The authors would like to thank Katrin Bennewitz and Marlene Hausner for zebrafish husbandry and technical assistance. The authors acknowledge the support of the Core Facility Live Cell Imaging Mannheim at the Centre for Biomedicine and Medical Technology Mannheim (DFG INST 91027/9-1). This study was supported by Deutsche Forschungsgemeinschaft (International Research Training Group 1874/1 &#34;DIAMICOM&#34;, project SP5 and SP9; Collaborative Research Centre SFB1118, project B1 and Collaborative Research Centre SFB/TR23 project Z5).

All procedures, including steps related to animal subjects, have been approved by the Animals Ethics Committee (Regierungspr&#228;sidium Karlsruhe) and follow the animal care guidelines of Heidelberg University.
1. Preparation of Fixative (4% PFA/PBS)
<ol>
	<li>Prepare fixative fresh every day to ensure optimal preservation of histological tissue integrity.</li>
	<li>Dissolve 0.2 g of sodium hydroxide (NaOH) in 90 mL double distilled water (ddH2O) while constantly stirring at room temperature.</li>
	<li>Add 4 g paraformaldehyde (PFA) and stir until the solution is completely clear for at least 5-10 min to assure depolymerization of macromolecules. 
	Caution: PFA is toxic, handle with care.</li>
	<li>Dissolve 0.84 g of sodium dihydrogen phosphate (NaH2PO4) in the solution and control pH at 7.2 via measurement.</li>
	<li>Adjust volume to 100 mL with ddH2O and filter the solution through grade 3 filter paper.</li>
	<li>Store 4% PFA/PBS on ice.</li>
</ol>
2. Preparation of Euthanasia Solution
NOTE: The following steps were done with ABTL wild-type zebrafish strain at a general age of 6-8 mon.
<ol>
	<li>Use ethyl 3-aminobenzoate methane sulfonate, also named &#34;tricaine&#34; or MS-222, in a concentration of 0.31 mg/mL for zebrafish euthanasia16. Utilize 1x egg water, used to raise zebrafish embryos, as solvent for the euthanasia solution. Generally, control the correct anesthesia depth before working with sedated fish by demonstrating loss of equilibrium and touch reaction loss.
	<ol>
		<li>To attain 1,000 mL of 10x egg water, add these salts to 1,000 mL double distilled water (ddH2O) while constantly stirring in the following order: 10 g NaCl, 0.3 g KCl, 0.4 g CaCl2*6H2O, 1.32 g MgSO4*6H2O.</li>
	</ol>
	</li>
</ol>
3. Fixation of Zebrafish Tissue
<ol>
	<li>Transfer 6 mL 4% PFA/PBS solution per sample into the wells of six-well plates in advance.</li>
	<li>Euthanize the adult zebrafish (previously maintained on a standard light-day cycle, e.g., 12 h: 12 h) 2 h after lights turn on in the morning, by placing them in the tricaine solution and wait until they have reached euthanasia. Following the cessation of opercular movement, which can take up to 10 min.</li>
	<li>Take the fish out of the tricaine euthanasia solution and put them on fresh paper towels. Dry the fish and use a scalpel to cut the heads behind the operculum (see Figure 2A). Transfer the heads directly into the prepped well plates, which contain the freshly prepared fixative.</li>
	<li>Store the well plates containing the fish heads at 4 &#176;C overnight for at least 24 h to assure the fixative penetrates the deeper retinal layers. 
	NOTE: Zebrafish heads can be stored for a maximum of 48 h in 1x phosphate buffered saline (PBS) at 4 &#176;C before preparation without a relevant loss of fluorescence signal strength. Increased latency can lead to loss of tissue integrity and reduced picture quality, and thus should be avoided.</li>
</ol>
4. Preparation of the Zebrafish Retinal Vasculature
<ol>
	<li>Fill a petri dish up to a third with heated 2% agarose and wait until the agar is firm. Cover the agar plate with 1x PBS to create a workspace to dissect the eyes. 
	NOTE: This will allow both preservation of preparation tweezers and low pressure on the tissue while dissecting, reducing the likelihood of structural damage. All further preparation steps should be performed in this workspace utilizing #5 straight forceps with fine tips under a dissecting microscope with additional epi-illumination.
	<ol>
		<li>While dissecting, use magnification between 4.0x and 6.0x to allow for both overview and detail-oriented assessment of the tissue.</li>
	</ol>
	</li>
	<li>Transfer the fixed sample into the petri dish and, by holding the head at the cut surface with one tweezer, insert another pinned-together tweezer below the eyeball at the orbital cavity. Slowly open the tweezer below the eye and grip the optic nerve, then carefully tear and detach the eyes (Figure 2B).</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56674/56674fig2.jpg" /> 
Figure 2: Removal of the adult zebrafish eye. Corneal side view with the eye still in the orbital cavity and intact optic nerve (A). Corneal side view with detached eye (B). Schematic depiction of the zebrafish eye at this step (C). <a href="//cloudfront.jove.com/files/ftp_upload/56674/56674fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>Remove any of the four rectus and two oblique extraocular muscles that are still connected to the eye as well as residual extraocular tissue connecting the eye to the orbital cavity (compare Figure 3 and Figure 4).
	<ol>
		<li>Achieve this by holding back the eye through a half-closed tweezer and, gripping the structure with the other tweezer, softly rip it off with a diametric movement. Since gripping the eyeball directly leads to internal vessel breakage, this technique is appropriately gentle to preserve the vasculature.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56674/56674fig3.jpg" /> 
Figure 3: Adult zebrafish eye with extraocular muscles in focus. Lateral view with attachment of an extraocular muscle to the left outer rim of the ocular globe in focus (A). Optic nerve side view with extraocular muscles symmetrically flanking the optic nerve (B). Schematic depiction of the zebrafish eye at this step (C). <a href="//cloudfront.jove.com/files/ftp_upload/56674/56674fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="4">
	<li>Use a 27G (0.4 x 19 mm) disposable needle to puncture the cornea (Figure 4C, red arrow) at the outer range. Through this opening hold the cornea with both tweezers and slightly tear it open. Afterwards carefully work to create a centered tear approximately the size of the respective pupil.</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56674/56674fig4.jpg" /> 
Figure 4: Adult zebrafish eye after removal of all extraocular muscles. Lateral view with the cleared outer rim of the ocular globe visible (A). Optic nerve side view with optic nerve in the middle. The light-reflecting sclera is not covering the whole area around the nerve (red dashed line) (B). Schematic depiction of the zebrafish eye at this step (C). <a href="//cloudfront.jove.com/files/ftp_upload/56674/56674fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="5">
	<li>Apply pressure on the corneal side of the ocular globe (bulbus oculi) at the outer corneal edge above the iris. This will create a small dent and push the lens to the height of the corneal tear. Run the tweezers under the lens and remove it (Figure 5A).</li>
</ol>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56674/56674fig5.jpg" /> 
Figure 5: Adult zebrafish eye in the process of lens removal. Corneal side view with the lens pushed through the corneal tear (A). Corneal side view with the lens beside the ocular globe (B). Schematic depiction of the zebrafish eye at this step (C). <a href="//cloudfront.jove.com/files/ftp_upload/56674/56674fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="6">
	<li>Turn the eye upside down to the optic nerve facing the researcher. Note that the sclera and cornea are connected and form the fibrous tunic of the eye bulb to protect the cup-shaped retina. (Figure 6).
	<ol>
		<li>This shell, consisting of the cornea and sclera, is also called &#34;corneosclera&#34; and spares out the area around the optic nerve (Figure 4B, red dashed line). Insert a needle at this discontinuation once to create an opening between the sclera and the retina.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56674/56674fig6.jpg" /> 
Figure 6: Corneosclera consisting of sclera and cornea, which is disconnected from the remaining intraocular tissue. Corneal side view showing the continuation of the translucent cornea into the pigmented sclera (A). Lateral view focused on the scleral part of the corneosclera (B). Schematic depiction of the removed corneosclera (C). <a href="//cloudfront.jove.com/files/ftp_upload/56674/56674fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="7">
	<li>Using this access with both tweezers, carefully rip the sclera axially into stripes to increase the opening around the optic nerve. Be careful to keep the corneosclera intact at the transition to the corneal side circumference (see Figure 6A).</li>
	<li>Hold the sclera with one tweezer and, by grabbing the optic nerve with the other and pulling away, wholly remove the corneosclera from the eye and discard it (Figure 6). Try to sever any connections before separating the corneosclera and the remaining intraocular tissue, as attachment to other structures will be a critical point; this step will provide a cup-shaped structure consisting of the uvea and retina containing the retinal vasculature (Figure 7).</li>
	<li>Create a rupture in the choroidal/RPE layer by holding a 27G needle sideways to the remaining cup while scraping the outer surface with the rim of the needle tip. Use the rupture as an access point to get a grip on the choroidal/RPE layer and rip it with both tweezers into stripes, but keep the connection to the iris intact.
	<ol>
		<li>Afterwards, run one tweezer under the iris and go around in a circuit from the outside while creating tension by pulling at the discontinued choroidal/RPE layer to achieve detachment of the combined structure. 
		NOTE: The iris must be disconnected, so as to not obstruct the fluorescence while visualizing the vessels (Figure 8B). Grabbing the iris directly to remove it can lead to extensive vessel damage, especially at the inner optic circle (IOC), since the iris is still connected to the surrounding tissue. The choroidal layer and the retinal pigmented epithelium (RPE) can be separated and often retain a connection to the iris, which allows for an easy secondary removal.</li>
		<li>If parts of the iris cannot be removed, utilize a natural breaking point the adult zebrafish eye exhibits inside the photoreceptor layer (PL), in a similar fashion to step 4.9, to remove the iris.</li>
		<li>Scrape over the outside of the remaining cup-shaped retina to induce discontinuations in the PL above the breaking point (Figure 11C, black arrow). Use the created access to remove the upper part of the layer while keeping the possible connection to the iris intact. Afterwards, run the tweezers under the iris and go around in a circuit as mentioned before to detach the combined structure. 
		NOTE: One should exercise patience as the whole step 4.9 can be difficult, but this care will achieve a better vascular outcome than straightforwardly grabbing the iris at the rim of the pupil or forcing an opening by destroying connections to the choroidal/RPE or photoreceptor layer without their respective removal. Following this step will thus preserve retinal vascular integrity, but lead to damage of the outer retinal layers.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/56674/56674fig7.jpg" /> 
Figure 7: Adult zebrafish eye after removal of the corneosclera. Lateral view shows the remaining intraocular tissue with intact iris and optic nerve (A). Corneal side view with iris in focus (B). Schematic depiction of the zebrafish eye at this step (C). <a href="//cloudfront.jove.com/files/ftp_upload/56674/56674fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="10">
	<li>Use a microdissection spring scissor with a straight 2.5 mm cutting edge to cut the optic nerve as close as possible to the retina; this will allow for a better flat-mounting of the tissue.</li>
</ol>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/56674/56674fig8.jpg" /> 
Figure 8: Adult zebrafish eye after removal of RPE/choroid and truncated optic nerve. Optic nerve side view shows the retina with a truncated optic nerve (A). Corneal side view with a direct look onto the most inner layer of the retina (B). Schematic depiction of the zebrafish eye at this step (C). <a href="//cloudfront.jove.com/files/ftp_upload/56674/56674fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Mounting of the Retinal Vasculature
<ol>
	<li>Wash the dissected retina two times for 5 minutes in 1x PBS. To transfer the retina after truncation of the optic nerve, utilize a lab spatula with a micro spoon end to avoid direct handling of the exposed tissue.</li>
	<li>Place a drop of PBS onto a glass slide and transfer the retina into the droplet. Use a tweezer to keep the tissue in place while cutting the cup-shaped structure with a scalpel to create a flat four-petal or five-petal shape depending on the retinal size (see Figure 9).</li>
	<li>Suck up the leftover PBS with a fine piece of paper. Be careful not to touch the retina.</li>
	<li>Coat the flat-mounted retina in mounting media and cover with a coverslip. Be attentive not to create foam, as air bubbles can distort visualization of the retinal vessels.</li>
	<li>Minimize movement of the coverslip and seal the cover with clear nail polish.</li>
</ol>
6. Visualization of the Retinal Vasculature
<ol>
	<li>Keep the time gap between preparation and visualization as short as possible to reduce image detail loss through fluorescence signal loss. Store the prepared retinal vasculature mounts at 4 &#176;C to reduce signal loss, if direct visualization cannot be achieved, as degradation of image quality slowly becomes visible 48 h after preparation.</li>
	<li>Visualize the retinal vasculature mounts via fluorescence microscopy or laser scanning confocal microscopy. 
	NOTE: Vessel visualization by fluorescence microscopy was achieved at 2.5x magnification with an exposure time of 2.0 s, gain of 6.0x and gamma settings of 1.67. For confocal microscopy, an Ar-laser (488 nm/20 mW) at 20% power was utilized in combination with a TD 488/543/633 excitation filter, a 20x/0.7 NA multi-immersion objective with ddH2O as immersion media, and emission filter settings of 505-560 nm. Confocal pictures are composed of 4x4, 4x5, or 5x5 singular images combined through a tile scan depending on the retinal size.</li>
</ol>
7. HE Sections of the Zebrafish Retina
NOTE: For HE sections of the zebrafish retina, prepare the tissue as described in the protocol until step 4.5 and afterwards skip directly to step 7.1.
<ol>
	<li>Wash the enucleated eyes (after step 4.5) two times for 5 minutes in 1x PBS. Afterwards, transfer the tissue into 70% ethanol (EtOH). At this point, prepared eyes can be stored at 4 &#176;C until paraffin embedding.</li>
	<li>Wash the tissue the following way to dehydrate it before paraffin embedding: 2x15 min in 80% ethanol, 2x15 min in 90% ethanol, 3x15 min in 96% ethanol, 3x15 min in 99% ethanol, 2x15 min in acetone/99% ethanol (1:2), 3x15 min in acetone. Keep the tissue in paraffin at 62 &#176;C overnight.</li>
	<li>Heat up fresh paraffin to 62 &#176;C and pour it into embedding molds. Transfer the tissue into the molds directly and, at a fast pace, control correct orientation of the eyes for the desired section. Afterwards, apply an embedding cassette to the mold and cover with additional heated paraffin.</li>
	<li>Remove the embedding molds after paraffin blocks have cooled down. Cut the paraffin blocks at a microtome into 10 &#181;m sections and float them out on a 45 &#176;C water bath on the water surface long enough for them to flatten completely.</li>
	<li>Catch the sections on a glass slide and drain them vertically to remove excess water before drying them overnight in an oven at 45 &#176;C.</li>
	<li>Process the sections further with the following schedule to deparaffinize and HE stain: 4x1 min in xylol, 1x1 min in 99% ethanol, 1x1 min in 96% ethanol, 1x1 min in 80% ethanol, 1x1 min in 70% ethanol, 1x1 min in ddH2O, 1x4 min in Mayer&#39;s hematoxylin, 1x10 min in ddH2O, 1x2min in 0.5% eosin, 1x30 s in ddH2O, 1x30 s in 80% ethanol, 2x30 s in 96% ethanol, 3x1 min in 99% ethanol, and 1x1 min in xylol.</li>
	<li>Take the glass slide out of the xylol and quickly cover the tissue with mounting media. Avoid to let the tissue dry out completely, then place a coverslip on top and seal the stained tissue.</li>
</ol>

<ol>
	<li>WHO. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Global report on diabetes. , 88 (2016).</li><li>Lim, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diabetic+nephropathy+-+complications+and+treatment.">Diabetic nephropathy - complications and treatment.</a> Int J Nephrol Renovasc Dis. 7, 361-381 (2014).</li><li>Yau, J. W., 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=Global+prevalence+and+major+risk+factors+of+diabetic+retinopathy.">Global prevalence and major risk factors of diabetic retinopathy.</a> Diabetes Care. 35 (3), 556-564 (2012).</li><li>Cheung, N., Mitchell, P., Wong, T. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diabetic+retinopathy.">Diabetic retinopathy.</a> The Lancet. 376 (9735), 124-136 (2010).</li><li>Raymond, N. T., 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=Higher+prevalence+of+retinopathy+in+diabetic+patients+of+South+Asian+ethnicity+compared+with+white+Europeans+in+the+community:+a+cross-sectional+study.">Higher prevalence of retinopathy in diabetic patients of South Asian ethnicity compared with white Europeans in the community: a cross-sectional study.</a> Diabetes Care. 32 (3), 410-415 (2009).</li><li>Heng, L. Z., 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=Diabetic+retinopathy:+pathogenesis,+clinical+grading,+management+and+future+developments.">Diabetic retinopathy: pathogenesis, clinical grading, management and future developments.</a> Diabet Med. 30 (6), 640-650 (2013).</li><li>Writing Committee for the Diabetic Retinopathy Clinical Research, N., 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=Panretinal+Photocoagulation+vs+Intravitreous+Ranibizumab+for+Proliferative+Diabetic+Retinopathy:+A+Randomized+Clinical+Trial.">Panretinal Photocoagulation vs Intravitreous Ranibizumab for Proliferative Diabetic Retinopathy: A Randomized Clinical Trial.</a> JAMA. 314 (20), 2137-2146 (2015).</li><li>Jao, L. E., Wente, S. R., Chen, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+multiplex+biallelic+zebrafish+genome+editing+using+a+CRISPR+nuclease+system.">Efficient multiplex biallelic zebrafish genome editing using a CRISPR nuclease system.</a> Proc Natl Acad Sci U S A. 110 (34), 13904-13909 (2013).</li><li>Sharma, K. R., 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=ELMO1+protects+renal+structure+and+ultrafiltration+in+kidney+development+and+under+diabetic+conditions.">ELMO1 protects renal structure and ultrafiltration in kidney development and under diabetic conditions.</a> Sci Rep. 6, 37172 (2016).</li><li>Baek, J. S., Fang, L., Li, A. C., Miller, Y. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ezetimibe+and+simvastatin+reduce+cholesterol+levels+in+zebrafish+larvae+fed+a+high-cholesterol+diet.">Ezetimibe and simvastatin reduce cholesterol levels in zebrafish larvae fed a high-cholesterol diet.</a> Cholesterol. , 564705 (2012).</li><li>Elo, B., Villano, C. M., Govorko, D., White, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Larval+zebrafish+as+a+model+for+glucose+metabolism:+expression+of+phosphoenolpyruvate+carboxykinase+as+a+marker+for+exposure+to+anti-diabetic+compounds.">Larval zebrafish as a model for glucose metabolism: expression of phosphoenolpyruvate carboxykinase as a marker for exposure to anti-diabetic compounds.</a> J Mol Endocrinol. 38 (4), 433-440 (2007).</li><li>Gut, P., Reischauer, S., Stainier, D. Y. R., Arnaout, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Little+Fish,+Big+Data:+Zebrafish+as+a+Model+for+Cardiovascular+and+Metabolic+Disease.">Little Fish, Big Data: Zebrafish as a Model for Cardiovascular and Metabolic Disease.</a> Physiol Rev. 97 (3), 889-938 (2017).</li><li>Kasprick, D. 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=Microanatomy+of+adult+zebrafish+extraocular+muscles.">Microanatomy of adult zebrafish extraocular muscles.</a> PLoS One. 6 (11), e27095 (2011).</li><li>Gestri, G., Link, B. A., Neuhauss, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+visual+system+of+zebrafish+and+its+use+to+model+human+ocular+diseases.">The visual system of zebrafish and its use to model human ocular diseases.</a> Dev Neurobiol. 72 (3), 302-327 (2012).</li><li>Lawson, N. D., Weinstein, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+imaging+of+embryonic+vascular+development+using+transgenic+zebrafish.">In vivo imaging of embryonic vascular development using transgenic zebrafish.</a> Dev Biol. 248 (2), 307-318 (2002).</li><li>Matthews, M., Varga, Z. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anesthesia+and+euthanasia+in+zebrafish.">Anesthesia and euthanasia in zebrafish.</a> ILAR J. 53 (2), 192-204 (2012).</li><li>Bell, B. 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=Retinal+vasculature+of+adult+zebrafish:+in+vivo+imaging+using+confocal+scanning+laser+ophthalmoscopy.">Retinal vasculature of adult zebrafish: in vivo imaging using confocal scanning laser ophthalmoscopy.</a> Exp Eye Res. 129, 107-118 (2014).</li><li>Avanesov, A., Malicki, J. <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+retina+in+the+zebrafish+model.">Analysis of the retina in the zebrafish model.</a> Methods Cell Biol. 100, 153-204 (2010).</li><li>Gramage, E., Li, J., Hitchcock, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+expression+and+function+of+midkine+in+the+vertebrate+retina.">The expression and function of midkine in the vertebrate retina.</a> Br J Pharmacol. 171 (4), 913-923 (2014).</li><li>Cao, R., Jensen, L. D., Soll, I., Hauptmann, G., Cao, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypoxia-induced+retinal+angiogenesis+in+zebrafish+as+a+model+to+study+retinopathy.">Hypoxia-induced retinal angiogenesis in zebrafish as a model to study retinopathy.</a> PLoS One. 3 (7), e2748 (2008).</li><li>Jorgens, K., Hillebrands, J. L., Hammes, H. P., Kroll, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish:+a+model+for+understanding+diabetic+complications.">Zebrafish: a model for understanding diabetic complications.</a> Exp Clin Endocrinol Diabetes. 120 (4), 186-187 (2012).</li><li>Alvarez, Y., 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=Predominant+cone+photoreceptor+dysfunction+in+a+hyperglycaemic+model+of+non-proliferative+diabetic+retinopathy.">Predominant cone photoreceptor dysfunction in a hyperglycaemic model of non-proliferative diabetic retinopathy.</a> Dis Model Mech. 3 (3-4), 236-245 (2010).</li><li>Gleeson, M., Connaughton, V., Arneson, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+hyperglycaemia+in+zebrafish+(Danio+rerio)+leads+to+morphological+changes+in+the+retina.">Induction of hyperglycaemia in zebrafish (Danio rerio) leads to morphological changes in the retina.</a> Acta Diabetol. 44 (3), 157-163 (2007).</li><li>Jorgens, K., 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=High+tissue+glucose+alters+intersomitic+blood+vessels+in+zebrafish+via+methylglyoxal+targeting+the+VEGF+receptor+signaling+cascade.">High tissue glucose alters intersomitic blood vessels in zebrafish via methylglyoxal targeting the VEGF receptor signaling cascade.</a> Diabetes. 64 (1), 213-225 (2015).</li><li>Lai, A. K., Lo, A. C. <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+of+diabetic+retinopathy:+summary+and+comparison.">Animal models of diabetic retinopathy: summary and comparison.</a> J Diabetes Res. 2013, 106594 (2013).</li></ol>

The authors have nothing to disclose.

Models of hypoxia-induced retinal neoangiogenesis show increased numbers of branching points, angiogenic sprouts, total vascular area and decreased intercapillary distance in zebrafish20. These findings support the idea that the zebrafish is susceptible to diabetic microvascular complications21, as key findings of later diabetic retinopathy include hypoxia-mediated neoangiogenesis, which is strongly linked to VEGF expression. Hyperglycemia-induced changes in the zebrafish retina lead to increased thickness of the vessels but maintain the overall patterning22. Alternating immersion in glucose solutions for 30 days also decreases the thickness of the IPL and INL23. The direct influence of glucose metabolites in zebrafish as metabolic proponents of vasculopathy was also already shown. Additional vascular hyperbranching was observed in the trunk vasculature of zebrafish embryos after incubation with methylglyoxal24. At the moment, there is no animal model which provides all the key criteria of diabetic retinopathy. Early changes are often found, but the progression to proliferative diabetic retinopathy is missing25. The zebrafish also falls into this definition, as we have only seen either hypoxia-mediated neoangiogenesis or hyperglycemia-induced changes until now. The current findings support the idea that the zebrafish is susceptible to hyperglycemia-mediated vascular changes and as an animal model may potentially show a progression to proliferative diabetic retinopathy. Depending on the strength and exposure to the hyperglycemia-mediated effect, the right experimental model could lead to ischemia in certain areas of the zebrafish retina and promote neoangiogenesis as the key criteria of proliferative diabetic retinopathy. However, as the zebrafish is a comparably new player in the field of modeling long-term microvascular pathologies, upcoming diabetes models in zebrafish will provide further information and clarify its importance in respect to other models and their pathologies. For example, an in-cross of tg(gata1a:DsRed) zebrafish with red-labelled erythrocytes into the tg(fli:EGFP) line could be used to simultaneously visualize potential intraocular bleedings in future zebrafish models showing microaneurysms as a predictor of progression to PDR.
Since the retinal vasculature progresses into a succession of arcades, evaluation of the intervascular distance, number of branching points and the total vascular area are related to the distance from the central optic artery. To avoid bias in the assessed vascular parameters, a point of orientation is required. The IOC is such a structure and is highly relevant since areas of vascular activity lie in direct spatial proximity. For consistent assessment, the retinal scan should be divided into multiple rectangular image sections with a consistent distance to the IOC. The entire retina should be analyzed and images numerically symmetrically distributed. The zebrafish retina shows areas with high and low capillary density and an unequal distribution of image sections can lead to additional bias.
<img alt="Figure 12" class="xfigimg" src="/files/ftp_upload/56674/56674fig12.jpg" /> 
Figure 12: Example presentation of vascular parameters as readout of retinal vasculature visualization. Measurement of intervascular distance (red double arrow) near the inner optic circle (IOC) (A). Three branching points (red circles) near the IOC (B). Visualization of zebrafish retinal vasculature with an enlarged area (red box) showing a sprouting vessel (C). Vessel diameter measured over a certain distance (white double arrow) from the central artery (white cross) (D). Vessel density is the percentage of retinal area occupied by overlaying vessels (diagonal red lines) (E). <a href="//cloudflare.jove.com/files/ftp_upload/56674/56674fig12large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
To measure the intervascular distance, one needs to set a certain distance to the IOC as standard (Figure 12A, white double arrow) and draw an imaginary horizontal line (Figure 12A, horizontal white line) parallel to the IOC. The distance between the vessels on this line added up and arithmetically averaged equals the intervascular distance. Branching points are counted inside each image whenever a vessel splits and more than one vascular lumen continue from the point of origin. This also includes horizontal connections between capillaries. It is important to stay consistent with analysis and decide which branching points to count and keep these rules throughout the whole experiment to reduce unneeded variation. Vascular sprouting, as demonstrated in Figure 12C, is another parameter that can be counted in each image to evaluate influence on the retinal vasculature. Angiogenic sprouts do not follow the arcade-like succession between central artery and IOC and focus near the outer parts of the retina. To assess certain vessel diameters, a point of orientation is always needed from which a set distance marks the measurement spot. The central artery&#39;s origin inside the retina provides such guidance for the main stem vessels (Figure 12D, white cross). The area occupied by vessels (Figure 12E, diagonal red lines) as a percentage of the whole retinal area is the vascular density and indirectly correlates to the avascular area.
A complete confocal scan of one sample should be composed of multiple high-detail pictures to allow visualization of small capillary hypersprouting. To optimize time and resources spent on this step, an automated tilling procedure should be used with a general scan depth for every tile. Uneven mounting of the retina can greatly increase the time spent to scan the vasculature with a confocal microscope. In an optimal approach one would like to only scan the vascular layer (Figure 13B, white box), but partial inclusion of the GCL is often necessary. Leaving an excess length of the optic nerve after truncation, as well as the cuts to achieve the flat-mount being too short, can lead to uneven mounting.
<img alt="Figure 13" class="xfigimg" src="/files/ftp_upload/56674/56674fig13.jpg" /> 
Figure 13: Comparison of HE staining and autofluorescence in the adult zebrafish retina. Retinal vasculature in focus above the GCL (A). Retinal vasculature (white box) showing green an EGFP signal and retinal layers exhibiting strong autofluorescence (B). <a href="//cloudflare.jove.com/files/ftp_upload/56674/56674fig13large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
As the proper preparation of the zebrafish retinal vasculature needs prior training, a small sample size with untrained dissectors and strongly varying preparation results are a main limitation of the technique. While the preparation of tg(fli:EGFP) zebrafish eyes gives fast insight into the state of the vasculature, the technique still utilizes around 20 min of work time per retina for an experienced researcher. All of these preparation steps for the retinal vasculature must be performed under a dissecting microscope and dissectors need to stay concentrated the whole time as a careless step can potentially induce vessel breakage. The dissector should practice regularly as prolonged absence from preparation reduces a dissector&#39;s likelihood of maintaining vessel integrity.
Furthermore, additional readout via immunohistochemistry (IHC) is still limited, as only a small number of antibodies validated on human and rodent tissue are working with zebrafish. Experimenters interested in IHC are advised to search for zebrafish-specific antibodies, especially when working with new targets. Alternatively, it is recommended to use additional zebrafish reporter lines which are useful to study cells beyond the vasculature in the eye. This strategy, however, is time consuming, as it takes a few months to generate adult zebrafish lines which carry multiple transgenic reporters.
Nonetheless, the zebrafish poses a unique array of advantages. They are relatively small and reproduce readily. They can grow to adult stages quickly and their embryos are optimal for drug screening. The field is readily growing and more literature is becoming accessible. With the ability to generate gene knockouts at a rapid speed and an abundance of transgene fluorescence reporter lines, the choice for zebrafish is only restrained by the chosen research question.

Diabetes mellitus is a metabolic disease defined by hyperglycemia resulting from dysfunctional insulin secretion or inadequate tissue response to secreted insulin. The WHO estimates that 422 million adults were living with diabetes mellitus in 20141 and the prevalence of diabetes worldwide is expected to increase to 8-10% of the population until 20352, making diabetes one of the key research fields in the next decades. Living with chronic high blood sugar leads to long-term microvascular complications, including diabetic retinopathy, nephropathy, and polyneuropathy. The management and prevention of these complications are difficult; indeed, diabetes is becoming the most frequent cause of end-stage renal disease (ESRD), which leads to dialysis2, and diabetes is the leading cause of blindness among middle-aged adults3.
The initial causes of microvascular damage in the diabetic eye are chronic hyperglycemia, metabolic alterations, as well as certain risk factors (e.g., hypertension, dyslipidemia), leading to vascular endothelial dysfunction, pericyte dropout, and capillary regression that results in acellular vascular sleeves. The resulting retinal ischemia is the cause of neovascularization and increased vascular permeability promoting the development of proliferative diabetic retinopathy (PDR)4. Diabetic retinopathy was generally detected in 37% of diabetes patients, while sight-threatening diabetic retinopathy was ascertained in 12% of the screened white European cohort in the UKADS study5. The current treatment can only prevent further complications and is not able to fully restore the already induced damage. Panretinal photocoagulation, in addition to glycemic control, is the standard therapy for proliferative diabetic retinopathy (PDR) but affects the adjacent healthy tissue as well. Anti-VEGF interventions show promising results as alternatives to laser treatment6,7, but ultimately, specialized, targeted therapy and new drugs are needed to manage the growing number of patients at risk of vision-loss.
The established animal research models of diabetic retinopathy do not share every aspect of human pathophysiology. Utilization of the correct species to address the specific requirements of the scientific research question is one of the most substantial parts of the experimental setup. The zebrafish embryo (Danio rerio) is already widely used in developmental research and provides an optimal practical background to knockdown or knockout specific genes via morpholinos or the CRISPR/Cas9 technique8. These methods can easily be utilized in zebrafish to investigate genes, which were identified by large scale genome-wide association studies (GWAS), generating insights into particular mechanisms of disease progression and susceptibility9. Short generational time, large amounts of offspring, easy and low-cost handling, and growing assay support have increased the relevance of the zebrafish model, especially given its great potential for modeling metabolic disease. Conservation of biological mechanisms in zebrafish has been shown as a basis for pharmacological therapy development. For example, the antidiabetic drug metformin and cholesterol-lowering simvastatin have been shown to &#34;treat&#34; conditions in models of cAMP/dexamethasone-induced high PEPCK expression and high-cholesterol diet-induced hypercholesterolemia10,11,12. This advancing insight into overarching conserved metabolic mechanisms is further supported by the growing number of zebrafish diabetes models, through experiments such as: alternating incubation in glucose solutions, Streptozotocin-induced ablation of beta cells, nitroreductase-mediated beta cell ablation using the prodrug metronidazole, monogenic diabetes mediated via pdx1 gene knockdown or knockout, as well as models of increased insulin resistance in skeletal muscle12. These already established protocols, the above mentioned specifics of the species, and the ability to efficiently manipulate the genome in a large number of samples all demonstrate the advantages of the zebrafish for studying the mechanisms driving complex disease processes as well as the ability to screen for pharmacological interventions.
A general understanding of the basic zebrafish eye anatomy (Figure 1) is necessary for the dissector in order to utilize zebrafish as a model for retinal angiopathy. The zebrafish eye has six extraocular muscles, four rectus, and two oblique muscles that insert on the outside of the globe at the sclera13. The cornea is the transparent tissue covering the lens and continues directly into the sclera, which forms the outer shell of the eye. The sclera is non-transparent, has a partially light reflecting surface, and is strongly pigmented. The lens itself is more ball-shaped than the human counterpart. The retina consists of three nuclear layers of neuronal cells while the oxygen-providing retinal vasculature is closely associated with the inner ganglion cell layer but does not form a subretinal network. Choroidal vessels, in contrast, lie in between the sclera and retina and are associated with the retinal pigmented epithelium (RPE). This capillary network supplies oxygen to the outer parts of the retina14.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56674/56674fig1.jpg" /> 
Figure 1: Schematic depiction of the adult zebrafish eye. <a href="//cloudflare.jove.com/files/ftp_upload/56674/56674fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Straightforward visualization of the retinal vasculature can be achieved by utilization of the transgenic tg(fli:EGFP) zebrafish line15. The green fluorescent protein expressed under control of the fli promotor in endothelial cells of the vasculature is the basis for the visualization via a laser scanning microscope in the later steps. This is achieved via dissection of the zebrafish retina and whole-mounting of the tissue. This transgenic model provides a fast vascular readout without any application of intravascular labeling or whole-mount immunohistochemistry. To analyze diabetic retinopathy in zebrafish, a step by step and standardized preparation routine should be used by every dissector. The following preparation protocol provides other research groups the option to easily evaluate vascular changes in the exposed vessels of the adult zebrafish eye and provide guidance to establish an optimized dissection routine for the zebrafish retina.

<table><tbody><tr><td>NaOH</td><td>Roth</td><td>6771.3</td><td></td></tr><tr><td>KCl</td><td>Merck</td><td>1.04936</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2*&lt;/sub&gt;6H&lt;sub&gt;2&lt;/sub&gt;0</td><td>Roth</td><td>5239.2</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4*&lt;/sub&gt;7H&lt;sub&gt;2&lt;/sub&gt;0</td><td>Merck</td><td>1.05886.0500</td><td></td></tr><tr><td>Paraformaldehyd</td><td>Roth</td><td>0335.3</td><td></td></tr><tr><td>Sodium dihydrogen phosphate</td><td>Roth</td><td>2370.1</td><td></td></tr><tr><td>Ethyl 3-aminobenzoate methanesulfonate salt (MS-222,Tricaine)</td><td>Sigma</td><td>A5040</td><td></td></tr><tr><td>PBS</td><td>Roth</td><td>9143.1</td><td></td></tr><tr><td>Agarose</td><td>Roth</td><td>2267.3</td><td></td></tr><tr><td>Fluoromount-G</td><td>Thermo-Fischer</td><td>00-4958-02</td><td></td></tr><tr><td>Petri dish</td><td>Greiner Bio one</td><td>633180</td><td></td></tr><tr><td>Six-well plate</td><td>StarLab</td><td>CC7682-7506</td><td></td></tr><tr><td>Needle</td><td>MSG Praxisbedarf</td><td>BD 300900</td><td></td></tr><tr><td>Micro Tweezer</td><td>World Precision Instruments</td><td>14095</td><td></td></tr><tr><td>Microdissection Scissor</td><td>World Precision Instruments</td><td>501778</td><td></td></tr><tr><td>Glass slide</td><td>Carl Roth</td><td>H872.1</td><td></td></tr><tr><td>Coverslip 22mmx22mm</td><td>neoLab</td><td>103512222</td><td></td></tr><tr><td>Scalpel</td><td>MSG Praxisbedarf</td><td>FEA111</td><td></td></tr><tr><td>Epi-Illumination</td><td>Leica</td><td>10446389</td><td></td></tr><tr><td>Fluorescence stereomicroscope MZ10 F</td><td>Leica</td><td>NA</td><td></td></tr><tr><td>Confocal laser-scanning microscope SP5 DS</td><td>Leica</td><td>NA</td><td></td></tr><tr><td>Stereomicroscope M80</td><td>Leica</td><td>NA</td><td></td></tr><tr><td>Zebrafish line: Tg(fli:EGFP), ABTL wildtype</td><td>NA</td><td>NA</td><td>see Reference 15</td></tr><tr><td>Mayer&amp;rsquo;s hematoxylin</td><td>Dr. K. Hollborn &amp;amp; S&amp;ouml;hne</td><td>0088663</td><td></td></tr><tr><td>0.5% eosin</td><td>Dr. K. Hollborn &amp;amp; S&amp;ouml;hne</td><td>NA</td><td></td></tr><tr><td>99,9% ethanol</td><td>Roth</td><td>9065.2</td><td></td></tr><tr><td>Paraffin</td><td>Merck</td><td>1,071,501,000</td><td></td></tr><tr><td>Xylol</td><td>Roth</td><td>4436.2</td><td></td></tr><tr><td>Acetone</td><td>Emsure</td><td>606-001-00-8</td><td></td></tr><tr><td>Microtome RM 2165</td><td>Leica</td><td>NA</td><td></td></tr></tbody></table>

studying diabetes through the eyes of a fish: microdissection, visualization, and analysis of the adult tg(fli:egfp) zebrafish retinal vasculature

Diabetic retinopathy is the leading cause of blindness among middle-aged adults. The rising prevalence of diabetes worldwide will make the prevention of diabetic microvascular complications one of the key research fields of the next decades. Specialized, targeted therapy and novel therapeutic drugs are needed to manage the increasing number of patients at risk of vision-loss. The zebrafish is an established animal model for developmental research questions with increasing relevance for modeling metabolic multifactorial disease processes. The advantages of the species allow for optimal visualization and high throughput drug screening approaches, combined with the strong ability to knock out genes of interest. Here, we describe a protocol which will allow easy analysis of the adult tg(fli:EGFP) zebrafish retinal vasculature as a fast read-out in settings of long-term vascular pathologies linked to neoangiogenesis or vessel damage. This is achieved via dissection of the zebrafish retina and whole-mounting of the tissue. Visualization of the exposed vessels is then achieved via confocal microscopy of the green EGFP reporter expressed in the adult retinal vasculature. Correct handling of the tissue will lead to better outcomes and less internal vessel breakage to assure the visualization of the unaltered vascular structure. The method can be utilized in zebrafish models of retinal vasculopathy linked to changes in the vessel architecture as well as neoangiogenesis.

Here we demonstrate two typical morphological examples of the retinal vasculature in adult tg(fli:EGFP) zebrafish: once visualized with a fluorescence microscope (Figure 9A) and once with a confocal laser scanning microscope (Figure 9B). The revealed retinal structure shows a highly-organized pattern. Strong autofluorescence is seen in the zebrafish retina, when visualized with a fluorescence microscope. Thus, a visualization of only the vascular layer via a confocal scan is advised to reduce background fluorescence and increase contrast. For faster picture acquisition or situations where qualitative data are enough, the fluorescence microscope is an attractive alternative.
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/56674/56674fig9.jpg" /> 
Figure 9: Representative pictures of adult tg(fli:EGFP) zebrafish retinal vasculature. Two typical morphological examples of the retinal vasculature are shown: The central optic artery spreads into 5-7 main vessels, which then branch into a succession of arcades. All further vessels drain into the circumferential vein (CV) limiting the outer part of each petal. Visualization via fluorescence microscopy at 2.5x magnification (A). Visualization of the retinal vasculature by confocal laser scanning microscopy through a combined 5x5 single image tile scan (B). <a href="https://cloudflare.jove.com/files/ftp_upload/56674/56674fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The optic artery penetrates the retina at the optic nerve head and, in most samples, spreads into 5-7 main vessels (Figure 10A). The main vessels then branch into a succession of arcades and connect to the inner optic circle (IOC), also called the circumferential vein (CV), encircling the optic disc in the periphery of the flat mounted retina (see Figure 10B). The IOC is the venous vessel that drains the arterial blood at the inner rim of the ocular globe (bulbus oculi). The zebrafish retinal vasculature also shows areas of high vascular activity with branching of the capillaries and angiogenic sprouting (Figure 10C). This vascular activity is mostly situated in near proximity to the IOC. It is important to notice that the same eye can demonstrate both high and low vascular activities in different regions of the retina (compare Figure 10B and Figure 10C). In general, the retinal vasculature of the zebrafish shows more avascular space and fewer capillaries in between the main branches if compared to, for example, the retina of mice17. Both eyes of each sample should be analyzed because the vasculature may vary in between and singular inspection can lead to bias.
<img alt="Figure 10" class="xfigimg" src="/files/ftp_upload/56674/56674fig10.jpg" /> 
Figure 10: Magnified areas of adult tg(fli:EGFP) zebrafish retinal vasculature visualization. The optic artery branching into main vessels (A). Retinal capillaries in an area of low vascular activity connecting to the inner optic circle (IOC) at the bottom of the picture (B). Retinal capillaries in an area of high vascular activity connecting to the IOC (C). <a href="https://cloudflare.jove.com/files/ftp_upload/56674/56674fig10large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
A highly-conserved architecture of neuronal layers is already present in zebrafish from approximately 72 h post fertilization (hpf) onwards18. The adult zebrafish retina shows the following layers from inside out (Figure 11): ganglion cell layer (GCL), inner plexiform layer (IPL), inner nuclear layer (INL), outer plexiform layer (OPL), outer nuclear layer (ONL), photoreceptor layer (PL), and the retinal pigmented epithelium (RPE). The estimation of vascular parameters from retinal vasculature visualization is shown in Figure 12. The inner retinal vasculature is situated on the ganglion cell layer (GCL) (Figure 13B, white box) while the choroidal capillaries would be associated with the retinal pigmented epithelium (RPE).
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/56674/56674fig11.jpg" /> 
Figure 11: HE staining of the adult zebrafish eye. Cross-sectional overview of the whole eye after removal of the lens (A). Retinal layers of the adult zebrafish eye19 (B). Indication (black arrow) of the natural breaking point in the PL (C). <a href="https://cloudflare.jove.com/files/ftp_upload/56674/56674fig11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tgfli:EGFP Zebrafish Retinal Vasculature at JoVE.com

Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tgfli:EGFP Zebrafish Retinal Vasculature

Here, we discuss a method protocol which will allow an easy analysis of the adult tg(fli:EGFP) zebrafish retinal vasculature as a fast read-out in settings of long-term vascular pathologies linked to neoangiogenesis and structural changes.

Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tg(fli:EGFP) Zebrafish Retinal Vasculature

Diabetic retinopathy is the leading cause of blindness among middle-aged adults. The rising prevalence of diabetes worldwide will make the prevention of ...

studying-diabetes-through-eyes-fish-microdissection-visualization

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

13.0K Views.  Heidelberg University. Here, we discuss a method protocol which will allow an easy analysis of the adult tg(fli:EGFP) zebrafish retinal vasculature as a fast read-out in settings of long-term vascular pathologies linked to neoangiogenesis and structural changes.

Video: Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tgfli:EGFP Zebrafish Retinal Vasculature

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Developmental Biology

Acute and Chronic Models of Hyperglycemia in Zebrafish: A Method to Assess the Impact of Hyperglycemia on Neurogenesis and the Biodistribution of Radiolabeled Molecules

Hyperglycemia is a major health issue that leads to cardiovascular and cerebral dysfunction. For instance, it is associated with increased neurological problems after stroke and is shown to impair neurogenic processes. Interestingly, the adult zebrafish has recently emerged as a relevant and useful model to mimic hyperglycemia/diabetes and to investigate constitutive and regenerative neurogenesis. This work provides methods to develop zebrafish models of hyperglycemia to explore the impact of hyperglycemia on brain cell proliferation under homeostatic and brain repair conditions. Acute hyperglycemia is established using the intraperitoneal injection of D-glucose (2.5 g/kg bodyweight) into adult zebrafish. Chronic hyperglycemia is induced by immersing adult zebrafish in D-glucose (111 mM) containing water for 14 days. Blood-glucose-level measurements are described for these different approaches. Methods to investigate the impact of hyperglycemia on constitutive and regenerative neurogenesis, by describing the mechanical injury of the telencephalon, dissecting the brain, paraffin embedding and sectioning with a microtome, and performing immunohistochemistry procedures, are demonstrated. Finally, the method of using zebrafish as a relevant model for studying the biodistribution of radiolabeled molecules (here,[18F]-FDG) using PET/CT is also described.

This work describes methods to establish acute and chronic hyperglycemia models in zebrafish. The aim is to investigate the impact of hyperglycemia on physiological processes, such as constitutive and injury-induced neurogenesis. The work also highlights the use of zebrafish to follow radiolabeled molecules (here, [18F]-FDG) using PET/CT.

Hyperglycemia is a major health issue that leads to cardiovascular and cerebral dysfunction. For instance, it is associated with increased neurological ...

A Zebrafish Model of Diabetes Mellitus and Metabolic Memory

Diabetes mellitus currently affects 346 million individuals and this is projected to increase to 400 million by 2030. Evidence from both the laboratory and large scale clinical trials has revealed that diabetic complications progress unimpeded via the phenomenon of metabolic memory even when glycemic control is pharmaceutically achieved. Gene expression can be stably altered through epigenetic changes which not only allow cells and organisms to quickly respond to changing environmental stimuli but also confer the ability of the cell to "memorize" these encounters once the stimulus is removed. As such, the roles that these mechanisms play in the metabolic memory phenomenon are currently being examined.
We have recently reported the development of a zebrafish model of type I diabetes mellitus and characterized this model to show that diabetic zebrafish not only display the known secondary complications including the changes associated with diabetic retinopathy, diabetic nephropathy and impaired wound healing but also exhibit impaired caudal fin regeneration. This model is unique in that the zebrafish is capable to regenerate its damaged pancreas and restore a euglycemic state similar to what would be expected in post-transplant human patients. Moreover, multiple rounds of caudal fin amputation allow for the separation and study of pure epigenetic effects in an in vivo system without potential complicating factors from the previous diabetic state. Although euglycemia is achieved following pancreatic regeneration, the diabetic secondary complication of fin regeneration and skin wound healing persists indefinitely. In the case of impaired fin regeneration, this pathology is retained even after multiple rounds of fin regeneration in the daughter fin tissues. These observations point to an underlying epigenetic process existing in the metabolic memory state. Here we present the methods needed to successfully generate the diabetic and metabolic memory groups of fish and discuss the advantages of this model.

Metabolic memory is the phenomenon by which diabetic complications persist and progress unimpeded even after euglycemia is achieved pharmaceutically. Here we describe a diabetes mellitus zebrafish model which is unique in that it allows for the examination of the mitotically transmissible epigenetic components of metabolic memory in vivo.

Diabetes mellitus currently affects 346 million individuals and this is projected to increase to 400 million by 2030. Evidence from both the laboratory ...

Microdissection of Zebrafish Embryonic Eye Tissues

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post fertilization (dpf), the larvae will show the first visual response. Many genes have been identified to control a proper eye development, but we are far from a complete understanding of the underlying genetic architecture. Whole genome gene expression profiling is a useful tool to elucidate genetic regulatory network for eye development. However, the small size of the embryonic eye in zebrafish makes it challenging to obtain intact and pure eye tissues for expression analysis. For example, the anterior-posterior length of the eye between day 2 and 3 is only approximately 200-300 &mu;m, while the diameter of the lens is less 100 &mu;m. Also, the retinal pigment epithelium (RPE) underlying the retina is just a single-layer epithelium. While gene expression profiles can be obtained from the whole embryo, they do not accurately represent the expression of these tissues. Therefore pure tissue must be obtained for a successful gene expression profiling of eye development. To address this issue, we have developed an approach to microdissect intact retina and retina with RPE attached from 1-3 dpf, which cover major stages of eye morphogenesis. All procedures can be done with fine forceps and general laboratory supplies under standard stereomicroscopes. For retinal dissection, the single-layer RPE is removed and peeled off by brushing action and the preferential adherence of the RPE remnants to the surface of the culture plate for dissection. For RPE-attached retinal dissection, the adherence of RPE to the dissection plate is removed before the dissection so that the RPE can be completely preserved with the retina. A careful lifting action of this tissue can efficiently separate the presumptive choroid and sclera. The lens can be removed in both cases by a chemically etched tungsten needle. In short, our approach can obtain intact eye tissues and has been successfully utilized to study tissue-specific expression profiles of zebrafish retina1, 2 and retinal pigment epithelium3.

This article describes an approach to microdissect zebrafish retinas with and without retinal pigment epithelium attached, from one to three days postfertilization embryos.

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post ...

Biology

Simultaneous Calcium Imaging and Glucose Stimulation in Living Zebrafish to Investigate In Vivo &#946;-Cell Function

The pancreatic &#946;-cells sustain systemic glucose homeostasis by producing and secreting insulin according to the blood glucose levels. Defects in &#946;-cell function are associated with hyperglycemia that can lead to diabetes. During the process of insulin secretion, &#946;-cells experience an influx of Ca2+. Thus, imaging the glucose-stimulated Ca2+ influx using genetically encoded calcium indicators (GECIs) provides an avenue to studying &#946;-cell function. Previously, studies showed that isolated zebrafish islets expressing GCaMP6s exhibit significant Ca2+ activity upon stimulation with defined glucose concentrations. However, it is paramount to study how &#946;-cells respond to glucose not in isolation, but in their native environment, where they are systemically connected, vascularized, and densely innervated. To this end, the study leveraged the optical transparency of the zebrafish larvae at early stages of development to illuminate &#946;-cell activity in vivo. Here, a detailed protocol for Ca2+ imaging and glucose stimulation to investigate &#946;-cell function in vivo is presented. This technique allows to monitor the coordinated Ca2+ dynamics in &#946;-cells with single-cell resolution. Additionally, this method can be applied to work with any injectable solution such as small molecules or peptides. Altogether, the protocol illustrates the potential of the zebrafish model to investigate islet coordination in vivo and to characterize how environmental and genetic components might affect &#946;-cell function.

Here, the study presents a protocol for calcium imaging and glucose stimulation of the pancreatic &#946;-cells of the zebrafish in vivo.

The pancreatic &#946;-cells sustain systemic glucose homeostasis by producing and secreting insulin according to the blood glucose levels. Defects in ...

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 ...

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding diseases, and future therapies hinge on fundamental understandings about how different retinal cells function normally. Gaining such information with biochemical methods has proven difficult because contributions of particular cell types are diminished in the retinal cell milieu. Live retinal imaging can provide a view of numerous biological processes on a subcellular level, thanks to a growing number of genetically encoded fluorescent biosensors. However, this technique has thus far been limited to tadpoles and zebrafish larvae, the outermost retinal layers of isolated retinas, or lower resolution imaging of retinas in live animals. Here we present a method for generating live ex vivo retinal slices from adult zebrafish for live imaging via confocal microscopy. This preparation yields transverse slices with all retinal layers and most cell types visible for performing confocal imaging experiments using perfusion. Transgenic zebrafish expressing fluorescent proteins or biosensors in specific retinal cell types or organelles are used to extract single-cell information from an intact retina. Additionally, retinal slices can be loaded with fluorescent indicator dyes, adding to the method&#39;s versatility. This protocol was developed for imaging Ca2+ within zebrafish cone photoreceptors, but with proper markers it could be adapted to measure Ca2+ or metabolites in M&#252;ller cells, bipolar and horizontal cells, microglia, amacrine cells, or retinal ganglion cells. The retinal pigment epithelium is removed from slices so this method is not suitable for studying that cell type. With practice, it is possible to generate serial slices from one animal for multiple experiments. This adaptable technique provides a powerful tool for answering many questions about retinal cell biology, Ca2+, and energy homeostasis.

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

Imaging retinal tissue can provide single-cell information that cannot be gathered from traditional biochemical methods. This protocol describes preparation of retinal slices from zebrafish for confocal imaging. Fluorescent genetically encoded sensors or indicator dyes allow visualization of numerous biological processes in distinct retinal cell types.

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding ...

Neuroscience

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

Alternate Immersion in Glucose to Produce Prolonged Hyperglycemia in Zebrafish

Zebrafish (Danio rerio) are an excellent model to investigate the effects of chronic hyperglycemia, a hallmark of Type II Diabetes Mellitus (T2DM). This alternate immersion protocol is a noninvasive, step-wise method of inducing hyperglycemia for up to eight weeks. Adult zebrafish are alternately exposed to sugar (glucose) and water for 24 hours each. The zebrafish begin treatment in a 1% glucose solution for 2 weeks, then a 2% solution for 2 weeks, and finally a 3% solution for the remaining 4 weeks. Compared to water-treated (stress) and mannitol-treated (osmotic) controls, glucose-treated zebrafish have significantly higher blood sugar levels. The glucose-treated zebrafish show blood sugar levels of 3-times that of controls, suggesting that after both four and eight weeks hyperglycemia can be achieved. Sustained hyperglycemia was associated with increased Glial Fibrillary Acidic Protein (GFAP) and increased nuclear factor Kappa B (NF-kB)&#160;levels in retina and decreased physiological responses, as well as cognitive deficits suggesting this protocol can be used to model disease complications.

This protocol noninvasively induces hyperglycemia in zebrafish for up to 8 weeks. Using this protocol, an in-depth study of the adverse effects of hyperglycemia can be made.

Zebrafish (Danio rerio) are an excellent model to investigate the effects of chronic hyperglycemia, a hallmark of Type II Diabetes Mellitus (T2DM). This ...

Zebrafish Maintenance and Husbandry

The zebrafish (Danio rerio) is a powerful vertebrate model system for studying development, modeling disease, and screening for novel therapeutics. Due to their small size, large numbers of zebrafish can be housed in the laboratory at low cost. Although zebrafish are relatively easy to maintain, special consideration must be given to both diet and water quality to in order to optimize fish health and reproductive success.
This video will provide an overview of zebrafish husbandry and maintenance in the lab. After a brief review of the natural zebrafish habitat, techniques essential to recreating this environment in the lab will be discussed, including key elements of fish facility water recirculation systems and the preparation of brine shrimp as part of the zebrafish diet. Additionally, the presentation will include information on how specific zebrafish strains are tracked in a laboratory setting, with specific reference to the collection of tail fin samples for DNA extraction and genotyping. Finally, experimental modifications of the zebrafish environment will be discussed as a means to further our understanding of these fish, and in turn, ourselves.

Maintenance and Husbandry of Danio rerio Zebrafish

The zebrafish (Danio rerio) is a powerful vertebrate model system for studying development, modeling disease, and screening for novel therapeutics. Due to ...

Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tg(fli:EGFP) Zebrafish Retinal Vasculature

Summary

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Chapters in this video

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

Acute and Chronic Models of Hyperglycemia in Zebrafish: A Method to Assess the Impact of Hyperglycemia on Neurogenesis and the Biodistribution of Radiolabeled Molecules

A Zebrafish Model of Diabetes Mellitus and Metabolic Memory

Microdissection of Zebrafish Embryonic Eye Tissues

Simultaneous Calcium Imaging and Glucose Stimulation in Living Zebrafish to Investigate In Vivo β-Cell Function

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

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Alternate Immersion in Glucose to Produce Prolonged Hyperglycemia in Zebrafish

Zebrafish Maintenance and Husbandry