Name: imaging dendritic spines in caenorhabditis elegans
Uploaded: 2021-09-27T13:30:00.000+00:00
Duration: 9 min 14 s
Description: Watch this Scientific Journal Video about Imaging Dendritic Spines in Caenorhabditis elegans at JoVE.com

Imaging and analysis on Imaris were performed in the Vanderbilt Cell Imaging Shared Resource (CIRS) supported by NIH (CA68485, DK20593, DK58404, DK59637, and EY08126). The LSM 880 is supported by grant 1S10OD201630. Imaging on a Nikon spinning disk was performed at the Nikon Center of Excellence. We thank Jenny Schafer, CISR director, and Bryan Millis for training and insightful discussions and members of the Burnette lab: Dylan Burnette, Aidan Fenix, and Nilay Taneja for advice. This work was supported by National Institutes of Health grants to DMM (R01NS081259 and R01NS106951) and an American Heart Association grant to ACC (18PRE33960581).

1. Determination of the structure of the DD dendritic spines
<ol>
	<li>Create transgenic worms to label DD spines
	<ol>
		<li>Use the flp-13 promoter to build an expression vector for the label of interest (e.g., cytoplasmic mCherry, MYR::mRuby, LifeAct::GFP, GFP::utrophin) (Figure 1). See the complete list of plasmids in Supplementary File 1.</li>
		<li>Use established methods to generate a transgenic line labeling DD spines8,9.</li>
	</ol>
	</li>
	<li>Prepare the sealant
	<ol>
		<li>Make &#65279;a 1:1 mixture of paraffin-based embedding medium10 (see Table of Materials).</li>
		<li>Heat the medium at 60 &#176;C until melting, then aliquot in 1.5 mL capped microcentrifuge tubes and maintain a heating block at 60-70 &#176;C. 
		NOTE: Sealant can last for 4 weeks in the heating block.</li>
	</ol>
	</li>
	<li>Prepare an anesthetic.
	<ol>
		<li>Make stock solutions in distilled H2O of 1% Tricaine and 1 M levamisole (see Table of Materials). Store at -20 &#176;C.</li>
		<li>Prepare a working solution of 0.05% Tricaine and 15 mM levamisole anesthetic, as described in steps 1.3.3-1.3.511.</li>
		<li>Mix 75 &#181;L of 1% Tricaine stock and 22.5 &#181;L of 1 M levamisole.</li>
		<li>Add M9 buffer to a final volume of 1.5 mL.</li>
		<li>Aliquot 10 &#181;L of 0.05% Tricaine, 15 mM levamisole into 0.5 mL microcentrifuge tubes and store at -20 &#176;C. 
		NOTE: The anesthetic mixture is sensitive to temperature, and individual aliquots of the working solution should not be refrozen after thawing for each experiment. See Supplementary File 2 for a recipe for M9 buffer.</li>
	</ol>
	</li>
	<li>Acquire high-resolution images
	<ol>
		<li>Prepare 10% agarose and maintain it in the water bath at 60 &#176;C. 
		NOTE: See the report by Monica Driscoll in WormBook12.</li>
		<li>Mount 15-20 young adults on 10% agarose pads and add 3 &#181;L of anesthetic (see step 3).</li>
		<li>Apply coverslip (worms are immobilized within 5 min).</li>
		<li>Seal the edges of the coverslip with melted adhesive sealant mixture (see Table of Materials).</li>
		<li>Acquire images
		<ol>
			<li>Super-resolution acquisition
			<ol>
				<li>Use a laser-scanning confocal microscope equipped for super-resolution microscopy with a 63x/1.40 Plan-Apochromat oil objective lens to achieve a small pixel size (e.g., &#60; 50 nm). Acquire Z-stacks using the step size recommended by the manufacturer&#39;s software (see Table of Materials).</li>
				<li>Collect a series of optical sections that span the total volume of the DD ventral process (e.g., 15-20 slices at 0.19 &#181;m step size or 2- 3&#181;m thick). Submit Z-stacks for image processing using the manufacturer&#39;s software and analyze images with a score higher than 7 (Figure 1B, Figure 2 and Figure 3).</li>
			</ol>
			</li>
			<li>Nyquist acquisition
			<ol>
				<li>Use a laser scanning confocal microscope to select the optimal pixel size for the wavelength of light and numerical aperture of the objective lens in use (e. g., 40x/1.4 Plan Fluor oil objective). 
				NOTE: The smaller pixel size will reveal the fine structure of DD spines.</li>
				<li>Submit stack for 3D deconvolution using Automatic algorithm (see Table of Materials) (Figure 3).</li>
			</ol>
			</li>
			<li>Use the smallest Z-step possible (e.g., determined by Piezo stage) because oversampling in Z can yield sharper images after 3D deconvolution13.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Image analysis
	<ol>
		<li>Use&#160;appropriate image processing software (see Table of Materials) to create maximum intensity projections of the Z-stacks14.</li>
		<li>Manually count the protrusions on the DD dendrite. 
		NOTE: Protrusions are perpendicular extensions from the main shaft (Figure 1B, arrowheads).</li>
		<li>Determine the length of the DD dendrite scored to calculate the density of spines per 10 &#181;m of DD dendrite (Figure 1C).</li>
		<li>Classify spines as thin/mushroom, filopodial, stubby or branched (Figure 2A). 
		NOTE: Thin/mushroom spines exhibit a narrow base (neck) and a broader tip (head). Filopodial spines do not display a constricted base (no neck) but have a constant width. Stubby spines have a wide base and tip. Branched spines are protrusions with more than one tip.</li>
	</ol>
	</li>
</ol>
2. Assessing activation of DD dendritic spines by presynaptic cholinergic signaling
<ol>
	<li>Create transgenic worms using conventional techniques (e.g., microinjection)8,9
	<ol>
		<li>Use the flp-13 promoter to drive expression of the Ca++ sensor, GCaMP6s, in DD neurons and the unc-4 promoter to drive expression of Chrimson, a red-shifted channelrhodopsin, in presynaptic VA neurons (Figure 4A). See the list of plasmids in Supplementary File 1.</li>
	</ol>
	</li>
	<li>Prepare All-trans Retinal (ATR) and control plates. 
	NOTE: ATR is a required cofactor for Chrimson to function as an optogenetically activated ion channel.
	<ol>
		<li>Prepare 100 mM ATR stock solution in ethanol (100%). Store at -20 &#176;C in 1 mL aliquots.</li>
		<li>Under a laminar flow hood, add 300 &#181;L of overnight OP50 bacterial culture and 0.25 &#181;L of ATR to each 60 mm NGM (Nematode Growth Medium) nutrient agar plate and spread with a sterile glass rod.</li>
		<li>For controls, add 300 &#181;L of OP50 bacteria and 0.25 &#181;L of ethanol (100%) to a separate group of NGM plates.</li>
		<li>Let plates sit in hood at room temperature for 24 h (protected from ambient light) to allow bacterial growth. 
		NOTE: Plates can be used after the initial 24 h incubation or maintained at 4 &#176;C to use within 5 days.</li>
	</ol>
	</li>
	<li>Setting-up the experiment
	<ol>
		<li>Place five NC3569 L4-stage larvae on OP50-seeded ATR or control plates that lack ATR and grow in darkness at 23&#176;C.</li>
		<li>Three days later, use a stereo dissecting microscope to confirm vulva development to pick L4 stage progeny from ATR and control plates for imaging as described in steps 2.4.1-2.4.3.</li>
		<li>On a microscope slide, place 2 &#181;L of 0.05 &#181;m polybeads (2.5% solids w/v) (see Table of Materials).</li>
		<li>Use a platinum wire (&#34;worm pick&#34;) to add a small globule of super glue to the solution and swirl gently to generate filamentous &#34;strands&#34; of glue. Then add 3 &#181;L of M9 buffer (Figure 4B).</li>
		<li>Place approximately ten L4 larvae in the solution and apply a coverslip. 
		NOTE: Glue fibers will randomly contact worms and immobilize them after the coverslip is applied. Worms that are embedded in large globules of glue appear desiccated and should not be imaged.</li>
		<li>Seal edges of coverslip as mentioned in step 1.4.4.</li>
	</ol>
	</li>
	<li>Recording of evoked Ca++ transients in dendritic spines.
	<ol>
		<li>Use a spinning disk confocal microscope equipped with a sensitive CCD camera, a 100x TIRF oil objective lens, and 488 nm and 561 nm laser lines (see Table of Materials).</li>
		<li>Adjust microscope stage to position DD spines in the focal plane.</li>
		<li>Set-up time-lapse acquisition to illuminate the sample with 488 nm laser line every frame (for detecting GCaMP6s fluorescence) and the 561 nm laser line at periodic intervals (for Chrimson excitation). 
		NOTE: For example, use 488 nm light to capture consecutive snapshots (200 ms) of GCaMP6s signal coupled with a 200 ms pulse of 561 nm light every 5th frame (Figure 4C-E). With this configuration, GCaMP levels before and after each 561 nm pulse are detected ~1 s apart (200 ms of 488 nm laser to detect GCaMP before VA activation, 200 ms of 561 nm pulse to activate VA and ~600 ms to switch between laser lines and emission filters. With this set-up, VA neurons are activated every 2.5 s.</li>
	</ol>
	</li>
	<li>Analysis of in vivo Ca++ imaging
	<ol>
		<li>Use 2D-deconvolution and image alignment to correct minor deviations arising from the worm movement during acquisition (see Table of Materials).</li>
		<li>Define the DD dendritic spine as the Region Of Interest (ROI in Figures 4C-D).</li>
		<li>Duplicate the ROI and relocate to a neighboring region inside the worm to collect background signal (i.e., noise).</li>
		<li>Use appropriate software (see Table of Materials) to export GCaMP6s&#160;intensities to excel for each time point. Subtract background fluorescence from spine ROI fluorescence.</li>
		<li>Determine the change in fluorescence by subtracting the GCaMP6s fluorescence in the frame immediately before 561 nm excitation (F0) from each time-point after excitation (&#916;F), then dividing by F0 to determine &#916;F/F0 (Figure 4E).</li>
		<li>Graph the normalized traces (See Table of Materials).</li>
		<li>Perform a paired statistical test for each measurement of GCaMP6s fluorescence before and after each pulse of 561 nm light. 
		NOTE: This approach effectively excludes random fluctuations in GCaMP6s fluorescence that otherwise reduce the statistical power of comparing the mean GCaMP6s signal from all measurements before and after 561 nm excitation (Figure 4F).</li>
		<li>For measurements that show a normal or Gaussian distribution, use a paired parametric ANOVA test and correct for multiple comparisons for each of the two groups (ATR before vs. after, no ATR before vs. after). Alternatively, for data that are not normally distributed, use a non-parametric ANOVA with posthoc correction for multiple testing. 
		NOTE: Worms grown on plates that lack ATR (&#34;no ATR&#34;) are necessary controls and should not show 561 nm-activated Ca++ transients because ATR is required for Chrimson function.</li>
	</ol>
	</li>
</ol>

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We declare no conflicts of interest.

The Airyscan detector was selected to acquire snapshots of DD spines because it affords a higher signal-to-noise ratio and better resolution than conventional confocal microscopes19,20. AiryScan imaging also allows the use of conventional fluorescent proteins (e.g., GFP, mCherry, etc.), now widely available for C. elegans. Although higher resolution images can be obtained with other super-resolution methods (e.g., STORM, STED, PALM), these methods require photo-activatable or photo-switchable fluorescent proteins21. As an alternative to Airyscan, conventional confocal microscopes are recommended. For example, imaging with Nyquist acquisition (Figure 3) achieves the pixel size using a 40x/1.3 objective of 123.9 nm, sufficient to distinguish the spine morphological types (Figure 2).
For determining spine density, using a cytosolic fluorescent protein such as (1) mCherry or GFP, (2) LifeAct to label the actin cytoskeleton, or (3) a myristoylated fluorescent protein (e.g., MYR::mRuby) to label the plasma membrane (Figure 1B) is recommended. In comparison, the F-actin binding protein Utrophin reduces spine density (Figure 1C), indicating a negative effect on spine morphogenesis when Utrophin is over-expressed.
The current imaging methods should help identify genetic variants that govern spine morphology1,16. DD spine morphology (i.e., thin/mushroom, filopodial, stubby, branched, see Figure 2) can be assessed from single 2D-projections of the ventral nerve cord lateral images since most DD spines adopt a characteristically ventrally-directed orientation. In these comparisons, it is essential to use the same fluorescent marker for each condition since apparent spine morphological types seem to be influenced by the labeling method (e.g., MYR::mRuby vs. LifeAct::GFP). In addition, it was noted that the spine shapes are dynamic and likely change shape in response to the external signals2,16. Thus, it is also essential to compare spine shapes between genotypes at similar developmental stages and under similar conditions.
The orientation of the C. elegans ventral cord is critically vital for accurate image acquisition. Both the ventral and dorsal cords on opposite sides of the animal should be visible in the same Z-plane, indicating that the worm is oriented on its side (Figure 1B). It is best not to collect images of worms moving or in contact with other worms or bubbles near the ventral cord, as this can degrade images of spines.
For in vivo calcium imaging, fresh slides need to be prepared immediately before each acquisition. It is best to image worms in contact with only thin glue fibers vs. &#34;globs&#34; of glue which tend to desiccate worms and degrade the image (Figure 4B). In the experiment shown in Figure 4, the pulse of 561 nm light activates the entire field of view. For increasing the temporal and spatial resolution to detect local Ca++ transients, for example, within individual DD spines, a galvo mini scanner set up for the 561 nm laser line can be used to stimulate a smaller region of interest17.

Dendritic spines are specialized cellular structures that receive input from neighboring neurons for synaptic transmission. Activation of neurotransmitter receptors elevates intracellular calcium and downstream signaling pathways in these&#160;characteristic neuronal protrusions1. Because of the fundamental importance of dendritic spines to neurotransmission and their misregulation in neurodevelopmental diseases1, the discovery of factors that modulate dendritic spine morphogenesis and function is of high interest to the field of neuroscience.
Recently, dendritic spines have been identified in the C. elegans nervous system based on key characteristics shared with mammalian spines2. This determination is crucial because it opens the possibility of exploiting the advantages of C. elegans to investigate spine biology. Dendritic spines on Dorsal D (DD) motor neurons receive input from cholinergic neurons (VA and VB) in the ventral nerve cord (Figure 1A)2,3,4. Here, imaging methods are presented to explore the structure of DD dendritic spines and their function in vivo in an intact nervous system that is readily accessible to live imaging and genetic analysis. For monitoring the shape of dendritic spines, (1) cytosolic fluorescent proteins, which fill the dendritic process and spines; (2) membrane-bound fluorescent proteins, which decorate the border of dendritic spines and dendrites; or (3) the actin markers, LifeAct5 or Utrophin6, which are enriched in dendritic spines are used, thus revealing their shape. To monitor the functionality of DD spines, GCaMP&#160; fluorescence is used to detect Ca++ transients evoked by activation of the red-shifted opsin, Chrimson, in presynaptic cholinergic neurons7. Both strategies are expected to facilitate the study of DD dendritic spines in wild-type and mutant animals.

<table><tbody><tr><td>All-trans retinal (ATR)</td><td>Sigma-Aldrich</td><td>R2500-100MG</td><td>Necessary cofactor for neuronal excitation with Chrimson</td></tr><tr><td>diH&lt;sub&gt;2&lt;/sub&gt;O</td><td>MilliQ</td><td></td><td>To prepare M9 buffer</td></tr><tr><td>Ethanol 100%</td><td>Sigma</td><td>64-17-5</td><td>To dilute ATR and make control plates for neuronal excitation</td></tr><tr><td>Ethyl 3-aminobenzoate methanesulfonate salt (tricaine)</td><td></td><td></td><td>To immobilize animals for imaging dendritic spines</td></tr><tr><td>ImageJ</td><td>NIH</td><td>(Schindelin J et al., 2012)</td><td>Open source image processing software</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fisher Bioreagents</td><td>7758-11-4</td><td>To prepare M9 buffer</td></tr><tr><td>Levamisole hydrochloride</td><td>Sigma</td><td>16595-80-5</td><td>To immobilize animals for imaging dendritic spines</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fisher Chemical</td><td>M63-500</td><td>To prepare M9 buffer</td></tr><tr><td>Microscope cover glass</td><td>Fisherbrand</td><td>12542B</td><td>To mount animals for microscopy acquisition</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fisher Scientific</td><td>S369-500</td><td>To prepare M9 buffer</td></tr><tr><td>NaCl</td><td>Fisher Chemical</td><td>S671-3</td><td>To prepare M9 buffer</td></tr><tr><td>NIS Elements version 05.21</td><td>Nikon</td><td></td><td>To analyze images and movies (e.g., Deconvolution, image alignment)</td></tr><tr><td>Polybeads carboxylate 0.05um microspheres</td><td>Polysciences, Inc</td><td>15913-10</td><td>To immobilize animals for imaging Ca++ transients</td></tr><tr><td>Prism</td><td></td><td></td><td>For statistical analysis and graphing normalized Ca++ transients</td></tr><tr><td>SeaKen ME agarose</td><td>Lonza</td><td>50014</td><td>To make agarose pads to mount animals for imaging</td></tr><tr><td>Super Glue</td><td>The gorilla company</td><td></td><td>To immobilize animals for imaging Ca++ transients</td></tr><tr><td>Superfrost microscope slides</td><td>Fisherbrand</td><td>22-034-980</td><td>To mount animals for microscopy acquisition</td></tr><tr><td>vaseline</td><td>Covidien</td><td>8884430300</td><td>To seal sample for confocal snapshots</td></tr><tr><td>Wax</td><td>Fisherbrand</td><td>23-021-399</td><td>Paraplast tissue embedding medium</td></tr><tr><td>&lt;strong&gt;Microscope for super-resolution imaging&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>LSM880</td><td>Zeiss</td><td></td><td></td></tr><tr><td>AiryScan detector</td><td>Zeiss</td><td></td><td></td></tr><tr><td>Plan Apochromat (oil) 63x/ 1.40 NA, WD = 0.19 mm</td><td></td><td></td><td></td></tr><tr><td>Laser lines</td><td></td><td></td><td></td></tr><tr><td>Stage controller</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Microscope for Nyquist image acquisition&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>A1R Confocal</td><td>Nikon</td><td></td><td></td></tr><tr><td>Plan Fluor (oil) 40x/1.3 NA, WD 0.24 mm</td><td></td><td></td><td></td></tr><tr><td>488 nm, 16mW</td><td></td><td></td><td></td></tr><tr><td>561 nm, 17mW</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Microscope to monitor evoked Ca++ transients in dendritic spines&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Spinning Disk Confocal</td><td>Nikon</td><td></td><td></td></tr><tr><td>Andor DU-897 EMCCD camera</td><td></td><td></td><td></td></tr><tr><td>Spinning disk Head CSU-X1</td><td>Yokogawa</td><td></td><td></td></tr><tr><td>Apo TIRF (oil) 100x/1.49 NA ,WD 0.12 mm</td><td></td><td></td><td></td></tr><tr><td>488 nm, 65mW</td><td></td><td></td><td></td></tr><tr><td>561 nm, 86mW</td><td></td><td></td><td></td></tr><tr><td>525 nm (+/- 18 nm)</td><td></td><td></td><td></td></tr><tr><td>605 nm (+/- 35 nm)</td><td></td><td></td><td></td></tr></tbody></table>

imaging dendritic spines in caenorhabditis elegans

Dendritic spines are specialized sites of synaptic innervation modulated by activity and serve as substrates for learning and memory. Recently, dendritic spines have been described for DD GABAergic neurons as the input sites from presynaptic cholinergic neurons in the motor circuit of Caenorhabditis elegans. This synaptic circuit can now serve as a powerful new in vivo model of spine morphogenesis and function that exploits the facile genetics and ready accessibility of C. elegans to live-cell imaging. 
This protocol describes experimental strategies for assessing DD spine structure and function. In this approach, a super-resolution imaging strategy is used to visualize the intricate shapes of actin-rich dendritic spines. To evaluate the DD spine function, the light-activated opsin, Chrimson, stimulates the presynaptic cholinergic neurons, and the calcium indicator, GCaMP, reports the evoked calcium transients in postsynaptic DD spines. Together, these methods comprise powerful approaches for identifying genetic determinants of dendritic spines in C. elegans that could also direct spine morphogenesis and function in the brain.

Measurements with three independent markers (cytosolic mCherry, LifeAct::GFP, MYR::mRuby) yielded an average density of 3.4 &#177; 1.03 DD dendritic spines per 10 &#181;m of DD dendrite in wild-type young adults (Figure 1B,C). For this analysis, the measurements obtained with the GFP::Utrophin marker that yielded a significantly lower spine density were excluded (2.4 &#177; 0.74, Figure 1)&#160;due to interactions of Utrophin with the actin cytoskeleton6 that&#160;potentially drives spine morphogenesis15. The measurements of spine density in the light microscope are comparable to the value of 4.2 spines/10 &#181;m dendrite obtained from the reconstruction of 12 spines from electron micrographs of the DD1 neuron2. The live-cell imaging approach confirmed that the thin/mushroom-shaped morphology of the DD spines predominates in the adult vs. alternative spine shapes (e.g., filopodial, stubby, branched) (Figure 2B), which is also typical for spines in the mature mammalian nervous system16.
An optogenetic strategy was used to ask if the presumptive dendritic spines detected by high-resolution light microscopy (Figure 1 and Figure 2) are responsive to neurotransmitter release from presynaptic sites, a characteristic hallmark of dendritic spines in mammalian neurons. Green light (561 nm) was used to activate a channelrhodopsin variant, Chrimson, in presynaptic cholinergic neurons and blue light (488 nm) to detect Ca++-dependent fluorescence emitted by a cytoplasmic GCaMP probe in postsynaptic DD dendritic spines. This experiment detected transient bursts of GCaMP signal in DD spines immediately after optogenetic activation of Chrimson in presynaptic VA neurons (Figure 3). The success of this experiment depends on the reliable expression of Chrimson in all presynaptic VA neurons. In this case, a chromosomal integrant17 of the Punc-4::Chrimson marker was used to ensure consistent VA expression. This experiment could also be conducted with an extrachromosomal array. Chrimson expression in a specific VA neuron can be independently confirmed, for example, by coupling the Chrimson transgene to an SL2 transpliced leader sequence with a downstream nuclear-localized GFP as a co-expression marker2. It is essential to perform a control experiment in the absence of ATR to confirm that the measured GCaMP signal depends on optogenetic activation of Chrimson, which is strictly ATR-dependent (Figure 4D). Finally, because evoked Ca++ signals are transient, it is crucial to adopt an imaging protocol that allows rapid switching (&#60;1 s) between 561 nm excitation and GCaMP signal acquisition with the 488 nm laser (Figure 4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62676/62676fig1v2.jpg" /> 
Figure 1: Labeling of DD dendritic spines. (A) (Top) Six Dorsal D (DD1-DD6) neurons in the ventral nerve cord of C. elegans. (Bottom) In adults, ventrally directed DD spines (arrowhead) contact presynaptic terminals of Ventral A (VA) and Ventral B (VB) motor neurons (magenta), and DD commissures extend to the dorsal nerve cord to provide GABAergic output to body muscles (arrow)18. This figure has been modified from Reference2. (B) Fluorescent micrographs (Airyscan) of DD spines labeled with cytosolic mCherry, myristoylated mRuby (MYR::mRuby), LifeAct::GFP and GFP::Utrophin in young adult worms. Gray arrowheads point to spines. Scale bar = 2 &#181;m. (C) Density (spines/10 &#181;m) of DD neuron dendritic spines labeled with cytosolic mCherry (3.77 &#177; 0.9), MYR::mRuby (3.09 &#177; 0.8), LifeAct::GFP (3.44 &#177; 1.1) or GFP::Utrophin (2.41 &#177; 0.8). All samples are normally distributed. One-Way ANOVA shows that spine densities for cytosolic mCherry, MYR::mRuby, and LifeAct::GFP are not significantly (NS) different, whereas spine density is reduced for GFP::Utrophin vs. cytosolically labeled mCherry (p = 0.0016) and LifeAct::GFP (p = 0.0082). The dashed red line represents the spine density of DD neurons assessed from 3D EM reconstruction (4.2 spines/10 &#181;m). This figure has been modified from Reference2. <a href="https://www.jove.com/files/ftp_upload/62676/62676fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62676/62676fig02.jpg" /> 
Figure 2: Imaging DD dendritic spines. (A) (Top) Schematic of spine shapes. (Bottom) Airyscan images of each type of spine (Scale bar = 500 nm) labeled with LifeAct::GFP (green) and 3D-reconstructions by serial electron micrographs of a high-pressure frozen adult (blue). (B)&#160;Spine frequency by type, visualized with LifeAct::GFP: Thin/Mushroom (55.5 &#177; 14.5%), Filopodial (10.3 &#177; 8.70%), Stubby (18.8 &#177; 10.7%), Branched (15.42 &#177; 6.01%). Spines frequency by type visualized with MYR::mRuby: Thin/Mushroom (52.2 &#177; 16.5%), Filopodial (5.68 &#177; 7.0%), Stubby (33.1 &#177; 14.8%), Branched (9.02 &#177; 9.6%). Unpaired T-test, Filopodial (p = 0.0339); Stubby (p = 0.0009) and Branched (p = 0.011) spines labeled with the MYR::mRuby marker are significantly different from LifeAct::GFP. This figure has been modified from Reference2. <a href="https://www.jove.com/files/ftp_upload/62676/62676fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62676/62676fig03.jpg" /> 
Figure 3: Strategies for acquiring high-resolution images of DD spines. (A-B) (Top) Fluorescent images of DD1 dendrites labeled with a cytosolic marker (mCherry) by (A) Airyscan detector and (B) Nyquist acquisition. (Bottom) DD dendrite (red) is depicted with an image analysis software (auto-path option of filament tracer), and DD spines (blue) are graphically illustrated using the semi-automated spines detection module. Arrow points to branched spine enlarged in C and D. Arrowheads denote neighboring thin/mushroom spines enlarged in C and D. Scale bar = 2 &#181;m.&#160;(C-D)&#160;Enlarged examples of (top) branched spine (arrow) and (bottom) two neighboring thin/mushroom spines (arrowheads) obtained with (C) Airyscan detector or by (D) Nyquist acquisition. Scale bar = 500 nm.&#160;Data reproduced from2. <a href="https://www.jove.com/files/ftp_upload/62676/62676fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62676/62676fig04.jpg" /> 
Figure 4: Assessing the function of DD spines. (A) DD motor neurons express the Ca++ indicator GCaMP6s (green), and VA motor neurons express the channelrhodopsin-variant, Chrimson (magenta)7. (B)&#160;Schematic depicting method for mounting worms for Ca++ measurements. (1) On a clean microscope slide, (2) place 2 &#181;L of 0.05 &#181;m poly beads, (3) use a platinum wire (&#34;worm pick&#34;) to add a small globule of super glue and (4) swirl into the solution to generate filamentous strands of glue. (5) Add 3&#181;L of M9 buffer. (6) Place approximately ten L4 larvae in the solution, (7) apply coverslip and seal edges with Vaseline/wax. (C-D) Activation of VA neurons correlates with Ca++ transients in DD1 spines. GCaMP6s fluorescence imaged (at 0.5 s intervals) with periodic light activation of Chrimson (2.5 s intervals) evokes Ca++ transients with (C) +ATR (n = 12) but not in (D) controls (-ATR, n = 12). Panels are snapshots over time (s), before and after pulse of 561 nm light (vertical pink line). Scale bars = 2 &#181;m. GCaMP6s signal is acquired from an ROI (Region of Interest) at the tip of each spine.&#160;(E) GCaMP6s fluorescence during the 10 s recording period plotted for +ATR (green) vs -ATR (control, gray) (n = 12 videos). Vertical pink bars denote 561 nm illumination (e.g., Chrimson activation). Each animal was stimulated 4 times with 561 nm light. Measurements were collected before and after each pulse of 561 nm light.&#160;(F) Plot of the GCaMP6s fluorescence before and after each pulse of 561 nm light. GCaMP6s fluorescence was measured 1 s after each pulse of 561 nm light. Because samples are not normally distributed, a paired non-parametric Friedman test was applied to correct for multiple comparisons of GCaMP6s fluorescence before vs. after 561 nm light stimulation for worms grown either with ATR (+ATR, green) (*** p = 0.0004, n = 48 measurements) or in the absence of ATR (-ATR, gray) (NS, Not Significant, p = 0.0962, n = 48 measurements). This figure has been modified from Reference2. <a href="https://www.jove.com/files/ftp_upload/62676/62676fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1: List of plasmids used in the study. <a href="https://www.jove.com/files/ftp_upload/62676/Supplementary File 1.docx" target="_blank">Please click here to download this File.</a>
Supplementary File 2: Composition and preparation of M9 buffer. <a href="https://www.jove.com/files/ftp_upload/62676/Supplementary File 2.docx" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Imaging Dendritic Spines in Caenorhabditis elegans at JoVE.com

Imaging Dendritic Spines in Caenorhabditis elegans

Dendritic spines are important cellular features of the nervous system. Here live imaging methods are described for assessing the structure and function of dendritic spines in C. elegans.&#160;These approaches support the development of mutant screens for genes that define dendritic spine shape or function.

Imaging Dendritic Spines in Caenorhabditis elegans

Dendritic spines are specialized sites of synaptic innervation modulated by activity and serve as substrates for learning and memory. Recently, dendritic ...

imaging-dendritic-spines-in-caenorhabditis-elegans

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Neuroscience

2.3K Views.  Vanderbilt University. Dendritic spines are important cellular features of the nervous system. Here live imaging methods are described for assessing the structure and function of dendritic spines in C. elegans. These approaches support the development of mutant screens for genes that define dendritic spine shape or function.

Video: Imaging Dendritic Spines in Caenorhabditis elegans

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

Longitudinal Two-Photon Imaging of Dorsal Hippocampal CA1 in Live Mice

Two-photon microscopy is a fundamental tool for neuroscience as it permits investigation of the brain of live animals at spatial scales ranging from subcellular to network levels and at temporal scales from milliseconds to weeks. In addition, two-photon imaging can be combined with a variety of behavioral tasks to explore the causal relationships between brain function and behavior. However, in mammals, limited penetration and scattering of light have limited two-photon intravital imaging mostly to superficial brain regions, thus precluding longitudinal investigation of deep-brain areas such as the hippocampus. The hippocampus is involved in spatial navigation and episodic memory and is a long-standing model used to study cellular as well as cognitive processes important for learning and recall, both in health and disease. Here, a preparation that enables chronic optical access to the dorsal hippocampus in living mice is detailed. This preparation can be combined with two-photon optical imaging at cellular and subcellular resolution in head fixed, anesthetized live mice over several weeks. These techniques enable repeated imaging of neuronal structure or activity-evoked plasticity in tens to hundreds of neurons in the dorsal hippocampal CA1. Furthermore, this chronic preparation can be used in combination with other techniques such as micro-endoscopy, head-mounted wide field microscopy or three-photon microscopy, thus greatly expanding the toolbox to study cellular and network processes involved in learning and memory.

This method describes a chronic preparation that allows optical access to the hippocampus of living mice. This preparation can be used to perform longitudinal optical imaging of neuronal structural plasticity and activity-evoked cellular plasticity over a period of several weeks.

Two-photon microscopy is a fundamental tool for neuroscience as it permits investigation of the brain of live animals at spatial scales ranging from ...

Behavior

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, ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Assessment of Dendritic Arborization in the Dentate Gyrus of the Hippocampal Region in Mice

Dendritic arborization has been shown to be a reliable marker for examination of structural and functional integrity of neurons. Indeed, the complexity and extent of dendritic arborization correlates well with the synaptic plasticity in these cells. A reliable method for assessment of dendritic arborization is needed to characterize the deleterious effects of neurological disorders on these structures and to determine the effects of therapeutic interventions. However, quantification of these structures has proven to be a formidable task given their complex and dynamic nature. Fortunately, sophisticated imaging techniques can be paired with conventional staining methods to assess the state of dendritic arborization, providing a more reliable and expeditious means of assessment. Below is an example of how these imaging techniques were paired with staining methods to characterize the dendritic arborization in wild type mice. These complementary imaging methods can be used to qualitatively and quantitatively assess dendritic arborization that span a rather wide area within the hippocampal region.

We describe two methods for visualization and quantification of dendritic arborization in the hippocampus of mouse models: real-time and extended depth of field imaging. While the former method allows sophisticated topographical tracing and quantification of the extent of branching, the latter allows speedy visualization of the dendritic tree.

Dendritic arborization has been shown to be a reliable marker for examination of structural and functional integrity of neurons. Indeed, the complexity ...

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized animals. Highly sensitive, genetically encoded fluorescent reporters of Ca2+ have revolutionized the recording of cell and synaptic activity using non-invasive optical approaches in behaving animals. When combined with genetic and optogenetic techniques, the molecular mechanisms that modulate cell and circuit activity during different behavior states can be identified.
Here we describe methods for ratiometric Ca2+ imaging of single neurons in freely behaving Caenorhabditis elegans worms. We demonstrate a simple mounting technique that gently overlays worms growing on a standard Nematode Growth Media (NGM) agar block with a glass coverslip, permitting animals to be recorded at high-resolution during unrestricted movement and behavior. With this technique, we use the sensitive Ca2+ reporter GCaMP5 to record changes in intracellular Ca2+ in the serotonergic Hermaphrodite Specific Neurons (HSNs) as they drive egg-laying behavior. By co-expressing mCherry, a Ca2+-insensitive fluorescent protein, we can track the position of the HSN within ~ 1 &#181;m and correct for fluctuations in fluorescence caused by changes in focus or movement. Simultaneous, infrared brightfield imaging allows for behavior recording and animal tracking using a motorized stage. By integrating these microscopic techniques and data streams, we can record Ca2+ activity in the C. elegans egg-laying circuit as it progresses between inactive and active behavior states over tens of minutes.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms. &#8195;

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized ...

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding these biological processes often requires long-term and repeated imaging of the same animal. Long recovery times associated with conventional immobilization methods done on agar pads have detrimental effects on animal health making it inappropriate to repeatedly image the same animal over long periods of time. This paper describes a microfluidic chip design, fabrication method, on-chip C. elegans culturing protocol, and three examples of long-term imaging to study developmental processes in individual animals. The chip, fabricated with polydimethylsiloxane and bonded on a cover glass, immobilizes animals on a glass substrate using an elastomeric membrane that is deflected using nitrogen gas. Complete immobilization of C. elegans enables robust time-lapse imaging of cellular and sub-cellular events in an anesthetic-free manner. A channel geometry with a large cross-section allows the animal to move freely within two partially sealed isolation membranes permitting growth in the channel with a continuous food supply. Using this simple chip, imaging of developmental phenomena such as neuronal process growth, vulval development, and dendritic arborization in the PVD sensory neurons, as the animal grows inside the channel, can be performed. The long-term growth and imaging chip operates with a single pressure line, no external valves, inexpensive fluidic consumables, and utilizes standard worm handling protocols that can easily be adapted by other laboratories using C. elegans.

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

The protocol describes a simple microfluidic chip design and microfabrication methodology used to grow C. elegans in presence of a continuous food supply for up to 36 h. The growth and imaging device also enables intermittent long-term high-resolution imaging of cellular and sub-cellular processes during development for several days.

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding ...

Bioengineering

Quantifying Levels of Dopaminergic Neuron Morphological Alteration and Degeneration in Caenorhabditis elegans

Dopamine neuron loss is involved in the pathology of Parkinson&#39;s Disease (PD), a highly prevalent neurodegenerative disorder affecting over 10 million people worldwide. Since many details about PD etiology remain unknown, studies investigating genetic and environmental contributors to PD are needed to discover methods of prevention, management, and treatment. Proper characterization of dopaminergic neuronal loss may be relevant not only to PD research, but to other increasingly prevalent neurodegenerative disorders.
There are established genetic and chemical models of dopaminergic neurodegeneration in the Caenorhabditis elegans model system, with easy visualization of neurobiology supported by the nematodes&#39; transparency and invariant neuronal architecture. In particular, hermaphroditic C. elegans&#39; dopaminergic neuron morphological changes can be visualized using strains with fluorescent reporters driven by cell-specific promotors such as the dat-1 dopamine transporter gene, which is expressed exclusively in their eight dopaminergic neurons.
With the capabilities of this model system and the appropriate technology, many laboratories have studied dopaminergic neurodegeneration. However, there is little consistency in the way the data is analyzed and much of the present literature uses binary scoring analyses that capture the presence of degeneration but not the full details of the progression of neuron loss. Here, we introduce a universal scoring system to assess morphological changes and degeneration in C. elegans&#39; cephalic neuron dendrites. This seven-point scale allows for analysis across a full range of dendrite morphology, ranging from healthy neurons to complete dendrite loss, and considering morphological details including kinks, branching, blebs, and breaks. With this scoring system, researchers can quantify subtle age-related changes as well as more dramatic chemical-induced changes. Finally, we provide a practice set of images with commentary that can be used to train, calibrate, and assess the scoring consistency of researchers new to this method. This should improve within- and between- laboratory consistency, increasing rigor and reproducibility.

Quantifying Levels of Dopaminergic Neuron Morphological Alteration and Degeneration in Caenorhabditis elegans

In this article, we showcase how to use a seven-point scoring system to consistently quantify changes to dopaminergic neuron dendrite morphology in C. elegans. This system is intended for analyses of dopaminergic neurodegeneration assays utilizing genetic, chemical, and age-based models of neurodegenerative disorders.

Dopamine neuron loss is involved in the pathology of Parkinson&#39;s Disease (PD), a highly prevalent neurodegenerative disorder affecting over 10 million ...

Protocol for Dendritic Spine Analysis Using Neurolucida 360

Dendritic Spine Quantification Using an Automatic Three-Dimensional Neuron Reconstruction Software

Synaptic connections allow for the exchange and processing of information between neurons. The post-synaptic site of excitatory synapses is often formed on dendritic spines. Dendritic spines are structures of great interest in research centered around synaptic plasticity, neurodevelopment, and neurological and psychiatric disorders. Dendritic spines undergo structural modifications during their lifespan, with properties such as total spine number, dendritic spine size, and morphologically defined subtype altering in response to different processes. Delineating the molecular mechanisms regulating these structural alterations of dendritic spines relies on morphological measurement. This mandates accurate and reproducible dendritic spine analysis to provide experimental evidence. The present study outlines a detailed protocol for dendritic spine quantification and classification using Neurolucida 360 (automatic three-dimensional neuron reconstruction software). This protocol allows for the determination of key dendritic spine properties such as total spine density, spine head volume, and classification into spine subtypes thus enabling effective analysis of dendritic spine structural phenotypes.

Author Spotlight: Optimizing Dendritic Spine Analysis for Balanced Manual and Automated Assessment in the Hippocampus CA1 Apical Dendrites

Dendritic spines are post-synaptic compartments of most excitatory synapses. Alterations to dendritic spine morphology occur during neurodevelopment, aging, learning, and many neurological and psychiatric disorders, underscoring the importance of reliable dendritic spine analysis. This protocol describes quantifying dendritic spine morphology accurately and reproducibly using automatic three-dimensional neuron reconstruction software.

Synaptic connections allow for the exchange and processing of information between neurons. The post-synaptic site of excitatory synapses is often formed ...

Imaging Dendritic Spines in Caenorhabditis elegans

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Quantifying Levels of Dopaminergic Neuron Morphological Alteration and Degeneration in Caenorhabditis elegans

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