We thank Bernard Kulemaka and members of the Smith lab for their helpful comments and reagent guidance, Sam Connell and Brent Redford of 3i for fielding imaging questions and Deborah Bang, Karen Heed and Kay Stewart for zebrafish care. This work was supported by the University of Notre Dame, the Elizabeth and Michael Gallagher Family, the Alfred P. Sloan Foundation, Center for Zebrafish Research at the University of Notre and Center of Stem Cells and Regenerative Medicine at the University of Notre Dame. All animal studies were done in compliance with University of Notre Dame IACUC to Dr. Cody Smith.

All animal studies were approved by the University of Notre Dame Institutional Animal Care and Use Committee.
1. Preparation of Zebrafish Specimen
<ol>
	<li>Place one adult male and one adult female Tg(convertible protein) into a mating chamber per standard procedures13. In this manuscript, use Tg(sox10:eos) fish9 because of access but other transgenic lines with photoconvertible protein can be equally used. Set up more than one chamber in case fish do not lay. Allow the fish to remain in the chamber overnight.</li>
	<li>The next morning, collect eggs in 100 mm Petri dishes. Allow eggs to mature to 24 h post-fertilization (hpf) before screening.</li>
	<li>If animals above were heterozygotes for the transgene, screen 24 hpf dishes for Tg(sox10:eos)+ embryos with a 488 nm light source and GFP filter sets on a dissecting microscope. Isolate Tg(sox10:eos)+ embryos and allow to mature to 48 hpf.</li>
	<li>Dechorionate embryos manually with a needle or tweezers.</li>
	<li>Prepare and microwave 5 mL of 0.8% low-melting point agarose solution.
	<ol>
		<li>Once agarose is cool to the touch, place 3-4 anesthetized Tg(sox10:eos)+48 hpf fish in the center of a 10 mm glass-coverslip bottom Petri dish.</li>
		<li>Add enough agarose to cover the surface of the coverslip, approximately 1 mL. Use a probe needle to arrange fish on their sides. Allow agarose to solidify to ensure mounting of animals. It may be necessary to continually re-arrange the zebrafish until the agar solidifies14. Solidification takes approximately 2 min.</li>
	</ol>
	</li>
	<li>After the agarose has solidified for 2 min, slowly add embryo medium containing 0.02% aminobenzoic acid ester (Tricaine) to dish until the bottom surface of the agar and dish is submerged. This should be approximately 3mL.</li>
</ol>
2. Microscope Mounting and Pre-conversion Imaging
<ol>
	<li>Open confocal software and select the capture and focus windows [Figure 1]. Under the capture window, select the lab-specific conversion imaging setting under the capture setting drop down tab [Figure 1]. Here, the lab-specific setting is called &#34;Fish Imaging.&#34;</li>
	<li>Place specimen on confocal scope and bring into focus using course and fine adjustment knobs.</li>
	<li>Open the focus window and locate the desired region of interest (i.e. dorsal root ganglia).</li>
	<li>Select the c488&#160; laser under the Filter Set menu. Set the exposure to 300 ms, laser power to 5, and intensify to 75 [Figure 2].</li>
	<li>Check the 3D box under the capture type section. In the 3D Capture section, select use current position and check range around current. In the same section, set the range to 35, the number of planes to 36, and the step size to 1. The range can be increased or decreased to accommodate for the depth of the imaging area. For the spinal cord a range value between 35-40 stacks typically is sufficient [Figure 5].</li>
	<li>Select current location [Figure 5].</li>
	<li>Click start at the bottom of the capture window to acquire image.</li>
</ol>
3. Single-cell Photoconversion
<ol>
	<li>Open confocal software and select the capture and focus windows [Figure 1]. Under the capture window, select the lab-specific conversion imaging setting under the capture setting drop down tab [Figure 1]. Here, the lab-specific setting is called &#34;Fish ablate full chip.&#34;</li>
	<li>Select the c488 and c541 laser under the Filter Set menu. If not using the same microscope software, find the menu to select different lasers and select the 488 nm and 541 nm lasers. Set the exposures to 300 ms, laser power to 5, and intensify to 75 [Figure 2]. These laser settings are selected based on producing enough fluorescent signal without causing photobleaching or toxicity. If toxicity or photobleaching is visualized, reduce laser power or exposure.</li>
	<li>Open the focus window and click on the photomanipulation tab in the focus window. Adjust laser parameters accordingly. Change the laser stack power to 2, and then click Go. Change the Raster block size to 1 and click Set. Change the Double-click size to 4. Change the laser line to v405 [Figure 3].</li>
	<li>Open the advanced capture settings in the capture window. Select the photomanipulation tab and change the Double-click repetitions to 2. Click OK [Figure 4].</li>
	<li>Select the XY tab in the focus window. Double check the laser parameters from step 2 in the photomanipulation tab [Figure 3, Figure 4]. Set laser settings to photoconvert the cell of interest without photoconversion of surrounding cells.
	<ol>
		<li>If photoconversion of adjacent cells is present, reduce laser power. If photoconversion of cells does not occur, laser powers can be increased. Optimally, set laser power to photoconvert only the region of interest and not surrounding areas.</li>
	</ol>
	</li>
	<li>Check the timelapse box under capture type. Then, click Start [Figure 6A].</li>
	<li>Once the live timelapse window opens, select the circle tool on the top toolbar [Figure 7]</li>
	<li>Draw a circle in the centermost region of the cell. Right click the drawn circle, select FRAP region, and wait 3 seconds. The selected area should become dimmer [Figure 8]. Click stop capture.</li>
	<li>If imaging more than one animal, go back to the XY tab in the focus menu and select position 2. Then, repeat steps 4.1-4.6.</li>
	<li>To convert a population of cells, follow the protocol and laser parameters for steps 4.1-4.4 except instead of drawing a circle to FRAP the region of interest, use the line tool. Draw a line on the region of interest and FRAP the region using the same parameters listed above.</li>
</ol>
4. Post-photoconversion Imaging
<ol>
	<li>Once all points are photoconverted. Select the lab-specific standard image stack setting described in the precoversion imaging step 3. This is found under the capture setting drop down tab in the capture window [Figure 5].</li>
	<li>Select the c488 laser and set the exposure to 300ms, laser power to 5, and intensify to 75 [Figure 2].</li>
	<li>Select the c541 laser found under the same menu as the c488 laser. Set the exposure to 500 ms, laser power to 10, and intensify to 75 [Figure 9].</li>
	<li>Check the 3D box under the capture type section. In the 3D Capture section, select use current position and check range around current. In the same section, set the range to 35, the number of planes to 36, and the step size to 1. The range number may increase or decrease to accommodate the desired imaging depth [Figure 5].</li>
	<li>If there are multiple points in the XY focus tab, select the multipoint list option in the capture window. If not, select current location.</li>
	<li>Click start at the bottom of the capture window to acquire image.</li>
</ol>

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F., Krisky, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Herpes+simplex+virus+vectors+for+gene+transfer+to+the+nervous+system.">Herpes simplex virus vectors for gene transfer to the nervous system.</a> J Neurovirol. 3 Suppl 1, S80-S88 (1997).</li><li>Boevink, P., Cruz, S., Hawes, C., Harris, N., Oparka, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Virus-mediated+delivery+of+the+green+fluorescent+protein+to+the+endoplasmic+reticulum+of+plant+cells.">Virus-mediated delivery of the green fluorescent protein to the endoplasmic reticulum of plant cells.</a> Plant J. 10 (5), 935-941 (1996).</li><li>Day, R. N., Davidson, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+fluorescent+protein+palette:+tools+for+cellular+imaging.">The fluorescent protein palette: tools for cellular imaging.</a> Chem Soc Rev. 38 (10), 2887-2921 (2009).</li><li>Wiedenmann, J., Ivanchenko, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EosFP,+a+fluorescent+marker+protein+with+UV-inducible+green-to-red+fluorescence+conversion.">EosFP, a fluorescent marker protein with UV-inducible green-to-red fluorescence conversion.</a> Proc Natl Acad Sci U S A. 101 (45), 15905-15910 (2004).</li><li>Maurel, D., Banala, S., Laroche, T., Johnsson, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photoactivatable+and+Photoconvertible+Fluorescent+Probes+for+Protein+Labeling.">Photoactivatable and Photoconvertible Fluorescent Probes for Protein Labeling.</a> ACS Chem Biol. 5 (5), 507-516 (2010).</li><li>Terskikh, A., Fradkov, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=&amp;#34;Fluorescent+timer&amp;#34;:+protein+that+changes+color+with+time.">&amp;#34;Fluorescent timer&amp;#34;: protein that changes color with time.</a> Science. 290 (5496), 1585-1588 (2000).</li><li>McGraw, H. 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The authors have nothing to disclose.

In complex tissues, distinct cell-types organize into specific domains. Recently techniques have been utilized to label individual cells within these large tissue structures1,2,3. Here we demonstrate two techniques that can similarly be utilized to visualize both single cell interactions and cell population interactions within complex tissues. The advantage of the photoconversion technique is the spatial control the scientist ha...

Multiple distinct cells interact to build and maintain complex animal and plant tissues. These cells are often intercalated and difficult to distinguish from neighbors at a single cell level without high resolution microscopy that require fixation of tissue. However, to understand how these tissues form, are maintained, and become diseased, it has been essential to investigate how single cells within the tissue are interacting over time. Ideally, these experiments require the labeling of single cells within a tissue in a non-invasive manner without the requirement of fixation. Scientists have now developed numerous techniques to accomplish this task1,2,3,4.
The discovery and implementation of the jellyfish green fluorescent protein (GFP) was one exciting approach that allowed for labeling of distinct cells in a tissue environment1. Using cell-specific promoters, it is possible to genetically select a subset of cells that are labeled1. Alternatively, viral induced expression of GFP can be utilized for user-selected expression of GFP3,4. Although quite useful, genetic mediated expression of GFP does not allow user-selected expression within a subset of cells in the tissue; and viral expression of GFP, although advantageous, can be invasive. With the advent of GFP derivatives and clever techniques like Brainbow to express distinct fluorescent proteins more sparsely within tissues, it has become possible to visualize single cells and the interactions among them in complex tissue2,5. However, these approaches label cells in a random fashion. If the desired experiment requires visualization of a single cell or population of cells that is defined by the experimenter, they are therefore limited. With such experiments, it would be advantageous to have a genetically expressed fluorescent protein that can be manipulated to distinguish, in a single cell fashion, it from other fluorescent and non-fluorescent cells.
To achieve this goal and visualize the cell biology of single cells within a complex living tissue, the scientific community uses single cell photoconversion of distinct fluorescent proteins6,7,8. Using genetically controlled expression of a photoconvertible protein (i.e., eos, kaede, etc.) that transitions from a green to red fluorescent state when exposed to UV (488 nm) light, we can distinguish a single cell from its fluorescently labeled neighbors6,7,8. This approach utilizes an apparatus attached to our confocal microscope which can direct light from a laser stack to a diffraction-limited region of interest. With this technique, we can either label single cells or larger populations in a user-defined manner9,10,11. The technique is minimally invasive compared to single cell injections of viral GFP. As a proof of concept, we show that we can photoconvert single cells within a ganglion in the peripheral nervous system and photoconvert larger populations like cells located on the ventral side of the spinal cord9,10,11,12. We then can visualize these photoconverted cell populations 24 h later to gain insight into their movement and differentiation during development.

<table><tbody><tr><td>&lt;font class=&quot;font5&quot;&gt;Tg(&lt;em&gt;sox10:eos&lt;/em&gt;)&lt;/font&gt;&lt;font class=&quot;font0&quot;&gt; zebrafish animals&lt;/font&gt;</td><td></td><td></td><td>Fish were obtained and crossed from the fish facility at the University of Notre Dame</td></tr><tr><td>100 X 15 mm petri dish</td><td>VWR</td><td>25384-302</td><td></td></tr><tr><td>Embryo medium</td><td></td><td></td><td>Embryo medium is made weekly and provided by the fish facility at the University of Notre Dame containing 5L RO water, 30uL methylene blue, and 200mL salt stock.</td></tr><tr><td>0.8% Low Melting Point Agarose</td><td>dot scientific inc</td><td>9012-36-6</td><td></td></tr><tr><td>35 X 10 mm glass-coverslip bottom petri dish</td><td>Ted Pella Inc.</td><td>14021-20</td><td></td></tr><tr><td>Needle dissecting probe</td><td></td><td></td><td></td></tr><tr><td>0.002% 3-aminobenzoic acid ester (Tricaine)</td><td>Fluka analytical</td><td>A5040-250G</td><td></td></tr><tr><td>Fluorescent Dissecting Microscope with GFP filters</td><td>Zeiss Axiozoom</td><td></td><td></td></tr><tr><td>Confocal microscope with lasers to excite GFP and RFP filter sets</td><td>3i spinning disk confocal</td><td></td><td></td></tr><tr><td>UV light source (laser) for photoconversion</td><td>405 nm laser</td><td></td><td></td></tr><tr><td>Slidebook software</td><td>3i</td><td></td><td></td></tr><tr><td>Methylene blue</td><td>Kordon</td><td></td><td></td></tr></tbody></table>

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.

Photoconversion of fluorescent proteins can be used to label distinct cells within a tissue6. To demonstrate this, Tg(sox10:eos) fish9 were used to express the photoconvertible protein Eos under the regulatory sequences of sox10. The Tg(sox10:eos) animals at 48 hpf were first mounted, and then imaged to detect any non-specific photoconversion that may have occurred. The Eos unconverted fluorescent signal with littl...

Watch this Scientific Journal Video about Single-cell Photoconversion in Living Intact Zebrafish at JoVE.com

Single-cell Photoconversion in Living Intact Zebrafish

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 ...

single-cell-photoconversion-in-living-intact-zebrafish

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

5.9K Views.  University of Notre Dame. 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. 

Video: Single-cell Photoconversion in Living Intact Zebrafish

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the field of developmental biology. The discovery and development of photoconvertible proteins have greatly aided our understanding of these dynamic changes by providing a method to optically highlight cells and tissues. However, while photoconversion, time-lapse microscopy, and subsequent image analysis have proven to be very successful in uncovering cellular dynamics in organs such as the brain or the eye, this approach is generally not used in the developing heart due to challenges posed by the rapid movement of the heart during the cardiac cycle. This protocol consists of two parts. The first part describes a method for photoconverting and subsequently tracking endocardial cells (EdCs) during zebrafish atrioventricular canal (AVC) and atrioventricular heart valve development. The method involves temporally stopping the heart with a drug in order for accurate photoconversion to take place. Hearts are allowed to resume beating upon removal of the drug and embryonic development continues normally until the heart is stopped again for high-resolution imaging of photoconverted EdCs at a later developmental time point. The second part of the protocol describes an image analysis method to quantify the length of a photoconverted or non-photoconverted region in the AVC in young embryos by mapping the fluorescent signal from the three-dimensional structure onto a two-dimensional map. Together, the two parts of the protocol allows one to examine the origin and behavior of cells that make up the zebrafish AVC and atrioventricular heart valve, and can potentially be applied for studying mutants, morphants, or embryos that have been treated with reagents that disrupt AVC and/or valve development.

This protocol describes a method for the photoconversion of Kaede fluorescent protein in endocardial cells of the living zebrafish embryo that enables the tracking of endocardial cells during atrioventricular canal and atrioventricular heart valve development.

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the ...

Culture and Transfection of Zebrafish Primary Cells

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in an intact and developing vertebrate. However, the detailed imaging of the morphologies of distinct cell types and subcellular structures is limited in whole mounts. Therefore, we established an efficient and easy-to-use protocol to culture live primary cells from zebrafish embryos and adult tissue.
In brief, 2 dpf zebrafish embryos are dechorionated, deyolked, sterilized, and dissociated to single cells with collagenase. After a filtration step, primary cells are plated onto glass bottom dishes and cultivated for several days. Fresh cultures, as much as long term differenciated ones, can be used for high resolution confocal imaging studies. The culture contains different cell types, with striated myocytes and neurons being prominent on poly-L-lysine coating. To specifically label subcellular structures by fluorescent marker proteins, we also established an electroporation protocol which allows the transfection of plasmid DNA into different cell types, including neurons. Thus, in the presence of operator defined stimuli, complex cell behavior, and intracellular dynamics of primary zebrafish cells can be assessed with high spatial and temporal resolution. In addition, by using adult zebrafish brain, we demonstrate that the described dissociation technique, as well as the basic culturing conditions, also work for adult zebrafish tissue.

We present an efficient and easy-to-use protocol for preparing primary cell cultures of zebrafish embryos for transfection and live cell imaging as well as a protocol to prepare primary cells from adult zebrafish brain.

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

Biology

Multicolor Time-lapse Imaging of Transgenic Zebrafish: Visualizing Retinal Stem Cells Activated by Targeted Neuronal Cell Ablation

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound transgenic fish which express different fluorescent reporters in neighboring cell types provide a means of following cellular interactions 2 and/or tissue-level responses to experimental manipulations over time. In this video, we demonstrate methods that can be used for imaging multiple transgenically labeled cell types serially in individual fish over time courses that can span from minutes to several days. The techniques described are applicable to any study seeking to correlate the "behavior" of neighboring cells types over time, including: 1) serial 'catch and release' methods for imaging a large number of fish over successive days, 2) simplified approaches for separating fluorophores with overlapping excitation/emission profiles (e.g., GFP and YFP), 3) use of hypopigmented mutant lines to extend the time window available for high-resolution imaging into late larval stages of development, 4) use of membrane targeted fluorescent reporters to reveal fine morphological detail of individual cells as well as cellular details in larger populations of cells, and 5) a previously described method for chemically-induced ablation of transgenically targeted cell types; i.e., nitroreductase (NTR) mediated conversion of prodrug substrates, such as metronidazole (MTZ), to cytotoxic derivatives 3,5.
As an example of these approaches, we will visualize the ablation and regeneration of a subtype of retinal bipolar neuron within individual fish over several days. Simultaneously we will monitor several other retinal cell types, including neighboring non-targeted bipolar cells and potential degeneration-stimulated retinal stem cells (i.e., M&#971;ller glia). This strategy is being applied in our lab to characterize cell- and tissue-level (e.g., stem cell niche) responses to the selective loss and regeneration of targeted neuronal cell types.

In this video, techniques for multicolor confocal time-lapse imaging and targeted cell ablation are provided. Time-lapse imaging is used to monitor the behavior of multiple cell types of interest in vivo. Targeted cell ablation facilitates the study neural circuit function and cell-specific neuronal regeneration paradigms.

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound ...

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

Cell Tracking Using Photoconvertible Proteins During Zebrafish Development

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of genetically-encoded fluorescent proteins has proven a valuable strategy for elucidating dynamic morphogenetic processes as they occur in the intact organism. During the past decade, the development of photoactivatable and photoconvertible fluorescent proteins has opened the possibility to investigate the fate of discrete subpopulations of tagged proteins1. Unlike photoactivatable proteins, photoconvertible fluorescent proteins (PCFPs) are readily tracked and imaged in their native emission state prior to photoconversion, making it easier to identify and select regions by optical inspection. PCFPs, such as Kaede2, KikGR3, Dendra4 and EosFP5, can be shifted from green to red upon exposure to UV or blue light due to a His-Tyr-Gly tripeptide sequence which forms a green chromophore that can be photoconverted to a red one by a light-catalyzed &beta;-elimination and subsequent extension of a &pi;-conjugated system3. PCFPs and their monomeric variants are useful tools for tracking cells6-10 and studying protein dynamics11-14, respectively. During recent years, PCFPs have been expressed in different animal model, such as zebrafish6, chicken7,8 and mouse9,10 for cell fate tracking. Here we report a protocol for cell-specific photoconversion of PCFPs in the living zebrafish embryo and further tracking of photoconverted proteins at later developmental stages. This methodology allows studying, in a tissue-specific manner, cell biological events underlying morphogenesis in the zebrafish animal model.

Here, we present a method for the photoactivated switch of photoconvertible fluorescent proteins (PCFPs) in the living zebrafish embryo and further tracking of photoconverted protein at specific time points during development. This methodology allows monitoring of cell biological events underlying different developmental processes in a live vertebrate organism.

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of ...