This work was supported by SERIKA FUND (KA), KAKENHI Grant numbers JP19K06933 (KA) and JP20H05345 (KA).

All fish work was conducted in accordance with the Guide for the Care and Use of Laboratory Animals of the Institutional Animal Care and Use Committee (approval identification number 24-2) of the National Institute of Genetics (Japan), which has an Animal Welfare Assurance on file (assurance number A5561-01) at the Office of Laboratory Animal Welfare of the National Institutes of Health (NIH, USA).
1. Construction of BACs for expression of optogenetic TDP-43 gene from the mnr2b promoter
<ol>
	<li>BAC preparation
	<ol>
		<li>Purchase a zebrafish BAC clone containing the zebrafish mnr2b locus (CH211-172N16, BACPAC Genomics). Purify the BAC DNA from a 5 mL overnight LB culture of DH10B Escherichia coli (E. coli) harboring CH211-172N16 as described in Warming et al.18.</li>
		<li>Transform CH211-172N16 into SW102 E. coli cells by electroporation, as described in Warming et al.18. 
		NOTE: Purification of the BAC DNA from the E. coli on the same day of electroporation usually gives a higher success rate of BAC transformation18. Otherwise, the purified BAC DNA is kept at -20 &#176;C until use.</li>
		<li>Introduce the iTol2-amp cassette for Tol2 transposon-mediated BAC transgenesis19 into the backbone of CH211-172N16 by electroporation, as described in Asakawa et al.20.
		<ol>
			<li>Make a glycerol stock of the E. coli cell clones carrying CH211-172N16 with the iTol2-amp cassette integration (CH211-172N16-iTol2A) after confirming the homologous recombination-mediated integration by polymerase chain reaction (PCR) (Ex Taq) using the primer pair Tol2-L-out (5&#39;-AAA GTA TCT GGC TAG AAT CTT ACT TGA-3&#39;) and pTARBAC-13371r (5&#39;-TAG CGG CCG CAA ATT TAT TA-3&#39;) and the following conditions: a denaturation step at 98 &#176;C for 1 min, followed by 25 cycles of denaturation at 95 &#176;C for 10 s, primer annealing at 55 &#176;C for 15 s, and primer extension at 72 &#176;C for 1 min, amplifying a PCR product of 354 base pairs (bp).</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>mnr2b-hs:opTDP-43h construction
	<ol>
		<li>Construct a plasmid carrying the expression cassette for the human wild-type TDP-43/TARDBP (TDP-43h) that is tagged with mRFP1 and CRY2olig at the N- and C-termini, respectively (hereafter, opTDP-43h)14. 
		&#8203;NOTE: The opTDP-43h fragment should be flanked with the zebrafish hsp70l gene promoter sequence (650 bp) and polyadenylation (polyA) signal sequence, followed by a kanamycin resistance gene (hsp70lp-opTDP-43h-polyA-Kan)14.</li>
		<li>Amplify the hsp70l-opTDP-43h-polyA-Kan cassette with primers that anneal hsp70l and Kan and contain 45 bp sequences of the upstream and downstream of the initiator codons of the mnr2b gene by PCR (GXL DNA Polymerase) using the primer pair mnr2b-hspGFF-Forward (5&#39;-tat cag cgc aat tac ctg caa ctc taa aca caa caa aag tgt tgc aGA ATT CAC TGG AGG CTT CCA GAA C-3&#39;) and Km-r (5&#39;-ggt tct tca gct aaa agg gcg tcg atc ctg aag ttc ttt gac ttt tcc atC AAT TCA GAA GAA CTC GTC AAG AA-3&#39;) with the following conditions: a denaturation step at 98 &#176;C for 1 min, followed by 30 cycles of denaturation at 95 &#176;C for 10 s, primer annealing at 55 &#176;C for 15 s, and primer extension at 68 &#176;C for 30 s, amplifying a PCR product of ~5.7 bp.</li>
		<li>Separate the PCR products by agarose gel electrophoresis (100 V) and purify the hsp70lp-opTDP-43h-polyA-Kan DNA band with a DNA column. Adjust the concentration of the purified hsp70lp-opTDP-43h-polyA-Kan cassette to 50 ng/&#181;L in Tris-EDTA buffer (TE) containing 10 mM Tris-HCl (pH 8.0) and 1 mM EDTA.</li>
		<li>Introduce the hsp70lp-opTDP-43h-polyA-Kan cassette into CH211-172N16-iTol2A by electroporation as described in Warming et al.18 and select ampicillin- and kanamycin-resistant transformants on LB agar plates. CH211-172N16-iTol2A carrying the hsp70lp-opTDP-43h-polyA-Kan cassette is designated as mnr2b-hs:opTDP-43h.</li>
		<li>Purify mnr2b-hs:opTDP-43h using a BAC purification kit and dissolve it at 250 ng/&#181;L in TE after phenol/chloroform extraction.</li>
	</ol>
	</li>
	<li>mnr2b-hs:EGFP-TDP-43z construction
	<ol>
		<li>Construct another plasmid carrying the zebrafish wild-type tardbp that is tagged with enhanced green fluorescent protein (EGFP) at its N-terminus (EGFP-TDP-43z) instead of opTDP-43h but is otherwise identical to the hsp70l-opTDP-43h-polyA-Kan construct (hsp70l-EGFP-TDP-43z-polyA-Kan)14. Use EGFP-TDP-43z as an internal control for the light stimulation of opTDP-43h.</li>
		<li>Construct CH211-172N16-iTol2A harboring the hsp70lp-EGFP-TDP-43z-polyA-Kan cassette in the mnr2b locus as described in 1.2.1-1.2.5. CH211-172N16-iTol2A carrying the hsp70lp-EGFP-TDP-43z-polyA-Kan cassette is designated as mnr2b-hs:EGFP-TDP-43z.</li>
	</ol>
	</li>
</ol>
2. Tol2 transposon-mediated BAC transgenesis in zebrafish
<ol>
	<li>Prepare the injection solution containing 40 mM KCl, phenol red (10% v/v), 25 ng/&#181;L of the mnr2b-hs:opTDP-43h DNA, and 25 ng/&#181;L Tol2 transposase mRNA19.</li>
	<li>Inject 1 nL of the injection solution (a droplet with a diameter of approximately 123 &#181;m calibrated in mineral oil) into the cytosol of wild-type zebrafish embryos at the one-cell stage. Screen the injected fish for the formation of red fluorescent protein (RFP)-positive aggregates in various embryonic tissues, including spinal motor neurons, under a fluorescence stereomicroscope at 2-3 days post-fertilization (dpf). Raise the RFP-positive fish to adulthood.</li>
	<li>Inject the mnr2b-hs:EGFP-TDP-43z DNA as described in 2.1 and 2.2. Raise EGFP-positive fish to adulthood.</li>
	<li>After a few months, put sexually matured injected fish and wild-type fish in pairs in standard 2 L mating cages to obtain F1 offspring. Screen F1 fish at 3 dpf for RFP (opTDP-43h) or EGFP (EGFP-TDP-43z) fluorescence in the spinal motor column using an epifluorescence microscope equipped with a Plan-Neofluar 5x/0.15 objective lens. Typically, one founder fish is identified from 10&#8722;20 injected fish (the germline transmission rate is 5%&#8722;10%).</li>
	<li>Isolate and compare multiple Tg[mnr2b-hs:opTDP-43h] and Tg[mnr2b-hs:EGFP-TDP-43z] inserts from different founder fish, as the intensity, but not the pattern, of opTDP-43h or EGFP-TDP-43z expression may vary between founder fish due to chromosomal position effects.</li>
	<li>Cross Tg[mnr2b-hs:opTDP-43h] and Tg[mnr2b-hs:EGFP-TDP-43z] fish lines to obtain offspring containing Tg[mnr2b-hs:opTDP-43h] Tg[mnr2b-hs:EGFP-TDP-43z] double-transgenic fish in a Mendelian ratio.</li>
	<li>Raise the fish in a plastic dish containing 30 mL of E3 buffer (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, 10-5% Methylene Blue) and add 0.003% (w/v) N-phenylthiourea at 8-10 h post-fertilization (hpf) to inhibit melanogenesis.</li>
	<li>Cover the plastic dish with aluminum foil after 30 hpf.</li>
</ol>
3. Preparation of LED for blue light illumination
<ol>
	<li>Turn on an LED panel by using the associated application installed on a tablet/phone. Put the probe of a spectrometer into an empty well of a 6 well dish and adjust the LED light to the wavelength peaking at ~456 nm through the application. Place the optical sensor of an optical power meter in the empty well and adjust the power of the LED light (~0.61&#160;mW/cm2). The LED light setting can be saved and is retrievable in the application.</li>
	<li>Introduce the dish/LED panel setting to the incubator at 28 &#176;C. Finish this step before imaging of fish starts at 48 hpf.</li>
</ol>
4. Imaging of zebrafish larvae expressing optogenetic TDP-43
<ol>
	<li>Select Tg[mnr2b-hs:opTDP-43h] Tg[mnr2b-hs:EGFP-TDP-43z] double-transgenic fish at least before 47 hpf, based on RFP (opTDP-43h) or EGFP (EGFP-TDP-43z) fluorescence in the spinal motor column using the epifluorescence microscope set-up described above.</li>
	<li>Dechorionate the Tg[mnr2b-hs:opTDP-43h] Tg[mnr2b-hs:EGFP-TDP-43z] double-transgenic fish.</li>
	<li>Preheat 1% low melting temperature agarose containing 250 &#181;g/mL of ethyl 3-aminobenzoate methanesulfonate salt at 42 &#176;C.</li>
	<li>Briefly anesthetize Tg[mnr2b-hs:opTDP-43h] Tg[mnr2b-hs:EGFP-TDP-43] double-transgenic fish at 48 hpf in E3 buffer containing the same concentration of Tricane.</li>
	<li>Put a drop of the preheated 1% low melting temperature agarose on the glass base dish at the room temperature. The diameter of the dome-shaped agarose drop on the glass dish is 8-10 mm.</li>
	<li>Using a Pasteur pipette, add the anesthetized fish to the low melting temperature agarose on the glass base dish, and then mix by pipetting a few times. Minimize the amount of the E3 buffer added to the agarose along with the fish.</li>
	<li>Maintain the fish on its side by using a syringe needle during the solidification of agarose (typically ~1 min) to ensure that the spinal cord is in an appropriate horizontal position. After the solidification, put a couple of drops of E3 buffer onto the dome-shaped agarose-mounted fish.</li>
	<li>Acquire serial confocal z-sections of the spinal cord by scanning with a confocal microscope equipped with a 20x water immersion objective lens with the numerical aperture 1.00, using a scan speed of 4.0 &#181;s per pixel (12 bits per pixel), a step size of 1.0 &#181;m per slice for the objective, and a combination of excitation/emission wavelengths: Channel 1) 473/510 nm for EGFP and Channel 2) 559/583 nm for mRFP1. 
	NOTE: The cloaca on the ventral side of the fish is included in the regions of interest (ROI) as a reference, which helps to identify and compare the spinal segments (levels 16-17) across the time points.</li>
	<li>Remove the fish from the agarose by carefully cracking the agarose with a syringe needle as soon as the imaging is complete. Keep the amount of time the fish is embedded in the agarose as short as possible, although the agarose embedding for &#60;30 min does not affect the viability of the fish.</li>
</ol>
5. Light stimulation of opTDP-43h-expressing fish by field illumination of a blue light-emitting diode (LED) light
<ol>
	<li>Add 7.5 mL of E3 buffer to the well and place the imaged Tg[mnr2b-hs:opTDP-43h] Tg[mnr2b-hs:EGFP-TDP-43z] double-transgenic fish into the well. Place the six-well dish on the LED panel by keeping the dish and LED panel 5 mm apart with a spacer (for example, with five slide glasses stacked).</li>
	<li>Turn on the blue LED light. Keep some of the Tg[mnr2b-hs:opTDP-43h] Tg[mnr2b-hs:EGFP-TDP-43z] double-transgenic fish in a separate six-well dish covered with aluminum foil when unilluminated control fish are necessary (i.e., in dark conditions)14.</li>
	<li>After the illumination (e.g., for 24 h at 72 hpf in Figure 3), image the spinal cord of the illuminated fish by repeating the steps 4.3 - 4.9.</li>
</ol>
6. Visualization of cytoplasmic relocation of optogenetic TDP-43 in the spinal motor neurons
<ol>
	<li>Open the image file in ImageJ/Fiji21 (Version: 2.1.0/1.53c), an open-source Java image processing program developed by NIH Image, which can be downloaded from https://imagej.net/Fiji/Downloads.</li>
	<li>Use the Z scrollbar to move through the focal planes. Create a maximum intensity projection of multiple slices by clicking Image | Stacks | Z project | Max Intensity and setting Start slice and Stop slice that cover the hemisegments of the spinal motor column.</li>
	<li>Split the multichannel image into two single-channel images by clicking Image | Color | Split Channels.</li>
	<li>Enhance the EGFP-TDP-43z signal to ensure that cytoplasmic EGFP-TDP-43z is visible clearly by clicking Image | Adjust | Brightness/Contrast and adjusting Minimum</li>
	<li>Open the ROI manager by clicking Analyze | Tools | ROI manager. Find the cells that are identifiable in both images at 48 hpf and 72 hpf, based on the relative positions of the cell bodies. Set the ROIs by outlining the contours of cell bodies of single spinal motor neurons visualized by the EGFP-TDP-43z signal using Freehand selections, and then clicking Add[t] in the ROI manager.</li>
	<li>Set a major axis of the soma by drawing a straight line using Straight and clicking Analyze | Plot Profile for EGFP-TDP-43z (C1) and opTDP-43h (C2) images at 48 and 72 hpf.</li>
	<li>Normalize the Plot Profile by dividing the values by the greatest value for each EGFP-TDP-43z and opTDP-43h signal. Plot the normalized values with x-y coordinates, where x and y represent the major axis of the soma and relative fluorescent intensities, respectively, using XY graph in a statistical software.</li>
</ol>
7. Ratiometric comparison between opTDP-43h and EGFP-TDP-43z signals using ImageJ/Fiji
<ol>
	<li>Open the image file in ImageJ/Fiji and set ROIs for single mnr2b-positive cells using a maximum intensity projection image of EGFP-TDP-43z as in 6.1-6.5. Add a ROI outside the ventral spinal cord (e.g., notochord) to represent the background signal (background ROI).</li>
	<li>For each EGFP-TDP-43z and opTDP-43 image, create a projection of multiple slices by clicking Image | Stacks | Z project | Sum Slices and set Start slice and Stop slice that cover the hemispinal motor column.</li>
	<li>Open the ROI manager created with the maximum intensity projection image. Display the ROIs on the Sum Slices image by selecting the image window for EGFP-TDP-43z and then clicking Show All in the ROI manager. Click Measure in the ROI manager to obtain mean values for each ROI.</li>
	<li>Acquire mean values for opTDP-43h following the same procedure provided in 7.3.</li>
	<li>For each ROI, after subtracting the mean for the background ROI, divide the subtracted mean for opTDP-43h by the subtracted mean for EGFP-TDP-43z to obtain the ratiometric value. The ratiometric values can be compared between different time points and presented using Column graph in Prism software.</li>
</ol>

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KA and KK are the inventors of the intellectual property described in this manuscript and provisional patents have been submitted by the National Institute of Genetics.

The mnr2b-BAC-mediated expression of opTDP-43h and EGFP-TDP-43z in zebrafish provides a unique opportunity for live imaging of TDP-43 phase transition in the spinal motor neurons. The optical transparency of body tissues of zebrafish larvae allows for the simple and noninvasive optogenetic stimulation of opTDP-43h. Comparisons between single spinal motor neurons over time demonstrated that the light-dependent oligomerization of opTDP-43h causes its cytoplasmic clustering, which is reminiscent of ALS pathology.</...

Ribonucleoprotein (RNP) granules control a myriad of cellular activities in the nucleus and cytoplasm by assembling membrane-less partitions via liquid-liquid phase separation (LLPS), a phenomenon in which a homogeneous fluid demixes into two distinct liquid phases1,2. The dysregulated LLPS of RNA-binding proteins that normally function as RNP granule components promote abnormal phase transition, leading to protein aggregation. This process has been implicated in neurodevelopmental and neurodegenerative diseases3,4,5. The precise evaluation of a causal relationship between aberrant LLPS of RNA-binding proteins and disease pathogenesis is crucial for determining whether and how LLPS can be exploited as an effective therapeutic target. LLPS of RNA-binding proteins is relatively easy to study in vitro and in unicellular models but is difficult in multicellular organisms, especially in vertebrates. A critical requirement for analyzing such LLPS in individual cells within a tissue environment is to stably express a probe for the imaging and manipulation of LLPS in a disease-vulnerable cell type of interest.
Amyotrophic lateral sclerosis (ALS) is an ultimately fatal neurological disorder in which motor neurons of the brain and spinal cord are selectively and progressively lost due to degeneration. To date, mutations in more than 25 genes have been associated with the heritable (or familial) form of ALS, which accounts for 5%-10% of total ALS cases, and some of these ALS-causing genes encode RNA-binding proteins consisting of RNPs, such as hnRNPA1, TDP-43, and FUS6,7. Moreover, the sporadic form of ALS, which accounts for 90%-95% of total ALS cases, is characterized by the cytoplasmic aggregation of TDP-43 deposited in degenerating motor neurons. A major characteristic of these ALS-associated RNA-binding proteins is their intrinsically disordered regions (IDRs) or low-complexity domains that lack ordered three-dimensional structures and mediate weak protein-protein interactions with many different proteins that drive LLPS7,8. The fact that ALS-causing mutations often occur in the IDRs has led to the idea that aberrant LLPS and phase transition of these ALS-related proteins may underlie ALS pathogenesis9,10.
Recently, the optoDroplet method, a Cryptochrome 2-based optogenetic technique that allows the modulation of protein-protein interactions by light, was developed to induce phase transition of proteins with IDRs11. As this technique has been extended successfully to TDP-43, it has begun to uncover the mechanisms underlying pathological phase transition of TDP-43 and its associated cytotoxicity12,13,14,15. In this protocol, we outline a genetic method to deliver an optogenetic TDP-43 to ALS-vulnerable cell types, namely, spinal motor neurons in zebrafish using the BAC for the mnr2b/mnx2b gene encoding a homeodomain protein for motor neuron specification16,17. The high translucency of zebrafish larvae allows for simple, noninvasive light stimulation of the optogenetic TDP-43 that triggers its phase transition in the spinal motor neurons. We also present a basic workflow for the live imaging of the zebrafish spinal motor neurons and image analysis using the freely available Fiji/ImageJ software to characterize the responses of the optogenetic TDP-43 to the light stimulation. These methods allow for an investigation of TDP-43 phase transition in an ALS-vulnerable cellular environment and should help to explore its pathological consequences at cellular and behavioral levels.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Confocal microscope</td>
			<td>Olympus</td>
			<td>FV1200</td>
			<td></td>
		</tr>
		<tr>
			<td>Epifluorescence microscope</td>
			<td>ZEISS</td>
			<td>Axioimager Z1</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence stereomicroscope</td>
			<td>Leica</td>
			<td>MZ16FA</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass base dish</td>
			<td>IWAKI</td>
			<td>3910-035</td>
			<td></td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td>MEE</td>
			<td>CN-25C</td>
			<td></td>
		</tr>
		<tr>
			<td>LED panel</td>
			<td>Nanoleaf Limited</td>
			<td>Nanoleaf AURORA smarter kit</td>
			<td></td>
		</tr>
		<tr>
			<td>Mupid-2plus</td>
			<td>TAKARA</td>
			<td>AD110</td>
			<td></td>
		</tr>
		<tr>
			<td>NucleoBond BAC100</td>
			<td>MACHEREY-NAGEL</td>
			<td>740579</td>
			<td></td>
		</tr>
		<tr>
			<td>NuSieve GTG Agarose</td>
			<td>LONZA</td>
			<td>50181</td>
			<td></td>
		</tr>
		<tr>
			<td>Objective lens</td>
			<td>Olympus</td>
			<td>XLUMPlanFL N 20&#215;/1.00</td>
			<td></td>
		</tr>
		<tr>
			<td>Objective lens</td>
			<td>ZEISS</td>
			<td>Plan-Neofluar 5x/0.15</td>
			<td></td>
		</tr>
		<tr>
			<td>Optical power meter</td>
			<td>HIOKI</td>
			<td>3664</td>
			<td></td>
		</tr>
		<tr>
			<td>Optical sensor</td>
			<td>HIOKI</td>
			<td>9742-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Phenol red solution 0.5%</td>
			<td>Merck</td>
			<td>P0290-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>PrimeSTAR GXL DNA Polymerase</td>
			<td>TAKARA</td>
			<td>R050A</td>
			<td></td>
		</tr>
		<tr>
			<td>QIAquick Gel Extraction Kit</td>
			<td>Qiagen</td>
			<td>28704</td>
			<td></td>
		</tr>
		<tr>
			<td>Six-well dish</td>
			<td>FALCON</td>
			<td>353046</td>
			<td></td>
		</tr>
		<tr>
			<td>Spectrometer probe BLUE-Wave</td>
			<td>StellerNet Inc.</td>
			<td>VIS-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe needle</td>
			<td>TERUMO</td>
			<td>NN-2725R</td>
			<td></td>
		</tr>
		<tr>
			<td>TaKaRa Ex Taq</td>
			<td>TAKARA</td>
			<td>RR001A</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricane</td>
			<td>Sigma-Aldrich</td>
			<td>A5040</td>
			<td></td>
		</tr>
		<tr>
			<td>Zebrafish BAC clone CH211-172N16</td>
			<td>BACPAC Genomics</td>
			<td>CH211-172N16</td>
			<td></td>
		</tr>
	</tbody>
</table>

optogenetic phase transition of tdp-43 in spinal motor neurons of zebrafish larvae

Abnormal protein aggregation and selective neuronal vulnerability are two major hallmarks of neurodegenerative diseases. Causal relationships between these features may be interrogated by controlling the phase transition of a disease-associated protein in a vulnerable cell type, although this experimental approach has been limited so far. Here, we describe a protocol to induce phase transition of the RNA/DNA-binding protein TDP-43 in spinal motor neurons of zebrafish larvae for modeling cytoplasmic aggregation of TDP-43 occurring in degenerating motor neurons in amyotrophic lateral sclerosis (ALS). We describe a bacterial artificial chromosome (BAC)-based genetic method to deliver an optogenetic TDP-43 variant selectively to spinal motor neurons of zebrafish. The high translucency of zebrafish larvae allows for the phase transition of the optogenetic TDP-43 in the spinal motor neurons by a simple external illumination using a light-emitting diode (LED) against unrestrained fish. We also present a basic workflow of live imaging of the zebrafish spinal motor neurons and image analysis with freely available Fiji/ImageJ software to characterize responses of the optogenetic TDP-43 to the light illumination. This protocol enables the characterization of TDP-43 phase transition and aggregate formation in an ALS-vulnerable cellular environment, which should facilitate an investigation of its cellular and behavioral consequences.

Live imaging of optogenetic and non-optogenetic TDP-43 proteins in the mnr2b+ spinal motor neurons of zebrafish larvae 
To induce TDP-43 phase transition in the spinal motor neurons in zebrafish, a human TDP-43h that is tagged with mRFP1 and CRY2olig22 at the N- and C-termini, respectively, was constructed and designated as opTDP-43h14 (Figure 1A). The opTDP-43h gene fragment was ...

Watch this Scientific Journal Video about Optogenetic Phase Transition of TDP-43 in Spinal Motor Neurons of Zebrafish Larvae at JoVE.com

Optogenetic Phase Transition of TDP-43 in Spinal Motor Neurons of Zebrafish Larvae

We describe a protocol to induce phase transition of TAR DNA-binding protein 43 (TDP-43) by light in the spinal motor neurons using zebrafish as a model.

Abnormal protein aggregation and selective neuronal vulnerability are two major hallmarks of neurodegenerative diseases. Causal relationships between ...

optogenetic-phase-transition-tdp-43-spinal-motor-neurons-zebrafish

Research

JoVE Journal

Neuroscience

5.9K Views.  Tokyo Medical University. We describe a protocol to induce phase transition of TAR DNA-binding protein 43 (TDP-43) by light in the spinal motor neurons using zebrafish as a model. 

Video: Optogenetic Phase Transition of TDP-43 in Spinal Motor Neurons of Zebrafish Larvae

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato

Larval zebrafish are emerging as a model for describing the development and function of simple neural circuits. Due to their external fertilization, rapid development, and translucency, zebrafish are particularly well suited for optogenetic approaches to investigate neural circuit function. In this approach, light-sensitive ion channels are expressed in specific neurons, enabling the experimenter to activate or inhibit them at will and thus assess their contribution to specific behaviors. Applying these methods in larval zebrafish is conceptually simple but requires the optimization of technical details. Here we demonstrate a procedure for expressing a channelrhodopsin variant in larval zebrafish somatosensory neurons, photo-activating single cells, and recording the resulting behaviors. By introducing a few modifications to previously established methods, this approach could be used to elicit behavioral responses from single neurons activated up to at least 4 days post-fertilization (dpf). Specifically, we created a transgene using a somatosensory neuron enhancer, CREST3, to drive the expression of the tagged channelrhodopsin variant, ChEF-tdTomato. Injecting this transgene into 1-cell stage embryos results in mosaic expression in somatosensory neurons, which can be imaged with confocal microscopy. Illuminating identified cells in these animals with light from a 473 nm DPSS laser, guided through a fiber optic cable, elicits behaviors that can be recorded with a high-speed video camera and analyzed quantitatively. This technique could be adapted to study behaviors elicited by activating any zebrafish neuron. Combining this approach with genetic or pharmacological perturbations will be a powerful way to investigate circuit formation and function.

Optogenetic techniques have made it possible to study the contribution of specific neurons to behavior. We describe a method in larval zebrafish for activating single somatosensory neurons expressing a channelrhodopsin variant (ChEF) with a diode-pumped solid state (DPSS) laser and recording the elicited behaviors with a high-speed video camera.

Larval  zebrafish are emerging as a model for describing the development and function  of simple neural circuits. Due to their  external fertilization, ...

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.
This extracellular technique has advantages. First, extracellular electrodes can stimulate and record neurons through the sheath5, so it does not need to be removed. Thus, neurons will be healthier in extracellular experiments than in intracellular ones. Second, if ganglia are rotated by appropriate pinning of the sheath, extracellular electrodes can access neurons on both sides of the ganglion, which makes it easier and more efficient to identify multiple neurons in the same preparation. Third, extracellular electrodes do not need to penetrate cells, and thus can be easily moved back and forth among neurons, causing less damage to them. This is especially useful when one tries to record multiple neurons during repeating motor patterns that may only persist for minutes. Fourth, extracellular electrodes are more flexible than intracellular ones during muscle movements. Intracellular electrodes may pull out and damage neurons during muscle contractions. In contrast, since extracellular electrodes are gently pressed onto the sheath above neurons, they usually stay above the same neuron during muscle contractions, and thus can be used in more intact preparations.
To uniquely identify motor neurons for a motor pool (in particular, the I1/I3 muscle in Aplysia) using extracellular electrodes, one can use features that do not require intracellular measurements as criteria: soma size and location, axonal projection, and muscle innervation4,6,7. For the particular motor pool used to illustrate the technique, we recorded from buccal nerves 2 and 3 to measure axonal projections, and measured the contraction forces of the I1/I3 muscle to determine the pattern of muscle innervation for the individual motor neurons.
We demonstrate the complete process of first identifying motor neurons using muscle innervation, then characterizing their timing during motor patterns, creating a simplified diagnostic method for rapid identification. The simplified and more rapid diagnostic method is superior for more intact preparations, e.g. in the suspended buccal mass preparation8 or in vivo9. This process can also be applied in other motor pools10,11,12 in Aplysia or in other animal systems2,3,13,14.

Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica

In animals with large identified neurons (e.g. mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.

In animals with large  identified neurons (e.g. mollusks), analysis of motor pools is done using  intracellular techniques1,2,3,4. Recently,  we developed ...

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the underlying neuronal components in the circuits1-6. Application of optogenetics7,8 in the larval neural circuit allows us to manipulate neuronal activity in spatially and temporally patterned ways9-13. Typically, specimens are broadly illuminated with a mercury lamp or LED, so specificity of the target neurons is controlled by binary gene expression systems such as the Gal4-UAS system14,15. In this work, to improve the spatial resolution to "sub-genetic resolution", we locally illuminated a subset of neurons in the ventral nerve cord using lasers implemented in a conventional confocal microscope. While monitoring the motion of the body wall of the semi-intact larvae, we interactively activated or inhibited neural activity with channelrhodopsin16,17 or halorhodopsin18-20, respectively. By spatially and temporally restricted illumination of the neural tissue, we can manipulate the activity of specific neurons in the circuit at a specific phase of behavior. This method is useful for studying the relationship between the activities of a local neural assembly in the ventral nerve cord and the spatiotemporal pattern of motor output.

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Here we describe a protocol for optogenetic manipulation of motoneuronal activity while monitoring changes in motor output (muscle contraction) in semi-intact Drosophila larvae using lasers within a conventional confocal microscope. This technique enables researchers to achieve local perturbation of neural activity in a few neuromeres to elucidate the dynamics of motor circuits.

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the ...

Automated High-throughput Behavioral Analyses in Zebrafish Larvae

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system measures spontaneous behaviors and the response to an aversive stimulus, which is shown to the larvae via a PowerPoint presentation. The recorded images are analyzed with an ImageJ macro, which automatically splits the color channels, subtracts the background, and applies a threshold to identify individual larvae placement in the lanes. We can then import the coordinates into an Excel sheet to quantify swim speed, preference for edge or side of the lane, resting behavior, thigmotaxis, distance between larvae, and avoidance behavior. Subtle changes in behavior are easily detected using our system, making it useful for behavioral analyses after exposure to environmental toxicants or pharmaceuticals.

Our laboratory developed a novel high-throughput automated imaging system that is useful for the detection of several different behaviors in 7-day-old zebrafish larvae. The system can be used for detecting subtle changes in behavior after the larvae have been exposed to environmental toxicants or pharmaceuticals.

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the surface of the cornea, electrical activity generated in response to light can be measured and used to assess the activity of retinal cells in vivo. This manuscript describes the use of the ERG to measure visual function in zebrafish. Zebrafish have long been utilized as a model for vertebrate development due to the ease of gene suppression by morpholino oligonucleotides and pharmacological manipulation. At 5-10 dpf, only cones are functional in the larval retina. Therefore, the zebrafish, unlike other animals, is a powerful model system for the study of cone visual function in vivo. This protocol uses standard anesthesia, micromanipulation and stereomicroscopy protocols that are common in laboratories that perform zebrafish research. The outlined methods make use of standard electrophysiology equipment and a low light camera to guide the placement of the recording microelectrode onto the larval cornea. Finally, we demonstrate how a commercially available ERG stimulator/recorder originally designed for use with mice can easily be adapted for use with zebrafish. ERG of larval zebrafish provides an excellent method of assaying cone visual function in animals that have been modified by morpholino oligonucleotide injection as well as newer genome engineering techniques such as Zinc Finger Nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9, all of which have greatly increased the efficiency and efficacy of gene targeting in zebrafish. In addition, we take advantage of the ability of pharmacological agents to penetrate zebrafish larvae to evaluate the molecular components that contribute to the photoresponse. This protocol outlines a setup that can be modified and used by researchers with various experimental goals.

We present a method for the electroretinographic (ERG) analysis of zebrafish larvae utilizing micromanipulation and electroretinography techniques. This is a simple and straightforward method for assaying visual function of zebrafish larvae in vivo.

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the ...

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.