The authors would like to thank A. Benham for assisting with the development of the protocol. This work was funded by NIH grant NS06747 to B. A. B.

1. Preparation of Worms for Imaging
This segment of the protocol is based on published C. elegans culture techniques1,2, and is outlined in Figure 1.
<ol>
	<li>Grow worms on high peptone NGM agar plates (10 cm) seeded with NA22 E. coli bacteria until nearly starved. If immunostaining, one plate will reliably produce enough worms for 1 individual stain, taking into account losses along the way, and the strict criteria used to select worms for imaging (see Part 2).</li>
	<li>Harvest worms by pouring a few ml of ddH20 onto plate, swirling briefly, and pouring liquid into a 15 ml conical tube (repeat if necessary to transfer most of the worms to the tube). Pellet in clinical centrifuge for 3 min, 1000x g. Wash pellet 1-3x with ddH2O until supernatant is clear to remove residual bacteria. Use polypropylene transfer pipettes to remove supernatants, as worms stick to glass (from this step onward it is critical to minimize time between steps to avoid losses in egg viability).</li>
	<li>After the last wash, resuspend worm pellet in 5 ml alkaline hypochlorite solution to lyse worms and release eggs. Rock gently, and examine tubes under a dissecting microscope about once per minute. When about 50% of worms are lysed (they will appear bent and broken open), terminate lysis by filling the tube to the top with Egg Buffer, inverting several times, and pelleting for 3 min, 1000x g. Lysis time must not exceed 5 min.</li>
	<li>Remove supernatant with transfer pipette, and wash pellet 3x with Egg Buffer.</li>
	<li>To separate eggs from debris, float them on a cushion of 30% sucrose in ddH2O: after the final wash from step 1.4, carefully remove supernatant and resuspend pellet in 5 ml ddH20. Add 5 ml sterile 60% sucrose in ddH2O, and mix well. Spin in clinical centrifuge 6 min, 1000x g. Eggs will collect at the meniscus (they will have a cloudy appearance), while debris will pellet.</li>
	<li>Using a plastic transfer pipette, suck up eggs in a minimal volume and transfer to unseeded NGM agar plates. Any eggs adhering to side of tube can be rinsed down gently and transferred as well. One 10 cm unseeded plate per strain analyzed is sufficient.</li>
	<li>Incubate the unseeded plates with the eggs overnight (~16 hr) at 20 &deg;C. Vent the plate for the first few hours to dry it by leaving the lid slightly off for a few hours, but be sure to cover overnight.</li>
	<li>After hatching, transfer the L1 larvae to a 15 ml conical tube in S Basal (add a few ml S Basal to plate, swirl, pour into tube). Pellet in clinical centrifuge (3 min, 1000x g), resuspend in a minimal volume of S Basal, and transfer to 10 cm NGM agar plates seeded with NA22 bacteria. Use 2 plates here for each original plate from step 1.1. Incubate until worms reach desired age.</li>
	<li>Monitor the cultures once or twice per day. If they are in danger of starving, transfer to fresh NA22 plates. For wild-type N2 worms, starvation is preceded by the formation of a line or wave of worms that moves across the plate as animals migrate en masse away from food-depleted zones. Once this forms, the food will be completely depleted within a few hours.</li>
	<li>Worms are now ready for immunostaining or live imaging. Fixing and staining worms in suspension is optimal because large numbers of intact worms can be imaged easily. Staining procedures based on the Finney and Ruvkun protocol3-6 are preferable to freeze-crack procedures because large numbers of worms are needed for imaging.</li>
</ol>
2. Confocal Imaging
<ol>
	<li>Mount worms on slides for confocal imaging. Adjust density of worms to a few hundred per microliter. Make a pad of 2% agarose in S-basal surrounded by a thin ring of Vaseline to prevent evaporation during imaging (applied via a 3 ml syringe fitted with a cut-off 25 gauge needle). Pipette a few microliters of the worm suspension onto a cover slip (use anaesthetic if imaging live worms), and lower the agarose pad onto the cover slip, spreading the worms evenly over the pad.</li>
	<li>Select worms for imaging. Choose worms where the synapses of interest are oriented toward the objective lens, with radial displacement of no more than &plusmn;45&deg; (where 0&deg; means oriented directly toward the objective). Perform this assessment quickly (a few seconds) to avoid photobleaching. The scale of the initial cultures is optimized to ensure sufficient worms will satisfy these geometric criteria.</li>
	<li>Complete the confocal imaging of the specimen. Relatively flat samples that can be encompassed within about 14 0.4 &mu;m thick sections produce the best results. High speed, low resolution images (512 x 512 pixels) are sufficient for rapid accurate data collection. Avoid saturating the detector with excessive signal strengths, as this will cause underestimation of fluorescence signals.</li>
</ol>
3. Automated Identification and Analysis of Individual Synaptic Clusters
<ol>
	<li>Open multi-TIF output files from the confocal microscope using Volocity 4.0 (or higher) software (PerkinElmer).</li>
	<li>Crop away areas of the image that do not contain the structure of interest (Image -&gt; Extended Focus -&gt; Select area of interest using 'Freehand ROI' tool -&gt; Actions -&gt; Crop to Selection; Figure 2A). Note, Extended Focus produces a z-plane projection on the computer screen to aid your analysis, but does not affect the underlying data; the same is true for brightness and contrast adjustments, if necessary).</li>
	<li>Measure the length of the area analyzed using the Line Tool. The length of the line will be calculated and added to the data table (Figure 2B).</li>
	<li>Identify objects using the 'Objects by Intensity' filter (Measurements mode). Specify a threshold such that synaptic clusters are highlighted and non-synaptic background is not (Figure 2C). Inclusion of an independent synaptic marker labeled with another color may help unambiguously identify synapses. Typical synapses range from a few to a few hundred voxels. Set the threshold once using a control sample, and use it for all subsequent samples. Thresholding each sample separately is unacceptable because it introduces the potential for experimenter bias. Objects are selected and tabulated automatically (Figure 2D).</li>
	<li>Arrange and Export the data. Eliminate objects of 2 or fewer voxels, since these are usually background specks. These can be eliminated by further filtering, or during subsequent analysis. To facilitate their removal downstream, re-sort the data file by 'object size' so they will all group together. Export the data in CSV format, which can be opened by Microsoft Excel or other spreadsheet programs. Each image produces one output file.</li>
	<li>If Volocity software is unavailable, alternative methods to analyze Z-plane projections of confocal data may be utilized (e.g. ref. 7, or manual analysis using ImageJ), although 3-dimensional information is lost (see Discussion).</li>
</ol>
4. Representative Results
The quantification method presented should be able to distinguish between cluster populations of different brightness and different volumes. Representative images and corresponding quantitative data presented in Figure 3 demonstrate examples of differentiation on the basis of these parameters. As a general rule the results should conform to what is evident to the eye. In the case of UNC-49 GABA receptor immunostaining the ventral nerve cord of C. elegans, all animals typically appeared very similar in size and maturity, and total synaptic fluorescence values for a group of five worms (normalized to the length of nerve cord analyzed) showed Standard Error values of about 10% of the mean 4. Genetic mutation and other experimental treatments (e.g. drug exposure) may alter not only the volume and intensity values and frequency distributions of synaptic clusters, but possibly the developmental time-course and synchrony of the worms, leading to higher variability. However the quantitative results should always reflect what can be appreciated visually and if not, inspection of the images and the objects identified by Volocity should reveal where the error occurred and suggest corrective action such as re-thresholding or removal of artefactual objects.
<img alt="Figure 1" src="/files/ftp_upload/4090/4090fig1.jpg" /> 
Figure 1. Preparation of synchronous C. elegans cultures. Synchronous cultures are obtained from this procedure because C. elegans development arrests and the L1 larval stage in the absence of food, and resumes when food is introduced.
<img alt="Figure 2" src="/files/ftp_upload/4090/4090fig2.jpg" /> 
Figure 2. Quantitation of synaptic puncta. 
<ol class="ualpha">
	<li>Extraneous regions of the image are cropped to reduce file sizes and increase specificity. Left hand panel shows starting image, middle panel shows selected region, right panel shows image after cropping. Red signal is UNC-49 GABA receptor immunofluorescence, green signal is synaptobrevin-GFP (a presynaptic marker) expressed in presynaptic GABA neurons 4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/4090/4090fig2alarge.jpg" target="_blank"> Click here to view larger figure</a>.</li>
	<li>A line is drawn the length of the nerve cord segment to be analyzed, and its length is automatically recorded in the results table. This information is necessary to normalize synaptic data due to the length of analyzable nerve cord, which can vary several fold if worms become fragmented during staining. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/4090/4090fig2blarge.jpg" target="_blank"> Click here to view larger figure</a>.</li>
	<li>A threshold is applied to identify individual synaptic clusters. Left panel shows image before thresholding; examples of different thresholding levels are shown in separate panels (green signal omitted). Each identified object is depicted on the image as a colored region. Note the correspondence between the visually evident synaptic clusters in the left-most panel and the colored regions on the right-most panel. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/4090/4090fig2clarge.jpg" target="_blank"> Click here to view larger figure</a>.</li>
	<li>Each identified object is listed separately in the results table. Individual objects may be selected, and are highlighted on the image for inspection if desired. Note that table contains information for the red ('rhodamine') and green ('EGFP') channels; the green channel information is potentially misleading since objects were identified based strictly on red fluorescence. This analysis may also be performed on single color images. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/4090/4090fig2dlarge.jpg" target="_blank"> Click here to view larger figure</a>.</li>
</ol>
<img alt="Figure 3" src="/files/ftp_upload/4090/4090fig3.jpg" /> 
Figure 3. Representative results. Representative micrographs from wild-type C. elegans stained for UNC-49 GABA receptors before (A) and after (B) treatment with muscimol, a GABA receptor agonist that causes receptors to become downregulated after long exposure. Panels show cropped images before (left) and after (right) thresholding and removal of small background specks. (C) Plots of quantitative synaptic parameters for the specimens shown in (A) and (B): total fluorescence normalized to nerve cord length (left), and cumulative probability histograms of individual synapse fluorescence content (middle) and synapse volume (right), demonstrating statistically significant reduction in synaptic content and volume induced by agonist exposure (n = 60 synapses for untreated, n = 115 synapses for muscimol-treated; p &lt; 0.001, Kolmogorov-Smirnov test <a href="http://www.physics.csbsju.edu/stats/KS-test.html" target="_blank">http://www.physics.csbsju.edu/stats/KS-test.html</a>).

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	<li>Christensen, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+primary+culture+system+for+functional+analysis+of+C.+elegans+neurons+and+muscle+cells.">A primary culture system for functional analysis of C. elegans neurons and muscle cells.</a> Neuron. 33, 503-514 (2002).</li><li>Lewis, J. A., Fleming, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+culture+methods.">Basic culture methods.</a> Methods Cell Biol. 48, 3-29 (1995).</li><li>Bettinger, J. C., Lee, K., Rougvie, A. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stage-specific+accumulation+of+the+terminal+differentiation+factor+LIN-29+during+Caenorhabditis+elegans+development.">Stage-specific accumulation of the terminal differentiation factor LIN-29 during Caenorhabditis elegans development.</a> Development. 122, 2517-2527 (1996).</li><li>Davis, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulated+lysosomal+trafficking+as+a+mechanism+for+regulating+GABAA+receptor+abundance+at+synapses+in+Caenorhabditis+elegans.">Regulated lysosomal trafficking as a mechanism for regulating GABAA receptor abundance at synapses in Caenorhabditis elegans.</a> Mol. Cell Neurosci. 44, 307-317 (2010).</li><li>Finney, M., Ruvkun, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+unc-86+gene+product+couples+cell+lineage+and+cell+identity+in+C.+elegans.">The unc-86 gene product couples cell lineage and cell identity in C. elegans.</a> Cell. 63, 895-905 (1990).</li><li>Rowland, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Presynaptic+terminals+independently+regulate+synaptic+clustering+and+autophagy+of+GABAA+receptors+in+Caenorhabditis+elegans.">Presynaptic terminals independently regulate synaptic clustering and autophagy of GABAA receptors in Caenorhabditis elegans.</a> J. Neurosci. 26, 1711-1720 (2006).</li><li>Burbea, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ubiquitin+and+AP180+regulate+the+abundance+of+GLR-1+glutamate+receptors+at+postsynaptic+elements+in+C.+elegans.">Ubiquitin and AP180 regulate the abundance of GLR-1 glutamate receptors at postsynaptic elements in C. elegans.</a> Neuron. 35, 107-120 (2002).</li><li>Miesenbock, G., De Angelis, D. A., Rothman, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+secretion+and+synaptic+transmission+with+pH-sensitive+green+fluorescent+proteins.">Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins.</a> Nature. 394, 192-195 (1998).</li><li>Oda, S., Tomioka, M., Iino, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+plasticity+regulated+by+the+insulin-like+signaling+pathway+underlies+salt+chemotaxis+learning+in+Caenorhabditis+elegans.">Neuronal plasticity regulated by the insulin-like signaling pathway underlies salt chemotaxis learning in Caenorhabditis elegans.</a> J. Neurophysiol. 106, 301-308 (2011).</li><li>Sankaranarayanan, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+pHluorins+for+optical+measurements+of+presynaptic+activity.">The use of pHluorins for optical measurements of presynaptic activity.</a> Biophys. J. 79, 2199-2208 (2000).</li><li>McDonald, N. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+and+functional+characterization+of+fluorescent,+N-terminally+tagged+CB1+receptor+chimeras+for+live-cell+imaging.">Generation and functional characterization of fluorescent, N-terminally tagged CB1 receptor chimeras for live-cell imaging.</a> Mol. Cell Neurosci. 35, 237-248 (2007).</li><li>Shakiryanova, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+neuropeptide+release+induced+by+octopamine+without+Ca2++entry+into+the+nerve+terminal.">Synaptic neuropeptide release induced by octopamine without Ca2+ entry into the nerve terminal.</a> Proc. Natl. Acad. Sci. U. S. A. 108, 4477-4481 (2011).</li></ol>

No conflicts of interest declared.

The method presented here is designed to extract quantitative multi-parameter data for large populations of synapses in C. elegans, while maximizing consistency within treatment groups. Three features contribute to these objectives. First, immunostaining is performed on synchronous worm populations to ensure that all animals are the same age. This step is critical because developmental regulation of expression levels may obscure effects of the experimental treatment (e.g. UNC-49 GABA receptor immunofluorescence varies several-fold during development (K. Davis et al., in preparation)). Additionally, permeabilization and fixation can be highly variable, making quantitative comparisons difficult. Using synchronized cultures minimizes this inherent variability, improving the reliability of the technique. Other ways to address this variability include using a second antibody to an unrelated target that can be independently visualized as an internal control for permeabilization, or imaging GFP-tagged proteins in live worms, which avoids the need for permeabilization altogether. However this latter approach introduces new problems because the endogenous protein is no longer being measured directly, so potential artefacts due to inconsistencies in transgene expression/copy number, structural alteration due to the fusion of GFP, or non-physiological transgene expression levels need to be considered. No single method is perfect; ideally, corroborating data can be obtained using multiple approaches. In this protocol, the size of the starter cultures has been optimized to result in sufficient numbers of fixed and stained worms for confocal analysis. Large numbers of worms are required because the geometric criteria used to select worms for imaging are strict, which is the second feature of the protocol to minimize variability. The same anatomical structure must be imaged in each animal (either the most biologically relevant site, or an arbitrarily chosen site in the case of broad tissue distribution of the protein of interest), and that structure must be oriented toward the objective lens with only slight deviation to either side. This criterion improves consistency by eliminating variability in the amount of intervening tissue that can scatter excitation and emission light, affecting signal strength. It is important that this is the ONLY criterion used to select worms, as it is easy to bias the results based on expectation of a certain outcome. Indeed experiments are best performed blind where possible. The third feature that contributes to the power of this technique is the automated identification and quantitative characterization of synaptic clusters in three dimensions using Volocity. This analysis is simple and rapid, capable of identifying and tabulating hundreds to thousands of individual synapses for each experimental condition. More importantly, Volocity includes data from all of the planes of multi-plane confocal data stacks, rather than discarding data to generate the 2-dimensional z-projections used in earlier protocols. Downstream of this analysis, experimental groups may be compared in terms of total synaptic fluorescence per animal, the spatial distribution of synapses, and frequency distributions of volumes, total fluorescence, and maximal fluorescence for individual synapses. These parameters can reveal key transitions indicating synaptic development events, the roles of developmental and regulatory proteins, and synaptic plasticity.
Quantitation of fluorescence signals using Volocity has applications in neurobiology beyond analysis of permeabilized fixed C. elegans samples, being useful for quantification of any fluorescent signal localized to high-contrast puncta. Therefore it is suitable for analysis of fixed tissue or cells from any organism. However, fixation and permeabilization leads to visualization of total synaptic protein populations regardless of whether they are surface-expressed, whereas for receptors at least, only the surface-expressed fraction contributes to synaptic strength. Volocity-based analysis can also be used to quantify surface-expressed populations, provided non-permeabilizing fixation conditions are used. Volocity analysis will also be useful to study protein dynamics in live cells. Many protein domains shuttle between vesicular and extracellular environments with different pH during neuronal signaling events (e.g. the carboxy terminus of synaptobrevin during synaptic vesicle release 8-10, or the amino terminal domains of neurotransmitter receptors during trafficking to the cell surface 11). Fusing pH sensitive ecliptic GFP tags to these regions can produce in vivo reporters of these transitions that can be studied quantitatively using a Volocity-based approach. Similarly, GFP-tagging of neuropeptide genes results in GFP-labeled dense-core vesicles that de-stain as these vesicles fuse with the plasma membrane 12; Volocity analysis may be useful in this context as an assay for specific neuropeptide release in vivo. The advantage of Volocity analysis in all of these settings is its ability to rapidly identify and quantify large numbers of synapses, enabling subtle differences in their individual properties or population distributions to be demonstrated with statistical rigor.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the software</td>
 <td>Company</td>
 <td>&nbsp;</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Volocity v4.0 or higher</td>
 <td>PerkinElmer/Improvision</td>
 <td>&nbsp;</td>
 <td>Check your local imaging core facility for access to this software. Demo software is available at the PerkinElmer website. This method requires only the Quantitation module of Volocity.</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Table 2. Specific reagents and equipment.</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td><table border="1">
 <tbody>
 <tr>
 <td>Egg buffer</td>
 <td>NaCl 
 KCl 
 CaCl2 
 MgCl2 
 HEPES 
 (Adjust pH to 7.3)</td>
 <td>118 mM 
 48 mM 
 2 mM 
 2 mM 
 24 mM</td>
 </tr>
 <tr>
 <td>Alkaline hypochlorite (for 5 ml)</td>
 <td>Fresh household bleach (discard bottle 30 days after opening) 
 10 N NaOH 
 ddH2O</td>
 <td>1.0 ml
 0.25 ml 
 3.75 ml</td>
 </tr>
 </tbody>
</table>
&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Table 1. Solutions.</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Automated Quantification of Synaptic Fluorescence in C. elegans at JoVE.com

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.

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 ...

automated-quantification-of-synaptic-fluorescence-in-c-elegans

Nanyang Technological University

Research

JoVE Journal

Neuroscience

10.2K Views.  University of Toledo. 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.

Video: Automated Quantification of Synaptic Fluorescence in C. elegans

Assaying &beta;-amyloid Toxicity using a Transgenic C. elegans Model

Accumulation of the &beta;-amyloid peptide (A&beta;) is generally believed to be central to the induction of Alzheimer's disease, but the relevant mechanism(s) of toxicity are still unclear. A&beta; is also deposited intramuscularly in Inclusion Body Myositis, a severe human myopathy. The intensely studied nematode worm Caenorhabditis elegans can be transgenically engineered to express human A&beta;. Depending on the tissue or timing of A&beta; expression, transgenic worms can have readily measurable phenotypes that serve as a read-out of A&beta; toxicity. For example, transgenic worms with pan-neuronal A&beta; expression have defects is associative learning (Dosanjh et al. 2009), while transgenic worms with constitutive muscle-specific expression show a progressive, age-dependent paralysis phenotype (Link, 1995; Cohen et al. 2006). One particularly useful C. elegans model employs a temperature-sensitive mutation in the mRNA surveillance system to engineer temperature-inducible muscle expression of an A&beta; transgene, resulting in a reproducible paralysis phenotype upon temperature upshift (Link et al. 2003). Treatments that counter A&beta; toxicity in this model [e.g., expression of a protective transgene (Hassan et al. 2009) or exposure to Ginkgo biloba extracts (Wu et al. 2006)] reproducibly alter the rate of paralysis induced by temperature upshift of these transgenic worms. Here we describe our protocol for measuring the rate of paralysis in this transgenic C. elegans model, with particular attention to experimental variables that can influence this measurement.

Assaying &#946;-amyloid Toxicity using a Transgenic C. elegans Model

The intensely studied nematode worm Caenorhabditis elegans can be transgenically engineered to express the human &beta;-amyloid peptide (A&beta;). Induced expression of A&beta; in C. elegans muscle leads to a rapid, reproducible paralysis phenotype that can be used to monitor treatments that modulate A&beta; toxicity.

Accumulation of the &beta;-amyloid peptide (A&beta;) is generally believed to be central to the induction of Alzheimer's disease, but the relevant ...

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

Lateral Diffusion and Exocytosis of Membrane Proteins in Cultured Neurons Assessed using Fluorescence Recovery and Fluorescence-loss Photobleaching

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This directed trafficking is of fundamental importance to deliver, maintain or remove synaptic proteins.
Super-ecliptic pHluorin (SEP) is a pH-sensitive derivative of eGFP that has been extensively used for live cell imaging of plasma membrane proteins1-2. At low pH, protonation of SEP decreases photon absorption and eliminates fluorescence emission. As most intracellular trafficking events occur in compartments with low pH, where SEP fluorescence is eclipsed, the fluorescence signal from SEP-tagged proteins is predominantly from the plasma membrane where the SEP is exposed to a neutral pH extracellular environment. When illuminated at high intensity SEP, like every fluorescent dye, is irreversibly photodamaged (photobleached)3-5. Importantly, because low pH quenches photon absorption, only surface expressed SEP can be photobleached whereas intracellular SEP is unaffected by the high intensity illumination6-10. 
FRAP (fluorescence recovery after photobleaching) of SEP-tagged proteins is a convenient and powerful technique for assessing protein dynamics at the plasma membrane. When fluorescently tagged proteins are photobleached in a region of interest (ROI) the recovery in fluorescence occurs due to the movement of unbleached SEP-tagged proteins into the bleached region. This can occur via lateral diffusion and/or from exocytosis of non-photobleached receptors supplied either by de novo synthesis or recycling (see Fig. 1). The fraction of immobile and mobile protein can be determined and the mobility and kinetics of the diffusible fraction can be interrogated under basal and stimulated conditions such as agonist application or neuronal activation stimuli such as NMDA or KCl application8,10. 
We describe photobleaching techniques designed to selectively visualize the recovery of fluorescence attributable to exocytosis. Briefly, an ROI is photobleached once as with standard FRAP protocols, followed, after a brief recovery, by repetitive bleaching of the flanking regions. This 'FRAP-FLIP' protocol, developed in our lab, has been used to characterize AMPA receptor trafficking at dendritic spines10, and is applicable to a wide range of trafficking studies to evaluate the intracellular trafficking and exocytosis.

This report describes the use of live cell imaging and photobleach techniques to determine the surface expression, transport pathways and trafficking kinetics of exogenously expressed, pH-sensitive GFP-tagged proteins at the plasma membrane of neurons.

Membrane proteins such as receptors and ion channels undergo active trafficking in neurons, which are highly polarised and morphologically complex. This ...

C. elegans Tracking and Behavioral Measurement

We have developed instrumentation, image processing, and data analysis techniques to quantify the locomotory behavior of C. elegans as it crawls on the surface of an agar plate. For the study of the genetic, biochemical, and neuronal basis of behavior, C. elegans is an ideal organism because it is genetically tractable, amenable to microscopy, and shows a number of complex behaviors, including taxis, learning, and social interaction1,2. Behavioral analysis based on tracking the movements of worms as they crawl on agar plates have been particularly useful in the study of sensory behavior3, locomotion4, and general mutational phenotyping5. Our system works by moving the camera and illumination system as the worms crawls on a stationary agar plate, which ensures no mechanical stimulus is transmitted to the worm. Our tracking system is easy to use and includes a semi-automatic calibration feature. A challenge of all video tracking systems is that it generates an enormous amount of data that is intrinsically high dimensional. Our image processing and data analysis programs deal with this challenge by reducing the worms shape into a set of independent components, which comprehensively reconstruct the worms behavior as a function of only 3-4 dimensions6,7. As an example of the process we show that the worm enters and exits its reversal state in a phase specific manner.

C. elegans Tracking and Behavioral Measurement

We have developed a video-rate tracking microscope system that can record and quantify C. elegans behavior at high resolution and high speeds. We have also developed computational methods to reduce the dimensionality of the worm images to a fundamental set of measurements that completely describe the shape of the worm.

We have developed instrumentation, image processing, and data analysis techniques to quantify the locomotory behavior of C. elegans as it crawls on the ...

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps shape attention and also protects cells from over-stimulation. Adaptation within the olfactory circuit of C. elegans was first described by Colbert and Bargmann1,2. Here, the authors defined parameters of the olfactory adaptation paradigm, which they used to design a genetic screen to isolate mutants defective in their ability to adapt to volatile odors sensed by the Amphid Wing cells type C (AWC) sensory neurons. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor3 for 30 min they will adapt their responsiveness to the odor and will then ignore the adapting odor in a chemotaxis behavioral assay for ~1 hr. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor for ~1 hr they will then ignore the adapting odor in a chemotaxis behavioral assay for ~3 hr. These two phases of olfactory adaptation in C. elegans were described as short-term olfactory adaptation (induced after 30 min odor exposure), and long-term olfactory adaptation (induced after 60 min odor exposure). Later work from L'Etoile et al.,4 uncovered a Protein Kinase G (PKG) called EGL-4 that is required for both the short-term and long-term olfactory adaptation in AWC neurons. The EGL-4 protein contains a nuclear localization sequence that is necessary for long-term olfactory adaptation responses but dispensable for short-term olfactory adaptation responses in the AWC4. By tagging EGL-4 with a green fluorescent protein, it was possible to visualize the localization of EGL-4 in the AWC during prolonged odor exposure. Using this fully functional GFP-tagged EGL-4 (GFP::EGL-4) molecule we have been able to develop a molecular readout of long-term olfactory adaptation in the AWC5. Using this molecular readout of olfactory adaptation we have been able to perform both forward and reverse genetic screens to identify mutant animals that exhibit defective subcellular localization patterns of GFP::EGL-4 in the AWC6,7. Here we describe: 1) the construction of GFP::EGL-4 expressing animals; 2) the protocol for cultivation of animals for long-term odor-induced nuclear translocation assays; and 3) the scoring of the long-term odor-induced nuclear translocation event and recovery (re-sensitization) from the nuclear GFP::EGL-4 state.

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

Here we describe a molecular readout of long-term olfactory adaptation in Caenorhabditis elegans. The Protein Kinase G, EGL-4, is necessary for stable adaptation responses in the primary sensory neuron pair called AWC. During prolonged odor exposure EGL-4 translocates from the cytosol to nucleus of the AWC.

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this protocol focuses on how to image dividing neuroblasts, which are found underneath the epidermal cells and may be important for epidermal morphogenesis. Tissue formation is crucial for metazoan development&#160;and relies on external cues from neighboring tissues. C. elegans is an excellent model organism to study tissue morphogenesis in vivo due to its transparency and simple organization, making its tissues easy to study via microscopy. Ventral enclosure is the process where the ventral surface of the embryo is covered by a single layer of epithelial cells. This event is thought to be facilitated by the underlying neuroblasts, which provide chemical guidance cues to mediate migration of the overlying epithelial cells. However, the neuroblasts are highly proliferative and also may act as a mechanical substrate for the ventral epidermal cells. Studies using this experimental protocol could uncover the importance of intercellular communication during tissue formation, and could be used to reveal the roles of genes involved in cell division within developing tissues.

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes how to image dividing cells within a tissue in Caenorhabditis elegans embryos. While several protocols describe how to image cell division in the early embryo, this protocol describes how to image cell division within a developing tissue during mid late embryogenesis.

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this ...

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of stress-induced plasticity is the dauer stage of C. elegans. Dauers are an alternative developmental larval stage formed under conditions of low concentrations of bacterial food and high concentrations of a dauer pheromone. Dauers display extensive developmental and behavioral plasticity. For example, a set of four inner-labial quadrant (IL2Q) neurons undergo extensive reversible remodeling during dauer formation. Utilizing the well-known environmental pathways regulating dauer entry, a previously established method for the production of crude dauer pheromone from large-scale liquid nematode cultures is demonstrated. With this method, a concentration of 50,000 - 75,000 nematodes/ml of liquid culture is sufficient to produce a highly potent crude dauer pheromone. The crude pheromone potency is determined by a dose-response bioassay. Finally, the methods used for in vivo time-lapse imaging of the IL2Qs during dauer formation are described.

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant &#8216;dauer&#8217; juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer.

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of ...