analysis, and performing confocal imaging to minimize variability between animals within treatment groups are also discussed. Although developed to analyze C. elegans postsynaptic receptors, this method is generally useful for any type of synaptically-localized protein, or indeed, any fluorescence signal that is localized to discrete clusters, puncta, or organelles.
The procedure is performed in three steps: 1) preparation of samples, 2) confocal imaging, and 3) image analysis. Steps 1 and 2 are specific to C. elegans, while step 3 is generally applicable to any punctate fluorescence signal in confocal micrographs.

Automated Quantification of Synaptic Fluorescence in C. elegans

The abundance of neurotransmitter receptors clustered at synapses strongly influences synaptic strength. This method quantifies fluorescently-labeled neurotransmitter receptors in three dimensions with single-synapse resolution in C. elegans, allowing hundreds of synapses to be rapidly characterized within a single sample without distortions introduced by z-plane projection.

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall ...

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Axonal transport and intraflagellar transport (IFT) are essential for axon and cilia morphogenesis and function. Kinesin superfamily proteins and dynein are molecular motors that regulate anterograde and retrograde transport, respectively. These motors use microtubule networks as rails. Caenorhabditis elegans (C. elegans) is a powerful model organism to study axonal transport and IFT in vivo. Here, I describe a protocol to observe axonal transport and IFT in living C. elegans. Transported cargo can be visualized by tagging cargo proteins using fluorescent proteins such as green fluorescent protein (GFP). C. elegans is transparent and GFP-tagged cargo proteins can be expressed in specific cells under cell-specific promoters. Living worms can be fixed by microbeads on 10% agarose gel without killing or anesthetizing the worms. Under these conditions, cargo movement can be directly observed in the axons and cilia of living C. elegans without dissection. This method can be applied to the observation of any cargo molecule in any cells by modifying the target proteins and/or the cells they are expressed in. Most basic proteins such as molecular motors and adaptor proteins that are involved in axonal transport and IFT are conserved in C. elegans. Compared to other model organisms, mutants can be obtained and maintained more easily in C. elegans. Combining this method with various C. elegans mutants can clarify the molecular mechanisms of axonal transport and IFT.

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Caenorhabditis elegans (C. elegans) is a good model to study axonal and intracellular transport. Here, I describe a protocol for in vivo recording and analysis of axonal and intraflagellar transport in C. elegans.

Axonal transport and intraflagellar transport (IFT) are essential for axon and cilia morphogenesis and function. Kinesin superfamily proteins and dynein ...

Developmental Biology

Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using "Cargo Mapping" Analysis

Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative analysis of motor/cargo associations at the single vesicle level. The goal of this protocol is to use quantitative fluorescence microscopy to correlate (&#8220;map&#8221;) the position and directionality of movement of live cargo to the composition and relative amounts of motors associated with the same cargo. &#8220;Cargo mapping&#8221; consists of live imaging of fluorescently labeled cargoes moving in axons cultured on microfluidic devices, followed by chemical fixation during recording of live movement, and subsequent immunofluorescence (IF) staining of the exact same axonal regions with antibodies against motors. Colocalization between cargoes and their associated motors is assessed by assigning sub-pixel position coordinates to motor and cargo channels, by fitting Gaussian functions to the diffraction-limited point spread functions representing individual fluorescent point sources. Fixed cargo and motor images are subsequently superimposed to plots of cargo movement, to &#8220;map&#8221; them to their tracked trajectories. The strength of this protocol is the combination of live and IF data to record both the transport of vesicular cargoes in live cells and to determine the motors associated to these exact same vesicles. This technique overcomes previous challenges that use biochemical methods to determine the average motor composition of purified heterogeneous bulk vesicle populations, as these methods do not reveal compositions on single moving cargoes. Furthermore, this protocol can be adapted for the analysis of other transport and/or trafficking pathways in other cell types to correlate the movement of individual intracellular structures with their protein composition. Limitations of this protocol are the relatively low throughput due to low transfection efficiencies of cultured primary neurons and a limited field of view available for high-resolution imaging. Future applications could include methods to increase the number of neurons expressing fluorescently labeled cargoes.

Intracellular transport of cargoes, such as vesicles or organelles, is carried out by molecular motor proteins that track on polarized microtubules. This protocol describes the correlation of the directionality of transport of individual cargo particles moving inside neurons, to the relative amount and type of associated motor proteins.

Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative ...

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study of this movement in situ was only possible using radioisotopic pulse-labeling, which permits analysis of axonal transport in whole nerves with a temporal resolution of days and a spatial resolution of millimeters. To study neurofilament transport in situ with higher temporal and spatial resolution, we developed a hThy1-paGFP-NFM transgenic mouse that expresses neurofilament protein M tagged with photoactivatable GFP in neurons. Here we describe fluorescence photoactivation pulse-escape and pulse-spread methods to analyze neurofilament transport in single myelinated axons of tibial nerves from these mice ex vivo. Isolated nerve segments are maintained on the microscope stage by perfusion with oxygenated saline and imaged by spinning disk confocal fluorescence microscopy. Violet light is used to activate the fluorescence in a short axonal window. The fluorescence in the activated and flanking regions is analyzed over time, permitting the study of neurofilament transport with temporal and spatial resolution on the order of minutes&#160;and microns, respectively. Mathematical modeling can be used to extract kinetic parameters of neurofilament transport including the velocity, directional bias and pausing behavior from the resulting data. The pulse-escape and pulse-spread methods can also be adapted to visualize neurofilament transport in other nerves. With the development of additional transgenic mice, these methods could also be used to image and analyze the axonal transport of other cytoskeletal and cytosolic proteins in axons.

We describe fluorescence photoactivation methods to analyze the axonal transport of neurofilaments in single myelinated axons of peripheral nerves from transgenic mice that express a photoactivatable neurofilament protein.

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study ...

In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons

In neurons, microtubule orientation has been a key assessor to identify axons that have plus-end out microtubules and dendrites that generally have mixed orientation. Here we describe methods to label, image, and analyze the microtubule dynamics and growth during the development and regeneration of touch neurons in C. elegans. Using genetically encoded fluorescent reporters of microtubule tips, we imaged the axonal microtubules. The local changes in microtubule behavior that initiates axon regeneration following axotomy can be quantified using this protocol. This assay is adaptable to other neurons and genetic backgrounds to investigate the regulation of microtubule dynamics in various cellular processes.

In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons

A protocol for imaging the dynamic microtubules in vivo using fluorescently labeled End binding protein has been presented. We described the methods to label, image, and analyze the dynamic microtubules in the Posterior lateral microtubule (PLM) neuron of C. elegans.

In neurons, microtubule orientation has been a key assessor to identify axons that have plus-end out microtubules and dendrites that generally have mixed ...

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal transport have devastating consequences for individual neurons and their networks, and contribute to a plethora of neurological disorders. As many of these conditions involve both cell autonomous and non-autonomous mechanisms, and often display a spectrum of pathology across neuronal subtypes, methods to accurately identify and analyze neuronal subsets are imperative.
This paper details protocols to assess in vivo axonal transport of signaling endosomes and mitochondria in sciatic nerves of anesthetized mice. Stepwise instructions are provided to 1) distinguish motor from sensory neurons in vivo, in situ, and ex vivo by using mice that selectively express fluorescent proteins within cholinergic motor neurons; and 2) separately or concurrently assess in vivo axonal transport of signaling endosomes and mitochondria. These complementary intravital approaches facilitate the simultaneous imaging of different cargoes in distinct peripheral nerve axons to quantitatively monitor axonal transport in health and disease.

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Using transgenic fluorescent mice, detailed protocols are described to assess in vivo axonal transport of signaling endosomes and mitochondria within motor and sensory axons of the intact sciatic nerve in live animals.

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal ...

Measuring Axonal Cargo Transport in Mouse Primary Cortical Cultured Neurons

Neuronal cells are highly polarized cells that stereotypically harbor several dendrites and an axon. The length of an axon necessitates efficient bidirectional transport by motor proteins. Various reports have suggested that defects in axonal transport are associated with neurodegenerative diseases. Also, the mechanism of the coordination of multiple motor proteins has been an attractive topic. Since the axon has uni-directional microtubules, it is easier to determine which motor proteins are involved in the movement. Therefore, understanding the mechanisms underlying the transport of axonal cargo is crucial for uncovering the molecular mechanism of neurodegenerative diseases and the regulation of motor proteins. Here, we introduce the entire process of axonal transport analysis, including the culturing of mouse primary cortical neurons, transfection of plasmids encoding cargo proteins, and directional and velocity analyses without the effect of pauses. Furthermore, the open-access software "KYMOMAKER" is introduced, which enables the generation of a kymograph to highlight transport traces according to their direction and allow easier visualization of axonal transport.

The present protocol describes the whole procedure of analysis for axonal transport. In particular, the calculation of transport velocity, not including the pausing, and the visualization method using open-access software "KYMOMAKER" are shown here.

Neuronal cells are highly polarized cells that stereotypically harbor several dendrites and an axon. The length of an axon necessitates efficient ...

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

Defining the cellular mechanisms underlying disease is essential for the development of novel therapeutics. A strategy frequently used to unravel these mechanisms is to introduce mutations in candidate genes and qualitatively describe changes in the morphology of tissues and cellular organelles. However, qualitative descriptions may not capture subtle phenotypic differences, might misrepresent phenotypic variations across individuals in a population, and are frequently assessed subjectively. Here, quantitative approaches are described to study the morphology of tissues and organelles in the nematode Caenorhabditis elegans using laser scanning confocal microscopy combined with commercially available bio-image processing software. A quantitative analysis of phenotypes affecting synapse integrity (size and integrated fluorescence levels), muscle development (muscle cell size and myosin filament length), and mitochondrial morphology (circularity and size) was performed to understand the effects of genetic mutations on these cellular structures. These quantitative approaches are not limited to the applications described here, as they could readily be used to quantitatively assess the morphology of other tissues and organelles in the nematode, as well as in other model organisms.

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

This study outlines quantitative measurements of synaptic size and localization, muscle morphology, and mitochondrial shape in C. elegans using freely available image processing tools. This approach allows future studies in&#160;C. elegans&#160;to quantitatively compare the extent of tissue and organelle structural changes as a result of genetic mutations.

Defining the cellular mechanisms underlying disease is essential for the development of novel therapeutics. A strategy frequently used to unravel these ...

Analyzing Mitochondrial Morphology in C. elegans

Imaging and Quantifying Mitochondrial Morphology in C. elegans During Aging

Mitochondria, important cellular organelles found in most eukaryotic cells, are major sites of energy production through aerobic respiration. Beyond this well-known role as the 'cellular powerhouse,' mitochondria are also involved in many other essential cellular processes, including the regulation of cellular metabolism, proliferation, immune signaling, and hormonal signaling. Deterioration in mitochondrial function during aging or under mitochondrial stress is often characterized by distinct changes in mitochondrial morphology and volume. The nematode C. elegans is an ideal model for studying these changes due to its transparent body and short lifespan, which facilitate live microscopy throughout its lifetime. However, even within the C. elegans field, numerous transgenic constructs and methods for mitochondrial imaging are available, each with its own limitations. Here, single-copy, matrix-localized GFP constructs are presented as a robust and reliable method for imaging mitochondrial morphology in C. elegans. This study specifically focuses on experimentally controllable factors to minimize errors and reduce variability between replicates and across studies when performing mitochondrial imaging during the aging process. Additionally, mitoMAPR is recommended as a robust method to quantify changes in mitochondrial morphology across tissue types during aging.

This protocol provides a standardized approach for imaging mitochondrial morphology in multiple tissues of C. elegans during aging.

Mitochondria, important cellular organelles found in most eukaryotic cells, are major sites of energy production through aerobic respiration. Beyond this ...

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

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