This work was supported by the German Research Foundation (DFG) Transregio 67 (project 387653785), the DFG SPP 2084 &#181;Bone (project KN 1102/2-1) to FK. This work was supported by the Light Microscopy Facility (DFG project 413875620), a Core Facility of the CMCB Technology Platform at TU Dresden. The work at the TU Dresden was co-financed with tax revenues based on the budget agreed by the Saxon Parliament (&#39;Landtag&#39;).

The study protocol received approval from the Landesdirektion Sachsen, permit numbers 25-5131/564/2, DD25-5131/450/4, 25-5131/496/56, DD25.1-5131/354/87. The zebrafish strains used were maintained according to national law and under standardized conditions as previously described27,28. The details of all the reagents and the equipment used in the study are listed in the Table of Materials.
1. Preparation of materials and solutions
<ol>
	<li>Maintain the embryos in 1x E3 media. Prepare the 1x E3 media by diluting 170 mL of a 60x E3 stock in 10 L of deionized water and adding 2 mL 0.1% methylene blue.
	<ol>
		<li>Prepare the 60x E3 media stock by mixing 17.2 g of NaCl, 0.76 g of KCl, 2.90 g of CaCl2&#183;2 H2O, 4.90 g of MgSO4&#183;7 H2O in 1 L of deionized autoclaved water.</li>
	</ol>
	</li>
	<li>For the larvae embedding, prepare aliquots of 1% low melting point agarose (LMA) in advance and keep them at room temperature.
	<ol>
		<li>To prepare aliquots, dissolve LMA in 1x E3 media, e.g., 500 mg of LMA in 50 mL of 1x E3 in an Erlenmeyer flask, heat the solution in the microwave until the agarose completely dissolves, and dispense it into 1 mL aliquots in 1.5 mL tubes.</li>
	</ol>
	</li>
</ol>
2. Zebrafish breeding and embryo collection
NOTE: Double transgenic zebrafish larvae reporting the presence of committed osteoblasts [osterix:nGFP = Tg(Ola.Sp7:NLS-GFP)zf132]29 and macrophages [mpeg:mCherry = Tg(mpeg:mCherry)gl23] were used here30 (Figure 1A).
<ol>
	<li>Set up parental zebrafish with the desired single transgenes in standard mating cages the evening before mating. 
	NOTE: Standard mating cages allow the eggs to pass to a lower compartment where the parental zebrafish cannot reach them.
	<ol>
		<li>Keep males and females separated until mating using a divider in the mating cage. Remove the divider in the morning of the next day to initiate mating.</li>
	</ol>
	</li>
	<li>At noon, at the latest, remove parental zebrafish from the breeding tanks and collect the embryos, straining the water from the mating cage through a sieve. 
	NOTE: In case the exact age of the larvae is decisive, harvest embryos in regular intervals of approximately 30 min. Clutch mates should be used in experiments with different experimental groups.</li>
	<li>Place the embryos in a 100 mm diameter dish containing 1x E3 media and keep them in an incubator at 28 &#176;C31.</li>
	<li>Remove unfertilized eggs and dead embryos from the dish using a plastic pipette with the help of a stereomicroscope and change 1x E3 media daily.</li>
	<li>Before the experiment, sort larvae positive for the reporter transgenes using a fluorescence stereomicroscope.</li>
</ol>
3. Larvae embedding in low-melting agarose
<ol>
	<li>Melt LMA aliquots in a thermoblock or water bath at 65 &#176;C. Once the LMA is melted, keep it at 42 &#176;C until embedding (Figure 1B). Add 50 &#181;L of 0.4% MS-222 to the melted 1 mL of LMA aliquot to achieve a 0.02% MS-222 concentration for anesthesia.</li>
	<li>Anaesthetize 4 dpf (days post fertilization) larvae (or older larvae) in 0.02% MS-222 E3 media.</li>
	<li>Transfer the&#160;anesthetized larvae to a glass-bottom microwell dish using a 3 mL plastic Pasteur pipette and remove the excess E3 media with the same pipette (Figure 1B). Avoid drying up of the larvae. 
	NOTE: More than one larva can be embedded in a dish simultaneously; it is recommended to embed 3 to 5 larvae to allow sufficient time for placement.</li>
	<li>Take the melted LMA agarose from the water bath/thermoblock and add a (cooling) drop of LMA to the dish, covering the glass bottom where the larvae were positioned. Pay attention that the LMA is not too hot but still liquid.</li>
	<li>Then, place the larvae into the desired lateral position before the agarose solidifies. Here, ensure a lateral position in which the opercle is as close as possible to the glass bottom of the dish as a preparation for the inverse microscope setup (Figure 1B).</li>
	<li>Let the agarose solidify and then add E3 media (containing a maximum of 0.02% MS-222) to the dish to avoid drying of the agarose (Figure 1B).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67279/67279fig01.jpg" /> 
Figure 1: Schematic representation of the embedding procedure. (A) Double transgenic Tg(osterix:nGFP; mpeg1:mCherry)29,30&#160;zebrafish larvae with green labeled opercular osteoblasts and macrophages labeled in red were used. (B) The workflow of the embedding procedure, as described in step 3 of the protocol. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67279/67279fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. In vivo imaging of the larvae and laser ablation
<ol>
	<li>To laser-ablate osteoblasts in the developing opercle of transgenic zebrafish Tg(osterix:nGFP; mpeg1:mCherry)29,30, use an inverted two-photon laser-scanning microscope with a 25x/0.95 water objective or an equivalent setup.</li>
	<li>Place the dish with the embedded larvae onto the microscopic stage at the inverted microscope and position the desired region for ablation in the center of the field of view. The opercle can be easily identified with the help of the GFP fluorescence signal.</li>
	<li>Once the opercle is focused and centered in the field of view of the microscope, choose the z-stack settings (often 1-2 &#181;m z-intervals are used) to image the entire breadth of the opercle.&#160;At the same time, choose the area in a specific z-plane to be ablated (usually in the posterior zone of the opercle). The two-photon laser ablation is only exerted in this plane. 
	NOTE: Higher resolution in the z dimension will lead to longer imaging times but will enable volumetric analyses of the acquired data later.&#160;</li>
	<li>Image the opercle area of interest before ablation. For the GFP detection, use a white-light laser at 488 nm and a hybrid detector ranging from 495 nm to 550 nm or equivalent. Image the macrophages using 555 nm (white-light laser) excitation, with a mCherry detection range of 595 to 780 nm on a hybrid detector (or equivalent).
	<ol>
		<li>Use the image format 512 x 512 for a good quality/time ratio (other formats can be used if higher resolution or alternatively higher speed are desired). 
		NOTE: A temperature-controlled incubator installed at the microscope can be used and adjusted, e.g., to 28 &#176;C.</li>
	</ol>
	</li>
	<li>Draw a circular region with a diameter of 25 &#181;m (other shapes and sizes can be selected depending on the experiment) in a selected 2D plane, focusing the opercle using the ROI tool.</li>
	<li>To perform the ablation, apply ablation power (measured without objective) at&#160;450 mW and expose the selected area to the two-photon laser at a scan speed of 400 Hz and a total duration of 10 s. During ablation, the area can be visualized using a hybrid detector in the range of 401-443 nm or equivalent.</li>
	<li>After ablation, confirm the ablation structurally by imaging again the same z-stack with the same settings as used prior to ablation. The ablated area does not contain fluorescent cells anymore.</li>
</ol>
5. Recovery of larvae and imaging to monitor immune cell recruitment
<ol>
	<li>If the imaging of immune cell recruitment needs to be performed with a delay, recover the larvae from the imaging and anesthesia. Remove the larvae carefully from the LMA using forceps or a dissection needle.
	<ol>
		<li>First, remove a layer of agarose covering the larvae. Second, remove the agarose in direct contact with the larvae. Omit this step if immediate imaging of immune cell recruitment is taking place.</li>
	</ol>
	</li>
	<li>Place the larvae back in a dish with pure E3 media and check their recovery. Recovery is evident by swimming of the larvae.</li>
	<li>At 4 or 6 hpl (hours post-lesion), image the opercle area again using the same settings at the same microscope to check for the effects of the ablation on macrophage recruitment. To do so, repeat the embedding (step 3) and imaging as performed before (step 4).</li>
	<li>After imaging, remove the larvae from the LMA. Sacrificing of the larvae can either be performed by incubation in ice water or with an overdose of MS-222 in E3, depending on the legal requirements. Specimens can then be used for follow-up studies or be disposed of.</li>
</ol>
6. Image analysis and statistical analysis
<ol>
	<li>Process the images using an image processing software such as Fiji32. Depending on the z-resolution of the acquired data, analyses can be performed in 2D or 3D.</li>
	<li>Quantify the macrophages manually by either including cells that are clearly separate, such as by discerning the main cell bodies, or alternatively quantifying cells with a nuclear label or by measuring the area in case cells are overlapping (next step).</li>
	<li>For quantification of the area, create a mask encompassing the macrophages. To do that, use the Image &#62; Adjust &#62; Threshold tool and create an ROI (Analyze &#62; Tools &#62; ROI Manager &#62; Add) in Fiji. Exclude small particles in the background for quantification.
	<ol>
		<li>Then, measure the area (Analyze &#62; Measure). Consider whether areas of overlapping cellular fluorescence need to be included in the measurement. 
		NOTE: Here, the circular ablation area was omitted from the area measurements since recruited macrophages cannot be distinguished in this area.</li>
	</ol>
	</li>
	<li>Generate graphs plotting the measurement results using suitable software. To compare the data before and after ablation, perform an appropriate statistical test (here, a paired one-tailed t-test was performed). 
	NOTE: The figures can be prepared using photo editing and graphic design software.</li>
</ol>

Laser Ablation Method to Study Bone Regeneration in Zebrafish Larvae

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laser-targeted+ablation+of+the+zebrafish+embryonic+ventricle:+a+novel+model+of+cardiac+injury+and+repair.">Laser-targeted ablation of the zebrafish embryonic ventricle: a novel model of cardiac injury and repair.</a> Int J Cardiol. 168 (4), 3913-3919 (2013).</li></ol>

The authors have nothing to disclose.

Laser ablation has been applied in various disciplines of biological research. In particular, it has been useful as a method to study tissue regeneration10,11,12. For example, the recruitment and phenotype changes of innate immune cells were recently analyzed over time using UV laser or two-photon laser ablation assays, along with the recovery of osteoblasts at the laser ablation site10. Live imaging of immune cell recruitment was carried out up to 12 h post-lesion (hpl); however, longer imaging periods can be envisioned with appropriate adjustments in anesthesia and approval from the respective authorities. In Geurtzen et al., analyses were carried out up to several days post-laser ablation and helped to define the inflammatory milieu at the ablation site, as well as to establish the supportive role of macrophages in bone regeneration10. Many other applications, such as studying the role of reactive oxygen species&#160;release33 upon cell death in bone tissue, are conceivable.
One of the advantages of using two-photon laser-mediated cell ablation is the possibility of combining it with confocal imaging. The software used here (see Table of Materials) can be set to a mode (Live Data Mode) in which imaging and ablation occur consecutively, speeding up the process. Another advantage of the technology is the accuracy of the ablation. The size and shape of the ablation can be easily adjusted in the software in a very precise manner. In the experimental setting presented here, a circular area with a diameter of 25 &#181;m was chosen for ablation. However, other ablation experiments in Drosophila employed 25 &#181;m linear ablations in order to reduce tissue tension in the embryonic epithelium by cutting neighbour-neighbour cellular junctions34. This accuracy allows for the ablation of a specific and controlled number of cells, single cells, or even cell membranes.
Notably, the type of lesion performed with a two-photon laser does not resemble injuries and cell death that naturally occur, which needs to be kept in&#160;mind in regeneration research but also other&#160;disciplines. In particular, laser ablations create very focused lesions by completely destroying (burning) the cells in the region of interest20 with minimal damage in surrounding tissues, while mechanical or other types of injuries tend to affect several tissue types and layers at a broader scale. This can lead to manifold consequences involving different types of cell death (e.g., necrosis, apoptosis, autophagy)35 and different regenerative outcomes. Moreover, reactions to laser ablation might differ in cellular and molecular terms. For example, it was shown that although cells surrounding the laser lesion site might lose plasma membrane integrity, they can survive the damage and restore integrity36. Laser ablation also leads to areas of the damaged extracellular matrix, which may result from &#34;thermal explosions&#34; in the very center of the applied laser beam and also strong heat exposure in the directly neighboring cells10,37. Nevertheless, cells surrounding laser ablation sites show an increased number of apoptotic bodies, as shown for laser-ablated heart tissue38, and an immune response is triggered, as shown here. In addition to the described differences, another limitation of laser-assisted ablation assays might be the depth at which the injury can be performed and the limited size of specimens. Structures hidden in thick specimens may not be appropriately ablated, and imaging will be difficult. This, however, will depend very much on the microscope and laser used, as well as the objective and embedding.
Embedding the larvae in a way so that the opercle is placed in close proximity to the glass bottom of the dish is essential in case an inverted setup is used, as in the presented protocol. Moreover, it is very important to test the temperature of the LMA before applying it to cover the larvae (step 3.4) since hot agarose will damage the larvae. To avoid that, take the LMA aliquot from the thermoblock early, e.g., while removing the leftover E3 media from the glass-bottom dish, and test the temperature, such as on the skin of the wrist. The LMA is supposed to be warm but not hot.
Once the larvae are covered with LMA, it is important to quickly put them into the correct position before the LMA starts to solidify. The lateral position is the easiest to achieve; in the presented study, the desired position was lateral with a slight tilt to the ventral side. In case the LMA solidifies before the larvae are positioned correctly, it is possible to remove the LMA (like in step 5.1) and repeat the embedding. Likewise, it is important that the area that is to be imaged (and laser ablated) is as close as possible to the glass bottom of the dish for maximum image quality and cell ablation, although this will also depend on the working distance of the objective.
It is possible to embed more than one larvae in the same dish for imaging. This will save time and make better use of the LMA. The number of larvae that can be embedded will depend on the experience of the experimenter, but 3-5 larvae are recommended.
Laser ablation can also be performed with upright microscopic setups employing dipping lenses. In this case, it is important to embed the larvae with a slightly ventral view (from above) to enable good imaging quality and laser ablation success of the opercle. In addition, only a thin layer of LMA should cover the larvae to allow good excitation and detection of fluorescent signals.
In this protocol, a two-photon laser-mediated ablation setup for osteoblasts is described as a methodology to perform lesions in developing bones, specifically in the opercle. This method can be adapted to ablate cells in other bones of developing zebrafish. Moreover, it is generally a useful approach to study the regeneration of injured tissues11,12. The ease of performing ablations and tissue cuts of different sizes and shapes allows its use in different experimental setups and research fields15,17,18,19,21,34. Altogether, two-photon laser-mediated ablation is a very adaptable method - for different tissues, organisms, and even&#160;in vitro settings - and it represents an&#160;important&#160;tool in biological research.

Zebrafish regenerate diverse organs such as the retina, brain, heart, and pancreas1. In addition, zebrafish regenerate skeletal elements, which is why they have been used to study the regeneration of fins, scales, and calvariae (skull caps)2. Different experimental paradigms have been used to study tissue and bone regeneration, such as fin resection (amputation), fin fracture, skull trepanation3,4, cryoinjury5,6, or genetic ablation7,8,9. Recently, laser ablation approaches have been widely used in zebrafish to study the healing and regeneration response after injury10,11,12,13,14.
Laser-mediated ablation has been used in different areas of biological research, such as mechanobiology, developmental biology, regeneration research, and tumor surgery10,11,12,15,16,17,18,19. There are various methodologies of laser ablation, such as using YAG (yttrium aluminum garnet), UV (ultraviolet), or 2p (two-photon) lasers20. Laser ablation allows the removal of single cells or larger portions of tissues in a very accurate manner, enabling the study of different processes, such as the response of the immune system to tumor ablation21 or during zebrafish opercle regeneration. In the latter example, ablation was confirmed by the disappearance of the nuclear and cytoplasmic fluorophore signal and necrosis staining10,22.
It is well known that the innate immune response and its regulated kinetics are essential for appropriate tissue regeneration. Neutrophils and macrophages are the first cells to be recruited to the injury site, where they perform different roles, such as cytokine and growth factor release, cellular debris removal, and extracellular matrix remodeling23. This recruitment and subsequent macrophage functionalization have also been observed in amputated zebrafish fins24 and the developing opercle, which was subjected to laser &#39;nanodissection&#39; leading to osteoblast ablation10. In the latter experiment, recovery of the osteoblast number, normal opercle development, and recruitment of innate immune cells (neutrophils, macrophages, and osteoclast-like cells) to the injured area was observed after UV laser and two-photon laser-mediated ablation10. Experiments in zebrafish in which the immune system was pharmacologically suppressed by using glucocorticoids showed an impairment of regeneration upon misregulated immunity, supporting the functional role of the immune system in tissue repair3,10,25,26.
Here, a two-photon laser-mediated ablation methodology to study the biology of bone regeneration in developing zebrafish bones is described. In particular, the effect of the osteoblast ablation approach in the opercle is shown in terms of the immune response, which is investigated by monitoring the recruitment of macrophages to the ablation site.

<table><tbody><tr><td>Calcium chloride dihydrate, CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Carl Roth</td><td>5239.1</td><td></td></tr><tr><td>Cell Culture Dish, PS, 100/20 mm</td><td>Greiner Bio-one</td><td>664160</td><td></td></tr><tr><td>Dumont #55 Foceps</td><td>FST</td><td>11295-51</td><td>Tip shape straight, 11 cm, 0.05 x 0.02 mm</td></tr><tr><td>Falcon 6-well plate</td><td>Corning</td><td>353502</td><td></td></tr><tr><td>Glass-bottom microwell dish</td><td>MatTek</td><td>P35G-1.5-14-C</td><td>35 mm Dish, No. 1.5 Coverslip, 14 mm Glass Diameter, Uncoated</td></tr><tr><td>Insight X3 multiphoton laser</td><td>Spectra-Physics</td><td></td><td></td></tr><tr><td>Leica Application Suite</td><td>LAS X, Leica Microsystems</td><td></td><td></td></tr><tr><td>Low melting agarose&amp;nbsp;</td><td>Biozyme</td><td>840101</td><td>Biozym Plaque Agarose</td></tr><tr><td>Magnesium sulfate heptahydrate, MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>M5921</td><td></td></tr><tr><td>Mating cages</td><td></td><td></td><td>many varieties, e.g. Tecniplast</td></tr><tr><td>Methyleneblue</td><td>Carl Roth</td><td>A514.1</td><td></td></tr><tr><td>MS-222</td><td>SIGMA Aldrich</td><td>A5040</td><td></td></tr><tr><td>Potassium Chloride, KCl</td><td>PanReac AppliChem</td><td>131494</td><td></td></tr><tr><td>Sodium chloride, NaCl</td><td>Carl Roth</td><td>3957.1</td><td></td></tr><tr><td>SP8 FALCON&amp;nbsp;</td><td>Leica Microsystems</td><td></td><td>Equipped with a Insight X3 multiphoton laser and Leica Application Suite software</td></tr><tr><td>Stainless Steel Dissect Needle</td><td>Bochem</td><td>12010</td><td>140 mm</td></tr><tr><td>Stereo Microscope System SZX16</td><td>Olympus</td><td></td><td>Equipped with a LED illumination base SZX2-ILLTQ</td></tr><tr><td>Thermostatic Cabinets TS - WTW</td><td>xylem</td><td>TS 608/2-i</td><td>For incubation (embryo)</td></tr><tr><td>Transfer pipette, 3.5 mL</td><td>SARSTEDT</td><td>861171</td><td>155 x 15 mm, LD-PE, transparent</td></tr><tr><td>Zeiss SteREO Discovery.V12 version 4.7.1.0.</td><td>Zeiss</td><td></td><td>Equipped with Axiocam MRm camera and AxioVision sofware</td></tr></tbody></table>

two-photon laser-mediated ablation of osteoblasts in developing zebrafish larvae

Zebrafish (Danio rerio) have an outstanding capacity to regenerate different organs and appendages. Bone regeneration in zebrafish has been studied using different methods such as fin amputation, scale plucking, skull trepanation, and microscopic approaches. Using a confocal laser scanning setup equipped with a two-photon laser, a laser ablation method was developed as a lesion paradigm to ablate bone-forming cells (osteoblasts) in the developing opercle of zebrafish larvae. The method described here allows the ablation of cells in a precise manner, as the area, shape, and depth can be finely adjusted. In addition, this method allows imaging of the area before and just after the ablation, so that short-term effects of the injury can be analyzed. In this experimental setup, the immune response after ablation of osteoblasts in the injured area was studied. An increase in the recruitment of macrophages was observed after ablation, indicating the relevance of their presence during bone regeneration.

Laser ablation was performed as indicated in the protocol above. The GFP signal of the osteoblasts in the ablated area disappeared instantaneously after ablation. To study the response of the osteoblast ablation in terms of the immune response, the presence of macrophages in 6 dpf larvae before and at 6 hpl was imaged. Before ablation, very few macrophages were observed in the opercle area10 (Figure 2). At 6 hpl, a strong accumulation of macrophages in the ablated opercle region and an increased number of macrophages in the field of view, including the opercle and the area surrounding the opercle, was detected10 (Figure 2).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67279/67279fig02v2.jpg" /> 
Figure 2: Macrophage recruitment at 6 hpl in 6 dpf zebrafish larvae. Opercle area without ablation and at 6 hpl. Opercular osteoblasts are labeled in green, macrophages in magenta, and the dashed circle indicates the laser-ablated area. Scale bar: 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67279/67279fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Macrophage recruitment was also analyzed in 4 dpf larvae with much smaller opercles at 4 hpl observing similar results (Figure 3). Quantification of the macrophage number (outside the immediate laser ablation site) showed an increase&#160;within the investigated&#160;time frame (4.00 &#177; 2.37 macrophages before&#160;ablation versus 7.00 &#177; 2.76 macrophages at 4&#160;hpl) (Figure 3B). Likewise,&#160;macrophage area (measured in &#181;m2, 785.8 &#181;m2 &#177; 723.1 &#181;m2 before ablation versus 1553 &#181;m2 &#177; 869.9 &#181;m2 at 4 hpl) increased (Figure 3C). As macrophages are sometimes difficult to tell apart, measuring the macrophage area instead of the macrophage number is recommended, as was&#160;done here.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67279/67279fig03.jpg" /> 
Figure 3: Macrophage recruitment at 4 hpl in 4 dpf zebrafish larvae. (A) Opercle area without ablation and at 4 hpl. Opercular osteoblasts are labeled in green, macrophages in magenta, and the dashed circle indicates the laser-ablated area. Scale bar: 50 &#181;m. (B) Quantification of the number of macrophages in the opercle area directly before (0 hpl) and after ablation (4 hpl). (C) Quantification of macrophage area (site of laser ablation excluded). Paired one-tailed t-tests. Each dot represents the data of one larva, and the mean &#177; SD is shown, n = 6. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67279/67279fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Author Spotlight: Immune-Mediated Bone Regeneration &#8212; The Critical Role of Macrophages and Controlled Immunomodulation in Restorative Processes

This protocol describes a two-photon laser ablation approach carried out in zebrafish larvae, which serves as a model to study bone regeneration and the effects of the immune response to ablation.

Two-Photon Laser-Mediated Ablation of Osteoblasts in Developing Zebrafish Larvae

Zebrafish (Danio rerio) have an outstanding capacity to regenerate different organs and appendages. Bone regeneration in zebrafish has been studied using ...

two-photon-laser-mediated-ablation-osteoblasts-developing-zebrafish

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

322 Views.  CRTD - Center for Regenerative Therapies TU Dresden. This protocol describes a two-photon laser ablation approach carried out in zebrafish larvae, which serves as a model to study bone regeneration and the effects of the immune response to ablation.

Video: Author Spotlight: Immune-Mediated Bone Regeneration — The Critical Role of Macrophages and Controlled Immunomodulation in Restorative Processes

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Hepatocyte-specific Ablation in Zebrafish to Study Biliary-driven Liver Regeneration

The liver has a great capacity to regenerate. Hepatocytes, the parenchymal cells of the liver, can regenerate in one of two ways: hepatocyte- or biliary-driven liver regeneration. In hepatocyte-driven liver regeneration, regenerating hepatocytes are derived from preexisting hepatocytes, whereas, in biliary-driven regeneration, regenerating hepatocytes are derived from biliary epithelial cells (BECs). For hepatocyte-driven liver regeneration, there are excellent rodent models that have significantly contributed to the current understanding of liver regeneration. However, no such rodent model exists for biliary-driven liver regeneration. We recently reported on a zebrafish liver injury model in which BECs extensively give rise to hepatocytes upon severe hepatocyte loss. In this model, hepatocytes are specifically ablated by a pharmacogenetic means. Here we present in detail the methods to ablate hepatocytes and to analyze the BEC-driven liver regeneration process. This hepatocyte-specific ablation model can be further used to discover the underlying molecular and cellular mechanisms of biliary-driven liver regeneration. Moreover, these methods can be applied to chemical screens to identify small molecules that augment or suppress liver regeneration.

To help assess the molecular mechanisms underlying zebrafish biliary-driven liver regeneration, we established a liver injury model in which the nitroreductase-expressing hepatocytes are genetically ablated upon metronidazole treatment. In this protocol, we describe how to adeptly manipulate, monitor and analyze hepatocyte ablation and biliary-driven liver regeneration.

The liver has a great capacity to regenerate. Hepatocytes, the parenchymal cells of the liver, can regenerate in one of two ways: hepatocyte- or ...

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Biology

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

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

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

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

Confocal Microscope-Based Laser Ablation and Regeneration Assay in Zebrafish Interneuromast Cells

Hair cells are mechanosensory cells that mediate the sense of hearing. These cells do not regenerate after damage in humans, but they are naturally replenished in non-mammalian vertebrates such as zebrafish. The zebrafish lateral line system is a useful model for characterizing sensory hair cell regeneration. The lateral line is comprised of hair cell-containing organs called neuromasts, which are linked together by a string of interneuromast cells (INMCs). INMCs act as progenitor cells that give rise to new neuromasts during development. INMCs can repair gaps in the lateral line system created by&#160;cell death. A method is described here for selective INMC ablation using a conventional laser-scanning confocal microscope and transgenic fish that express green fluorescent protein in INMCs. Time-lapse microscopy is then used to monitor INMC regeneration and determine the rate of gap closure. This represents an accessible protocol for cell ablation that does not require specialized equipment, such as a high-powered pulsed ultraviolet laser. The ablation protocol may serve broader interests, as it could be useful for the ablation of additional cell types, employing a tool set that is already available to many users. This technique will further enable the characterization of INMC regeneration under different conditions and from different genetic backgrounds, which will advance the understanding of sensory progenitor cell regeneration.

Laser ablation is a widely applicable technology for studying regeneration in biological systems. The presented protocol describes use of a standard laser-scanning confocal microscope for laser ablation and subsequent time-lapse imaging of regenerating interneuromast cells in the zebrafish lateral line.

Hair cells are mechanosensory cells that mediate the sense of hearing. These cells do not regenerate after damage in humans, but they are naturally ...

Deep and Spatially Controlled Volume Ablations using a Two-Photon Microscope in the Zebrafish Gastrula

Morphogenesis involves many cell movements to organize cells into tissues and organs. For proper development, all these movements need to be tightly coordinated, and accumulating evidence suggests this is achieved, at least in part, through mechanical interactions. Testing this in the embryo requires direct physical perturbations. Laser ablations are an increasingly used option that allows relieving mechanical constraints or physically isolating two cell populations from each other. However, many ablations are performed with an ultraviolet (UV) laser, which offers limited axial resolution and tissue penetration. A method is described here to ablate deep, significant, and spatially well-defined volumes using a two-photon microscope. Ablations are demonstrated in a transgenic zebrafish line expressing the green fluorescent protein in the axial mesendoderm and used to sever the axial mesendoderm without affecting the overlying ectoderm or the underlying yolk cell. Cell behavior is monitored by live imaging before and after the ablation. The ablation protocol can be used at different developmental stages, on any cell type or tissue, at scales ranging from a few microns to more than a hundred microns.

Embryonic development requires large-scale coordination of cell motion. Two-photon excitation mediated laser ablation allows the spatially controlled 3-dimensional ablation of large groups of deep cells. In addition, this technique can probe the reaction of collectively migrating cells in vivo to perturbations in their mechanical environment.

Morphogenesis involves many cell movements to organize cells into tissues and organs. For proper development, all these movements need to be tightly ...

Laser Ablation of the Zebrafish Pronephros to Study Renal Epithelial Regeneration

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in renal failure1. AKI can be caused by a number of factors including ischemia, drug-based toxicity, or obstructive injury1. This results in an inability to maintain fluid and electrolyte homeostasis. While AKI has been observed for decades, effective clinical therapies have yet to be developed. Intriguingly, some patients with AKI recover renal functions over time, a mysterious phenomenon that has been only rudimentally characterized1,2. Research using mammalian models of AKI has shown that ischemic or nephrotoxin-injured kidneys experience epithelial cell death in nephron tubules1,2, the functional units of the kidney that are made up of a series of specialized regions (segments) of epithelial cell types3. Within nephrons, epithelial cell death is highest in proximal tubule cells. There is evidence that suggests cell destruction is followed by dedifferentiation, proliferation, and migration of surrounding epithelial cells, which can regenerate the nephron entirely1,2. However, there are many unanswered questions about the mechanisms of renal epithelial regeneration, ranging from the signals that modulate these events to reasons for the wide variation of abilities among humans to regenerate injured kidneys.
The larval zebrafish provides an excellent model to study kidney epithelial regeneration as its pronephric kidney is comprised of nephrons that are conserved with higher vertebrates including mammals4,5. The nephrons of zebrafish larvae can be visualized with fluorescence techniques because of the relative transparency of the young zebrafish6. This provides a unique opportunity to image cell and molecular changes in real-time, in contrast to mammalian models where nephrons are inaccessible because the kidneys are structurally complex systems internalized within the animal. Recent studies have employed the aminoglycoside gentamicin as a toxic causative agent for study of AKI and subsequent renal failure: gentamicin and other antibiotics have been shown to cause AKI in humans, and researchers have formulated methods to use this agent to trigger kidney damage in zebrafish7,8. However, the effects of aminoglycoside toxicity in zebrafish larvae are catastrophic and lethal, which presents a difficulty when studying epithelial regeneration and function over time. Our method presents the use of targeted cell ablation as a novel tool for the study of epithelial injury in zebrafish. Laser ablation gives researchers the ability to induce cell death in a limited population of cells. Varying areas of cells can be targeted based on morphological location, function, or even expression of a particular cellular phenotype. Thus, laser ablation will increase the specificity of what researchers can study, and can be a powerful new approach to shed light on the mechanisms of renal epithelial regeneration. This protocol can be broadly applied to target cell populations in other organs in the zebrafish embryo to study injury and regeneration in any number of contexts of interest.

Acute kidney injury (AKI) in humans is a common clinical problem caused by damage to the epithelial cells that comprise kidney nephrons, and AKI is associated with high mortality rates of 50-70%1. Following epithelial cell destruction, nephrons have a limited ability to regenerate, though the mechanisms and limitations that guide this phenomenon remain poorly understood. In this video article, we describe our technique for targeted laser ablation of kidney nephron cells in the zebrafish embryo kidney, or pronephros. Our new method can be used to complement nephrotoxicity-induced models of AKI and gain a high-resolution understanding of the cell and molecular alterations that are associated with epithelial regeneration in the kidney nephron.

Acute kidney injury (AKI) is characterized by high mortality rates from deterioration of renal function over a period of hours or days that culminates in ...

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ablation of neurons is an effective method for this purpose when neurons are selectively labeled with fluorescent probes. In the present study, protocols for laser ablating a subpopulation of neurons using a two-photon microscope and testing of its functional and behavioral consequences are described. In this study, prey capture behavior in zebrafish larvae is used as a study model. The pretecto-hypothalamic circuit is known to underlie this visually-driven prey catching behavior. Zebrafish pretectum were laser-ablated, and neuronal activity in the inferior lobe of the hypothalamus (ILH; the target of the pretectal projection) was examined. Prey capture behavior after pretectal ablation was also tested.

Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ...

Neuroscience

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections (bupivacaine, cardiotoxin or notexin), muscle transplantations (denervation-devascularization induced regeneration), intensive exercise, but also murine muscular dystrophy models such as the mdx mouse (for a review of these approaches see 1). In zebrafish, genetic approaches include mutants that exhibit muscular dystrophy phenotypes (such as runzel2 or sapje3) and antisense oligonucleotide morpholinos that block the expression of dystrophy-associated genes4. Besides, chemical approaches are also possible, e.g. with Galanthamine, a chemical compound inhibiting acetylcholinesterase, thereby resulting in hypercontraction, which eventually leads to muscular dystrophy5. However, genetic and pharmacological approaches generally affect all muscles within an individual, whereas the extent of physically inflicted injuries are more easily controlled spatially and temporally1. Localized physical injury allows the assessment of contralateral muscle as an internal control. Indeed, we recently used laser-mediated cell ablation to study skeletal muscle regeneration in the zebrafish embryo6, while another group recently reported the use of a two-photon laser (822 nm) to damage very locally the plasma membrane of individual embryonic zebrafish muscle cells7.
Here, we report a method for using the micropoint laser (Andor Technology) for skeletal muscle cell injury in the zebrafish embryo. The micropoint laser is a high energy laser which is suitable for targeted cell ablation at a wavelength of 435 nm. The laser is connected to a microscope (in our setup, an optical microscope from Zeiss) in such a way that the microscope can be used at the same time for focusing the laser light onto the sample and for visualizing the effects of the wounding (brightfield or fluorescence). The parameters for controlling laser pulses include wavelength, intensity, and number of pulses.
Due to its transparency and external embryonic development, the zebrafish embryo is highly amenable for both laser-induced injury and for studying the subsequent recovery. Between 1 and 2 days post-fertilization, somitic skeletal muscle cells progressively undergo maturation from anterior to posterior due to the progression of somitogenesis from the trunk to the tail8, 9. At these stages, embryos spontaneously twitch and initiate swimming. The zebrafish has recently been recognized as an important vertebrate model organism for the study of tissue regeneration, as many types of tissues (cardiac, neuronal, vascular etc.) can be regenerated after injury in the adult zebrafish10, 11.

The method presented here comprises the precise injury of live zebrafish embryos with high-energy laser pulses and the subsequent analysis of these injuries and their recovery with time. We also show how genetically labeled single or groups of skeletal muscle cells can be tracked during and after laser light induced damage.

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections ...

Two-Photon Laser Axotomy: A Method to Injure Axons in Zebrafish Embryos and Observe Axonal Recovery

Source: O&#39;Brien G. S., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/1129/" target="_blank">Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos</a>. J. Vis. Exp. (2009).
This video describes the method to injure axons in zebrafish embryos using two photon laser axotomy and observing axonal recovery from injury.

Source: O&#39;Brien G. S., et al. Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos. J. Vis. Exp. (2009).
This video describes ...