Name: building finite element models to investigate zebrafish jaw biomechanics
Uploaded: 2016-12-03T14:00:00.000+00:00
Duration: 14 min 11 s
Description: Watch this Scientific Journal Video about Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics at JoVE.com

LHB was funded by the Wellcome Trust Dynamic Cell PhD programme; KAR was funded by MRC project grant MR/L002566/1 (awarded to EJR and CLH) and CLH was funded by ARUK grant 19479. We would also like to thank the Wolfson Bioimaging facility for imaging advice.

All steps within the protocol follow the University of Bristol&#39;s animal care and welfare guidelines and those of the U.K. Home Office.
1. Visualization of Musculoskeletal Anatomy
NOTE: To visualize the shape of the skeletal elements, to quantify muscle and to identify the exact placement of the muscle attachments, immunostain (section 1.1) fish at the appropriate age for skeletal myosin (which reveals muscle) and type II collagen (to visualize cartilage). Alternatively, visualize the musculoskeletal anatomy using transgenic fluorescent reporter lines such as the collagen a1 reporter col2a1:mCherry 13,14 to visualize cartilage and the slow myosin heavy chain reporter smyhc:GFP 15 to visualize the position of muscle attachments (Section 1.2).
Alternative lines that mark cartilage and muscle could work equally well.
<ol>
	<li>Fluorescent Immunostaining
	<ol>
		<li>Fix larva in excess 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) for 1 hr. Wash in PBS with 0.1% Tween 20 (PBT) and dehydrate in 50% methanol (MeOH) in PBT and 100% MeOH for 5 min respectively. 
		Caution: PFA is toxic and should be handled in accordance with the material safety sheet. 
		NOTE: Larva can be stored in 100% MeOH until required.</li>
		<li>Rehydrate larva in 50% MeOH in PBT for 5 min. Wash in PBT for 5 min.</li>
		<li>Permeabilize larva in 0.25% trypsin in PBT on ice for 5-6 min. Wash in 4x PBT for 5 min each.</li>
		<li>Block for 2-3 hr in 5% serum in PBT.</li>
		<li>Incubate larva in the recommended dilution of rabbit anti-type 2 collagen and mouse anti-myosin antibodies in 5% serum in PBT for 1 hr at room temp or overnight at 4 &#176;C. 
		NOTE: The recommended dilution range is normally on the antibody data sheet. Choose antibodies that are raised against different species to one another and that are also different to the tissue.</li>
		<li>Wash larva 6x for 15 mins in PBT.</li>
		<li>Block for 1-2 hr in 5% serum in PBT.</li>
		<li>Incubate in secondary antibodies in the dark. Use fluorescently labelled anti-mouse (550) and anti-rabbit (488) secondary antibodies at an appropriate dilution in 5% serum and PBT for the specific antibody.</li>
		<li>Wash 6X for 10 minutes each in PBT and image on a 10X confocal microscope as soon as possible.</li>
	</ol>
	</li>
	<li>Imaging Musculoskeletal Geometry
	<ol>
		<li>Mount the larvae ventrally on a coverslip in tepid 0.3-0.5% low melting point (LMP) agarose in Danieau&#39;s solution 16. 
		NOTE: Transgenic fish will need to be sedated in 0.02% MS222 (Tricaine methanesulfonate, pH 7) before mounting and during imaging.</li>
		<li>Take a confocal image stack of the region of interest using the 10X objective lens and the approximately 2.5X digital zoom. Prepare images of the green and red channel using the 488 nm and 561 nm lasers respectively. Image at a 512 x 512 pixel resolution with an interval between z planes of 1.3 &#181;m and 3 line averages. The resulting stack will comprise of approximately 100 z slices.</li>
		<li>Export the data as a tiff series. Maximum projections of the muscle and cartilage elements from a 5dpf zebrafish larva are shown in Figure 1.</li>
	</ol>
	</li>
</ol>
2. Generating a 3D Surface
<ol style="margin-left: 40px;">
	<li>Choose a representative dataset for each time point at 3, 4 and 5 dpf (choose after visualization of multiple samples).</li>
	<li>Open 3-dimensional tiff stack and select all channels in analysis software. Right click on the cartilage channel and select image filter and smoothing:Gaussian (Figure 2B).</li>
	<li>In project view right click on filtered image and select &#39;image segmentation&#39; and then &#39;edit new label&#39;. Create a new label for each material, i.e., cartilage and joint. Select the cartilage region of the image (Figure 2C, white signal, purple outline) using the magic wand tool. Use the brush tool to remove noise from the outlines. 
	NOTE: If using magic wand tool, click &#39;All slices&#39;.</li>
	<li>Select the joint region with the brush tool and assign to a joint component (Figure 2C, blue outline)</li>
	<li>Smooth multiple slices at once by selecting segmentation in the top menu and smooth labels. Right click on the image and select generate surface to produce a 3D surface render of the component (Figure 2D).</li>
	<li>Click on the surface and save data as a hmascii file for import into meshing software.</li>
</ol>
3. Calculating the Muscle Forces to Be Used in the FE Model
<ol style="margin-left: 40px;">
	<li>Count the number of muscle fibers from confocal images of smyhc:GFP transgenic zebrafish (Figure 1A, arrowhead, 1C) and measure the diameter of the fibers to calculate their cross sectional area (&#960;r2).</li>
	<li>Identify suitable force per muscle unit area from the literature. The maximal muscle force generated per unit area for larval zebrafish skeletal muscles (40 nN/&#181;m2) were used 17.</li>
	<li>Calculate the forces for each anatomical muscle group by multiplying the number of fibers and their area by the force per unit area. See Table 1.</li>
</ol>
4. Generating a Mesh
<ol style="margin-left: 40px;">
	<li>Import the 3D model generated in section 2 (above) into a software package capable of generating a finite element mesh.</li>
	<li>Generate a2D mesh of the cartilage and joint surfaces by using the shrink wrap tool under the 2D menu. Choose an appropriate element size. 
	Note: Use an element size between 1.5-2.5. If necessary, generate a range of differently sized 2D surface meshes to carry out 3D mesh optimization (Section 4.4).</li>
	<li>Carry out mesh quality checks found under the &#39;2D&#62;Tools&#62;Check elements&#39; panel to check for duplicated elements, insertions and penetrations in the mesh. Fix dihedral angles by using the utility tab in model tree.</li>
	<li>Generate a 3D mesh from the 2D surface meshes of differing element size using the 3D&#62;Tetramesh subpanel. 
	NOTE: Compare the results of different mesh sizes and select the FE model with the lowest mesh size that converges after further simulations and does not compromise feature definition. The example in Figure 3 contains 1.5 million tetrahedral elements for the lower jaw cartilages and had a 2D element size of 2.0.</li>
	<li>Transform the mesh so jaw model is to scale as per confocal stack using the Geometry&#62;Distance subpanel. 
	NOTE: Ensure the cartilage and joint components are connected in the model by exporting a merged model or by using ties.</li>
</ol>
5. Finite Element Model Construction
<ol>
	<li>Using commercial finite element (FE) software, create a FE model. Using the 3D muscle and cartilage labeled confocal stacks generated in section 1 as a guide, assign nodes that correspond to muscle attachment points. Create a vector between two nodes representing the origin and insertion of each muscle (Figure 3).</li>
	<li>Create a load collector of type &#39;history&#39; to apply a &#39;C load&#39; for each muscle. Specify the magnitude in Newtons (calculated in step 3.3) and assign the associated vector. Figure 3 shows the attachment points for the adductor mandibulae (AM), protractor hyoideus (PH) and intermandibularis (IM). 
	NOTE: For these jaw muscles, maximum contractile force is distributed between the origin and insertion so that only 50% of each load is applied at each site.</li>
	<li>Assign appropriate elastic isotropic material properties as determined by the literature. Young&#39;s modulus for cartilage and the interzone in this model were 1.1 MPa and 0.25 MPa respectively and Poisson&#39;s ratio was 0.25 for both 18,19.</li>
	<li>Create a load collector of type &#39;boundary&#39; to apply initial constraints on the model. Go to the tab Analysis&#62;Constraints and in the create subpanel, pick the nodes on the model you wish to constrain. Select the degrees of freedom (DOF) that limit movement of the model to the best approximation of its natural range of motion. 
	NOTE: The model in Figure 3 was constrained in all axes of motion (DOF: 1, 2, 3 represent x, y and z, respectively) at the ceratohyal to anchor it in space at a mid-point in the model and in the y and z axis at the point where the palatoquadrate connects to the rest of the zebrafish skull (Figure 3, Table 1). The model must be constrained in all three DOF in a least one node.</li>
	<li>Create a &#39;Load step&#39;, for each type of movement you wish to simulate (i.e., opening, closing), under the analysis menu and select all the relevant loads (made in Section 5.2) and constraints (made in Section 5.4) to simulate this movement. Select &#39;Static&#39; from the drop down menu when it appears.</li>
	<li>Export model including the mesh, loads, constraints and material properties in an appropriate file format, in this case &#34;.inp&#34; format.</li>
	<li>Load model into FE analysis software. Create and execute a job for the model using the Job module.</li>
	<li>Analyze output for stress, strain, displacement, etc. found in the results tab and visualization menu (Figure 4 and 5).</li>
</ol>
6. Validation of Jaw Deformation/Displacement Distances
<ol>
	<li>Select 3-6 Tg(Col2a1aBAC:mcherry) transgenic zebrafish.</li>
	<li>Lightly anaesthetize the larvae with 0.02% MS222 until they cease to respond to touch but their hearts are still beating.</li>
	<li>Mount the larvae laterally (while anaesthetized) on coverslips in tepid 1% LMP agarose (made up in Danieau&#39;s solution).</li>
	<li>Remove the agarose from around the head and jaw with forceps.</li>
	<li>Flush fresh Danieau&#39;s solution (with no MS222) over the head of the larva to remove anesthesia using a Pasteur pipette until normal mouth movements resume.</li>
	<li>Use movie capture software to take bright-field high-speed videos of mouth movements. Take movies of around a minute duration at the highest frame rate, or sufficient to record multiple cycles of jaw opening.</li>
	<li>Choose frames that show the jaw open to its maximum displacement. Measure the distance between the anterior tip of the Meckel&#39;s cartilage and the upper jaw (tip of ethmoid plate) in &#181;m.</li>
	<li>Calculate the average displacement from multiple fish larvae.</li>
	<li>Extract displacement data from the model. Use the average displacement calculated in 6.8 to verify model displacement behavior (Figure 4). If the model deviates from expected, run sensitivity analysis by sequentially altering material properties and muscle loads.</li>
</ol>

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Finite+element+modelling+predicts+changes+in+joint+shape+and+cell+behaviour+due+to+loss+of+muscle+strain+in+jaw+development.">Finite element modelling predicts changes in joint shape and cell behaviour due to loss of muscle strain in jaw development.</a> J Biomech. 48, 3112-3122 (2015).</li><li>Roddy, K. A., Prendergast, P. J., Murphy, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+influences+on+morphogenesis+of+the+knee+joint+revealed+through+morphological,+molecular+and+computational+analysis+of+immobilised+embryos.">Mechanical influences on morphogenesis of the knee joint revealed through morphological, molecular and computational analysis of immobilised embryos.</a> PloS one. 6, e17526 (2011).</li><li>Cuff, A. R., Bright, J. 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The authors have nothing to disclose.
Some data in Figs 3-5 has been reprinted from J.Biomech, 48 (12), Brunt et al., Finite element modeling predicts changes in joint shape and cell behavior due to loss of muscle strain in jaw development, 3112-22., 2015, with permission from Elsevier15.

Finite Element models have been used to relate the areas of skeletal elements that are under strain with those undergoing bone formation 10, as well as to map the areas under strain during endochondral ossification and joint morphogenesis 8,12,21. Other studies have also been able to apply theoretical growth models to replicate changes during joint development 11,12. Here we show the protocol for building FE models for a relatively simple system, the zebrafish jaw 20. Unlike alternative methods of collecting raw images for the FE models, such as CT scanning 22, confocal imaging of transgenic lines or immunostained zebrafish allows for multiple tissues to be studied. It can, therefore, provide direct information on muscle attachment points in relation to cartilage. Among vertebrate models zebrafish are particularly amenable to genetic and pharmacological manipulation. The generation of FE models for zebrafish craniofacial cartilage now opens up the possibility of further study of the interplay between biomechanics and genetics in joint morphogenesis.
There are a number of critical steps to the process of creating an FE model; the first is generating an accurate three-dimensional representation of the system. This requires imaging at high enough resolution to clearly define boundaries. Note that even with high-resolution imaging to make a good surface one may have to smooth out some regions. Another critical step is defining the correct placement of the load and correct constraints. An insufficiently constrained model will fail to solve and incorrect placement of the loads will cause abnormal movement.
Some processing of the raw data (Figure 2) is necessary as a surface generated from the raw data would be difficult to mesh (Figure 2B). We filtered the data using a Gaussian filter (Figure 2C) and we carried out some manual smoothing of the curves to produce a set of clean outlines that can be converted into a 3D surface. Too much smoothing can produce a &#34;melted&#34; surface that has lost many of its features. Choosing the correct element size is an iterative process as choosing too small an element size creates too large a mesh which is computationally intensive. However, choosing too large an element size will produce a mesh which fails to recapitulate the correct shape of the structure. The correct mesh had the smallest element size that captured the correct shape of the jaw and converged on a correct solution, verified using the jaw displacement. It may also be necessary to modify the material properties or load calculations to better emulate the correct displacement as different ages and species will have substantially different properties.
It is important to remember that there are always limitations to a hypothetical model and assumptions made to run FE models. When only modeling one or a small number of samples it is critical to ensure that a representative sample is chosen as there are likely to be small variations between individuals. As only some of the jaw elements and muscles were included, the model is a simplified version of the zebrafish craniofacial musculoskeletal system. Therefore, constraints had to be positioned to account for where the modelled jaw elements would connect with the rest of the skull and the model was artificially constrained in the center to fix it in &#39;space&#39;. This artificial constraint did not impact on the interpretation drawn from the models as the ceratohyal itself was not analyzed. The inclusion of more of the craniofacial structure, especially other jaw opening muscles such as the sternohyals and its attached cartilage 23, could have added to the model, but limitations include the ability of larger models to run in the Finite Element software.
Another limitation is that we have not modeled ligament insertion, though this could be achieved by the insertion of springs 8. One other assumption made in this case was that the model would behave linearly. The magnitudes of strains on the models were comparable to those in published models and applied to in vitro cells 10,24, with strains being below +3,500 and above -5,000 &#181;&#603; apart from constraint and muscle attachment points. Therefore, the strains at the relevant regions of the model were deemed within a range acceptable for a linear model. Cartilage does not behave entirely as a linear material and has previously been modelled as a poroelastic material, which enabled analysis of the fluid behavior in the model 25. Spreading the muscle attachment points amongst a cluster of local nodes would distribute the peak forces and more accurately represent the muscle insertion for certain muscles.
Use of FE allows an assessment of the strains and stresses acting on a structure. As a technique it is frequently used in many bioscience disciplines including orthopedics, paleontology and more recently developmental biology. Here we describe how to build FEs for the zebrafish lower jaw. In the future these models could be extended to look at the whole jaw, including the palate. Similar techniques could be used to model spinal biomechanics in fish, which to date have mostly been studied by kinematic means.

Finite Element (FE) modelling is an engineering technique that can computationally calculate and map the magnitude and location of strains acting on a structure 1. The model consists of the 3D structure, represented by a mesh of &#34;Finite Elements&#34;, and the end result of the analysis is governed by a number of factors including the structure and number of elements in the mesh, the magnitude and location of the mechanical loads and the material properties. Material properties describe certain aspects of a material&#39;s behavior under a given type of load; Young&#39;s modulus (E) describes the elasticity of the material while Poisson&#39;s ratio describes the proportional decrease in the width of a material to its length when a sample is stretched. FE modelling can be used to calculate a variety of variables including displacement, stress, pressure and strain acting on the model by taking into account the unique input data about the structure&#39;s shape, location and magnitude of loads and the specific material properties.
FE modeling is widely used in engineering 2 and increasingly for orthopedic 3 and paleontological applications4. In development biomechanical forces are known to act as a stimulus in many cells to activate cell responses 5-8 and it is useful to predict both the relative positions and magnitudes of mechanical stimuli within developing organ systems, however, currently FE modeling has been little used for zebrafish development.
Both cartilage and bone have been shown to be mechanosensitive materials. For example, in vitro compression has been found to activate chondrogenic pathways, whilst tension has been shown to be necessary for bone formation 9. FE analysis (FEA) has been exploited to model strains acting on biological specimens, including those acting on skeletal elements during bone formation 10. Other development applications include its use to predict the shape of a joint after it has been exposed to theoretical biomechanical forces 11,12 and to show the pattern of strains present during chick knee joint morphogenesis 8.
This protocol is aimed at sharing the experience of generating 3-dimensional surfaces, meshes and Finite Element models from confocal images with a view to understanding the mechanics of developing tissues. We also show ways of validating the FE models though capturing real joint displacement information in vivo. While we use the zebrafish jaw as an exemplar the same techniques could be used on any small biological system for which 3D information on the structure of the musculoskeletal system can be obtained by confocal or multiphoton imaging.

<table><tbody><tr><td>Coll2</td><td>Abcam</td><td>ab34712</td><td>Type II collagen antibody - stains all cartilage</td></tr><tr><td>A4.1025 / MF20</td><td>Developmental studies hybridoma bank</td><td>A4.1025</td><td>Skeletal mysoin antibody - marks all skeletal muscle&amp;nbsp;</td></tr><tr><td>Low melt agarose</td><td>Sigma&amp;nbsp;</td><td>A9414-5G</td><td>For mounting zebrafish</td></tr><tr><td>MS222 (Ethyl 3-aminobenzoate methanesulfonate )</td><td>Sigma</td><td>E10521-10G</td><td>To make anesthetic</td></tr><tr><td>Trypsin</td><td>Fisher</td><td>T/3760/48</td><td>sample permeablilisation</td></tr><tr><td>Dylight 488 Mouse IgG</td><td>Thermofisher</td><td>35502</td><td>Secondary antibody</td></tr><tr><td>Dylight 550 Rabbit IgG</td><td>Thermofisher&amp;nbsp;</td><td>84541</td><td>Secondary antibody</td></tr><tr><td>SP8/SP5 or SPE confocal</td><td>Leica&amp;nbsp;</td><td></td><td>For imaging&amp;nbsp;</td></tr><tr><td>LAS Leica capture software</td><td>Leica</td><td></td><td>Imaging software</td></tr><tr><td>Aviso (version 7.0.0)</td><td>FEI Visualization Science Group</td><td></td><td>3D image analysis software (Section 2)</td></tr><tr><td>Hypermesh part of the Hyperworks package (version 10)</td><td>Altair Engineering</td><td></td><td>FE model generating software (Section 4-5)</td></tr><tr><td>Abaqus (version 6.14)</td><td>SIMULIA</td><td></td><td>FE analysis software (Section 5.7-5.8)</td></tr></tbody></table>

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.

Immunostaining for muscle (Figure 1A) and cartilage (Figure 1B) or imaging of transgenic reporters (Figure 1C) allows the 3D structure of the jaw to be visualized, along with the associated musculature. By imaging at a high resolution it was possible to build a model that captures both the three-dimensional shape of the jaw (Figure 2) and the location and placement of loads (Figure 3). Utilizing in vivo displacements seen through high speed video capture (Figure 4) we verified that the range of motion in the model was within a realistic range.
The FE models once run can be used to display a range of data, such as stress (Figure 5A), minimum and maximum principal strain (Figure 5B-K). These results are three-dimensional so the model can be magnified to see fine patterns of detail (Figure 5E, 5I) rotated to obtain relevant views (Figure 5F, 5G, 5J, 5K) and digitally sectioned (Figure 5E&#39;, 5E&#39;&#39;, 5I&#39;, 5I&#39;&#39;) to show how the patterns of stress, strain or pressure change throughout the model. It is also possible to extract quantitative data from the model (not shown). By verifying the model and using the most accurate material properties, loads and mesh shape the FE model can be used to explore the best estimate of the mechanical environment being experienced by cells during that window of development. The results of the model can be directly compared to changes in cellular behavior and gene expression 20.
<img alt="Figure 1" src="/files/ftp_upload/54811/54811fig1.jpg" /> 
Figure 1: Representative images of the musculoskeletal elements of the zebrafish lower jaw at 5 dpf. Representative confocal stacks of the lower jaw of 5dpf larvae all shown with anterior to top (A) Immunostaining for A4.1025 which stains all skeletal myosin (B) Immunostaining for Type II collagen which marks all cartilage (C) Stack from a live larva expressing the transgenic reporters col2a1:mCherry marking cartilage (red) and smyhc:GFP slow muscle (green). IA: intermandibularis anterior, PH: protractor hyoideus, AM:adductor mandibulae, HI: hyoideus inferior, HI: hyoideus superior, CH: sternohyoideus, MC: Meckel&#39;s cartilage, PQ: Palatoquadrate, CH: ceratohyal. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54811/54811fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54811/54811fig2.jpg" /> 
Figure 2: Generation of a 3D surface from confocal data. Images showing the transition from confocal data into a 3D surface for the zebrafish lower jaw with higher magnification of the joint region. (A) Raw confocal data; (B) Dataset after application of a Gaussian filter; (C) Filtered outline; (D) 3D surface. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54811/54811fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54811/54811fig3.jpg" /> 
Figure 3: Representative meshes showing constraints and force vectors. Representative meshes for a 5 dpf larva for (A) mouth closing and (B) mouth opening. White dots denote places where the model is constrained and in which dimensions (e.g., x and y or x, y and z). White lines denote muscle positions, with the vector of muscle force denoted by white arrows. Red shows cartilage and Yellow the interzone. This figure has been modified from supplementary material previously published in Brunt et al. 15. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54811/54811fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54811/54811fig4.jpg" /> 
Figure 4: Sensitivity testing. FE-model simulating jaw displacement in 5dpf zebrafish for different cartilage and interzone Young&#39;s moduli. Jaw displacement (open to closed in &#181;m) is marked on the jaw; recorded using the color key. Each model (A-L) has a different combination of cartilage (c = 1.1, 3.1, or 6.1 MPa) or interzone (i = of 0.25, 0.5, 0.75, or 1 MPa) properties. Horizontal black arrow highlights the value of jaw displacement at the tip of the Meckel&#39;s cartilage (represented by the vertical black arrow). M and N stills from videos of 5 dpf larvae showing minimum, i.e., jaw closed (M) and maximum, i.e., jaw fully open (N) with the two superimposed (O) - white line on O represents the displacement (of 43 &#181;m). In this case relative cartilage properties of 1.1 with an interzone of 0.25 (A) best match the displacements seen in live fish (O). Panels A-L of this figure have been previously published in Brunt et al. 15. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54811/54811fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54811/54811fig5.jpg" /> 
Figure 5: Representative data from the FE models. FE-model simulation of all muscles applied in a 5 dpf larva (A-C). (A) Von Mises (EMaxmin) (B) Minimum Principal strain (E Min. P, &#181;&#603;) (C) Maximum principal strain (E Max. P., &#181;&#603;). FE-model simulation of maximum and minimum principal strains during jaw opening. (D-K): Maximum principal strain (E Max. P., &#181;&#603;) in (D) ventral jaw view and (E) ventral joint view (E) shows location of proximal-distal sections through the Meckel&#39;s cartilage joint and the interzone in (E&#39;) and (E&#39;&#39;), respectively. (F): lateral jaw view. (G): lateral joint view. (H-K): Minimum Principal strain (E Min. P, &#181;&#603;) in (H) ventral jaw view and (I) ventral joint view. (I) shows location of the proximal-distal sections through the Meckel&#39;s cartilage joint and interzone in (I&#39;) and (I&#39;&#39;) respectively (J): lateral jaw view. (K): lateral joint view. This figure has been previously published in Brunt et al. 15. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54811/54811fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Number of muscle fibers</td>
			<td>Muscle fiber area (&#181;m2)</td>
			<td>Muscle group area (&#181;m2)</td>
			<td>Force (N)</td>
		</tr>
		<tr>
			<td>5 dpf intermandibularis anterior</td>
			<td>5</td>
			<td>23.8</td>
			<td>119</td>
			<td>4.76e-6</td>
		</tr>
		<tr>
			<td>5 dpf protractor hyoideus</td>
			<td>6</td>
			<td>23.8</td>
			<td>142.8</td>
			<td>5.71e-6</td>
		</tr>
		<tr>
			<td>5 dpf adductor mandibulae</td>
			<td>9</td>
			<td>23.8</td>
			<td>214.2</td>
			<td>8.57e-6</td>
		</tr>
	</tbody>
</table>
Table 1: Muscle quantification. Calculated average muscle forces for the Intermandibularis Anterior, Adductor Mandibulae and Protractor Hyoideus at 5 dpf using 40 nN/&#181;m2 (value per unit area taken from reference 17). (Lorga et al., 2011) (n = 3).

Watch this Scientific Journal Video about Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics at JoVE.com

Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics | Developmental Biology | Video

Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics

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 ...

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9.1K Views.  University of Bristol. 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. 

Video: Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Drug Treatment and In Vivo Imaging of Osteoblast-Osteoclast Interactions in a Medaka Fish Osteoporosis Model

Bone-forming osteoblasts interact with bone-resorbing osteoclasts to coordinate the turnover of bone matrix and to control skeletal homeostasis. Medaka and zebrafish larvae are widely used to analyze the behavior of bone cells during bone formation, degeneration, and repair. Their optical clarity allows the visualization of fluorescently labeled bone cells and fluorescent dyes bound to the mineralized skeletal matrix. Our lab has generated transgenic medaka fish that express the osteoclast-inducing factor Receptor Activator of Nuclear-factor &#954;B Ligand (RANKL) under the control of a heat&#160;shock-inducible promoter. Ectopic expression of RANKL results in the excess formation of activated osteoclasts, which can be visualized in reporter lines with nlGFP expression under the control of the cathepsin K (ctsk) promoter. RANKL induction and ectopic osteoclast formation leads to severe osteoporosis-like phenotypes. Compound transgenic medaka lines that express ctsk:nlGFP in osteoclasts, as well as mCherry under the control of the osterix (osx) promoter in premature osteoblasts, can be used to study the interaction of both cell types. This facilitates the in vivo observation of cellular behavior under conditions of bone degeneration and repair. Here, we describe the use of this system to test a drug commonly used in human osteoporosis therapy and describe a protocol for live imaging. The medaka model complements studies in cell culture and mice, and offers a novel system for the in vivo analysis of drug action in the skeletal system.

Drug Treatment and In Vivo Imaging of Osteoblast-Osteoclast Interactions in a Medaka Fish Osteoporosis Model

Small laboratory fish have become popular models for bone research on the mechanisms underlying human bone disorders and for the screening of bone-modulating drugs. In this report, we describe a protocol to assess the effect of alendronate on bone cells in medaka larvae with osteoporotic lesions.

Bone-forming osteoblasts interact with bone-resorbing osteoclasts to coordinate the turnover of bone matrix and to control skeletal homeostasis. Medaka ...

Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

Zebrafish mutant phenotypes are often incompletely penetrant, only manifesting in some mutants. Interesting phenotypes that inconsistently appear can be difficult to study, and can lead to confounding results. The protocol described here is a straightforward breeding paradigm to increase and decrease penetrance in lethal zebrafish skeletal mutants. Because lethal mutants cannot be selectively bred directly, the classic selective breeding strategy of progeny testing is employed. This method also includes protocols for Kompetitive Allele Specific PCR (KASP) genotyping zebrafish and staining larval zebrafish cartilage and bone. Applying the husbandry strategy described here can increase the penetrance of an interesting skeletal phenotype enabling more reproducible results in downstream applications. In addition, decreasing the mutant penetrance through this selective breeding strategy can reveal the developmental processes that most crucially require the function of the mutated gene. While the skeleton is specifically considered here, we propose that this methodology will be useful for all zebrafish mutant lines.

The goal of this protocol is to alter the penetrance of lethal skeletal mutant phenotypes in zebrafish by selective breeding. Lethal mutants cannot be grown to adulthood and bred themselves, therefore this protocol describes a method for tracking and selecting penetrance through multiple generations by progeny testing.

Zebrafish mutant phenotypes are often incompletely penetrant, only manifesting in some mutants. Interesting phenotypes that inconsistently appear can be ...

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.

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 ...

Methods for the Study of the Zebrafish Maxillary Barbel

Barbels are skin sensory appendages found in fishes, reptiles and amphibians. The zebrafish, Danio rerio, develops two pairs of barbels- a short nasal pair and a longer maxillary pair. Barbel tissue contains cells of ectodermal, mesodermal and neural crest origin, including skin cells, glands, taste buds, melanocytes, circulatory vessels and sensory nerves. Unlike most adult tissue, the maxillary barbel is optically clear, allowing us to visualize the development and maintenance of these tissue types throughout the life cycle. 
This video shows early development of the maxillary barbel (beginning approximately one month post-fertilization) and demonstrates a surgical protocol to induce regeneration in the adult appendage (&gt;3 months post-fertilization). Briefly, the left maxillary barbel of an anesthetized fish is elevated with sterile forceps just distal to the caudal edge of the maxilla. A fine, sterile spring scissors is positioned against the forceps to cut the barbel shaft at this level, establishing an anatomical landmark for the amputation plane. Regenerative growth can be measured with respect to this plane, and in comparison to the contralateral barbel. Barbel tissue regenerates rapidly, reaching maximal regrowth within 2 weeks of injury.
Techniques for analyzing the regenerated barbel include dissecting and embedding matched pairs of barbels (regenerate and control) in the wells of a standard DNA electrophoresis gel. Embedded specimens are conveniently photographed under a stereomicroscope for gross morphology and morphometry, and can be stored for weeks prior to downstream applications such as paraffin histology, cryosectioning, and/or whole mount immunohistochemistry. These methods establish the maxillary barbel as a novel in vivo tissue system for studying the regenerative capacity of multiple cell types within the genetic context of zebrafish.

The zebrafish maxillary barbel is an integumentary sense organ containing ectodermal, mesodermal and neural crest derivatives. Importantly, the adult barbel can regenerate after proximal amputation. This video introduces maxillary barbel development and demonstrates a surgical protocol to induce regeneration, followed by collection, embedding and downstream imaging of barbel specimens.

Barbels are skin sensory appendages found in fishes, reptiles and amphibians.  The zebrafish, Danio rerio, develops two pairs of barbels- a short nasal ...

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish with fluorescently-marked plasma membranes can be scanned in a confocal laser scanning microscope in order to determine the volume of muscle tissue and the number of muscle fibers present. Efficient approaches for the measurement of cell number and size in live animals over time are described and validated against more arduous segmentation methods. Methods are described that permit the control of muscle electrical, and thus contractile, activity. Loss of skeletal muscle contractile activity greatly reduced muscle growth. In larvae, a protocol is described that allows reintroduction of patterned electrical-evoked contractile activity. The described methods minimize the effect of inter-individual variability and will permit analysis of the effect of electrical, genetic, drug, or environmental stimuli on a variety of cellular and physiological growth parameters in the context of the living organism. Long-term follow-up of the measured effects of a defined early-life intervention on individuals can subsequently be performed.

Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish ...

Removing Zebrafish Scales for Culturing and Bone Biology Studies

In Vitro Culturing Technique for Studying Cellular Dynamics in Zebrafish Scales

Zebrafish scales offer a variety of advantages for use in standard laboratories for teaching and research purposes. Scales are easily collected without the need for euthanasia, regenerate within a couple of weeks, and are translucent and small, allowing them to be viewed using a standard microscope. Zebrafish scales are especially useful in educational environments, as they provide a unique opportunity for students to engage in hands-on learning experiences, particularly in understanding cellular dynamics and in vitro culturing methods. The main objective of this protocol is to describe a method for collecting and maintaining zebrafish scales in culture for use in a variety of biological studies using basic laboratory equipment. Additionally, the protocol details their use in understanding bone homeostasis by examining the activity of bone cells involved in bone resorption and deposition. It also includes additional protocols for general techniques, such as the visualization of nuclei and apoptotic cells. The in vitro culturing protocol produces reliable results with minimal reagents and equipment. This article discusses the benefits of using in vitro cultures of zebrafish scales to foster scientific inquiry and outlines the resources needed to support their integration into educational settings.

Author Spotlight: Advancing Bone Cell Activity Research with Zebrafish Scales 

This protocol offers a method to study cellular dynamics using a simple in vitro culture technique. It provides an opportunity for zebrafish researchers and educators to study cellular processes, such as those related to bone homeostasis and basic cell biology, by visualizing fluorescent nuclei and apoptotic cells within the scales.

Zebrafish scales offer a variety of advantages for use in standard laboratories for teaching and research purposes. Scales are easily collected without ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is an excellent model for developmental biology research. During embryogenesis, zebrafish develop with a yolk mass, which presents three-dimensional challenges for sample observation and analysis. This protocol describes how to create two-dimensional flat mount preparations of whole mount in situ (WISH) stained zebrafish embryo specimens.

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model ...

A Manual Small Molecule Screen Approaching High-throughput Using Zebrafish Embryos

Zebrafish have become a widely used model organism to investigate the mechanisms that underlie developmental biology and to study human disease pathology due to their considerable degree of genetic conservation with humans. Chemical genetics entails testing the effect that small molecules have on a biological process and is becoming a popular translational research method to identify therapeutic compounds. Zebrafish are specifically appealing to use for chemical genetics because of their ability to produce large clutches of transparent embryos, which are externally fertilized. Furthermore, zebrafish embryos can be easily drug treated by the simple addition of a compound to the embryo media. Using whole-mount in situ hybridization (WISH), mRNA expression can be clearly visualized within zebrafish embryos. Together, using chemical genetics and WISH, the zebrafish becomes a potent whole organism context in which to determine the cellular and physiological effects of small molecules. Innovative advances have been made in technologies that utilize machine-based screening procedures, however for many labs such options are not accessible or remain cost-prohibitive. The protocol described here explains how to execute a manual high-throughput chemical genetic screen that requires basic resources and can be accomplished by a single individual or small team in an efficient period of time. Thus, this protocol provides a feasible strategy that can be implemented by research groups to perform chemical genetics in zebrafish, which can be useful for gaining fundamental insights into developmental processes, disease mechanisms, and to identify novel compounds and signaling pathways that have medically relevant applications.

The zebrafish is an excellent experimental organism to study vertebrate developmental processes and model human disease. Here, we describe a protocol on how to perform a manual high-throughput chemical screen in zebrafish embryos with a whole-mount in situ hybridization (WISH) read-out.

Zebrafish have become a widely used model organism to investigate the mechanisms that underlie developmental biology and to study human disease pathology ...

Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics

Summary

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Chapters in this video

Microbead Implantation in the Zebrafish Embryo

Drug Treatment and In Vivo Imaging of Osteoblast-Osteoclast Interactions in a Medaka Fish Osteoporosis Model

Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

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

Methods for the Study of the Zebrafish Maxillary Barbel

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

In Vitro Culturing Technique for Studying Cellular Dynamics in Zebrafish Scales

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

A Manual Small Molecule Screen Approaching High-throughput Using Zebrafish Embryos