Name: eye removal in living zebrafish larvae to examine innervation-dependent growth and development of the visual system
Uploaded: 2022-02-11T14:00:00
Description: Watch this Scientific Journal Video about Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System at JoVE.com

Funding for this work was supported primarily by start-up funds from Reed College to KLC, Helen Stafford Research Fellowship funds to OLH, and a Reed College Science Research Fellowship to YK. This project began in Steve Wilson&#39;s lab as a collaboration with HR, who was supported by a Wellcome Trust Studentship (2009-2014). We thank M&#225;t&#233; Varga, Steve Wilson, and other members of the Wilson lab for initial discussions about this project, and we especially thank Florencia Cavodeassi and Kate Edwards, who were the first to teach KLC how to mount embryos in agarose and perform zebrafish brain dissections. We also thank Greta Glover and Jay Ewing for help with assembling our tungsten needle-sharpening device.

The methods in this paper were conducted in accordance with guidelines and approval of the Institutional Animal Care and Use Committees of Reed College and University College London. See the Table of Materials for details about zebrafish strains used in this study.
1. Prepare materials and tools
<ol>
	<li>Make solutions.
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		<li>Make embryo medium (E314) by diluting a 60x stock (300 mM NaCl, 10.2 mM KCl, 20 mM CaCl2-dih...

<ol>
	<li>Butler, A. B., Hodos, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optic+tectum.">Optic tectum.</a> Comparative Vertebrate Neuroanatomy: Evolution and Adaptation. , 311-340 (2005).</li><li>Cang, J., Feldheim, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+mechanisms+of+topographic+map+formation+and+alignment.">Developmental mechanisms of topographic map formation and alignment.</a> Annual Review of Neuroscience. 36 (1), 51-77 (2013).</li><li>Basso, M. A., Bickford, M. E., Cang, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unraveling+circuits+of+visual+perception+and+cognition+through+the+superior+colliculus.">Unraveling circuits of visual perception and cognition through the superior colliculus.</a> Neuron. 109 (6), 918-937 (2021).</li><li>Burrill, J. D., Easter, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+retinofugal+projections+in+the+embryonic+and+larval+zebrafish+(Brachydanio+rerio).">Development of the retinofugal projections in the embryonic and larval zebrafish (Brachydanio rerio).</a> The Journal of Comparative Neurology. 346 (4), 583-600 (1994).</li><li>Regeneration in the goldfish visual system. Webvision: The Organization of the Retina and Visual System Available from: <a target="_blank" href="https://webvision.med.utah.edu/book/part-x-repair-and-regeneration-in-the-visual-system/regeneration-in-the-goldfish-visual-system/">https://webvision.med.utah.edu/book/part-x-repair-and-regeneration-in-the-visual-system/regeneration-in-the-goldfish-visual-system/</a> (2021)</li><li>Regeneration in the visual system of adult mammals. Webvision: The Organization of the Retina and Visual System Available from: <a target="_blank" href="https://webvision.med.utah.edu/book/part-x-repair-and-regeneration-in-the-visual-system/regeneration-in-the-visual-system-of-adult-mammals/">https://webvision.med.utah.edu/book/part-x-repair-and-regeneration-in-the-visual-system/regeneration-in-the-visual-system-of-adult-mammals/</a> (2021)</li><li>Cerveny, K. L., Varga, M., Wilson, S. 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R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+lens+has+a+specific+influence+on+optic+nerve+and+tectum+development+in+the+blind+cavefish+Astyanax.">The lens has a specific influence on optic nerve and tectum development in the blind cavefish Astyanax.</a> Developmental Neuroscience. 26 (5-6), 308-317 (2004).</li><li>White, E. 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Modulation of cell proliferation by retinal fiber input.</a> The Journal of Neuroscience. 3 (5), 1092-1099 (1983).</li><li>Sato, Y., Yano, H., Shimizu, Y., Tanaka, H., Ohshima, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optic+nerve+input-dependent+regulation+of+neural+stem+cell+proliferation+in+the+optic+tectum+of+adult+zebrafish.">Optic nerve input-dependent regulation of neural stem cell proliferation in the optic tectum of adult zebrafish.</a> Developmental Neurobiology. 77 (4), 474-482 (2017).</li><li>Hall, Z. 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J., Bracewell, T. G., Hawkins, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anatomical+dissection+of+zebrafish+brain+development.">Anatomical dissection of zebrafish brain development.</a> Brain Development. 1082, 197-214 (2014).</li><li>Brady, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple+technique+for+making+very+fine,+durable+dissecting+needles+by+sharpening+tungsten+wire+electrolytically.">A simple technique for making very fine, durable dissecting needles by sharpening tungsten wire electrolytically.</a> Bulletin of the World Health Organization. 32 (1), 143-144 (1965).</li><li>. ZFIN Tricaine recipe Available from: <a target="_blank" href="https://zfin.atlassian.net/wiki/spaces/prot/pages/362220023/TRICAINE">https://zfin.atlassian.net/wiki/spaces/prot/pages/362220023/TRICAINE</a> (2018)</li><li>Zhang, L., Leung, Y. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microdissection+of+zebrafish+embryonic+eye+tissues.">Microdissection of zebrafish embryonic eye tissues.</a> Journal of Visualized Experiments: JoVE. (40), e2028 (2010).</li><li>. ZFIN protocols Available from: <a target="_blank" href="https://zfin.atlassian.net/wiki/spaces/prot/overview">https://zfin.atlassian.net/wiki/spaces/prot/overview</a> (2021)</li><li>Engerer, P., Plucinska, G., Thong, R., Trov&#242;, L., Paquet, D., Godinho, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+subcellular+structures+in+the+living+zebrafish+embryo.">Imaging subcellular structures in the living zebrafish embryo.</a> Journal of Visualized Experiments: JoVE. (110), e53456 (2016).</li><li>ImageJ with batteries included. Fiji Available from: <a target="_blank" href="https://figi.sc/">https://figi.sc/</a> (2021)</li><li>O'Brien, J., Hayder, H., Peng, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+quantification+and+analysis+of+cell+counting+procedures+using+ImageJ+plugins.">Automated quantification and analysis of cell counting procedures using ImageJ plugins.</a> Journal of Visualized Experiments: JoVE. (117), e54719 (2016).</li><li>Poggi, L., Vitorino, M., Masai, I., Harris, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influences+on+neural+lineage+and+mode+of+division+in+the+zebrafish+retina+in+vivo.">Influences on neural lineage and mode of division in the zebrafish retina in vivo.</a> The Journal of Cell Biology. 171 (6), 991-999 (2005).</li><li>Karlstrom, R. O., 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=Zebrafish+mutations+affecting+retinotectal+axon+pathfinding.">Zebrafish mutations affecting retinotectal axon pathfinding.</a> Development. 123 (1), 427-438 (1996).</li><li>Harvey, B. M., Baxter, M., Granato, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optic+nerve+regeneration+in+larval+zebrafish+exhibits+spontaneous+capacity+for+retinotopic+but+not+tectum+specific+axon+targeting.">Optic nerve regeneration in larval zebrafish exhibits spontaneous capacity for retinotopic but not tectum specific axon targeting.</a> PLOS ONE. 14 (6), 0218667 (2019).</li><li>Robles, E., Filosa, A., Baier, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precise+lamination+of+retinal+axons+generates+multiple+parallel+input+pathways+in+the+tectum.">Precise lamination of retinal axons generates multiple parallel input pathways in the tectum.</a> Journal of Neuroscience. 33 (11), 5027-5039 (2013).</li><li>Vargas, M. E., Barres, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Why+Is+Wallerian+degeneration+in+the+CNS+so+slow.">Why Is Wallerian degeneration in the CNS so slow.</a> Annual Review of Neuroscience. 30 (1), 153-179 (2007).</li><li>Hughes, A. N., Appel, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microglia+phagocytose+myelin+sheaths+to+modify+developmental+myelination.">Microglia phagocytose myelin sheaths to modify developmental myelination.</a> Nature Neuroscience. 23 (9), 1055-1066 (2020).</li><li>de Calbiac, H., Dabacan, A., Muresan, R., Kabashi, E., Ciura, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behavioral+and+physiological+analysis+in+a+zebrafish+model+of+epilepsy.">Behavioral and physiological analysis in a zebrafish model of epilepsy.</a> Journal of Visualized Experiments: JoVE. (176), e58837 (2021).</li><li>Adams, S. L., Zhang, T., Rawson, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+external+medium+composition+on+membrane+water+permeability+of+zebrafish+(Danio+rerio)+embryos.">The effect of external medium composition on membrane water permeability of zebrafish (Danio rerio) embryos.</a> Theriogenology. 64 (7), 1591-1602 (2005).</li><li>Fredj, N. B., 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=Synaptic+activity+and+activity-dependent+competition+regulates+axon+arbor+maturation,+growth+arrest,+and+territory+in+the+retinotectal+projection.">Synaptic activity and activity-dependent competition regulates axon arbor maturation, growth arrest, and territory in the retinotectal projection.</a> Journal of Neuroscience. 30 (32), 10939-10951 (2010).</li><li>Alberio, L., 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=A+light-gated+potassium+channel+for+sustained+neuronal+inhibition.">A light-gated potassium channel for sustained neuronal inhibition.</a> Nature Methods. 15 (11), 969-976 (2018).</li><li>Kay, J. N., Finger-Baier, K. C., Roeser, T., Staub, W., Baier, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Retinal+ganglion+cell+genesis+requires+lakritz,+a+zebrafish+atonal+homolog.">Retinal ganglion cell genesis requires lakritz, a zebrafish atonal homolog.</a> Neuron. 30 (3), 725-736 (2001).</li><li>Gnuegge, L., Schmid, S., Neuhauss, S. C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+activity-deprived+zebrafish+mutant+macho+reveals+an+essential+requirement+of+neuronal+activity+for+the+development+of+a+fine-grained+visuotopic+map.">Analysis of the activity-deprived zebrafish mutant macho reveals an essential requirement of neuronal activity for the development of a fine-grained visuotopic map.</a> The Journal of Neuroscience. 21 (10), 3542-3548 (2001).</li><li>Jeffery, W. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astyanax+surface+and+cave+fish+morphs.">Astyanax surface and cave fish morphs.</a> EvoDevo. 11 (1), 14 (2020).</li><li>Sieger, D., Peri, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+for+studying+microglia:+The+first,+the+popular,+and+the+new.">Animal models for studying microglia: The first, the popular, and the new.</a> Glia. 61 (1), 3-9 (2013).</li><li>Svahn, A. 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The techniques described in this paper illustrate one of many approaches for studying vertebrate visual system development in zebrafish. Other researchers have published methods to dissect the embryonic retina and perform gene expression analyses19 or visualize neuronal activity in the optic tectum30. This paper provides an approach for exploring how differential retinal input may influence cell behaviors in the optic tectum.
To ensure successful...

The goal of this method is to investigate how retinal input influences the growth and development of the optic tectum, the visual processing center in the zebrafish brain. By removing one eye and then comparing the two sides of the optic tectum, tectal changes within the same specimen can be observed and normalized, enabling comparison across multiple specimens. Modern molecular approaches combined with this technique will yield insights into the mechanisms underlying visual system growth and development, as well as axonal degeneration and regeneration.
Sensory systems - visual, auditory, and somatosensory - gather information from external organs and relay that information to the central nervous system, generating &#34;maps&#34; of the external world across the midbrain1,2. Vision is the dominant sensory modality for nearly all vertebrates, including many fishes. The retina, the neural tissue in the eye, gathers information with a neuronal circuit consisting primarily of photoreceptors, bipolar cells, and retinal ganglion cells (RGCs), the projection neurons of the retina. RGCs have long axons that find their way across the inner surface of the retina to the optic nerve head, where they fasciculate and travel together through the brain, ultimately terminating in the visual processing center in the dorsal midbrain. This structure is called the optic tectum in fish and other non-mammalian vertebrates and is homologous to the superior colliculus in mammals3.
The optic tectum is a bilaterally symmetric multilayered structure in the dorsal midbrain. In zebrafish and most other fishes, each lobe of the optic tectum receives visual input solely from the contralateral eye, such that the left optic nerve terminates in the right tectal lobe and the right optic nerve terminates in the left tectal lobe4 (Figure 1). Like its mammalian counterpart, the superior colliculus, the optic tectum integrates visual information with other sensory inputs, including audition and somatosensation, controlling shifts in visual attention and eye movements such as saccades1,5,6. However, unlike the mammalian superior colliculus, the optic tectum continuously generates new neurons and glia from a specialized stem cell niche near the medial and caudal edges of the tectal lobes called the tectal proliferation zone7. Maintenance of proliferative progenitors in the optic tectum and other regions of the central nervous system contributes, in part, to the remarkable regenerative capacity documented in zebrafish8.
Previous work examining the brains of blind or one-eyed fishes revealed that optic tectum size is directly proportional to the amount of retinal innervation it receives9,10,11. In adult cave fish, whose eyes degenerate in early embryogenesis, the optic tectum is noticeably smaller than that of closely related, sighted surface fish9. Cave fish eye degeneration can be blocked by replacing the endogenous lens with a lens from a surface fish during embryogenesis. When these one-eyed cave fish are reared to adulthood, the innervated tectal lobe contains approximately 10% more cells than the non-innervated tectal lobe9. Similarly, in larval killifish that were incubated with chemical treatments to generate eyes of different sizes within the same individual, the side of the tectum with more innervation was larger and contained more neurons10. Evidence from optic nerve crush experiments in adult goldfish indicates that innervation promotes proliferation, with tectal cell proliferation decreasing when innervation was disrupted11.
Confirming and extending these classical studies, several recent reports provide data suggesting that proliferation in response to innervation is modulated, at least in part, by the BDNF-TrkB pathway12,13. Many open questions about optic tectum growth and development remain, including how a developing sensory system copes with injury and axon degeneration, which cellular and molecular signals enable retinal input to regulate optic tectum growth, when these mechanisms become active, and whether innervation-linked proliferation and differentiation enable the retina and its target tissue to coordinate growth rates and ensure accurate retinotopic mapping. In addition, there are much larger questions about activity-dependent development that can be addressed by interrogating the zebrafish visual system with surgical approaches such as the one described below.
To investigate the cellular and molecular mechanisms by which neural activity, specifically from visual input, alters cell survival and proliferation, the described approach directly compares innervated and denervated tectal lobes (Figure 1) within individual zebrafish larvae. This method allows for the documentation of RGC axon degeneration in the optic tectum and confirmation that the number of mitotic cells correlates with innervation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63509/63509fig01.jpg" /> 
Figure 1: Sketches of zebrafish larvae before and after unilateral eye removal. (A) Drawing of 5 dpf larvae as viewed under a dissecting microscope. Each larva is embedded in low-melting-point agarose and oriented laterally before a tungsten needle with a sharp, hooked tip is used to scoop out the eye facing up (left eye in this example). (B) Drawing of the dorsal view of a 9 dpf larva resulting from the surgery depicted in A. Only three highly schematized RGC axons from the right eye are shown defasciculating and connecting with neurons in the left tectal lobe. Abbreviations: dpf = days post fertilization; dps = days post surgery; RGC = retinal ganglion cells. <a href="https://www.jove.com/files/ftp_upload/63509/63509fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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			<td>Equipment and supplies:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Breeding boxes</td>
			<td>Aquaneering</td>
			<td>ZHCT100</td>
			<td></td>
		</tr>
		<tr>
			<td>Dow Corning high vacuum grease</td>
			<td>Sigma or equivalent supplier</td>
			<td>Z273554</td>
			<td></td>
		</tr>
		<tr>
			<td>Erlenmeyer flasks (125 mL)</td>
			<td></td>
			<td></td>
			<td>For making Marc&#39;s Modified Ringers (MMR) with antibiotics for post-surgery incubation</td>
		</tr>
		<tr>
			<td>Fine forceps - Dumont #5</td>
			<td>Fine Science Tools (FST)</td>
			<td>11252-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Pasteur pipettes</td>
			<td>DWK Lifescience</td>
			<td>63A53 &#38; 63A53WT</td>
			<td>For pipetting embryos and larvae</td>
		</tr>
		<tr>
			<td>Glass slides for microscopy</td>
			<td>VWR or equivalent supplier</td>
			<td>48311-703</td>
			<td>Standard glass microscope slides can be ordered from many different laboratory suppliers.</td>
		</tr>
		<tr>
			<td>Glassware including graduated bottles and graduated cylinders</td>
			<td></td>
			<td></td>
			<td>For making and storing solutions</td>
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			<td>2-part epoxy resin</td>
			<td>ACE Hardware or other equivalent supplier of Gorilla Glue or equivalent</td>
			<td>0.85 oz syringe</td>
			<td>https://www.acehardware.com/departments/paint-and-supplies/tape-glues-and-adhesives/glues-and-epoxy/1590793</td>
		</tr>
		<tr>
			<td>Microcentrifuge tube (1.7 mL)</td>
			<td>VWR or equivalent supplier</td>
			<td>22234-046</td>
			<td></td>
		</tr>
		<tr>
			<td>Nickel plated pin holder (17 cm length)</td>
			<td>Fine Science Tools (FST)</td>
			<td>26018-17</td>
			<td>To hold tungsten wire while sharpening and performing surgeries/dissections.</td>
		</tr>
		<tr>
			<td>Nylon mesh tea strainer or equivalent</td>
			<td>Ali Express or equivalent</td>
			<td></td>
			<td>For harvesting zebrafish eggs after spawning; https://www.aliexpress.com/item/1005002219569756.html</td>
		</tr>
		<tr>
			<td>Paper clip</td>
			<td></td>
			<td></td>
			<td>For Tungsten needle sharpening device.</td>
		</tr>
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			<td>Petri dishes 100 mm</td>
			<td>Fischer Scientific or equivalent supplier</td>
			<td>50-190-0267</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes 35 mm</td>
			<td>Fischer Scientific or equivalent supplier</td>
			<td>08-757-100A</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette pump</td>
			<td>SP Bel-Art or equivalent</td>
			<td>F37898-0000</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium hydroxide (KOH)</td>
			<td>Sigma</td>
			<td>909122</td>
			<td>For Tungsten needle sharpening device. Make a 10% w/v solution of KOH in the hood by adding pellets to deionized water.</td>
		</tr>
		<tr>
			<td>Power supply (variable voltage)</td>
			<td></td>
			<td></td>
			<td>For Tungsten needle sharpening device. Any power supply with variable voltage will work (even one used for gel electrophoresis).</td>
		</tr>
		<tr>
			<td>Sylgard 184 Elastomer kit</td>
			<td>Dow Corning</td>
			<td>3097358</td>
			<td></td>
		</tr>
		<tr>
			<td>Tungsten wire (0.125 mm diameter)</td>
			<td>World Precision Instruments (WPI)</td>
			<td>TGW0515</td>
			<td>Sharpen to remove eye and dissect larvae.</td>
		</tr>
		<tr>
			<td>Variable temperature heat block</td>
			<td>The Lab Depot or equivalent supplier</td>
			<td>BSH1001 or BSH1002</td>
			<td>Set to 40-42 &#176;C ahead of experiments.</td>
		</tr>
		<tr>
			<td>Wide-mouth glass jar with lid (e.g., clean jam or salsa jar)</td>
			<td></td>
			<td></td>
			<td>For Tungsten needle sharpening device.</td>
		</tr>
		<tr>
			<td>Wires with alligator clip leads</td>
			<td></td>
			<td></td>
			<td>For Tungsten needle sharpening device.</td>
		</tr>
		<tr>
			<td>Microscopes:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td></td>
			<td></td>
			<td>Any type will work but having adjustable transmitted light on a mirrored base is preferred.</td>
		</tr>
		<tr>
			<td>Laser scanning confocal microscope</td>
			<td></td>
			<td></td>
			<td>High NA, 20-25x water dipping objective lens is recommended. 
			Microscope control and image capture software (Elements) is used here but any confocal microscope will work.</td>
		</tr>
		<tr>
			<td>Reagents for surgeries and dissections:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma</td>
			<td>C7902</td>
			<td>For Marc&#39;s Modified Ringers (MMR) and embryo medium (E3).</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H7006</td>
			<td>For Marc&#39;s Modified Ringers (MMR).</td>
		</tr>
		<tr>
			<td>Low melting point agarose</td>
			<td>Invitrogen</td>
			<td>16520-050</td>
			<td>Make 1% in embryo medium (E3) or Marc&#39;s Modified Ringers (MMR).</td>
		</tr>
		<tr>
			<td>Magnesium chloride hexahydrate</td>
			<td>Sigma</td>
			<td>1374248</td>
			<td>For embryo medium (E3).</td>
		</tr>
		<tr>
			<td>Magnesium sulfate</td>
			<td>Sigma</td>
			<td>M7506</td>
			<td>For Marc&#39;s Modified Ringers (MMR).</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>19210</td>
			<td>Dilute 8% (w/v) stock with 2x concentrated PBS (diluted from 10x PBS stock).</td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin</td>
			<td>Sigma</td>
			<td>P4333-20ML</td>
			<td>Dilute 1:100 in Marc&#39;s Modified Ringers.</td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS) tablets</td>
			<td>Diagnostic BioSystems</td>
			<td>DMR E404-01</td>
			<td>Make 10x stock in deionized water, autoclave and store at room temperature. Dilute to 1x working concentration.</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma</td>
			<td>P3911</td>
			<td>For Marc&#39;s Modified Ringers (MMR) and embryo medium (E3).</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma</td>
			<td>S9888</td>
			<td>For Marc&#39;s Modified Ringers (MMR) and embryo medium (E3).</td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>Sigma</td>
			<td>S5881</td>
			<td>Make 10 M and use to adjust pH of MMR to 7.4.</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma</td>
			<td>S9378</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine-S</td>
			<td>Pentair</td>
			<td>100G #TRS1</td>
			<td>Recipe: https://zfin.atlassian.net/wiki/spaces/prot/pages/362220023/TRICAINE</td>
		</tr>
		<tr>
			<td>Reagents for immunohistochemistry:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Alexafluor 568 tagged Secondary antibody to detect rabbit IgG</td>
			<td>Invitrogen</td>
			<td>A-11011</td>
			<td>Use at 1:500 dilution for wholemount immunohistochemistry.</td>
		</tr>
		<tr>
			<td>DAPI or ToPro3</td>
			<td>Invitrogen</td>
			<td>1306 or T3605</td>
			<td>Make up 1 mg/mL solutions in DMSO; 1:5,000 dilution for counterstaining.</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO)</td>
			<td>Sigma</td>
			<td>D8418</td>
			<td>A component of immunoblock buffer.</td>
		</tr>
		<tr>
			<td>Methanol (MeOH)</td>
			<td>Sigma</td>
			<td>34860</td>
			<td>Mixing MeOH with aqueous solutions like PBST is exothermic. Make the MeOH/PBST solutions at least several hours ahead of time or cool them on ice before using.</td>
		</tr>
		<tr>
			<td>Normal goat serum</td>
			<td>ThermoFisher Scientific</td>
			<td>50-062Z</td>
			<td>A component of immunoblock buffer. Can be aliquoted in 1-10 mL volumes and stored at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>Primary antibody to detect phosphohistone H3</td>
			<td>Millipore</td>
			<td>06-570</td>
			<td>Use at 1:300 dilution for wholemount immunohistochemistry.</td>
		</tr>
		<tr>
			<td>Primary antibody to detect Red Fluorescent Protein (RFP; detects dsRed derivatives)</td>
			<td>MBL International</td>
			<td>PM005</td>
			<td>Use at 1:500 dilution for wholemount immunohistochemistry.</td>
		</tr>
		<tr>
			<td>Proteinase K (PK)</td>
			<td>Sigma</td>
			<td>P2308-10MG</td>
			<td>Make up 10 mg/mL stock solutions in PBS and use at 10 &#181;g/mL.</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma</td>
			<td>T8787</td>
			<td>Useful to make a 20% (v/v) stock solution in PBS.</td>
		</tr>
		<tr>
			<td>Software for data analysis</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ (Fiji)</td>
			<td></td>
			<td></td>
			<td>freeware for image analysis; https://imagej.net/software/fiji/</td>
		</tr>
		<tr>
			<td>Rstudio</td>
			<td></td>
			<td></td>
			<td>freeware for statistical analysis and data visualization; https://www.rstudio.com/products/rstudio/download/</td>
		</tr>
		<tr>
			<td>Adobe Photoshop or GIMP</td>
			<td></td>
			<td></td>
			<td>Proprietary image processing software (Adobe Photoshop and Illustrator) are often used to compose figures). A freeware alternative is Gnu Image Manipulation Program (GIMP; https://www.gimp.org/)</td>
		</tr>
		<tr>
			<td>Zebrafish strains</td>
			<td></td>
			<td></td>
			<td>available from the&#160; Zebrafish International Resource Centers in the US (https://zebrafish.org/home/guide.php) or in Europe (https://www.ezrc.kit.edu/). Specialized transgenic strains that have not yet been deposited in either resource center can be requested from individual labs after publication.</td>
		</tr>
	</tbody>
</table>

eye removal in living zebrafish larvae to examine innervation-dependent growth and development of the visual system

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis support continuous growth of the entire visual system, both in the eye and the brain. Coordinated growth between the retinae and the optic tectum ensures accurate retinotopic mapping as new neurons are added in the eyes and brain. To address whether retinal axons provide crucial information for regulating tectal stem and progenitor cell behaviors such as survival, proliferation, and/or differentiation, it is necessary to be able to compare innervated and denervated tectal lobes within the same animal and across animals. 
Surgical removal of one eye from living larval zebrafish followed by observation of the optic tectum achieves this goal. The accompanying video demonstrates how to anesthetize larvae, electrolytically sharpen tungsten needles, and use them to remove one eye. It next shows how to dissect brains from fixed zebrafish larvae. Finally, the video provides an overview of the protocol for immunohistochemistry and a demonstration of how to mount stained embryos in low-melting-point agarose for microscopy.

To confirm whether eye removal was complete and assess how the optic tectum changes, surgeries were performed in the Tg[atoh7:RFP] strain, which labels all RGCs with a membrane-targeted RFP and, thus, all axons that project from the retina and form the optic nerve24. Although using this strain is not absolutely necessary, it enables straightforward observation and visualization of the optic nerve termini in the optic tectum neuropil. Other approaches for labeling the optic nerve, such as ...

Watch this Scientific Journal Video about Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System at JoVE.com

Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

The article explains how to surgically remove eyes from living zebrafish larvae as the first step toward investigating how retinal input influences optic tectum growth and development. In addition, the article provides information about larval anesthetization, fixation, and brain dissection, followed by immunohistochemistry and confocal imaging.

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis ...

So my lab is interested in understanding how the eye and brain grow together. This technique allows us to study how retinal input influences brain growth and development. Surgical removal of one eye from the living larval zebrafish, followed by observation of the optic tectum allows us to compare innervated and denervated tectal lobes within the same animal and across animals.Combining this technique with modern molecular approaches can give new insight into mechanisms underlying neural development, regeneration, and degeneration. To begin, connect the cathode and anode wires to the power supply. Attach the alligator clip at the end of the cathode wire to a partially straighten paperclip and insert the paperclip into the potassium hydroxide solution, attaching it to the side of the jar.Attach the alligator clip of the anode wire to the neck of the needle holder. Turn the power to approximately 20 volts and dip the tungsten wire into the potassium hydroxide solution, pulling the wire out of the solution at an angle to electrolytically sharpen the wire into a fine tip. Check the tip of the needle under the dissecting microscope to ensure that it is sharp enough.The curve needle tip results when the sharp needle is bent by gently touching it to the Petri dish. On the day of surgery, use a wide-bore glass pasture pipette to transfer 10 to 15 larvae to a 35 millimeter Petri dish filled with fresh E3.Add three to five drops of anesthetic to the larva. To determine if the larvae are adequately anesthetized, look for lack of touch response.Add two to three more drops of the anesthetic if the larvae are still responsive to touch after three minutes and reassess. To immobilize the larva for surgery, take one larva up into a narrow-bore glass pasture pipette with only a small amount of E3.Next, take up approximately 200 microliters of melted 1%low melting point or LMP agarose into the pipette containing the larva and mix to ensure that the larva is suspended in the LMP agarose. Place the lid of a 35 millimeter Petri dish faced up under the dissecting microscope, then squirt the larva and the agarose onto the upside down Petri dish lid.Spread the agarose so that the larva is in the center of the drop. Using a dull tungsten needle, quickly but gently maneuver the larva so that it is lateral with one eye facing upwards. Wait several minutes for the agarose to set.Following the edge of the eye orbit, use the tip of a fine electrolytically sharpened tungsten needle to pierce the skin around the eye. Then slide the edge of the needle under the eye from the temporal-ventral side of the eye. Use controlled pressure to release the eye from the socket.Keep pressing the eye dorsally and anteriorly with the side of the needle, eventually slicing through the optic nerve and releasing the eye. Alternatively, you can use very fine surgical forceps to remove the eye by pinching the optic nerve and pushing the eye from medial to lateral. After successful eye removal, cover the agarose with MMR solution, liberate each larva from the agarose by gently brushing a tungsten needle around their head and then around their body, while stabilizing the Petri dish lid with forceps.After the surgeries, place the larva in MMR solution supplemented with antibiotics until the following day, then return the larva to E3 and rear them until the endpoint of the experiment. After larva are terminally anesthetized, fix them with 4%paraformaldehyde. Suspend the fixed larvae in PBS droplets on a sealguard plate under a dissecting microscope.Secure them laterally by placing two tungsten pins through the notochord with one pin posterior to the pigmented area covering the AGM region and another in line with the end of the yolk extension. Remove the eye with a sharp tungsten needle and fine forceps. Sometimes it is possible to poke the needle through the jaw to the opposite side and remove the other eye.Use the tungsten needle to scratch from the temporal to the ventral side of the ear. In the same action in motion, bring the needle posterior to the jaw and gently pull anteriorly until the ear and jaw are removed. Remove the other eye and ear by poking the needle through the jaw.Use forceps to pull out the ventral organs and remaining yolk. Finally, make a shallow incision in the dorsal cranial skin near the junction between the hind brain and spinal cord. Lift the skin with the forceps and pull it anteriorly and around the telencephalon.Remove any remaining tissue with forceps. Unpin the larva and transfer to PBS into 1.5 milliliter microcentrifuge tube. Secure a chambered slide into the lid of a 100 millimeter Petri dish with vacuum grease.Transfer the immunostained and processed larva to a well plate or depression slide to view them with a dissecting microscope. Mount the larva on the chambered slide in 1%LMP agarose columns. Using a glass past your pipette, put one larva on the chambered slide, depositing as little PBS as possible.Cover the larva with melted and warm 1%LMP agarose using the same pipette. Pipette the LMP agarose into a column and then position the larva as symmetrically as possible with the dorsal surface visible. After eye removal, progressive degeneration of retinal axons was observed in the optic tectum neuropil.By two days postsurgery, RFP labeled axons exhibited hallmarks of rapid Wallerian degeneration, such as blabbing and fragmentation. RFP positive puncta were noted both within and outside the neuropil on the right side of the tectum as indicated by the arrows. By four days post-surgery, fragmented axons and RFP labeled axonal debris were considerably reduced in the right tectal lobe, indicating relatively rapid clearance of the dying and degenerating axons.The brain was exposed by dissecting the eyes, jaw, ears, and skullcap of the skin and connective tissues. Ideally, the brain remains intact during this procedure. However, parts of the fore brain, particularly the olfactory bulb, were likely damaged or completely removed when the skull cap of the skin or jaw was removed.Moreover, sometimes the lateral ledge of the tectum gets sliced when piercing or pinching the skin to pull it off the brain. No two samples are identical. So you need to be able to make subtle adaptations through the eye removal and brain dissection as needed.Patients and sharp tools are essential for success. This technique can be followed up with cellular and molecular approaches, like in vivo live cell imaging or RNA sequencing to gain new insights into the molecular and cellular mechanisms of optic tectum growth and development.

eye-removal-living-zebrafish-larvae-to-examine-innervation-dependent

Research

JoVE Journal

Developmental Biology

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 ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

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 ...

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to ...

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 ...

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in zebrafish during different developmental stages. To investigate the role of endoglin(eng) in vessel formation during the development of hereditary hemorrhagic telangiectasia&#160;(HHT), morpholino-mediated targeted knockdown of eng in zebrafish are used to study its temporal expression and associated functions. Here, whole-mount in situ RNA hybridization (WISH) is employed for the analysis of eng and its downstream genes in zebrafish embryos. Also, tube formation assays are performed in HHT patient-derived induced pluripotent stem cell-differentiated&#160;endothelial cells (iPSC-ECs; with eng mutations). A specific signal amplifying system using the whole amount In Situ Hybridization &#8211; WISH provides higher resolution and lower background results compared to traditional methods. To obtain a better signal, the post-fixation time is adjusted to 30 min after probe hybridization. Because fluorescence staining is not sensitive in zebrafish embryos, it is replaced with diaminobezidine (DAB) staining here. In this protocol, HHT&#160;patient-derived iPSC lines containing an eng mutation are differentiated into endothelial cells. After coating a plate with basement membrane matrix for 30 min at 37 &#176;C, iPSC-ECs are seeded as a monolayer into wells and kept at 37 &#176;C for 3 h. Then, the tube length and number of branches are calculated using microscopic images. Thus, with this improved WISH protocol, it is shown that reduced eng expression affects endothelial progenitor formation in zebrafish embryos. This is further confirmed by tube formation assays using iPSC-ECs derived from a patient with HHT. These assays confirm the role for eng in early vascular development.

Presented here is a protocol for whole-mount in situ RNA hybridization analysis in zebrafish and tube formation assay in patient-derived induced pluripotent stem cell-derived endothelial cells to study the role of endoglin in vascular formation.

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...