Name: dissection and flat-mounting of the threespine stickleback branchial skeleton
Uploaded: 2016-05-07T15:00:00.000+00:00
Duration: 8 min 2 s
Description: Watch this Scientific Journal Video about Dissection and Flat-mounting of the Threespine Stickleback Branchial Skeleton at JoVE.com

This work was funded in part by NIH R01 #DE021475 to CTM and an NSF Graduate Research Fellowship to NAE. Thanks to Miles Johnson for assistance with imaging and Priscilla Erickson for critical reading of the manuscript.

All fish work was approved by the Institutional Animal Care and Use Committee of the University of California-Berkeley (protocol number R330). Euthanasia was performed using immersion in 0.025% Tricaine-S buffered with 0.1% sodium bicarbonate39. All steps are performed at room temperature.
1. Preparation&#160;
Note: Perform steps 1.1-1.5 in conical tubes or scintillation vials that can seal tightly and be laid horizontally. Fish do not need to be constantly shaken, but try to mix the solution as often as possible by gently inverting or shaking the rack of tubes or vials to expose all sides of the fish to the staining solution and allow stain to penetrate the tissue evenly. Do not place a large batch of fish on a platform shaker, as the heavy weight of the liquid will break the shaker.
<ol>
	<li>Fix either freshly euthanized fish or fish stored in ethanol with 10% Neutral Buffered Formalin (NBF) overnight. Alternatively, use 4% paraformaldehyde in 1x PBS solution instead of 10% NBF. 
	Note: If extracting DNA, clip a small portion of the caudal or pectoral fins prior to fixation and store in ethanol.</li>
	<li>Dispose of fix properly in a chemical hood and replace with tap water (that is ~pH 7.0) for 2 hr. Avoid using de-ionized water as it can often be acidic and can decalcify bone.</li>
	<li>Remove water and stain fish with 0.008% Alizarin Red S in 1% KOH in water for 24 hr. For fish less than 20 mm standard length, use 0.004% Alizarin Red S. (Make a 100x (0.8%) stock solution of Alizarin Red S which can then be diluted).</li>
	<li>Remove stain (putting in appropriate waste container in hood) and place fish in tap water for a few hr. Change water as needed until water rinse is mostly clear.</li>
	<li>Remove water and place fish into 50% glycerol, 0.25% KOH for mild clearing and subsequent dissection. 
	Note: This staining protocol is modified from previously described methods40,41.</li>
</ol>
2. Dissection&#160;
Note: See Figure 1 for a review of relevant head skeletal morphology.
<img alt="Figure 1" src="/files/ftp_upload/54056/54056fig1.jpg" /> 
Figure 1: Stickleback head skeletal morphology. Alizarin Red stained threespine stickleback head imaged with fluorescence under a rhodamine B filter set. Useful morphology is labeled: Op = opercle, Subop = subopercle, BSRs = branchiostegal rays, Preop = preopercle, Infraorb 1-3 = infraorbital 1-3 (also called circumorbitals or suborbitals), Dent = dentary, Premax = premaxilla, Max = maxilla, Nas = nasal, Lat. ethm = lateral ethmoid, Psph = parasphenoid, Fron = frontal bone. For a more detailed description of the stickleback head skeleton, see Anker (1974)31. <a href="https://www.jove.com/files/ftp_upload/54056/54056fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Lay the fish flat (Figure 2A) and insert sharp #5 watchmaker&#39;s forceps into the side of the eye at a ~45&#176; angle to puncture the membrane covering the eye.</li>
	<li>Peel the membrane away from the eye, similar to peeling a yogurt lid (Figure 2B).</li>
	<li>Insert open forceps behind the eye, grab hold of the optic nerve behind the eye, and remove the eye (Figure 2C). Do not puncture the eye as it will leak melanin. If punctured, melanin can be washed away during later steps.</li>
	<li>Repeat on other side.</li>
	<li>Starting from the posterior, place one small dissecting scissor blade under the opercle flap, drag scissor blade dorsally above opercle, then cut soft tissue through to the eye socket (Figure 2D). Cut dorsal to the opercle bone.</li>
	<li>Cut the frontal bone (dorsal to the eye socket) (Figure 2E).</li>
	<li>Cut the midline parasphenoid bone around the center of the eye sockets (Figure 2F).</li>
	<li>Repeat opercle cut on the opposing side.</li>
	<li>Insert forceps under the opercle and slowly peel the face away from the body, trimming any soft tissue still attached (Figure 2G-H). Take care not to disrupt the first row of gill rakers.
	<ol>
		<li>With forceps, detach the ceratohyals on both sides from the midline basihyal while peeling away and removing the anterior craniofacial skeleton (the entire jaw including the dentary, premaxilla, and maxilla; the entire hyoid skeleton including the exterior dermal opercle, preopercle, subopercle, and branchiostegal rays and the underlying dorsal and ventral endochondral elements; and the anterior part of the skull including the nasal, lateral ethmoid, and infraorbital bones, see Figures 1 and 2I).</li>
		<li>Pelvic spines can be folded out from the body, and can serve as a handle for forceps to grab hold of when present. Spines lock into place. To unlock, gently pull spine with forceps directly away from fish body, then gently bend posteriorly to press spine flat against fish.</li>
	</ol>
	</li>
	<li>Insert closed forceps posterior and ventral to the branchial skeleton (just below gut tube) and drag forceps anteriorly, teasing apart the remaining muscles and ligaments attached to the branchial skeleton (Figure 2J-K).</li>
	<li>Using the tips of closed forceps, scrape away the muscles attaching the dorsal branchial skeleton to the ventral braincase in a posterior to anterior direction (Figure 2L).</li>
	<li>Repeat 2.9 and 2.10 on the opposing side.</li>
	<li>Grasp the base of the gut tube and pull anteriorly to remove the branchial skeleton and gut tube (Figure 2M-N).</li>
	<li>Separate the gut tube by making a perpendicular cut posterior to the end of the fifth ceratobranchial (Figure 2O).</li>
	<li>After removing any remaining bone fragments from the braincase on the dorsal side of the branchial skeleton, insert scissors into the branchial basket to make a dorsal cut (cutting anterior to posterior) between the bilateral sets of dorsal tooth plates (Figure 3A-D). Ensure cut is centered to avoid damaging the dorsal tooth plates.</li>
	<li>Make two shallow lateral cuts in the rubbery gut lumen at the posterior end of the branchial skeleton (anterior end of gut tube) to assist with opening the branchial skeleton (Figure 3E).</li>
	<li>Place fish and all tissue pieces into a jar and place the branchial skeleton into a microcentrifuge tube with 50% glycerol, 0.25% KOH to continue gentle clearing, or 100% glycerol if no further clearing is required. Label jars and tubes with a unique identifier so they can be tracked. The amount of clearing required is largely a function of the size of the fish, large adult fish (over 40 mm in standard length) typically require additional clearing.</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/54056/54056fig2.jpg" /> 
Figure 2: Stickleback branchial skeleton dissection. Alizarin Red stained threespine stickleback fish ready for dissection. The eye is depigmented from extensive clearing. Blue arrows indicate the direction of motion. (A) Lateral view of stickleback head, anterior is to the right. (B) Removal of membrane covering the eye. (C) Removal of the eye. (D) Dorsal cut above the opercle. (E) Frontal bone cut. (F) Parasphenoid cut. (G-I) Removal of the facial skeleton. (J) Removal of ventral branchial skeleton soft tissue connections. (K-L) Removal of dorsal branchial skeleton connections. (M-N) Removal of the branchial skeleton. (O) Separate the gut tube from the branchial skeleton. See steps 2.1 through 2.16 for more details. Scale bar = 5 mm. <a href="https://www.jove.com/files/ftp_upload/54056/54056fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Branchial Skeleton Re-staining (If Necessary)
<ol>
	<li>To stain the branchial skeleton darker or clear tissue more, remove 50% glycerol, 0.25% KOH solution and wash with 1% KOH twice (one five minute wash followed by a second 24 hr wash while shaking horizontally on a platform shaker).</li>
	<li>Remove 1% KOH and re-stain with 0.008% Alizarin Red S in 1% KOH for 24 hr.</li>
	<li>Remove stain and replace with 1% KOH for 24 hr.</li>
	<li>Remove KOH solution and replace with 50% glycerol, 0.25% KOH.</li>
</ol>
4. Mounting Branchial Skeleton
<ol>
	<li>Remove branchial skeleton from 50% glycerol, 0.25% KOH or 100% glycerol and place near the bottom of a 22 mm x 60 mm glass cover slip with the dorsal side facing up (Figure 3F). Add a few drops of 50% glycerol, 0.25% KOH or 100% glycerol on top of branchial skeleton. If transitioning from 50% glycerol, 0.25% KOH to 100% glycerol, change solution in a microcentrifuge tube and shake for &#62;5 min prior to mounting to equilibrate tissue.</li>
	<li>Roll out two small balls of modeling clay and place on either end of the cover slip to act as spacers.</li>
	<li>Loosely place a second coverslip on top with enough pressure to flatten the anterior branchial skeleton (Figure 3G).</li>
	<li>Peel open the left dorsal flap including dorsal tooth plates, flatten, and slide between the coverslips (Figure 3H).</li>
	<li>Repeat technique with right dorsal flap and push entire branchial skeleton away from the edge of the coverslip (Figure 3I).
	<ol>
		<li>Alternatively, hold both dorsal flaps open with forceps and carefully place the coverslip on, flattening the branchial skeleton in one smooth motion.</li>
		<li>Alternatively, mount the branchial skeleton upside-down on one cover slip, splaying each dorsal side out laterally so gravity does not allow the branchial skeleton to close back up. Then cover with second 22 mm x 60 mm glass cover slip and invert prep. 
		Note: Different mounting techniques tend to work better or worse for each individual. Try each and see what feels most comfortable.</li>
	</ol>
	</li>
	<li>Lightly press on top coverslip to flatten clay balls enough to keep the branchial skeleton mounted flat, but take care not to crush the specimen.
	<ol>
		<li>During the mounting process, ceratobranchials may rotate and obscure a row of rakers. Remedy this by sliding forceps between the coverslips and re-orienting the ceratobranchials or entire branchial skeleton.</li>
	</ol>
	</li>
	<li>Store preps flat in slide trays at room temperature. Mounted in 100% glycerol, preps can be stored between bridged coverslips for at least a decade. Clean forceps and scissors with isopropanol or ethanol and cover tips.</li>
</ol>
<img alt="Figure 3" src="/files/ftp_upload/54056/54056fig3.jpg" /> 
Figure 3: Flat mounting the branchial skeleton. Manipulation and mounting of the branchial skeleton is shown. Blue arrows indicate the direction of motion. (A) Branchial skeleton dorsal side up. (B-D) Rotation and incision between the dorsal tooth plates. (E) Lateral cut in soft tissue to further open the base of the gut tube. (F) Branchial skeleton placed at the bottom of a coverslip ready for mounting. (G) Second coverslip placed on the anterior half of the branchial skeleton (above dorsal tooth plates). (H-I) Flat mounting of the branchial skeleton by opening dorsal tooth plate flaps and sliding between two cover slips. See steps 4.1 through 4.6 for more details. Scale bar = 5 mm. <a href="https://www.jove.com/files/ftp_upload/54056/54056fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolved+tooth+gain+in+sticklebacks+is+associated+with+a+cis-regulatory+allele+of+Bmp6.">Evolved tooth gain in sticklebacks is associated with a cis-regulatory allele of Bmp6.</a> Proc. Natl. Acad. Sci. 111 (38), 13912-13917 (2014).</li><li>Ellis, N. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+developmental+and+genetic+mechanisms+underlie+convergently+evolved+tooth+gain+in+sticklebacks.">Distinct developmental and genetic mechanisms underlie convergently evolved tooth gain in sticklebacks.</a> Development. (142), 2442-2451 (2015).</li><li>Tucker, A. S., Fraser, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolution+and+developmental+diversity+of+tooth+regeneration.">Evolution and developmental diversity of tooth regeneration.</a> Semin. Cell Dev. Biol. 25-26, 71-80 (2014).</li><li>Erickson, P. A., Glazer, A. M., Cleves, P. A., Smith, A. S., Miller, C. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+developmentally+temporal+quantitative+trait+loci+underlie+convergent+evolution+of+increased+branchial+bone+length+in+sticklebacks.">Two developmentally temporal quantitative trait loci underlie convergent evolution of increased branchial bone length in sticklebacks.</a> Proc. R. Soc. B. 281, (2014).</li><li>Leary, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> AVMA Guidelines for the Euthanasia of Animals. , (2013).</li><li>Bell, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolutionary+phenetics+and+genetics.">Evolutionary phenetics and genetics.</a> Evol. Genet. Fishes. , 431-528 (1984).</li><li>Taylor, W. R., Van Dyke, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+procedures+for+staining+and+clearing+small+fishes+and+other+vertebrates+for+bone+and+cartilage+study.">Revised procedures for staining and clearing small fishes and other vertebrates for bone and cartilage study.</a> Cybium. 9 (2), 107-119 (1985).</li><li>Erickson, P. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+190+base+pair,+TGF-&#946;+responsive+tooth+and+fin+enhancer+is+required+for+stickleback+Bmp6+expression.">A 190 base pair, TGF-&#946; responsive tooth and fin enhancer is required for stickleback Bmp6 expression.</a> Dev. Biol. 401 (2), 310-323 (2015).</li><li>Miller, C. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=cis-Regulatory+changes+in+Kit+ligand+expression+and+parallel+evolution+of+pigmentation+in+sticklebacks+and+humans.">cis-Regulatory changes in Kit ligand expression and parallel evolution of pigmentation in sticklebacks and humans.</a> Cell. 131 (6), 1179-1189 (2007).</li><li>Aigler, S. R., Jandzik, D., Hatta, K., Uesugi, K., Stock, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selection+and+constraint+underlie+irreversibility+of+tooth+loss+in+cypriniform+fishes.">Selection and constraint underlie irreversibility of tooth loss in cypriniform fishes.</a> Proc. Natl. Acad. Sci. U. S. A. 111 (21), 7707-7712 (2014).</li><li>Pasco-Viel, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolutionary+trends+of+the+pharyngeal+dentition+in+Cypriniformes+(Actinopterygii+Ostariophysi).">Evolutionary trends of the pharyngeal dentition in Cypriniformes (Actinopterygii Ostariophysi).</a> PLoS One. 5 (6), e11293 (2010).</li></ol>

The authors have nothing to disclose.

The branchial skeleton is a complex set of bones in the throat of a fish that manipulates, filters, and masticates food items on their way to the esophagus. Many interesting trophic traits including the patterning of gill rakers, pharyngeal teeth, and branchial bones vary across and within species. The majority of these traits are difficult to near impossible to accurately measure with the branchial skeleton in situ (e.g., gill raker length, branchial bone length). This flat-mounting protocol places all...

An incredible amount of diversity exists in the head skeleton among vertebrates, especially among fishes. In many cases this diversity facilitates different feeding strategies1-4, and can involve major changes to both external and internal craniofacial patterning. The branchial skeleton is located internally in the throat of a fish and surrounds most of the buccal cavity. The branchial skeleton is comprised of 5 serially homologous segments, the anterior four of which support the gills. Together these five segments function as an interface between fish and their food5. Variation in a multitude of traits including gill rakers, pharyngeal teeth, and branchial bones contribute to efficient foraging on different types of food.
Sticklebacks have undergone an adaptive radiation after ancestral oceanic forms colonized freshwater lakes and creeks throughout the northern hemisphere. The shift in diet from small zooplankton in the ocean to larger prey in freshwater has resulted in dramatic trophic variation in several craniofacial traits6. While many studies have focused on external craniofacial differences in sticklebacks7-13, important craniofacial changes evolve repeatedly in the internal branchial skeleton. The ability to create fertile hybrids between morphologically distinct stickleback populations provides an excellent opportunity to map the genetic basis of evolved changes to the branchial skeleton.
One trophic trait of ecological significance is the patterning of gill rakers, periodic dermal bones that line the anterior and posterior faces of the branchial bones and are used to filter prey items. Fish that typically feed on small prey items tend to have longer and more densely spaced gill rakers compared to fish that feed on larger prey14,15. Variation in gill rakers has been reported both within and between species14-19, and aspects of gill raker patterning contribute to trophic niches and fitness16. Decades of research have extensively documented gill raker number and length variation in threespine sticklebacks17-21; however, these studies typically focus on the first row of gill rakers. Recent work has shown modularity in the genetic control of gill raker number across the branchial skeleton22,23 and across a single row in gill raker spacing23 and length24 highlighting the importance of studying more than row one or a single gill raker to understand the developmental genetic basis of gill raker reduction.
A second trophic trait of both ecological and biomedical significance is the patterning of pharyngeal teeth. Teeth in fishes can be located in both the oral jaw and in the branchial skeleton, known as pharyngeal teeth. Oral teeth are used primarily for prey capture while pharyngeal teeth are used for mastication and prey manipulation25-27. Both sets form via shared developmental mechanisms and are considered developmentally homologous28. Interesting modularity occurs whereby some species, such as zebrafish, lack oral and dorsal pharyngeal teeth29 while other species have multiple toothed ceratobranchials, pharyngobranchials, and sometimes toothed basihyal and hypobranchials30. In sticklebacks, pharyngeal teeth are found ventrally on the fifth ceratobranchial and dorsally on the anterior and posterior pharyngobranchials31. Kinematics on stickleback feeding show the oral jaw is used primarily for prey capture and facilitating suction feeding9 leaving mastication to the pharyngeal jaw. In cichlids, lower pharyngeal jaw morphology varies dramatically32,33 and has been shown to be adaptive and correlated with trophic niche34. Multiple freshwater stickleback populations have evolved dramatic increases in ventral pharyngeal tooth number23,35,36. Recent work has demonstrated that the developmental genetic basis of this evolved tooth gain is largely distinct in two independently derived populations of freshwater sticklebacks36. Unlike mammalian teeth, fish regenerate their teeth continuously throughout adult life37. Both of these previously described high toothed freshwater populations have evolved an accelerated tooth replacement rate, providing a rare vertebrate system to study the genetic basis of regeneration36.
A third trophic trait that has evolved repeatedly in freshwater sticklebacks is longer epibranchial and ceratobranchial bones, the branchial arch segmental homologs of the upper and lower jaw, respectively38. Longer branchial bones confer a larger buccal cavity and likely are adaptive for allowing larger prey items to be consumed. Furthermore, in other fish, epibranchial bones are important for depression of the dorsal pharyngeal tooth plates25. Like gill rakers and pharyngeal teeth, the branchial bones are internal and thus, difficult to easily visualize or quantify.
Here we present a detailed protocol to dissect and flat-mount the branchial skeleton, allowing easy visualization and quantification of a variety of important craniofacial traits. While this protocol describes a stickleback dissection, this same method works on a variety of other fishes.

<table><tbody><tr><td>Potassium Hydroxide (KOH)</td><td>EMD</td><td>PX1480-1</td><td></td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>G7893-4L</td><td></td></tr><tr><td>10% Neutral Buffered Formalin (NBF)</td><td>Azer Scientific</td><td>NBF-4-G</td><td></td></tr><tr><td>Alizarin Red S</td><td>EMD</td><td>AX0485-3</td><td></td></tr><tr><td>Microscope Cover Glasses 22 mm x 60 mm</td><td>VWR</td><td>16004-350</td><td></td></tr><tr><td>100 mm x 10 mm Glass Petri Dish</td><td>Kimble Chase</td><td>23064-10010</td><td>To dissect samples on</td></tr><tr><td>Sylgard 184 Silicone Elastomer Kit</td><td>Ellsworth Adhesives</td><td>184 SIL ELAST KIT 0.5KG</td><td>Can be poured into glass or plastic Petri dishes to make dissecting plates</td></tr><tr><td>Modeling Clay</td><td>Sargent Art</td><td>22-4000</td><td>1 lb cream</td></tr><tr><td>Scintillation Vials (case of 500)</td><td>Wheaton</td><td>986586</td><td>Borosilicate Glass with Screw Cap</td></tr><tr><td>Forceps-Dumont #5 Inox (Biologie tip)</td><td>FST</td><td>11252-20</td><td>Dumostars are an alternative</td></tr><tr><td>Dissecting Scissors&amp;nbsp;</td><td>FST</td><td>15003-08</td><td>Alternate sizes are available depending on size of sample</td></tr><tr><td>Dissecting Microscope</td><td>Leica</td><td>S6E with KL300 LED</td><td>Many other models work nicely, having a flat base helps</td></tr><tr><td>Microcentrifuge Tubes 1.7 ml</td><td>Denville</td><td>C2170</td><td></td></tr><tr><td>Cardboard slide tray</td><td>Fisher</td><td>12-587-10</td><td></td></tr></tbody></table>

dissection and flat-mounting of the threespine stickleback branchial skeleton

The posterior pharyngeal segments of the vertebrate head give rise to the branchial skeleton, the primary site of food processing in fish. The morphology of the fish branchial skeleton is matched to a species' diet. Threespine stickleback fish (Gasterosteus aculeatus) have emerged as a model system to study the genetic and developmental basis of evolved differences in a variety of traits. Marine populations of sticklebacks have repeatedly colonized countless new freshwater lakes and creeks. Adaptation to the new diet in these freshwater environments likely underlies a series of craniofacial changes that have evolved repeatedly in independently derived freshwater populations. These include three major patterning changes to the branchial skeleton: reductions in the number and length of gill raker bones, increases in pharyngeal tooth number, and increased branchial bone lengths. Here we describe a detailed protocol to dissect and flat-mount the internal branchial skeleton in threespine stickleback fish. Dissection of the entire three-dimensional branchial skeleton and mounting it flat into a largely two-dimensional prep allows for the easy visualization and quantification of branchial skeleton morphology. This dissection method is inexpensive, fast, relatively easy, and applicable to a wide variety of fish species. In sticklebacks, this efficient method allows the quantification of skeletal morphology in genetic crosses to map genomic regions controlling craniofacial patterning.

This protocol results in a dissected and flat mounted branchial skeleton (Figure 4) where a variety of important trophic traits can be quantified. From a dorsal view, all rows of gill rakers, all pharyngeal tooth plates, and nearly all branchial bones can be easily visualized and quantified22-24,35,36,38,42. Alizarin Red S also fluoresces on a rhodamine or similar red filter allowing double labeling with other markers (e.g., trans...

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Dissection and Flat-mounting of the Threespine Stickleback Branchial Skeleton

The branchial skeleton, including gill rakers, pharyngeal teeth, and branchial bones, serves as the primary site of food processing in most fish. Here we describe a protocol to dissect and flat-mount this internal skeleton in threespine sticklebacks. This method is also applicable to a variety of other fish species.

The posterior pharyngeal segments of the vertebrate head give rise to the branchial skeleton, the primary site of food processing in fish. The morphology ...

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Research

JoVE Journal

Biology

9.7K Views.  University of California, Berkeley. The branchial skeleton, including gill rakers, pharyngeal teeth, and branchial bones, serves as the primary site of food processing in most fish. Here we describe a protocol to dissect and flat-mount this internal skeleton in threespine sticklebacks. This method is also applicable to a variety of other fish species. 

Video: Dissection and Flat-mounting of the Threespine Stickleback Branchial Skeleton

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the cephalochordate amphioxus, Branchiostoma lanceolatum. We use constructs for membrane and nuclear targeted mCherry and eGFP that have been modified to accommodate amphioxus codon usage and Kozak consensus sequences. We describe the type of injection needles to be used, the immobilization protocol for the unfertilized oocytes, and the overall injection set-up. This technique generates fluorescently labeled embryos, in which the dynamics of cell behaviors during early development can be analyzed using the latest in vivo imaging strategies. The development of a microinjection technique in this amphioxus species will allow live imaging analyses of cell behaviors in the embryo as well as gene-specific manipulations, including gene overexpression and knockdown. Altogether, this protocol will further consolidate the basal chordate amphioxus as an animal model for addressing questions related to the mechanisms of embryonic development and, more importantly, to their evolution.

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here the robust and efficient expression of fluorescent proteins after mRNA injection into unfertilized oocytes of Branchiostoma lanceolatum. The development of the microinjection technique in this basal chordate will pave the way for far-reaching technical innovations in this emerging model system, including in vivo imaging and gene-specific manipulations.

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the ...

Developmental Biology

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

Building Finite Element Models to Investigate Zebrafish Jaw Biomechanics

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

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

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

Characterization Of Multi-layered Fish Scales (Atractosteus spatula) Using Nanoindentation, X-ray CT, FTIR, and SEM

The hierarchical architecture of protective biological materials such as mineralized fish scales, gastropod shells, ram&#8217;s horn, antlers, and turtle shells provides unique design principles with potentials for guiding the design of protective materials and systems in the future. Understanding the structure-property relationships for these material systems at the microscale and nanoscale where failure initiates is essential. Currently, experimental techniques such as nanoindentation, X-ray CT, and SEM provide researchers with a way to correlate the mechanical behavior with hierarchical microstructures of these material systems1-6. However, a well-defined standard procedure for specimen preparation of mineralized biomaterials is not currently available. In this study, the methods for probing spatially correlated chemical, structural, and mechanical properties of the multilayered scale of A. spatula using nanoindentation, FTIR, SEM, with energy-dispersive X-ray (EDX) microanalysis, and X-ray CT are presented.

Characterization Of Multi-layered Fish Scales Atractosteus spatula Using Nanoindentation, X-ray CT, FTIR, and SEM

This paper presents the methods used for probing spatially correlated chemical, structural, and mechanical properties of the multilayered scale of Atractosteus spatula (A. spatula) using nanoindentation, Fourier transform infrared (FTIR) spectroscopy, scanning electron microscopy (SEM), and X-ray computed tomography (X-ray CT). The experimental results have been used to investigate the design principles of protective biological materials.

The hierarchical architecture of protective biological materials such as mineralized fish scales, gastropod shells, ram&#8217;s horn, antlers, and turtle ...

Bioengineering

Sequential Photo-bleaching to Delineate Single Schwann Cells at the Neuromuscular Junction

Sequential photo-bleaching provides a non-invasive way to label individual SCs at the NMJ. The NMJ is the largest synapse of the mammalian nervous system and has served as guiding model to study synaptic structure and function. In mouse NMJs motor axon terminals form pretzel-like contact sites with muscle fibers. The motor axon and its terminal are sheathed by SCs. Over the past decades, several transgenic mice have been generated to visualize motor neurons and SCs, for example Thy1-XFP1 and Plp-GFP mice2, respectively.
Along motor axons, myelinating axonal SCs are arranged in non-overlapping internodes, separated by nodes of Ranvier, to enable saltatory action potential propagation. In contrast, terminal SCs at the synapse are specialized glial cells, which monitor and promote neurotransmission, digest debris and guide regenerating axons. NMJs are tightly covered by up to half a dozen non-myelinating terminal SCs - these, however, cannot be individually resolved by light microscopy, as they are in direct membrane contact3.
Several approaches exist to individually visualize terminal SCs. None of these are flawless, though. For instance, dye filling, where single cells are impaled with a dye-filled microelectrode, requires destroying a labelled cell before filling a second one. This is not compatible with subsequent time-lapse recordings3. Multi-spectral "Brainbow" labeling of SCs has been achieved by using combinatorial expression of fluorescent proteins4. However, this technique requires combining several transgenes and is limited by the expression pattern of the promoters used. In the future, expression of "photo-switchable" proteins in SCs might be yet another alternative5. Here we present sequential photo-bleaching, where single cells are bleached, and their image obtained by subtraction. We believe that this approach - due to its ease and versatility - represents a lasting addition to the neuroscientist's technology palette, especially as it can be used in vivo and transferred to others cell types, anatomical sites or species6.
In the following protocol, we detail the application of sequential bleaching and subsequent confocal time-lapse microscopy to terminal SCs in triangularis sterni muscle explants. This thin, superficial and easily dissected nerve-muscle preparation7,8 has proven useful for studies of NMJ development, physiology and pathology9. Finally, we explain how the triangularis sterni muscle is prepared after fixation to perform correlated high-resolution confocal imaging, immunohistochemistry or ultrastructural examinations.

Visualizing individual cells in densely packed tissues, such as terminal Schwann cells (SCs) at neuromuscular junctions (NMJs), is challenging. "Sequential photo-bleaching" allows delineating single terminal SCs, for instance in the triangularis sterni muscle explant, a convenient nerve-muscle preparation, where sequential bleaching can be combined with time-lapse imaging and post-hoc immunostainings.

Sequential photo-bleaching provides a non-invasive way to label individual SCs at the NMJ. The NMJ is the largest synapse of the mammalian nervous system ...

Neuroscience

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Complete Spinal Cord Injury and Brain Dissection Protocol for Subsequent Wholemount In Situ Hybridization in Larval Sea Lamprey

After a complete spinal cord injury, sea lampreys at first are paralyzed below the level of transection. However, they recover locomotion after several weeks, and this is accompanied by short distance regeneration (a few mm) of propriospinal axons and spinal-projecting axons from the brainstem. Among the 36 large identifiable spinal-projecting neurons, some are good regenerators and others are bad regenerators. These neurons can most easily be identified in wholemount CNS preparations. In order to understand the neuron-intrinsic mechanisms that favor or inhibit axon regeneration after injury in the vertebrates CNS, we determine differences in gene expression between the good and bad regenerators, and how expression is influenced by spinal cord transection. This paper illustrates the techniques for housing larval and recently transformed adult sea lampreys in fresh water tanks, producing complete spinal cord transections under microscopic vision, and preparing brain and spinal cord wholemounts for in situ hybridization. Briefly, animals are kept at 16&#160;&#176;C and anesthetized in 1% Benzocaine in lamprey Ringer. The spinal cord is transected with iridectomy scissors via a dorsal approach and the animal is allowed to recover in fresh water tanks at 23 &#176;C. For in situ hybridization, animals are reanesthetized and the brain and cord removed via a dorsal approach.

Complete Spinal Cord Injury and Brain Dissection Protocol for Subsequent Wholemount In Situ Hybridization in Larval Sea Lamprey

Lampreys recover locomotion after a complete spinal cord injury. However, some spinal-projecting neurons are good regenerators and others are not. This paper illustrates the techniques for housing sea lamprey larvae (and recently transformed adults), producing complete spinal cord transections and preparing wholemount brains and spinal cords for in situ hybridization.

After a complete spinal cord injury, sea lampreys at first are paralyzed below the level of transection. However, they recover locomotion after several ...

A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

Vertebrate embryos pattern their major body axis as repetitive somites, the precursors of vertebrae, muscle, and skin. Somites progressively segment from the presomitic mesoderm (PSM) as the tail end of the embryo elongates posteriorly. Somites form with regular periodicity and scale in size. Zebrafish is a popular model organism as it is genetically tractable and has transparent embryos that allow for live imaging. Nevertheless, during somitogenesis, fish embryos are wrapped around a large, rounding yolk. This geometry limits live imaging of PSM tissue in zebrafish embryos, particularly at higher resolutions that require a close objective working distance. Here, we present a flattened 3-D tissue culture method for live imaging of zebrafish tail explants. Tail explants mimic intact embryos by displaying a proportional slowdown of axis elongation and shortening of rostrocaudal somite lengths. We are further able to stall axis elongation speed through explant culture. This, for the first time, enables us to untangle the chemical input of signaling gradients from the mechanistic input of axial elongation. In future studies, this method can be combined with a microfluidic setup to allow time-controlled pharmaceutical perturbations or screening of vertebrate segmentation without any drug penetration concerns.

Here, we present the protocol for 3-D tissue culture of the zebrafish posterior body axis, enabling live study of vertebrate segmentation. This explant model provides control over axis elongation, alteration of morphogen sources, and subcellular resolution tissue-level live imaging.

Vertebrate embryos pattern their major body axis as repetitive somites, the precursors of vertebrae, muscle, and skin. Somites progressively segment from ...

Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a wide range of techniques encompassing ex vivo and in vitro approaches. However, most studies still rely on the analysis of two-dimensional (2D) imaging data, which limits proper evaluation of a developmental process like axial extension and somitogenesis involving highly dynamic interactions in a complex 3D space. Here we describe techniques that allow mouse live imaging acquisition, dataset processing, visualization and analysis in 3D and 4D to study the cells (e.g., neuromesodermal progenitors) involved in these developmental processes. We also provide a step-by-step protocol for optical projection tomography and whole-mount immunofluorescence microscopy in mouse embryos (from sample preparation to image acquisition) and show a pipeline that we developed to process and visualize 3D image data. We extend the use of some of these techniques and highlight specific features of different available software (e.g., Fiji/ImageJ, Drishti, Amira and Imaris) that can be used to improve our current understanding of axial extension and somite formation (e.g., 3D reconstructions). Altogether, the techniques here described emphasize the importance of 3D data visualization and analysis in developmental biology, and might help other researchers to better address 3D and 4D image data in the context of vertebrate axial extension and segmentation. Finally, the work also employs novel tools to facilitate teaching vertebrate embryonic development.

Here, we describe computational tools and methods that allow visualization and analysis of three and four-dimensional image data of mouse embryos in the context of axial elongation and segmentation, obtained by in toto optical projection tomography, and by live imaging and whole-mount immunofluorescence staining using multiphoton microscopy.

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a ...

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 ...

Dissection and Flat-mounting of the Threespine Stickleback Branchial Skeleton

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A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

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