This work was supported by grants from the National Institutes of Health [HD089934, DK108495].

The procedures described throughout this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) at Harvard Medical School.
1. Breeding
NOTE: There are several published methods for breeding15,16,17,20 that could also be used at this step. Prior to breeding, adult fish are maintained on a 10:14 light:dark cycle at 23 &#176;C and fed a pellet diet (see Table of Materials) once daily. Fish can be bred in a recirculating system with mechanical filtration and UV sterilization. Breeding can also be accomplished in static (non-recirculating) tanks, but fish should not be left in a static tank for more than 3 days, as water quality quickly degrades.
<ol>
	<li>Fill a 5 gal tank with fish-ready water (dechlorinated water adjusted to: pH = 7.1 +/- 2, conductivity = 900 +/- 150 &#181;S, temperature = 23 &#176;C).</li>
	<li>Place plastic mesh (see Materials) in the bottom of the tank. If breeding in static tanks, affix a water heater to the side of the tank. If breeding in a recirculating system, place a water heater in the system sump. 
	NOTE: The plastic mesh prevents adults from consuming eggs.</li>
	<li>Place one female and two male A. mexicanus fish of greater than 1 year old into the tank. Allow the fish to acclimate for 30 min.</li>
	<li>Set the temperature of the heater to 24 &#176;C (or 1 &#176;C warmer than the starting temperature).</li>
	<li>After 24 h, increase the temperature by 1 &#176;C.</li>
	<li>After 24 h, increase the temperature by 1 &#176;C. Check the tanks daily for eggs by shining a flashlight into the bottom of the tank. Withhold the fish from food during this 3-day period.</li>
	<li>If spawning has not been induced after 3 days, turn off the heater and allow the water to return to room temperature (RT) before moving the fish to their original tank.</li>
</ol>
2. Hatching Fertilized Eggs
<ol>
	<li>Once eggs are identified in a breeding tank, remove the adults and plastic mesh, and reduce the water to a depth of 10 cm using a beaker or cup. 
	NOTE: To estimate time of spawning, use a transfer pipette to place several eggs into a Petri dish and view them with a stereomicroscope to determine the stage14 and estimate the time of fertilization.</li>
	<li>Move the tank to a convenient work surface and remove any opaque eggs or feces, leaving only the translucent, fertile eggs in the tank. 
	NOTE: The number of fertile eggs can be recorded at this time.</li>
	<li>Fill the tank with fish-ready water (see step 1.1) and add 6-7 drops of methylene blue to the tank as the water fills. 
	NOTE: The final concentration of methylene blue is approximately 1.5 ppm.</li>
	<li>Add a heater and aquarium bubbler (attached to an air pump, with a regulator) to the tank.</li>
	<li>Set the heater to 24 &#176;C and adjust the airstream regulator to produce a gentle stream of bubbles.</li>
	<li>Place a cover on the tank to help maintain the water temperature. 
	NOTE: The eggs should begin hatching within 24 h of the spawning time.</li>
</ol>
3. Transfer of Hatched Larvae to Nursery Containers
<ol>
	<li>Add 20 g of salt (see Materials) to 8 L of fish-ready water (see step 1.1) and stir until dissolved. Fill each 1.5 L nursery container (see Materials) with 1 L of the prepared water.</li>
	<li>Use a transfer pipette to move hatched larvae into prepared nursery containers at a density of 20 fish per container. 
	NOTE: A headlamp can be useful to locate and transfer the hatched larvae.</li>
	<li>After all the visible hatched larvae have been removed, agitate the water in the tank by stirring, and/or blowing water jets into the edges and corners of the tank with the pipette. 
	NOTE: This will help reveal larvae that were missed on the first pass.</li>
	<li>Dispose of unused larvae at this stage using the guidelines of the Office of Laboratory Welfare (OLAW). Add sodium hypochlorite to the tank to achieve a final concentration of 6.15%. Wait at least 5 min before pouring down the sink.</li>
	<li>Label each nursery container with the date and time of fertilization. View nursery containers daily and continue to remove any dead larvae.</li>
</ol>
4. Preparation of Rotifer-based Fish Food
<ol>
	<li>See Table of Materials for information on obtaining rotifers. Follow the referenced protocol to set up, maintain, and harvest rotifers22.</li>
	<li>Prepare fish food by adding 3 mL of algae mixture (see Table of Materials) to 1 L of harvested rotifers. This mixture will be added directly to the nursery containers as a food supply.</li>
</ol>
5. Feeding of Post-larval Fish
<ol>
	<li>When the fish are 5 days post fertilization (dpf), add 3 mL of fish food (prepared in step 4.2) to each nursery container. At the optimal density, rotifers should be visible in dense groups at the corners of the nursery containers, and less apparent at the center of the container. Add additional rotifer mixture until the appropriate density has been reached.</li>
	<li>Check the containers daily for the presence of rotifers and add more if the concentration gets depleted. Continue to remove any dead larvae.</li>
	<li>When fish reach 14 dpf, move them to a tank fit with a recirculating system at a density of 5 fish/L of water. 
	NOTE: The number of surviving post-larval fish can be recorded at this time.</li>
</ol>
6. Whole-mount Immunohistochemistry of Post-larval Fish
<ol>
	<li>Remove post-larval fish of the desired stage from food for 24 h by pouring the nursery container containing the fish through a nylon mesh strainer and placing the strainer into a container with clean fish-ready water (see step 1.1). 
	NOTE: It is essential to remove all of the food. Even small amounts of food in the gut are auto-fluorescent and will influence imaging.</li>
	<li>Collect and euthanize the fish. 
	NOTE: The euthanasia protocol should follow OLAW guidelines and be approved by your Institutional Animal Care and Use Committee. We have noted that tricaine alone does not euthanize post-larval fish, and the fish begin to move when transferred from tricaine to fixative if the fish are not also kept on ice.
	<ol>
		<li>Prepare tricaine solution in a beaker by adding 0.4 g of tricaine-S and 0.8 g of sodium bicarbonate to 1 L of deionized water, and place it on ice.</li>
		<li>Pour the water containing the fish through a nylon mesh strainer to collect the fish. Gently submerge the strainer in ice-cold Tricaine solution and leave it on ice for 10 min.</li>
	</ol>
	</li>
	<li>Fix the euthanized fish.
	<ol>
		<li>Use a transfer pipette with a cut tip to transfer the fish to a conical tube.</li>
		<li>Remove the Tricaine solution with a transfer pipette and replace with fixative. Incubate with rocking. 
		NOTE: Fixative and fixation time must be determined based on the antibody used. 10% formalin solution (4% formaldehyde, see materials) overnight at 4 &#176;C is adequate for the antibodies listed in the Table of Materials. 
		CAUTION: Formalin is toxic and flammable. Wear personal protective equipment (gloves, lab coat, and splash goggles) and handle in a chemical hood. Solutions containing formalin should be disposed as hazardous waste.</li>
		<li>Use a transfer pipette to carefully remove the fixative without disturbing the fish. Add 3 mL of phosphate buffered saline-triton solution [PBS with 0.1% Triton (PBST), see Materials] and incubate for 15 min at RT with rocking.</li>
		<li>Remove PBST and replace with fresh PBST and incubate for 15 min. Repeat this &#8220;washing&#8221; one additional time. 
		NOTE: Fish can be stored in PBS containing 0.02% sodium azide at 4 &#176;C for several weeks.</li>
	</ol>
	</li>
	<li>Perform whole-mount immunostaining.
	<ol>
		<li>Prepare 50 mL of blocking solution [PB-0.5% Triton X, 0.2% bovine serum albumin (BSA), 1% dimethyl sulfoxide (DMSO), 0.02% sodium azide, 5% donkey serum]. 
		CAUTION: Sodium azide and DMSO are toxic. Personal protective equipment (gloves, lab coat, and splash goggles) should be used when handling. Any solutions should be disposed as hazardous waste.</li>
		<li>Use a transfer pipette to transfer the fish to a 4 mL glass vial with screw-top cap. Use a transfer pipette to remove the PBST and add 3 mL blocking solution. Incubate for 1 h at RT with rocking.</li>
		<li>Use a transfer pipette to remove the blocking solution and add primary antibody diluted in blocking solution. Incubate overnight at room temperature with agitation. 
		NOTE: For example, add 1:250 dilution of anti-HuC/HuD Neuronal Protein Mouse Monoclonal Antibody (see Table of Materials for a list of antibodies that have been successfully used in A. mexicanus). A set of fish with no primary antibody added should be included at this step. Volume of antibody should be enough to cover the fish and allow for agitation.</li>
		<li>Wash the fish 3 times with PBST as described in step 6.3.4. Replace PBST with secondary antibody diluted in blocking solution and incubate overnight at RT with agitation. 
		NOTE: Optimal primary and secondary antibody concentrations, incubation time, and incubation temperature should be determined for each antibody. Overnight at RT was effective for the antibodies listed in Table of Materials.</li>
		<li>Wash the fish 3 times with PBST, with each wash lasting 15 min as described in step 6.3.4.</li>
		<li>Transfer the fish to PBS for short-term storage before proceeding with dissecting, mounting, or sectioning the fish.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Comparing gene activity between surface and cave A. mexicanus requires carefully controlled environmental parameters and methods that can be replicated across laboratories. Our protocol for raising A. mexicanus provides consistent nutritional content during post-larval development. Following this feeding regime, gene function can be confidently compared between populations using the robust immunohistochemistry protocol we present. Here we discuss the significance of this method as well as its limitations and future applications.
To achieve density-matched growth, we found that post-larval fish can be raised without recirculating water for two weeks in 1.5 L containers on a diet of rotifers. This protocol can also be used to raise fish on a recirculating system; however, rotifers must be added daily to compensate for those lost through the tank outflow. Newly hatched Artemia nauplii are commonly used as a food source in aquaculture, but we found that using rotifers has considerable advantages, including: reduced price, improved biosecurity, consistent nutrition, and better water quality (see below).
First, the weekly cost in consumables is 4 dollars for rotifers, compared to 14 dollars for Artemia. Regarding biosecurity, rotifers are raised in the laboratory under controlled conditions, while Artemia are collected from the wild and subject to natural variation in microbial or pathogen contents23. Additionally, the nutrient composition of Artemia nauplii are environmentally determined and therefore inconsistent. Nauplii thrive on their own energy stores after hatching; they quickly loose nutritional value as they develop and should optimally be fed to the fish within several hours. Artemia begin feeding at 12 house post-hatching, representing the first time that they could be nutritionally enriched; however, at this stage they have become too large for 5 dpf fish to consume. In comparison, rotifers continuously feed on marine microalgae resulting in high nutrient content regardless of when the rotifers are harvested. Rotifers are much smaller than Artmeia nauplii (160 vs. 400 microns), making them easier for the fish to capture and swallow. Post-larval surface fish and cavefish consume rotifers in similar quantities suggesting no difference in preference or ability to capture the rotifers10.
Finally, Artemia nauplii begin dying in fresh water several hours after being introduced. Uneaten nauplii will decay, rapidly decreasing the water quality if they are not manually removed. Removing dead Artemia is time-consuming and dangerous for post-larval fish that are not much bigger than Artemia and may be accidently removed or injured. Rotifers can live indefinitely in the nursery containers and provide food to the fish at all times without significantly impacting water quality.
While using rotifers as a food source has considerable benefits, to maintain the rotifer stock, algae must be added to the culture system daily. This can be achieved with an automatic feeder that dispenses liquid algae into the rotifer culture container (see Table of Materials). Rotifers must also be harvested from this set-up every 24-48 hours to maintain the health of the culture. Researchers that breed fish very infrequently (once a year, for example) and are not concerned with making comparisons between the populations at post-larval stages may prefer Artemia as a food source, since the encysted embryos can be hatched at any time.
We recommend tracking the number of hatched and surviving larvae to monitor the success of hatching and growth. If most of the embryos or larvae die, it may be due to bacterial or fungal contamination. It is recommended to monitor the water quality of the fish-ready water and sterilize any equipment with 70% ethanol. Nursery containers can be re-used after they are cleaned and sterilized. To minimize risk of disease, it is also critical to remove any dead fish from the nursery containers and not add rotifers before 5 dpf, when the fish begin to eat.
A. mexicanus are sexually mature at approximately one year old. This is a limitation for generating transgenic A. mexicanus compared to Danio rerio (zebrafish) that are able to breed at 10-12 weeks24. Immunohistochemistry (IHC) is an alternative method to investigate gene expression and protein localization. The protocol described here can be adapted at each step and used to detect antigens in any tissue of interest. It is important to note, however, that some tissues may be more difficult to visualize in surface fish due to pigmentation (a barrier that does not exist in unpigmented cavefish), which may influence the interpretation of comparative studies. To address this potential problem, surface fish pigment can be bleached after fixation using 3% hydrogen peroxide.
IHC requires successful tissue fixation, blocking, and antibody penetration. Methods for each vary depending on the tissue and protein of interest. The fixative and fixation time must preserve cell architecture while maintaining the antigen epitope. For this protocol, we use a cross-linking fixative (paraformaldehyde) and omit a denaturing fixative (such as methanol or acetone). We found that incubation in acetone diminished antibody signal for neuronal and pancreatic markers. The blocking step is essential to prevent antibodies from binding to non-target proteins in the tissue. This method uses a combination of normal serum (5%) and BSA (0.2%) in the blocking solution. The blocking solution contains antibodies and proteins that bind to reactive sites on proteins in the tissue, diminishing non-specific binding of the primary and secondary antibodies.
To achieve antibody penetration, the tissue must be permeabilized. This can be achieved with detergents or denaturing solvents but must be optimized to preserve the antigen epitope. Our protocol uses a combination of Triton and dimethyl sulfoxide (DMSO). Triton and DMSO are included during the blocking and antibody incubation steps at concentrations of 0.5% and 1%, respectively. Using this concentration, we have observed staining in the brain, pancreas, intestine, and muscle, suggesting that it is likely effective for penetration of all tissues. Fish size may also influence penetration. This protocol has not been tested on fish that are greater than 14 days old (approximately 7 mm in length). To troubleshoot staining, it is recommended to alter the fixation, blocking, and antibody penetration steps. It is also important to examine the sequence conservation of the immunogen with the A. mexicanus protein of interest using the available genome25.
A. mexicanus is an excellent model to investigate evolution&#160;as populations of the same species that have evolved in dramatically different environments can be directly compared in the laboratory. Standard husbandry protocols, both within and among laboratories, are essential to understanding the biological differences between surface fish and cavefish. Our article provides a method to examine development and gene activity in post-larval fish exposed to consistent growth parameters.

The Mexican tetra, Astyanax mexicanus, is a single species of fish that exists as river-dwelling populations (surface fish) and a number of cave-dwelling populations (cavefish) named for the caves they inhabit (i.e., Tinaja, Molino, Pach&#243;n). A growing number of researchers are using A. mexicanus to investigate the genetic and developmental bases of behavioral1,2,3,4, metabolic5,6,7,8, and morphological evolution9,10,11. Available resources for studies of A. mexicanus include a sequenced and annotated genome12; transcriptome13; developmental staging table14; and methods for breeding15,16,17, creating transgenics18, and editing genes19. Dissemination of additional tools and updated standard protocols will accelerate growth of the cavefish research community (see this methods collection20).
Our goal is to add to the existing repertoire of tools by providing a robust method for assessing gene activity in situ in post-larval A. mexicanus, in a manner comparable between laboratories. There are two challenges to achieving this goal. First, there is a need for standardized regimes for hatching and raising the fish between laboratories, as differences in parameters such as feeding and density affect growth and maturation, thereby impacting gene activity. Second, there is a need for a standardized yet adaptable method for examining patterns of gene activity in the post-larval fish. We address these issues here, establishing standard practices for raising fish to post-larval stages and introducing a robust whole-mount immunohistochemistry (IHC) protocol for assessing gene expression in A. mexicanus.
We first demonstrate how to breed the fish through natural spawning and identify fertilized eggs. Described next is how to hatch fertilized eggs (larvae) and transfer them to nursery containers, where they are maintained at a density of 20 fish per container for two weeks without recirculating or changing the water. At 5-days post fertilization, the fish have developed to post-larval stages (no longer having a yolk supply) and are provided algae-fed Brachionus plicatilis (rotifers) as a nutrient-rich food source that do not require daily replenishment. This method provides consistent growth parameters for larval and post-larval development.
To assess gene function, we demonstrate how to remove the fish from nursery containers and perform whole-mount IHC. The IHC method presented is adapted from protocols developed for use with Danio rerio21 and is effective for examining antigens in all A. mexicanus tissues tested, including the brain, intestine, and pancreas. IHC is a faster alternative to generating transgenic animals for examination of gene expression and protein localization. This protocol will be useful for studies aimed at growing A. mexicanus and comparing the phenotypes of surface fish and cavefish at post-larval stages.

<table><tbody><tr><td>methylene blue</td><td>Kordon</td><td>B016CBHZUS</td><td>antifungal</td></tr><tr><td>heater</td><td>Finnex</td><td>4711457836017</td><td>100W Digital Control Heater</td></tr><tr><td>airstone</td><td>Lee&#039;s Aquarium &amp;amp; Pet Products</td><td>10838125202</td><td>disposable air stone</td></tr><tr><td>salt</td><td>Instant Ocean</td><td>51378014021</td><td>Sea Salt</td></tr><tr><td>nursery container</td><td>IPC</td><td>21545-002</td><td>40 oz or 1.5 L clear containers</td></tr><tr><td>transfer pipette</td><td>VWR</td><td>414004-002</td><td>plastic bulb pipettes</td></tr><tr><td>compact culture system(CCS) starter kit with Brachionus plicatilis (L-type) rotifers</td><td>Reed Mariculture</td><td>na</td><td>fish food</td></tr><tr><td>RGcomplete</td><td>APBreed</td><td>817656016572</td><td>32 oz bottle of rotifer food</td></tr><tr><td>Programmable Auto Dosing Pump DP-4</td><td>Jebao</td><td>DP-4</td><td>automatic feeder for rotifers</td></tr><tr><td>Tricane-S</td><td>Western Chemical</td><td>MS 222</td><td>fish anesthetic</td></tr><tr><td>sodium bicarbonate</td><td>Sigma-Aldrich</td><td>S5761-500G</td><td>for tricane solution</td></tr><tr><td>nylon mesh strainer</td><td>HIC (Harold Import Co.)</td><td>735343476235</td><td>3-inch diameter</td></tr><tr><td>Formalin solution, neutral buffered, 10%</td><td>Sigma-Aldrich</td><td>HT501128-4L</td><td>fixative</td></tr><tr><td>10X PBS</td><td>Invitrogen</td><td>AM9625</td><td>buffer, dilute to 1X using distilled water</td></tr><tr><td>Triton-x 100</td><td>Sigma-Aldrich</td><td>T8787-250ML</td><td>detergent</td></tr><tr><td>sodium azide</td><td>Sigma-Aldrich</td><td>S2002-25G</td><td>anti-bacterial</td></tr><tr><td>bovine serum albumin</td><td>Sigma-Aldrich</td><td>A9647-100G</td><td>blocking reagent</td></tr><tr><td>glass vial with screw-top cap 4mL</td><td>Wheaton</td><td>224742</td><td>staining vial</td></tr><tr><td>plastic mesh screen for breeding tank</td><td>Pentair</td><td>N1670</td><td>Cut into a rectangle 6mm larger on all edges than the dimensions of the bottom of the breeding tank. Cut a 6mm square from each corner of the rectangle. Bend the edges of the screen down along all four edges.Place a pair of 6mm vinyl-coated disk magnets on either side (top and bottom) of the mesh on each corner. The screen should be as snug as possible to the sides of the tank. The screen can be removed from the tank with a metal fish net.</td></tr><tr><td>vinyl-coated disk magnets</td><td>Kjmagnets</td><td>D84PC-AST</td><td></td></tr><tr><td>New Life Spectrum Thera-A pellet fish food</td><td>New Life International</td><td>na</td><td>Adult fish food. A list of retailers for this product is available on the company website</td></tr><tr><td>&lt;strong&gt;Antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>insulin antibody from guinea pig</td><td>Dako</td><td>A0564</td><td>1:200</td></tr><tr><td>glucagon antibody from sheep</td><td>Abcam</td><td>ab36215</td><td>1:200</td></tr><tr><td>acetylated tubulin antibody from mouse</td><td>Sigma</td><td>T6793</td><td>1:500</td></tr><tr><td>HuD/HuC antibody from mouse</td><td>Life Technologies</td><td>A-21271</td><td>1:500</td></tr><tr><td>nitric oxide synthase (nNOS) antibody from rabbit</td><td>Abcam</td><td>ab106417</td><td>5&amp;mu;g/mL</td></tr><tr><td>choline acetyltransferase (ChAT) from rabbit</td><td>Abcam</td><td>ab178850</td><td>1:2000</td></tr><tr><td>seratonin (5HT) from rabbit</td><td>Immunostar</td><td>20080</td><td>1:500</td></tr></tbody></table>

raising the mexican tetra astyanax mexicanus for analysis of post-larval phenotypes and whole-mount immunohistochemistry

River and cave-adapted populations of Astyanax mexicanus show differences in morphology, physiology, and behavior. Research focused on comparing adult forms has revealed the genetic basis of some of these differences. Less is known about how the populations differ at post-larval stages (at the onset of feeding). Such studies may provide insight into how cavefish survive through adulthood in their natural environment. Methods for comparing post-larval development in the laboratory require standardized aquaculture and feeding regimes. Here we describe how to raise fish on a diet of nutrient-rich rotifers in non-recirculating water for up to two-weeks post fertilization. We demonstrate how to collect post-larval fish from this nursery system and perform whole-mount immunostaining. Immunostaining is an attractive alternative to transgene expression analysis for investigating development and gene function in A. mexicanus. The nursery method can also be used as a standard protocol for establishing density-matched populations for growth into adults.

Table 1 shows success during one year of breeding surface fish and Tinaja, Molino, and Pach&#243;n cavefish in static breeding tanks. Surface and Pach&#243;n spawns with fertilized embryos always produced hatched larvae, while Molino and Tinaja were unsuccessful some of the time (2/6 and 2/18 spawning events did not produce hatched larvae, respectively). There is variation in clutch size that does not appear to be attributed to age of the parent fish. Table 2 shows the total number of hatched larvae resulting from some of the spawning events, and the age of the parent fish. In general, we found that surface fish produce the greatest number of larvae per spawn (average 1,550 &#177; 894, n = 5), followed by Pach&#243;n (average 879 &#177; 680, n = 6), Tinaja (average 570 &#177; 373, n = 11), and Molino (average 386 &#177; 276, n =3). The number of larvae produced is typically more than are needed per experiment or for growth into adults. We typically set up 6-18 nursery containers (120-360 larvae) and euthanize the remaining fish.
To measure the success of the nursery protocol we recorded the number of hatched larvae and surviving post-larval fish from successful spawning events. Table 3 shows data from 1 month of breeding in recirculating tanks and includes the number of larvae transferred to nursery containers that survived to 14 dpf. During this month, the survival rate ranged from 41-81 percent, resulting in 65-293 fish available per population for experiments or growth into adults.
To determine if whole-mount immunostaining is successful, we compared the fluorescence of samples incubated with primary antibody to those incubated with secondary antibody only. The fluorescent signal is only visible in the fish incubated with primary antibody. We have used this protocol to successfully label neurons10 (Figure 1) and pancreatic cells6 at stages up to 12.5 dpf in both surface and cave morphotypes.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58972/58972fig1.jpg" /> 
Figure 1: Neuron labeling. Whole-mount immunostaining of A. mexicanus. Image of 12.5 dpf surface fish (a) and Pach&#243;n cavefish (b). Image of the mid-body region [hatched yellow outline shown in (a) and (b)] of surface fish (c) and Pach&#243;n cavefish (d) stained with pan-neuronal antibody (Hu). (e) Confocal image of a region of the surface fish intestine showing enteric neurons (Hu) and their projections (acetylated tubulin). For this image, the intestine was dissected out and mounted in medium containing DAPI to stain the nuclei. <a href="https://www.jove.com/files/ftp_upload/58972/58972fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>population</td>
			<td>attempts</td>
			<td>spawning events</td>
			<td>clutches</td>
		</tr>
		<tr>
			<td>surface</td>
			<td>94</td>
			<td>23</td>
			<td>23</td>
		</tr>
		<tr>
			<td>Molino</td>
			<td>110</td>
			<td>6</td>
			<td>4</td>
		</tr>
		<tr>
			<td>Pach&#243;n</td>
			<td>167</td>
			<td>13</td>
			<td>13</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>242</td>
			<td>18</td>
			<td>16</td>
		</tr>
	</tbody>
</table>
Table 1: Summary of data from one year of breeding A. mexicanus in static tanks. Number of breeding attempts, resulting spawning events, and number of spawning events that produced hatched larvae.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>population&#160;</td>
			<td>parent age</td>
			<td>hatched larvae</td>
		</tr>
		<tr>
			<td>surface</td>
			<td>1 year</td>
			<td>989</td>
		</tr>
		<tr>
			<td>surface</td>
			<td>1 year</td>
			<td>1050</td>
		</tr>
		<tr>
			<td>surface</td>
			<td>3 years</td>
			<td>2214</td>
		</tr>
		<tr>
			<td>surface</td>
			<td>3 years</td>
			<td>432</td>
		</tr>
		<tr>
			<td>surface</td>
			<td>3 years</td>
			<td>1768</td>
		</tr>
		<tr>
			<td>surface</td>
			<td>4 years</td>
			<td>2852</td>
		</tr>
		<tr>
			<td>Pach&#243;n</td>
			<td>1 year</td>
			<td>1194</td>
		</tr>
		<tr>
			<td>Pach&#243;n</td>
			<td>1 year</td>
			<td>1933</td>
		</tr>
		<tr>
			<td>Pach&#243;n</td>
			<td>1.5 years</td>
			<td>371</td>
		</tr>
		<tr>
			<td>Pach&#243;n</td>
			<td>3 years</td>
			<td>480</td>
		</tr>
		<tr>
			<td>Pach&#243;n</td>
			<td>4 years</td>
			<td>1190</td>
		</tr>
		<tr>
			<td>Pach&#243;n</td>
			<td>4 years</td>
			<td>110</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>9 months</td>
			<td>259</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>9 months</td>
			<td>253</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>10 months</td>
			<td>1100</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>11 months</td>
			<td>857</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>1 year</td>
			<td>713</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>1 year</td>
			<td>853</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>1 year</td>
			<td>542</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>1.5 years</td>
			<td>360</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>1.5 years</td>
			<td>58</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>1.5 years</td>
			<td>1100</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>4 years</td>
			<td>185</td>
		</tr>
		<tr>
			<td>Molino</td>
			<td>2.5 years</td>
			<td>460</td>
		</tr>
		<tr>
			<td>Molino</td>
			<td>2.5 years</td>
			<td>619</td>
		</tr>
		<tr>
			<td>Molino</td>
			<td>3 years</td>
			<td>81</td>
		</tr>
	</tbody>
</table>
Table 2: Approximate female age and number of hatched larvae from individual spawning events from the indicated populations of A. mexicanus.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>population</td>
			<td>attempts</td>
			<td>spawning events</td>
			<td>clutches</td>
			<td>clutch size</td>
			<td>larvae transferred to nursery cups</td>
			<td>post-larval fish at 14dpf</td>
			<td>survival (%)</td>
		</tr>
		<tr>
			<td>surface</td>
			<td>4</td>
			<td>3</td>
			<td>2</td>
			<td>576 &#38; 1728</td>
			<td>360</td>
			<td>174</td>
			<td>48</td>
		</tr>
		<tr>
			<td>Molino</td>
			<td>4</td>
			<td>2</td>
			<td>1</td>
			<td>228</td>
			<td>159</td>
			<td>65</td>
			<td>41</td>
		</tr>
		<tr>
			<td>Tinaja</td>
			<td>4</td>
			<td>1</td>
			<td>1</td>
			<td>1952</td>
			<td>175</td>
			<td>93</td>
			<td>53</td>
		</tr>
		<tr>
			<td>Pach&#243;n</td>
			<td>4</td>
			<td>1</td>
			<td>1</td>
			<td>1696</td>
			<td>360</td>
			<td>293</td>
			<td>81</td>
		</tr>
	</tbody>
</table>
Table 3: Summary of data from one month of breeding A. mexicanus in a recirculating system and raising the larvae. Number of breeding attempts, resulting spawning events, clutches that produced hatched larvae, average number of larvae per clutch, larvae transferred to the nursery containers, and post-larval fish present in the nursery containers after 14 days.

Watch this Scientific Journal Video about Raising the Mexican Tetra Astyanax mexicanus for Analysis of Post-larval Phenotypes and Whole-mount Immunohistochemistry at JoVE.com

Raising the Mexican Tetra Astyanax mexicanus for Analysis of Post-larval Phenotypes and Whole-mount Immunohistochemistry

In this protocol, we demonstrate how to breed Astyanax mexicanus adults, raise the larvae, and perform whole-mount immunohistochemistry on post-larval fish to compare the phenotypes of surface and cave morphotypes.

Raising the Mexican Tetra Astyanax mexicanus for Analysis of Post-larval Phenotypes and Whole-mount Immunohistochemistry

River and cave-adapted populations of Astyanax mexicanus show differences in morphology, physiology, and behavior. Research focused on comparing adult ...

raising-mexican-tetra-astyanax-mexicanus-for-analysis-post-larval

Nanyang Technological University

Research

JoVE Journal

Genetics

8.7K Views.  Harvard Medical School. In this protocol, we demonstrate how to breed Astyanax mexicanus adults, raise the larvae, and perform whole-mount immunohistochemistry on post-larval fish to compare the phenotypes of surface and cave morphotypes. 

Video: Raising the Mexican Tetra Astyanax mexicanus for Analysis of Post-larval Phenotypes and Whole-mount Immunohistochemistry

Manipulation of Gene Function in Mexican Cavefish

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, including eye degeneration, albinism, sleep loss, hyperphagia, and sensory processing. Species of cavefish from around the world display a convergent evolution of morphological and behavioral traits due to shared environmental pressures between different cave systems. Diverse cave species have been studied in the laboratory setting. The Mexican tetra, Astyanax mexicanus, with sighted and blind forms, has provided unique insights into biological and molecular processes underlying the evolution of complex traits and is well-poised as an emerging model system. While candidate genes regulating the evolution of diverse biological processes have been identified in A. mexicanus, the ability to validate a role for individual genes has been limited. The application of transgenesis and gene-editing technology has the potential to overcome this significant impediment and to investigate the mechanisms underlying the evolution of complex traits. Here, we describe a different methodology for manipulating gene expression in A. mexicanus. Approaches include the use of morpholinos, Tol2 transgenesis, and gene-editing systems, commonly used in zebrafish and other fish models, to manipulate gene function in A. mexicanus. These protocols include detailed descriptions of timed breeding procedures, the collection of fertilized eggs, injections, and the selection of genetically modified animals. These methodological approaches will allow for the investigation of the genetic and neural mechanisms underlying the evolution of diverse traits in A. mexicanus.&#160;

We describe approaches for the manipulation of genes in the evolutionary model system Astyanax mexicanus. Three different techniques are described: Tol2-mediated transgenesis, targeted manipulation of the genome using CRISPR/Cas9, and knockdown of expression using morpholinos. These tools should facilitate the direct investigation of genes underlying the variation between surface- and cave-dwelling forms.

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, ...

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval body wall contains several cell types that are models to study cell and tissue morphogenesis. For example tracheae are used to examine tube morphogenesis1, and the dendritic arborization (DA) sensory neurons of the Drosophila larva have become a primary system for the elucidation of general and neuron-class-specific mechanisms of dendritic differentiation2-5 and degeneration6.
The shape of dendrite branches can vary significantly between neuron classes, and even among different branches of a single neuron7,8. Genetic studies in DA neurons suggest that differential cytoskeletal organization can underlie morphological differences in dendritic branch shape4,9-11. We provide a robust immunological labeling method to assay in vivo microtubule organization in DA sensory neuron dendrite arbor (Figures 1, 2, Movie 1). This protocol illustrates the dissection and immunostaining of first instar larva, a stage when active sensory neuron dendrite outgrowth and branching organization is occurring 12,13.
In addition to staining sensory neurons, this method achieves robust labeling of microtubule organization in muscles (Movies 2, 3), trachea (Figure 3, Movie 3), and other body wall tissues. It is valuable for investigators wishing to analyze microtubule organization in situ in the body wall when investigating mechanisms that control tissue and cell shape.

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how complex cell shapes, such as neuronal dendrites, are achieved during development, it is important to be able to accurately assay microtubule organization. Here we describe a robust immunohistological labeling method to examine microtubule organization of dendritic arborization neuron sensory dendrites, trachea, muscle, and other Drosophila larva body wall tissues.

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval ...

Neuroscience

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods are described here to analyze distinct populations of MTs in the developing neural tube of the zebrafish embryo. While the focus is on neural tissue, this methodology is broadly applicable to other tissues. The procedures are optimized for early to mid-somitogenesis-stage embryos (1 somite to 12 somites), however they can be adapted to a range of other stages with relatively minor adjustments. The first protocol provides a method to assess the spatial distribution of stable and dynamic MTs and perform a quantitative analysis of these populations with image-processing software. This approach complements existing tools to image microtubule dynamics and distribution in real-time, using transgenic lines or transient expression of tagged constructs. Indeed, such tools are very useful, however they do not readily distinguish between dynamic and stable MTs. The ability to image and analyze these distinct microtubule populations has important implications for understanding mechanisms underlying cell polarization and morphogenesis. The second protocol outlines a technique to analyze nascent MTs specifically. This is accomplished by capturing the de novo growth properties of MTs over time, following microtubule depolymerization with the drug nocodazole and a recovery period after drug washout. This technique has not yet been applied to the study of MTs in zebrafish embryos, but is a valuable assay for investigating the in vivo function of proteins implicated in microtubule assembly.

Immunolabeling methods to analyze distinct populations of microtubules in the developing zebrafish brain are described here, which are broadly applicable to other tissues. The first protocol outlines an optimized method for immunolabeling stable and dynamic microtubules. The second protocol provides a method to image and quantify nascent microtubules specifically.

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods ...

Developmental Biology

Behavioral Tracking and Neuromast Imaging of Mexican Cavefish

Cave-dwelling animals have evolved a series of morphological and behavioral traits to adapt to their perpetually dark and food-sparse environments. Among these traits, foraging behavior is one of the useful windows into functional advantages of behavioral trait evolution. Presented herein are updated methods for analyzing vibration attraction behavior (VAB: an adaptive foraging behavior) and imaging of associated mechanosensors of cave-adapted tetra, Astyanax mexicanus. In addition, methods are presented for high-throughput tracking of a series of additional cavefish behaviors including hyperactivity and sleep-loss. Cavefish also show asociality, repetitive behavior and higher anxiety. Therefore, cavefish serve as an animal model for evolved behaviors. These methods use free-software and custom-made scripts that can be applied to other types of behavior. These methods provide practical and cost-effective alternatives to commercially available tracking software.

Here, we present methods for high-throughput study of a series of the Mexican cavefish behaviors and vital staining of a mechanosensory system. These methods use free-software and custom-made scripts, providing a practical and cost-effective method for the studies of behaviors.

Cave-dwelling animals have evolved a series of morphological and behavioral traits to adapt to their perpetually dark and food-sparse environments. Among ...

Behavior

Mass-Rearing and Molecular Studies in Tortricidae Pest Insects

Tortricidae (Lepidoptera), commonly known as tortrix or leafroller moths, comprises many agricultural and forestry pests, which cause serious agricultural losses. To understand the biology of such pest moths, fundamental techniques have been in high demand. Here, methods for mass-rearing, observations, and molecular studies are developed using two tea tortrix, Homona magnanima and Adoxophyes honmai (Lepidoptera: Tortricidae). Insects were mass-reared with sliced artificial diet and maintained by inbreeding for over 100 generations by considering their biological characteristics. Insects have various sex dimorphisms; hence it is difficult to distinguish the sex during the developing stages, which have prevented subsequent assays. The present work highlighted that the sex of tortricids larvae could be determined by observing testes or lactic-acetic orcein staining to visualize the female-specific W chromosome. Moreover, using the sex determination methods, the present study enabled nucleic acid extractions from sex determined embryos and application toward high throughput sequencing. These tips are applicable for other pest insects and will facilitate further morphological and genetic studies.

The present protocol describes the rearing method of tortricid pest insects in the laboratories. The procedures to distinguish insects' sex and extract nucleic acids for high throughput sequencing are established using two tortricid pests.

Tortricidae (Lepidoptera), commonly known as tortrix or leafroller moths, comprises many agricultural and forestry pests, which cause serious agricultural ...

Environment

Wholemount In Situ Hybridization for Astyanax Embryos

In recent years, a draft genome for the blind Mexican cavefish (Astyanax mexicanus) has been released, revealing the sequence identities for thousands of genes. Prior research into this emerging model system capitalized on comprehensive genome-wide investigations that have identified numerous quantitative trait loci (QTL) associated with various cave-associated phenotypes. However, the ability to connect genes of interest to the heritable basis for phenotypic change remains a significant challenge. One technique that can facilitate deeper understanding of the role of development in troglomorphic evolution is whole-mount in situ hybridization. This technique can be implemented to directly compare gene expression between cave- and surface-dwelling forms, nominate candidate genes underlying established QTL, identify genes of interest from next-generation sequencing studies, or develop other discovery-based approaches. In this report, we present a simple protocol, supported by a flexible checklist, that can be widely adapted for use well beyond the presented study system. It is hoped that this protocol can serve as a broad resource for the Astyanax community and beyond.

Wholemount In Situ Hybridization for Astyanax Embryos

This protocol enables visualization of gene expression in embryonic Astyanax cavefish. This approach has been developed with the goal of maximizing gene expression signal, while minimizing non-specific background staining.

In recent years, a draft genome for the blind Mexican cavefish (Astyanax mexicanus) has been released, revealing the sequence identities for thousands of ...

Incremental Temperature Changes for Maximal Breeding and Spawning in Astyanax mexicanus

The Mexican tetra, Astyanax mexicanus, is an emerging model system for studies in development and evolution. The existence of eyed surface (surface fish) and blind cave (cave fish) morphs in this species presents an opportunity to interrogate the mechanisms underlying morphological and behavioral evolution. Cave fish have evolved novel constructive and regressive traits. The constructive changes include increases in taste buds and jaws, lateral line sensory organs, and body fat. The regressive changes include loss or reduction of eyes. melanin pigmentation, schooling behavior, aggression, and sleep. To experimentally interrogate these changes, it is crucial to obtain large numbers of spawned embryos. Since the original A. mexicanus surface fish and cave fish were collected in Texas and Mexico in the 1990s, their descendants have been routinely stimulated to breed and spawn large numbers of embryos bimonthly in the Jeffery laboratory. Although breeding is controlled by food abundance and quality, light-dark cycles, and temperature, we have found that incremental temperature changes play a key role in stimulating maximal spawning. The gradual increase of temperature from 72 &#176;F to 78 &#176;F in the first three days of a breeding week provides two-three consecutive spawning days with maximal numbers of high-quality embryos, which is then followed by a gradual decrease of temperature from 78 &#176;F to 72 &#176;F during the last three days of the spawning week. The procedures shown in this video outline the workflow before and during a laboratory breeding week for incremental temperature stimulated spawning.

Incremental Temperature Changes for Maximal Breeding and Spawning in Astyanax mexicanus

This article outlines the basic laboratory conditions and protocols for an incremental temperature regime to stimulate maximal spawning in the Mexican tetra Astyanax mexicanus, which is an emerging model for developmental and evolutionary studies.

The Mexican tetra, Astyanax mexicanus, is an emerging model system for studies in development and evolution. The existence of eyed surface (surface fish) ...

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of specific mRNA transcripts within whole mount specimen. This technique (adapted from Albertson and Yelick, 2005) was used in an upper level undergraduate Comparative Vertebrate Biology laboratory classroom at Syracuse University. The first two thirds of the Comparative Vertebrate Biology lab course gave students the opportunity to study the embryology and gross anatomy of several organisms representing various chordate taxa primarily via traditional dissections and the use of models. The final portion of the course involved an innovative approach to teaching anatomy through observation of vertebrate development employing molecular techniques in which WISH was performed on zebrafish embryos. A heterozygous fibroblast growth factor 8 a (fgf8a) mutant line, ace, was used. Due to Mendelian inheritance, ace intercrosses produced wild type, heterozygous, and homozygous ace/fgf8a mutants in a 1:2:1 ratio. RNA probes with known expression patterns in the midline and in developing anatomical structures such as the heart, somites, tailbud, myotome, and brain were used. WISH was performed using zebrafish at the 13 somite and prim-6 stages, with students performing the staining reaction in class. The study of zebrafish embryos at different stages of development gave students the ability to observe how these anatomical structures changed over ontogeny. In addition, some ace/fgf8a mutants displayed improper heart looping, and defects in somite and brain development. The students in this lab observed the normal development of various organ systems using both external anatomy as well as gene expression patterns. They also identified and described embryos displaying improper anatomical development and gene expression (i.e., putative mutants).
For instructors at institutions that do not already own the necessary equipment or where funds for lab and curricular innovation are limited, the financial cost of the reagents and apparatus may be a factor to consider, as will the time and effort required on the part of the instructor regardless of the setting. Nevertheless, we contend that the use of WISH in this type of classroom laboratory setting can provide an important link between developmental genetics and anatomy. As technology advances and the ability to study organismal development at the molecular level becomes easier, cheaper, and increasingly popular, many evolutionary biologists, ecologists, and physiologists are turning to research strategies in the field of molecular biology. Using WISH in a Comparative Vertebrate Biology laboratory classroom is one example of how molecules and anatomy can converge within a single course. This gives upper level college students the opportunity to practice modern biological research techniques, leading to a more diversified education and the promotion of future interdisciplinary scientific research.

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) was used in an upper level undergraduate Comparative Vertebrate Biology course in addition to vertebrate dissections. This gave students the opportunity to study gene expression patterns as well as gross anatomy, linking the study of molecular and organismal biology within one course.

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of ...

Biology

Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases (TALENs)

Identifying alleles of genes underlying evolutionary change is essential to understanding how and why evolution occurs. Towards this end, much recent work has focused on identifying candidate genes for the evolution of traits in a variety of species. However, until recently it has been challenging to functionally validate interesting candidate genes. Recently developed tools for genetic engineering make it possible to manipulate specific genes in a wide range of organisms. Application of this technology in evolutionarily relevant organisms will allow for unprecedented insight into the role of candidate genes in evolution. Astyanax mexicanus (A. mexicanus) is a species of fish with both surface-dwelling and cave-dwelling forms. Multiple independent lines of cave-dwelling forms have evolved from ancestral surface fish, which are interfertile with one another and with surface fish, allowing elucidation of the genetic basis of cave traits. A. mexicanus has been used for a number of evolutionary studies, including linkage analysis to identify candidate genes responsible for a number of traits. Thus, A. mexicanus is an ideal system for the application of genome editing to test the role of candidate genes. Here we report a method for using transcription activator-like effector nucleases (TALENs) to mutate genes in surface A. mexicanus. Genome editing using TALENs in A. mexicanus has been utilized to generate mutations in pigmentation genes. This technique can also be utilized to evaluate the role of candidate genes for a number of other traits that have evolved in cave forms of A. mexicanus.

Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases TALENs

Gene-targeting mutagenesis is now possible in a wide range of organisms using genome editing techniques. Here, we demonstrate a protocol for targeted gene mutagenesis using transcription activator like effector nucleases (TALENs) in Astyanax mexicanus, a species of fish that includes surface fish and cavefish.

Identifying alleles of genes underlying evolutionary change is essential to understanding how and why evolution occurs.  Towards this end, much recent ...

Gamete Collection and In Vitro Fertilization of Astyanax mexicanus

Astyanax mexicanus is emerging as a model organism for a variety of research fields in biological science. Part of the recent success of this teleost fish species is that it possesses interfertile cave and river-dwelling populations. This enables the genetic mapping of heritable traits that were fixed during adaptation to the different environments of these populations. While this species can be maintained and bred in the lab, it is challenging to both obtain embryos during the daytime and create hybrid embryos between strains. In vitro fertilization (IVF) has been used with a variety of different model organisms to successfully and repeatedly breed animals in the lab. In this protocol, we show how, by acclimatizing A. mexicanus to different light cycles coupled with changes in water temperature, we can shift breeding cycles to a chosen time of the day. Subsequently, we show how to identify suitable parental fish, collect healthy gametes from males and females, and produce viable offspring using IVF. This enables related procedures such as the injection of genetic constructs or developmental analysis to occur during normal working hours. Furthermore, this technique can be used to create hybrids between the cave and surface-dwelling populations, and thereby enable the study of the genetic basis of phenotypic adaptations to different environments.

Gamete Collection and In Vitro Fertilization of Astyanax mexicanus

In vitro fertilization is a commonly used technique with a variety of model organisms to maintain lab populations and produce synchronized embryos for downstream applications. Here, we present a protocol that implements this technique for different populations of the Mexican tetra fish, Astyanax mexicanus.

Astyanax mexicanus is emerging as a model organism for a variety of research fields in biological science. Part of the recent success of this teleost fish ...

Raising the Mexican Tetra Astyanax mexicanus for Analysis of Post-larval Phenotypes and Whole-mount Immunohistochemistry

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Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases (TALENs)

Gamete Collection and In Vitro Fertilization of Astyanax mexicanus