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><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&amp;times;/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 ...

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Universidad de Cartagena UDEC

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

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the cells are located. Specifically, they have been developed to analyze the electrical activity of excitable cells, such as spinal neurons. In such a scenario, it is crucial to preserve the integrity of the spinal cell, but it is also important to preserve the anatomy and physiological shape of the systems involved. Indeed, the comprehension of the manner in which the nervous system-and other complex systems-works must be based on a systemic approach. For this reason, the live zebrafish embryo was chosen as a model system, and the spinal neuron membrane voltage changes were evaluated without interfering with the physiological conditions of the embryos. 
Here, an approach combining the employment of zebrafish embryos with a FRET-based biosensor is described. Zebrafish embryos are characterized by a very simplified nervous system and are particularly suited for imaging applications thanks to their transparency, allowing for the employment of fluorescence-based voltage indicators at the plasma membrane during zebrafish development. The synergy between these two components makes it possible to analyze the electrical activity of the cells in intact living organisms, without perturbing the physiological state. Finally, this non-invasive approach can co-exist with other analyses (e.g., spontaneous movement recordings, as shown here).

Protocols described here allow for the study of the electrical properties of excitable cells in the most non-invasive physiological conditions by employing zebrafish embryos in an in vivo system together with a fluorescence resonance energy transfer (FRET)-based genetically encoded voltage indicator (GEVI) selectively expressed in the cell type of interest.

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the ...

Developmental Biology

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

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, ...

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

The nervous system is often described as a hard-wired component of the body even though it is a considerably fluid organ system that reacts to external stimuli in a consistent, stereotyped manner, while maintaining incredible flexibility and plasticity. Unlike the central nervous system (CNS), the peripheral nervous system (PNS) is capable of significant repair, but we have only just begun to understand the cellular and molecular mechanisms that govern this phenomenon. Using zebrafish as a model system, we have the unprecedented opportunity to couple regenerative studies with in vivo imaging and genetic manipulation. Peripheral nerves are composed of axons surrounded by layers of glia and connective tissue. Axons are ensheathed by myelinating or non-myelinating Schwann cells, which are in turn wrapped into a fascicle by a cellular sheath called the perineurium. Following an injury, adult peripheral nerves have the remarkable capacity to remove damaged axonal debris and re-innervate targets. To investigate the roles of all peripheral glia in PNS regeneration, we describe here an axon transection assay that uses a commercially available nitrogen-pumped dye laser to axotomize motor nerves in live transgenic zebrafish. We further describe the methods to couple these experiments to time-lapse imaging of injured and control nerves. This experimental paradigm can be used to not only assess the role that glia play in nerve regeneration, but can also be the platform for elucidating the molecular mechanisms that govern nervous system repair.

Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

The  nervous system is often described as a hard-wired component of the body even  though it is a considerably fluid organ system that reacts to external ...

Light-Induced GFP Expression in Zebrafish Embryos using the Optogenetic TAEL/C120 System

Inducible gene expression systems are an invaluable tool for studying biological processes. Optogenetic expression systems can provide precise control over gene expression timing, location, and amplitude using light as the inducing agent. In this protocol, an optogenetic expression system is used to achieve light-inducible gene expression in zebrafish embryos. This system relies on an engineered transcription factor called TAEL based on a naturally occurring light-activated transcription factor from the bacterium E. litoralis. When illuminated with blue light, TAEL dimerizes, binds to its cognate regulatory element called C120, and activates transcription. This protocol uses transgenic zebrafish embryos that express the TAEL transcription factor under the control of the ubiquitous ubb promoter. At the same time, the C120 regulatory element drives the expression of a fluorescent reporter gene (GFP). Using a simple LED panel to deliver activating blue light, induction of GFP expression can first be detected after 30 min of illumination and reaches a peak of more than 130-fold induction after 3 h of light treatment. Expression induction can be assessed by quantitative real-time PCR (qRT-PCR) and by fluorescence microscopy. This method is a versatile and easy-to-use approach for optogenetic gene expression.

Optogenetics is a powerful tool with wide-ranging applications. This protocol demonstrates how to achieve light-inducible gene expression in zebrafish embryos using the blue light-responsive TAEL/C120 system.

Inducible gene expression systems are an invaluable tool for studying biological processes. Optogenetic expression systems can provide precise control ...

Investigating Development with Optogenetic Signaling Manipulation in Zebrafish

Optogenetic Signaling Activation in Zebrafish Embryos

Signaling pathways orchestrate fundamental biological processes, including development, regeneration, homeostasis, and disease. Methods to experimentally manipulate signaling are required to understand how signaling is interpreted in these wide-ranging contexts. Molecular optogenetic&#160;tools can provide reversible, tunable manipulations of signaling pathway activity with a high degree of spatiotemporal control and have been applied in vitro, ex vivo, and in vivo. These tools couple light-responsive protein domains, such as the blue light homodimerizing light-oxygen-voltage sensing (LOV) domain, with signaling effectors to confer light-dependent experimental control over signaling. This protocol provides practical guidelines for using the LOV-based bone morphogenetic protein (BMP) and Nodal signaling activators bOpto-BMP and bOpto-Nodal in the optically accessible early zebrafish embryo. It describes two control experiments: A quick phenotype assay to determine appropriate experimental conditions, and an immunofluorescence assay to directly assess signaling. Together, these control experiments can help establish a pipeline for using optogenetic tools in early zebrafish embryos. These strategies provide a powerful platform to investigate the roles of signaling in development, health, and physiology.

Author Spotlight: Manipulating Signaling in Zebrafish Embryos to Decode Cell Fate Decisions 

Optogenetic manipulation of signaling pathways can be a powerful strategy to investigate how signaling is decoded in development, regeneration, homeostasis, and disease. This protocol provides practical guidelines for using light-oxygen-voltage sensing domain-based Nodal and bone morphogenic protein (BMP) signaling activators in the early zebrafish embryo.

Signaling pathways orchestrate fundamental biological processes, including development, regeneration, homeostasis, and disease. Methods to experimentally ...

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 ...

Protein Phase Separation Assay: An Optogenetic Method for Mutant RNA-Binding Protein Phase Separation in Spinal Motor Neurons of Zebrafish Larvae

Source: Asakawa, K. et al. <a href="https://jove.unicartagenaproxy.elogim.com/v/62932">Optogenetic Phase Transition of TDP-43 in Spinal Motor Neurons of Zebrafish Larvae.</a> J. Vis. Exp. (2022)
In this video, we describe a phase separation assay, wherein intracellular proteins comprising intrinsically disordered regions fused to a photosensitive oligomerization domain are induced via blue-light exposure to self-associate into membrane-less liquid-like condensates.

Source: Asakawa, K. et al. Optogenetic Phase Transition of TDP-43 in Spinal Motor Neurons of Zebrafish Larvae. J. Vis. Exp. (2022)
In this video, we ...

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Zebrafish Optogenetics: Activating Genetically Modified Somatosensory Neuron to Study Larval Behavioral Responses

Source: Palanca A. M. S., et al. <a href="https://jove.unicartagenaproxy.elogim.com/v/50184/" target="_blank">Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato</a>. J. Vis. Exp. (2013).
This video describes optogenetic techniques by activating genetically modified somatosensory neurons and recording elicited behavioral response with a high-speed video camera.

Source: Palanca A. M. S., et al. Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato. J. Vis. Exp. (2013).
This video describes ...

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

Summary

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Chapters in this video

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

Single-cell Photoconversion in Living Intact Zebrafish

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

Light-Induced GFP Expression in Zebrafish Embryos using the Optogenetic TAEL/C120 System

Optogenetic Signaling Activation in Zebrafish Embryos

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

Protein Phase Separation Assay: An Optogenetic Method for Mutant RNA-Binding Protein Phase Separation in Spinal Motor Neurons of Zebrafish Larvae

Zebrafish Optogenetics: Activating Genetically Modified Somatosensory Neuron to Study Larval Behavioral Responses