Department of Biological Sciences, University of Toledo
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Sturt, B. L., Bamber, B. A. Automated Quantification of Synaptic Fluorescence in C. elegans. J. Vis. Exp. (66), e4090, doi:10.3791/4090 (2012).
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.
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.
2. Confocal Imaging
3. Automated Identification and Analysis of Individual Synaptic Clusters
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.
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.
Figure 2. Quantitation of synaptic puncta.
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 < 0.001, Kolmogorov-Smirnov test http://www.physics.csbsju.edu/stats/KS-test.html).
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.
No conflicts of interest declared.
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.
|Volocity v4.0 or higher||PerkinElmer/Improvision||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.|
|Table 2. Specific reagents and equipment.|
|Table 1. Solutions.|