This study was supported by a grant of the Center of Regenerative Therapies Dresden (&#34;Zebrafish as a model to unravel the mechanisms of glucocorticoid-induced bone loss&#34;) and additionally by a grant of the Deutsche Forschungsgemeinschaft (Transregio 67, project 387653785) to FK. We are very grateful to Jan Kaslin and Avinash Chekuru for their guidance and assistance on performing trepanation of the calvariae and fractures in bony fin rays. Experiments were designed, performed and analyzed by KG and FK. FK wrote the manuscript. We would also like to thank Katrin Lambert,&#160;Nicole Cudak, and other members of the Knopf and Brand labs for technical assistance and discussion. Our thanks also goes to Marika Fischer and Jitka Michling for excellent fish care and to Henriette Knopf and Josh Currie for proofreading the manuscript.

All methods described here were approved by the Landesdirektion Dresden (Permit numbers: AZ 24D-9168.11-1/2008-1, AZ 24-9168.11-1/2011-52, AZ 24-9168.11-1/2013-5, AZ 24-9168.11-1/2013-14, AZ DD24.1- 5131/354/87).
1. Preparation of Materials and Solutions
NOTE: Prednisolone, like other glucocorticoids, leads to immunosuppression. Thus, precaution must be taken to prevent infection in treated animals during the experiment. To this end, autoclave glass ware and &#39;fish water&#39; (i.e., the water that is used to rear adult zebrafish) before starting the experiment.
<ol>
	<li>Calculate the number of zebrafish to be used in the experiment. Be sure to include the individuals that serve as controls.</li>
	<li>Autoclave 600 mL glass beakers that are covered with aluminum foil with autoclave tape. Zebrafish will be treated individually, thus 1 glass beaker is required per zebrafish.</li>
	<li>Calculate the amount of fish water needed for the experiment. 300 mL are required per individual and day. Take into consideration the length of the experiment (e.g., 1 week/7 days).</li>
	<li>Fill glass bottles with fish water, close them and autoclave the bottles with autoclave tape. Ideally, use 2 L bottles, but other bottles like 1 L or 5 L can be used as well. 
	NOTE: If experiments are long (e.g., 4 weeks/28 days) and the amount of glass bottles is limited, make sure to prepare bottles for a minimum of several days at a time and repeat the process (1.3 to 1.4) throughout the experiment.</li>
	<li>Prepare a prednisolone stock solution of sterile-filtered 100 mM prednisolone in dimethylsulfoxide (DMSO). Freeze the stock in 1 mL or 2 mL microcentrifuge tube aliquots. 
	CAUTION: Always wear protective clothing when working with prednisolone (protective gloves, safety goggles, lab coat).</li>
	<li>On the first day of the experiment, prepare incubation solutions with prednisolone and DMSO, respectively. To this end, thaw the required amount of 100 mM prednisolone stock solution and add prednisolone to autoclaved fish water at a final concentration of 50 &#181;M.
	<ol>
		<li>For the control treatments, add the corresponding volume of DMSO to autoclaved fish water. As an example, to produce 2 L prednisolone containing fish water, add 1 mL of 100 mM prednisolone stock to 2 L fish water. In another 2 L bottle of fish water, add 1 mL DMSO, correspondingly.</li>
		<li>Work in an appropriate place that is designated to perform drug treatments, ideally in a room at 27 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Generation of Injuries in Zebrafish Fins
NOTE: To injure bone in zebrafish fins, perform resection of the fin (amputation, usually in the caudal fin) or fracture individual bony fin rays (fracture model). To this end anaesthetize adult zebrafish first.
<ol>
	<li>Amputation
	<ol>
		<li>To anaesthetize adult zebrafish, prepare a 100 mm diameter Petri dish containing 0.02% MS-222 (Ethyl-3-aminobenzoate methanesulfonate) in fish water (this water does not need to be autoclaved).
		<ol>
			<li>Transfer one zebrafish at a time to the MS-222 containing Petri dish by using a fish net and incubate it until it lies on its side without movement and does not respond to touch. Test the latter by touching the anal fin with blunt forceps.</li>
			<li>Wait until level 4 of anesthesia is reached which is the appropriate level for fin resection12. At this stage, the rate of opercular movement is decreased, muscle tone is lost and the fish does not move upon touch12.</li>
		</ol>
		</li>
		<li>Take blunt forceps and transfer the anaesthetized zebrafish carefully to the inversed lid of the 100 mm Petri dish. Place the zebrafish on its lateral side. Make sure the zebrafish always lie in the same orientation, for example on their right body side. With a scalpel or a razor blade, resect 50% of the fin (Figure 1A, 2A). 
		NOTE: The speed of zebrafish fin regeneration depends on the amount of resected fin tissue, i.e., the more fin tissue is resected, the faster the fin regenerates13. Thus, to minimize variations in fin regeneration make sure to always resect the fin at the same level in all zebrafish.</li>
		<li>Transfer the injured zebrafish to an autoclaved glass beaker containing 300 mL of prednisolone or DMSO containing autoclaved fish water. Cover the glass beaker with aluminum foil to avoid escape of the zebrafish.</li>
		<li>Repeat steps 2.1.2 to 2.1.3 until the required number of zebrafish for the experiment is reached.</li>
	</ol>
	</li>
	<li>Fin fracture
	<ol>
		<li>Prepare an agarose-coated 100 mm Petri dish by heating a 2% agarose solution in fish water or E3 (embryo media 3, 50 mM sodium chloride, 0.17 mM potassium chloride, 0.33 mM calcium chloride, 0.33 mM magnesium sulfate). Carefully pour roughly 10&#8211;15 mL of the solution into the Petri dish. The bottom of the dish should be completely covered. Let the agarose solidify.</li>
		<li>Anaesthetize zebrafish as described in section 2.1.</li>
		<li>Take blunt forceps and transfer the anaesthetized zebrafish carefully to the agarose-coated 100 mm Petri dish. Place the zebrafish on its lateral side. Under the dissection microscope, spread the caudal fin completely to be able to identify distinct bony fin rays and fin ray segments. Make sure the zebrafish always lie in the same orientation, for example on their right body side.</li>
		<li>With an injection needle (0.3 mm x 13 mm) introduce a fracture in the center of a bony fin ray segment (Figure 1B). To do this, the needle is pushed slightly onto the segment until a crack appears (arrow in Figure 2B). While gentle pressure leads to fracture of only one of the two opposing hemirays stronger pressure will lead to fracture of both hemiray segments.
		<ol>
			<li>Do not introduce too many fractures in fins, since this will greatly impair stability of the fin. As a suggestion, apply fractures only in 3 to 5 fin rays.</li>
			<li>Always produce fractures in corresponding segments across different fin rays and zebrafish, for example in dorsal fin rays 3, 7 (or 8) and ventral fin ray 3. The proximal-distal level of fracture is ideal at 3 segments proximal to the fin rays&#39; bifurcation point 11.</li>
		</ol>
		</li>
		<li>Transfer the injured zebrafish to an autoclaved glass beaker containing 300 mL of prednisolone or DMSO containing autoclaved fish water. Cover the glass beaker with aluminum foil to avoid escape of the zebrafish.</li>
		<li>Repeat steps 2.2.2 to 2.2.5 until the required number of zebrafish for the experiment is reached.</li>
	</ol>
	</li>
</ol>
3. Generation of Calvarial Skull Injuries (Trepanation)
NOTE: The calvariae in zebrafish are homologous to calvarial bones in mammals. Thus, these exoskeletal bones14 represent a tissue of special interest when studying the pathogenesis of GIO. To injure the skull, trepanation is performed by drilling a hole in the Os frontale and/or Os parietale (Figure 1C, 2C) with the help of a microdrill11.
<ol>
	<li>Anaesthetize the zebrafish according to section 2.1.</li>
	<li>Hold the anesthetized zebrafish upright with forceps, so that the calvarial bones are well visible under a dissection microscope from above (Figure 1C).</li>
	<li>Locate the rotating microdrill on the center of the frontal bone (Os frontale) and touch the bone. The produced hole is of the size of the microdrill burr (500 &#181;m, Figure 2C, Figure 3B). 
	NOTE: Be sure to not move the microdrill laterally while producing the injury. Also stop immediately when the resistance of the bone suddenly drops. Do not drill any further, otherwise the brain will be damaged 15.</li>
	<li>Transfer the injured zebrafish to a cage filled with fish water until it fully recovers from anesthesia. Watch it carefully.</li>
	<li>Transfer the injured zebrafish to a fish water plus prednisolone or DMSO containing glass beaker.</li>
</ol>
4. Treatment of Zebrafish During Incubation
NOTE: During application of prednisolone/DMSO, the drug containing fish water needs to be changed daily, and zebrafish need to be fed regularly.
<ol>
	<li>Water changes
	<ol>
		<li>To exchange the fish water, thaw 100 mM prednisolone stock and prepare the required amount of 50 &#181;M prednisolone in autoclaved fish water fresh every day of the treatment period. Prepare DMSO containing fish water for the control zebrafish accordingly.</li>
		<li>Take a glass beaker containing a single housed zebrafish in its incubation solution and transfer the solution including the zebrafish into an appropriate container, for example a temporal housing cage. 
		NOTE: Always use separate interim containers for prednisolone and DMSO treated zebrafish, respectively, to make sure DMSO control zebrafish do not get exposed to prednisolone and vice versa.</li>
		<li>Refill the glass beaker with fresh drug containing fish water (300 mL).</li>
		<li>Transfer the zebrafish from the interim container to the glass beaker containing fresh incubation solution using a fish net. Put back the foil to avoid escape of the zebrafish.</li>
		<li>If the time of the experiment exceeds 2 weeks, exchange the glass beakers.</li>
	</ol>
	</li>
	<li>Feeding
	<ol>
		<li>During short treatments (for up to 2 days) do not feed zebrafish. During longer experiments, feed zebrafish. Take special care to avoid pollution of the incubation solution. Thus, feed with Artemia ssp. rather than with flake food, and only on every second day. 
		NOTE: Artemia are regularly used to feed young zebrafish, and should be available in the zebrafish haltering unit.
		<ol>
			<li>Roughly 2 h before the fish water is exchanged, feed zebrafish with Artemia ssp in their glass beakers. To do so, use a plastic pipette and feed 0.5 to 1 mL of hatched Artemia solution per glass beaker.</li>
			<li>Wait for 2 h and exchange the incubation solution with fresh drug containing fish water according to section 4.1.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
5. Analyses of Samples
NOTE: After incubation of injured zebrafish in prednisolone and DMSO containing fish water, respectively, either perform bone mineralization/calcification analyses (5.1 to 5.3) or carry out live imaging of zebrafish under the dissection microscope (5.4)10,11,16. Use live imaging to determine fin regenerate length and to detect differences in reporter gene expression in transgenic zebrafish.
<ol>
	<li>Fixation of tissues of interest
	<ol>
		<li>For fixation of fins and/or skulls prepare 4% paraformaldehyde (PFA) in PBS and freeze at -20 &#176;C for storage. Use freshly prepared 4% PFA in PBS or thaw the required amount before use. Alternatively, 2% PFA in PBS can be used. 
		CAUTION: PFA is toxic and should be handled carefully under a fume hood. Wear protective clothing.</li>
		<li>For fixation of fins prepare a small Petri dish (e.g., 30 mm) with cold 4% PFA in PBS and make sure that the bottom of the dish is fully covered. For fixation of skulls, prepare a little glass jar with lid or a tube with cold 4% PFA in PBS. 
		CAUTION: Handle PFA containing solutions only under the fume hood as it is toxic. Wear protective clothing.</li>
		<li>Harvest the tissue of interest.
		<ol>
			<li>To harvest regenerating fin tissue or fins with fractures anaesthetize the zebrafish according to section 2.1 or euthanize the zebrafish according to section 5.1.5.</li>
			<li>Transfer the anaesthetized zebrafish to the lid of a Petri dish. By using a scalpel or razor blade, harvest the fin regenerate. Make sure to include stump tissue in regenerating fins to facilitate handling of the harvested tissue.</li>
		</ol>
		</li>
		<li>Fix fins flat in the PFA containing Petri dish at 4 &#176;C overnight. Flat fixation avoids curling up of the fin tissue during fixation.
		<ol>
			<li>To fix flat, grab the fin tissue with one fine forceps at the dorsal or ventral edge of the stump and immerse it in the fixation solution. Grab the opposite edge of the stump with a second fine forceps and slightly depress the tissue onto the bottom of the dish.</li>
			<li>Hold for 10 to 20 s before releasing. The fin tissue should not curl up anymore. If it does, repeat.</li>
		</ol>
		</li>
		<li>Euthanize zebrafish in order to harvest injured skull tissue. Prepare a 100 mm diameter Petri dish containing 0.1% MS-222 in fish water. Transfer one zebrafish at a time to the MS-222 containing Petri dish by using a fishing net. When level 6 of anesthesia (medullary collapse) is reached wait at least 10 min to assure death of the specimen12.</li>
		<li>Using a scalpel cut off the head plus a little trunk tissue from the rest of the body. Immerse the harvested tissue in the prepared 4% PFA solution in the glass jar or plastic tube. Fix for 24 h at 4 &#176;C with gentle rocking.</li>
		<li>To store samples long-term after fixation, wash them with PBS (3 x 20 min at RT) and methanol-treat in an increasing methanol series (1x 30%, 1x 50%, 1x 70%, 2x 100% methanol in PBS for 20 min each).
		<ol>
			<li>Store them in 100% methanol at -20 &#176;C. Otherwise, directly proceed with bone staining or other analyses after washing with PBS.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Bone staining techniques I (alizarin red staining) 
	NOTE: This staining is performed according to van Eeden et al. 199617 and Walker &#38; Kimmel 200718 with modifications.
	<ol>
		<li>To perform alizarin red staining on adult fins or skulls, harvest and fix tissue as described in sections 5.1.</li>
		<li>If samples were stored in methanol at -20 &#176;C, rehydrate samples by doing an inverse methanol series (70%, 50% and 30% in PBS, each 1x20 min) and wash twice 20 min with PBS. Wash samples for 5 min minimum with deionized water.</li>
		<li>To do alizarin red staining in skulls, treat samples with 50 mg/mL trypsin in 30% saturated Na2B4O7 at room temperature overnight. Rock samples during this time. Rinse samples in 30% saturated Na2B4O7 solution. Wash samples with deionized water 2 times for 5 min each. This step is not performed for fin tissue.</li>
		<li>Add alizarin red solution (Part B: 0.5% alizarin red S powder, 10% glycerol in deionized water) and rock the samples during incubation at RT. Check for staining after 1, 2, 4 h or overnight.</li>
		<li>To clear samples treat them with a decreasing 1% potassium hydroxide:glycerol series (3:1 overnight, 1:1 overnight, 1:3 overnight).</li>
		<li>Store the samples in a 1:1 solution of 0.1% potassium hydroxide: 80% glycerol and mount them with 80% glycerol for imaging. 
		NOTE: For combined Alcian blue/alizarin red staining treat samples with Alcian blue staining solution (Part A: final 0.02% Alcian blue, 50 mM MgCl2, 70% ethanol) after rehydration (section 5.2.2) for 2 h at RT. Treat fins with a decreasing ethanol series in 50 mM MgCl2 (70%, 50%, 30%, each 1x 15 min) and wash them for a minimum of 1h (3x 20 min) in deionized water. Proceed with alizarin red staining (section 5.2).</li>
	</ol>
	</li>
	<li>Bone staining techniques II (calcein staining) 
	NOTE: To detect calcium deposition in fin fractures and skull injuries, live fish are incubated in calcein containing solution.
	<ol>
		<li>Prepare the required amount of 0.2% calcein in deionized water (pH 7.5). Make sure the pH is neutral. Use 1 M NaOH to adjust the pH.</li>
		<li>Incubate up to 3 zebrafish simultaneously in 100 mL of calcein solution for 20 min in a glass beaker covered with aluminum foil.</li>
		<li>Transfer the zebrafish to a beaker with fish water. Let them swim briefly before transferring them to a new beaker with fish water. Let them swim in this new beaker for 20 min to wash. Take pictures (see section&#160;5.4).</li>
	</ol>
	</li>
	<li>Live imaging of zebrafish
	<ol>
		<li>Use this approach to acquire images of regenerating bone tissue such as in the fin or skull throughout the experiment without sacrificing the zebrafish. Anaesthetize the zebrafish according to section 2.1.</li>
		<li>To acquire images of fin regenerates, transfer the anaesthetized zebrafish onto an agarose-coated Petri dish (see section 2.2.1 and Figure 1B).</li>
		<li>To acquire images of regenerating skull bones, place the zebrafish upright in a sponge (Figure 1D) or place it in an agarose-coated dish that has been prepared to hold the trunk of the zebrafish.</li>
		<li>Acquire images at the desired magnification and resolution using a stereomicroscopic setup.</li>
		<li>Return the zebrafish to a container with fresh or drug containing fish water.</li>
	</ol>
	</li>
</ol>

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M., 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=Retinoic+acid+and+Cyp26b1+are+critical+regulators+of+osteogenesis+in+the+axial+skeleton.">Retinoic acid and Cyp26b1 are critical regulators of osteogenesis in the axial skeleton.</a> Development. 135 (22), 3765-3774 (2008).</li><li>Gioia, 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=The+chaperone+activity+of+4PBA+ameliorates+the+skeletal+phenotype+of+Chihuahua,+a+zebrafish+model+for+dominant+osteogenesis+imperfecta.">The chaperone activity of 4PBA ameliorates the skeletal phenotype of Chihuahua, a zebrafish model for dominant osteogenesis imperfecta.</a> Human Molecular Genetics. 26 (15), 2897-2911 (2017).</li><li>Fiedler, I. A. 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=Severely+impaired+bone+material+quality+in+Chihuahua+zebrafish+resembles+classical+dominant+human+osteogenesis+imperfecta.">Severely impaired bone material quality in Chihuahua zebrafish resembles classical dominant human osteogenesis imperfecta.</a> Journal of Bone and Mineral Research. , (2018).</li><li>Fisher, S., Jagadeeswaran, P., Halpern, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Radiographic+analysis+of+zebrafish+skeletal+defects.">Radiographic analysis of zebrafish skeletal defects.</a> Developmental Biology. 264 (1), 64-76 (2003).</li><li>Hayes, A. J., 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=Spinal+deformity+in+aged+zebrafish+is+accompanied+by+degenerative+changes+to+their+vertebrae+that+resemble+osteoarthritis.">Spinal deformity in aged zebrafish is accompanied by degenerative changes to their vertebrae that resemble osteoarthritis.</a> PLoS One. 8 (9), e75787 (2013).</li><li>Mitchell, R. E., 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=New+tools+for+studying+osteoarthritis+genetics+in+zebrafish.">New tools for studying osteoarthritis genetics in zebrafish.</a> Osteoarthritis Cartilage. 21 (2), 269-278 (2013).</li><li>Fleming, A., Sato, M., Goldsmith, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-throughput+in+vivo+screening+for+bone+anabolic+compounds+with+zebrafish.">High-throughput in vivo screening for bone anabolic compounds with zebrafish.</a> Journal of Biomolecular Screening. 10 (8), 823-831 (2005).</li><li>de Vrieze, E., Zethof, J., Schulte-Merker, S., Flik, G., Metz, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+novel+osteogenic+compounds+by+an+ex+vivo+sp7:luciferase+zebrafish+scale+assay.">Identification of novel osteogenic compounds by an ex vivo sp7:luciferase zebrafish scale assay.</a> Bone. 74, 106-113 (2015).</li></ol>

The authors have nothing to disclose.

Zebrafish have proven useful in skeletal research in many regards. Selected mutants mimic aspects of human disease such as osteogenesis imperfecta or osteoarthritis23,24,25,26,27, and larvae as well as scales are being used to identify bone anabolic compounds in small molecule screens7,28,29. Treatments of zebrafish with drugs that are applied in clinical practice are furthermore ideally suited to study putative adverse effects, for example in bone tissue. In this context, the presence of rapidly regenerating bone is advantageous to investigate the underlying mechanisms of medication-induced bone deficiencies. Here, we present a protocol in which drug treatment with the glucocorticoid prednisolone, a commonly used anti-inflammatory drug with adverse effects in bone tissue, is combined with adult bone regeneration regimes. This protocol has successfully been used to induce immunosuppressive and bone inhibitory effects in regenerative tissues of zebrafish, and can also be adapted for studies on the impact of prednisolone and other drugs in adult tissue such as the spine. The following details should be taken into account when immunosuppressive drug treatments are performed in zebrafish undergoing fin or skull regeneration.
Reproducibility of injury assays
During fin regeneration, regeneration speed (i.e., regrowth of fin tissue, including bone, per time unit) depends very much on the amount of fin tissue that is being resected13. In order to avoid unwanted variability of fin regeneration, make sure to always resect equivalent amounts of fin tissue in all specimens.
Execution of bony fin ray fractures is carried out by slightly pushing an injection needle onto the center of a chosen bony fin ray segment. As bony fin rays consist of 2 opposing hemirays, the amount of pressure used determines whether only one or both hemirays are fractured. We prefer to apply low pressure to fracture only one hemiray, because fracture of both hemirays tends to destabilize the fin ray, which as a result may detach during the following day(s).
Calvarial injuries of the skull by microdrilling should be performed cautiously. If damage is done to the brain (i.e., the cerebellum) postural and locomotory deficits will become apparent by erratic swimming of the specimen15. Zebrafish with cerebellar injury may show adverse reactions to drug treatment and should not be used to study bone regeneration.
Considerations regarding drug treatments
To produce a consistent immunosuppressive and anti-regenerative effect in adult zebrafish we employ a dose of 50 &#181;M prednisolone in fish water. We identified this dose during initial dose-response experiments in larval and adult zebrafish. Thus, dose-response experiments should be carried out to identify the required dose of other immunosuppressive agents that might be applied. For adults, we recommend combining these initial experiments with the fin regeneration regime, as fin regenerate length is a highly sensitive and easy-to-detect readout for tissue alterations.
Immunosuppressive treatment predisposes treated specimens to microbial infections. Therefore, single housing in autoclaved beakers containing autoclaved fish water is important. Although these conditions are more cumbersome to carry out and are not sterile, they help to minimize infection in treated zebrafish. We did not pretreat (&#39;sterilize&#39;) Artemia eggs. However, this might be an additional measure to prevent infection in zebrafish, if necessary. Furthermore, antiseptic substances such as methylene blue (1%) can be added to fish water before use.
Experiments with prednisolone, both short- and long-term, require daily changes of fish water. We have treated adult zebrafish for up to 8 weeks. Treatment of a large number of individuals for such a long time can lead to a certain experimental &#39;burden&#39;, and should be planned carefully. It is pivotal to always have the required amounts of autoclaved fish water and glass ware ready. Although this has not been tested in zebrafish (to our knowledge), implantation of slow release pellets for drugs of interest might represent a valuable alternative for long term drug exposure in zebrafish.
Analyses in transgenic zebrafish
Here, we present 2 methods to stain for mineralization/calcification of bone tissues by alizarin red and calcein staining. Additionally, we show how to image regenerating fin and skull tissue in vivo with the help of a stereomicroscope. The latter technique is very useful, if transgenic zebrafish reporting the number or activity of selected cells, such as osteoblasts or immune cells, are being imaged. For example, before sacrificing injured and prednisolone-treated zebrafish to perform alizarin red staining, fin regenerates or trepanated skulls undergoing repair can be photographed to look for the presence, number and activity of bone forming osteoblasts in the transgenic reporter line Tg (osterix:nGFP)22. Likewise, epidermal wound closure, which occurs within a day, can be visualized in transgenic fish, which express a fluorophore in epidermal tissue. Also, accumulation of immune cells at the site of injury (or their absence in prednisolone treated individuals) can be monitored easily with a stereomicroscope equipped with the required light source and filters.
In sum, the protocol presented here can be used to study effects of immunosuppressive agents and other drugs after systemic administration in zebrafish that are undergoing bone regeneration either in the fin or skull. This will be useful to delineate the pathogenesis of GIO and to investigate the mechanisms underlying successful bone regeneration.

Zebrafish represent a powerful animal model to study vertebrate development and disease. This is due to the fact that they are small animals that breed extremely well and that their genome is fully sequenced and amenable to manipulation1. Other advantages include the option to perform continued live imaging at different stages, including in vivo imaging of adult zebrafish2, and the ability to perform high throughput drug screens in zebrafish larvae3. Additionally, zebrafish possess a high regenerative capacity in a variety of organs and tissues including bone, and thus serve as a useful system to study skeletal disease and repair4,5.
Glucocorticoid-induced osteoporosis (GIO) is a disease that results from long term treatment with glucocorticoids, for example in the course of autoimmune disease treatment such as of asthma or rheumatoid arthritis. GIO develops in about 30% of glucocorticoid-treated patients and represents a major health issue6; therefore, it is important to investigate the impact of immunosuppression on bone tissue in great detail. In recent years a variety of zebrafish models dealing with the pathogenesis of GIO have been developed. Glucocorticoid-mediated bone loss has been induced in zebrafish larvae, for example, which led to the identification of counteractive compounds increasing bone mass in a drug screen7. Furthermore, glucocorticoid-induced bone inhibitory effects have been mimicked in zebrafish scales both in vitro and in vivo8,9. These assays are very convincing approaches, especially when it comes to the identification of novel immunosuppressive and bone anabolic drugs. However, they only partly take into account the endoskeleton and have not been performed in a regenerative context. Thus, they do not allow the investigation of glucocorticoid-mediated effects during rapid modes of adult, regenerative bone formation.
Here, we present a protocol enabling researchers to study glucocorticoid-mediated effects on adult zebrafish bones undergoing regeneration. Injury models include partial amputation of the zebrafish caudal fin, trepanation of the skull, as well as the creation of fin ray fractures (Figure 1A-1C), and are combined with glucocorticoid exposure via incubation (Figure 1E). We have recently used a portion of this protocol to describe the consequences of exposure to prednisolone, one of the commonly prescribed corticosteroid drugs, on adult zebrafish regenerating fin and skull bone10. In zebrafish, prednisolone administration leads to decreased osteoblast proliferation, incomplete osteoblast differentiation and rapid induction of apoptosis in the monocyte/macrophage lineage10. In this protocol, we also describe how fractures can be introduced into single bony fin ray segments11, as this approach may be useful when studying glucocorticoid-mediated effects on bone occuring during fracture repair. The methods presented here will help to further address underlying mechanisms of glucocorticoid action in rapidly regenerating bone and may also be employed in other settings of systemic drug administration in the context of zebrafish tissue regeneration.

<table>
	<tbody>
		<tr>
			<td>Prednisolone</td>
			<td>Sigma-Aldrich</td>
			<td>P6004</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethylsulfoxid (DMSO)</td>
			<td>Sigma-Aldrich</td>
			<td>D8418</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethyl-3-aminobenzoate methanesulfonate (MS-222)</td>
			<td>Sigma-Aldrich</td>
			<td>A5040</td>
			<td></td>
		</tr>
		<tr>
			<td>Blunt forceps</td>
			<td>Aesculap</td>
			<td>BD027R</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>Dumont</td>
			<td>91150-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Braun</td>
			<td>5518059</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Biozym</td>
			<td>840004</td>
			<td></td>
		</tr>
		<tr>
			<td>Injection needle (0.3x13 mm)</td>
			<td>BD Beckton Dickinson</td>
			<td>30400</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro drill</td>
			<td>Cell Point Scientific</td>
			<td>67-1000</td>
			<td>distributed e.g. by Harvard Apparatus</td>
		</tr>
		<tr>
			<td>Steel burrs (0.5 &#181;m diameter)</td>
			<td>Fine Science tools</td>
			<td>19007-05</td>
			<td></td>
		</tr>
		<tr>
			<td>Artemia ssp.</td>
			<td>Sanders</td>
			<td>425GR</td>
			<td></td>
		</tr>
		<tr>
			<td>Pasteur pipette (plastic, Pastette)</td>
			<td>Alpha Labs</td>
			<td>LW4111</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>158127</td>
			<td></td>
		</tr>
		<tr>
			<td>Alizarin red S powder</td>
			<td>Sigma-Aldrich</td>
			<td>A5533</td>
			<td></td>
		</tr>
		<tr>
			<td>Alcian blue 8 GX</td>
			<td>Sigma-Aldrich</td>
			<td>A5268</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcein</td>
			<td>Sigma-Aldrich</td>
			<td>C0875</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Sigma-Aldrich</td>
			<td>T7409</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Leica</td>
			<td>MZ16 FA</td>
			<td>with QIMAGING RETIGA-SRV camera</td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Olympus</td>
			<td>MVX10</td>
			<td>with Olympus DP71 or DP80 camera</td>
		</tr>
	</tbody>
</table>

adult zebrafish injury models to study the effects of prednisolone in regenerating bone tissue

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows within roughly two weeks after injury. Furthermore, zebrafish are able to heal bone rapidly after trepanation of the skull, and repair fractures that can be easily introduced into zebrafish bony fin rays. These injury assays represent feasible experimental paradigms to test the effect of administered drugs on rapidly forming bone. Here, we describe the use of these 3 injury models and their combined use with systemic glucocorticoid treatment, which exerts bone inhibitory and immunosuppressive effects. We provide a workflow on how to prepare for immunosuppressive treatment in adult zebrafish, illustrate how to perform fin amputation, trepanation of calvarial bones, and fin fractures, and describe how the use of glucocorticoids affects both bone forming osteoblasts and cells of the monocyte/macrophage lineage as part of innate immunity in bone tissue.

The protocol presented here has been used repeatedly to induce rapid bone formation in the course of regeneration of the zebrafish fin and skull10,11,16. In combination with the presented method of prednisolone administration, studies on prednisolone&#39;s effects during bone regeneration can be pursued. For example, studies on bone formation and mineralization in the regenerate can be performed. Prednisolone, as other glucocorticoids19,20, leads to overall inhibition of fin regeneration, including bone formation, as detected by alizarin red staining on fixed caudal fin tissue (Figure 3A). Similarly, prednisolone has a delaying effect on (calvarial) skull injury closure, which can be illustrated by alizarin red (Figure 3B) or in vivo calcein staining. In addition, prednisolone exerts a profound anti-inflammatory effect in both fin and skull tissue, by triggering apoptosis in the monocyte/macrophage lineage. Reduced macrophage numbers can be detected by immunohistochemistry on frozen tissue sections, e.g., by using an anti-mcherry antibody in transgenic mpeg1:mCherry zebrafish (Figure 3C)10,21. Similarly, the number, distribution, proliferation and apoptosis of other cell types of interest both in the exo- and the endoskeleton (e.g., vertebrae) can be analyzed with the help of immunohistochemistry.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58429/58429fig1.jpg" /> 
Figure 1: Procedures carried out in zebrafish. (A)&#160;Resection of the caudal fin (amputation) in anaesthetized zebrafish with the help of a scalpel. The red dashed line indicates the amputation level. (B)&#160;Fin ray fracture in anaesthetized zebrafish carried out with an injection needle under the stereomicroscope. The fish is laying on an agarose-coated Petri dish during the procedure. Agarose-coated Petri dishes are also used to acquire images of zebrafish fins during regeneration or fracture repair. (C)&#160;Trepanation of the calvariae (skull injury) performed in anaestetized zebrafish with the help of a microdrill under the stereomicroscope. (D)&#160;Image aquisition of regenerating skull bone after injury. The anaesthetized zebrafish is placed upright in a sponge and images are acquired with the help of a stereomicroscope. The Petri dish is filled with fish water containing 0.02% MS-222. (E)&#160;Incubation of zebrafish in prednisolone or DMSO containing fish water. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58429/58429fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58429/58429fig2.jpg" /> 
Figure 2: Microscopic live view of injuries at 0 h post injury hpi).&#160;(A)&#160;Amputated caudal fin. Scale bar = 200 &#181;m. (B)&#160;Fractured fin ray. The fracture is indicated by the red arrow. Scale bar = 100 &#181;m. (C)&#160;Trepanated skull. The injury site is indicated by the white arrowhead. Scale bar = 500 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58429/58429fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58429/58429fig3.jpg" /> 
Figure 3: Representative results of prednisolone treatment in adult zebrafish.&#160;(A)&#160;Alizarin red stained fin regenerates at 14 days post amputation (dpa) and days of treatment (dt). Prednisolone treated fin regenerates are shorter (not shown), and a smaller domain of alizarin red positive bone matrix is detected. Scale bar = 500 &#181;m. (B)&#160;Alizarin red stained skulls at 7 days post injury (dpi) and dt. Scale bar = 100 &#181;m. This figure has been modified with permission10. (C)&#160;Cryosection view of uninjured skull and brain tissue of treated (prednisolone) and untreated (DMSO) Tg (mpeg1:mCherry) x Tg (osterix:nGFP) transgenic reporter fish, in which macrophages (innate immune cells) are labeled in red 21 and bone forming osteoblasts are labeled in green22. The number of macrophages significantly drops in prednisolone treated zebrafish. Immunohistochemistry was performed as described in Geurtzen et al.10. Nuclei are labeled in blue (DAPI). Scale bar = 20 &#181;m. BF: bright field, epid: epidermis, bn: bone. This figure has been modified with permission10. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58429/58429fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Adult Zebrafish Injury Models to Study the Effects of Prednisolone in Regenerating Bone Tissue at JoVE.com

Adult Zebrafish Injury Models to Study the Effects of Prednisolone in Regenerating Bone Tissue

Here, we describe 3 adult zebrafish injury models and their combined use with immunosuppressive drug treatment. We provide guidance on imaging of regenerating tissues and on detecting bone mineralization therein.

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows ...

adult-zebrafish-injury-models-to-study-effects-prednisolone

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

8.8K Views.  TU Dresden. Here, we describe 3 adult zebrafish injury models and their combined use with immunosuppressive drug treatment. We provide guidance on imaging of regenerating tissues and on detecting bone mineralization therein.

Video: Adult Zebrafish Injury Models to Study the Effects of Prednisolone in Regenerating Bone Tissue

Kidney Regeneration in Adult Zebrafish by Gentamicin Induced Injury

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is the nephron. Mammals are able to repair existing nephrons after injury, but lose the ability to form new nephrons soon after birth. In contrast to mammals, adult fish produce new nephrons (neonephrogenesis) throughout their lives in response to growth requirements or injury. Recently, lhx1a has been shown to mark nephron progenitor cells in the adult zebrafish kidney, however mechanisms controlling the formation of new nephrons after injury remain unknown. Here we show our method for robust and reproducible injury in the adult zebrafish kidney by intraperitoneal (i.p.) injection of gentamicin, which uses a noninvasive visual screening process to select for fish with strong but nonlethal injury. Using this method, we can determine optimal gentamicin dosages for injury and go on to demonstrate the effect of higher temperatures on kidney regeneration in zebrafish.

Here we present a reliable method to study adult kidney regeneration by inducing acute kidney injury by gentamicin injection. We show that injury is dependent on gentamicin dosage and environmental temperature using in situ hybridization to label lhx1a+ developing new nephrons.

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is ...

Mechanical Vessel Injury in Zebrafish Embryos

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes. External development and optical transparency make these embryos especially amenable to microscopy, and numerous transgenic lines that label specific cell types with fluorescent proteins are available, making the zebrafish embryo an ideal system for visualizing the interaction of vascular, hematopoietic, and other cell types during injury and repair in vivo. Forward and reverse genetics in zebrafish are well developed, and pharmacological manipulation is possible. We describe a mechanical vascular injury model using micromanipulation techniques that exploits several of these features to study responses to vascular injury including hemostasis and blood vessel repair. Using a combination of video and timelapse microscopy, we demonstrate that this method of vascular injury results in measurable and reproducible responses during hemostasis and wound repair. This method provides a system for studying vascular injury and repair in detail in a whole animal model.

This article describes a method for creating a mechanical vessel injury in zebrafish embryos. This injury model provides a platform for studying hemostasis, injury-related inflammation, and wound healing in an organism ideally suited for real-time microscopy.

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes.  External development ...

Use of the TetON System to Study Molecular Mechanisms of Zebrafish Regeneration

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough understanding of the molecular and cellular mechanisms involved requires experimental tools that allow for inducible, tissue-specific manipulation of gene expression or signaling pathways. Therefore, we and others have recently adapted the TetON system for use in zebrafish. The TetON system facilitates temporally and spatially-controlled gene expression and we have recently used this tool to probe for tissue-specific functions of Wnt/beta&#8211;catenin signaling during zebrafish tail fin regeneration. Here we describe the workflow for using the TetON system to achieve inducible, tissue-specific gene expression in the adult regenerating zebrafish tail fin. This includes the generation of stable transgenic TetActivator and TetResponder lines, transgene induction and techniques for verification of tissue-specific gene expression in the fin regenerate. Thus, this protocol serves as blueprint for setting up a functional TetON system in zebrafish and its subsequent use, in particular for studying fin regeneration.

Here we outline the workflow for using the TetON system to achieve tissue-specific gene expression in the adult regenerating zebrafish tail fin.

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough ...

Protocols for Analyzing the Role of Paneth Cells in Regenerating the Murine Intestine using Conditional Cre-lox Mouse Models

The epithelial surface of the mammalian intestine is a dynamic tissue that renews every 3 - 7 days. Understanding this renewal process identified a population of rapidly cycling intestinal stem cells (ISCs) characterized by their expression of the Lgr5 gene. These are supported by a quiescent stem cell population, marked by Bmi-1 expression, capable of replacing them in the event of injury. Investigating the interactions between these populations is crucial to understanding their roles in disease and cancer. The ISCs exist within crypts on the intestinal surface, these niches support the ISC in replenishing the epithelia. The interaction between active and quiescent ISCs likely involves other differentiated cells within the niche, as it has previously been demonstrated that the &#8216;&#8216;stemness&#8217;&#8217; of the Lgr5 ISC is closely tied to the presence of their neighboring Paneth cells. Using conditional cre-lox mouse models we tested the effect of deleting the majority of active ISCs in the presence or absence of the Paneth cells. Here we describe the techniques and analysis undertaken to characterize the intestine and demonstrate that the Paneth cells play a crucial role within the ISC niche in aiding recovery following substantial insult.

Protocols for Analyzing the Role of Paneth Cells in Regenerating the Murine Intestine using Conditional Cre-lox Mouse Models

Intestinal epithelial stem cells (ISCs) are intermingled with Paneth cells. These cells are differentiated progeny of the ISC, which support the ISCs and provide antibacterial protection. Here we demonstrate how we used transgenic conditional mouse models to establish that Paneth cells play a crucial role in maintaining the intestinal epithelia.

The epithelial surface of the mammalian intestine is a dynamic tissue that renews every 3 - 7 days. Understanding this renewal process identified a ...

Combining Intravital Fluorescent Microscopy (IVFM) with Genetic Models to Study Engraftment Dynamics of Hematopoietic Cells to Bone Marrow Niches

Increasing evidence indicates that normal hematopoiesis is regulated by distinct microenvironmental cues in the BM, which include specialized cellular niches modulating critical hematopoietic stem cell (HSC) functions1,2. Indeed, a more detailed picture of the hematopoietic microenvironment is now emerging, in which the endosteal and the endothelial niches form functional units for the regulation of normal HSC and their progeny3,4,5. New studies have revealed the importance of perivascular cells, adipocytes and neuronal cells in maintaining and regulating HSC function6,7,8. Furthermore, there is evidence that cells from different lineages, i.e. myeloid and lymphoid cells, home and reside in specific niches within the BM microenvironment. However, a complete mapping of the BM microenvironment and its occupants is still in progress.
Transgenic mouse strains expressing lineage specific fluorescent markers or mice genetically engineered to lack selected molecules in specific cells of the BM niche are now available. Knock-out and lineage tracking models, in combination with transplantation approaches, provide the opportunity to refine the knowledge on the role of specific &#34;niche&#34; cells for defined hematopoietic populations, such as HSC, B-cells, T-cells, myeloid cells and erythroid cells. This strategy can be further potentiated by merging the use of two-photon microscopy of the calvarium. By providing in vivo high resolution imaging and 3-D rendering of the BM calvarium, we can now determine precisely the location where specific hematopoietic subsets home in the BM and evaluate the kinetics of their expansion over time. Here, Lys-GFP transgenic mice (marking myeloid cells)9 and RBPJ knock-out mice (lacking canonical Notch signaling)10 are used in combination with IVFM to determine the engraftment of myeloid cells to a Notch defective BM microenvironment.

Combining Intravital Fluorescent Microscopy IVFM with Genetic Models to Study Engraftment Dynamics of Hematopoietic Cells to Bone Marrow Niches

Intravital fluorescence microscopy (IVFM) of the calvarium is applied in combination with genetic animal models to study the homing and engraftment of hematopoietic cells into bone marrow (BM) niches.

Increasing evidence indicates that normal hematopoiesis is regulated by distinct microenvironmental cues in the BM, which include specialized cellular ...

In Vitro and In Vivo Models to Study Corneal Endothelial-mesenchymal Transition

Corneal endothelial cells (CECs) play a crucial role in maintaining corneal clarity through active pumping. A reduced CEC count may lead to corneal edema and diminished visual acuity. However, human CECs are prone to compromised proliferative potential. Furthermore, stimulation of cell growth is often complicated by gradual endothelial-mesenchymal transition (EnMT). Therefore, understanding the mechanism of EnMT is necessary for facilitating the regeneration of CECs with competent function. In this study, we prepared a primary culture of bovine CECs by peeling the CECs with Descemet's membrane from the corneal button and demonstrated that bovine CECs exhibited the EnMT process, including phenotypic change, nuclear translocation of &#946;-catenin, and EMT regulators snail and slug, in the in vitro culture. Furthermore, we used a rat corneal endothelium cryoinjury model to demonstrate the EnMT process in vivo. Collectively, the in vitro primary culture of bovine CECs and in vivo rat corneal endothelium cryoinjury models offers useful platforms for investigating the mechanism of EnMT.

In Vitro and In Vivo Models to Study Corneal Endothelial-mesenchymal Transition

A primary culture of bovine corneal endothelial cells was used to investigate the mechanism of corneal endothelial-mesenchymal transition. Furthermore, a rat corneal endothelium cryoinjury model was used to demonstrate corneal endothelial-mesenchymal transition in vivo.

Corneal endothelial cells (CECs) play a crucial role in maintaining corneal clarity through active pumping. A reduced CEC count may lead to corneal edema ...

Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics

Skeletal morphogenesis occurs through tightly regulated cell behaviors during development; many cell types alter their behavior in response to mechanical strain. Skeletal joints are subjected to dynamic mechanical loading. Finite element analysis (FEA) is a computational method, frequently used in engineering that can predict how a material or structure will respond to mechanical input. By dividing a whole system (in this case the zebrafish jaw skeleton) into a mesh of smaller 'finite elements', FEA can be used to calculate the mechanical response of the structure to external loads. The results can be visualized in many ways including as a 'heat map' showing the position of maximum and minimum principal strains (a positive principal strain indicates tension while a negative indicates compression. The maximum and minimum refer the largest and smallest strain). These can be used to identify which regions of the jaw and therefore which cells are likely to be under particularly high tensional or compressional loads during jaw movement and can therefore be used to identify relationships between mechanical strain and cell behavior. This protocol describes the steps to generate Finite Element models from confocal image data on the musculoskeletal system, using the zebrafish lower jaw as a practical example. The protocol leads the reader through a series of steps: 1) staining of the musculoskeletal components, 2) imaging the musculoskeletal components, 3) building a 3 dimensional (3D) surface, 4) generating a mesh of Finite Elements, 5) solving the FEA and finally 6) validating the results by comparison to real displacements seen in movements of the fish jaw.

Finite Element Analysis is a frequently used tool to investigate the mechanical performance of structures under load. Here we apply its use to modeling the biomechanics of the zebrafish jaw.

Skeletal morphogenesis occurs through tightly regulated cell behaviors during development; many cell types alter their behavior in response to mechanical ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Histological Analyses of Acute Alcoholic Liver Injury in Zebrafish

Alcoholic Liver Disease (ALD) refers to damage to the liver due to acute or chronic alcohol abuse. It is among the leading causes of alcohol-related morbidity and mortality and affects more than 2 million people in the United States. A better understanding of the cellular and molecular mechanisms underlying alcohol-induced liver injury is crucial for developing effective treatment for ALD. Zebrafish larvae exhibit hepatic steatosis and fibrogenesis after just 24 h of exposure to 2% ethanol, making them useful for the study of acute alcoholic liver injury. This work describes the procedure for acute ethanol treatment in zebrafish larvae and shows that it causes steatosis and swelling of the hepatic blood vessels. A detailed protocol for Hematoxylin and Eosin (H&#38;E) staining that is optimized for the histological analysis of the zebrafish larval liver, is also described. H&#38;E staining has several unique advantages over immunofluorescence, as it marks all liver cells and extracellular components simultaneously and can readily detect hepatic injury, such as steatosis and fibrosis. Given the increasing usage of zebrafish in modeling toxin and virus-induced liver injury, as well as inherited liver diseases, this protocol serves as a reference for the histological analyses performed in all these studies.

This protocol describes histological analyses of the livers from zebrafish larvae that have been treated with 2% ethanol for 24 h. Such acute ethanol treatment results in hepatic steatosis and swelling of the hepatic vasculature.

Alcoholic Liver Disease (ALD) refers to damage to the liver due to acute or chronic alcohol abuse. It is among the leading causes of alcohol-related ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...