We thank S. Yokoyama for assistance with collecting fish. We are also grateful to W. Kawamura for helpful advice about rearing methods. This study was supported by the Japan Society for the Promotion of Science (KAKENHI) through grants (No. 24370006 and 16K07507) awarded to T.S.

1. Collecting and Transporting Live Fish
NOTE: This protocol describes how to collect fish that possess a gas bladder, from a depth of &#8805;15 m to the surface. Rapid conveyance to the surface will induce expansion of the gas bladder by a change of pressure, which can seriously harm or kill the fish. Caution is warranted, as damage caused to the fish during this first step will negatively effect their survival and later behavior.
<ol>
	<li>Prior to diving with SCUBA, gather the following materials: a hand net suitable for capturing the targeted species underwater; double polyethylene bags that are large enough for the fish; rubberbands (&#248; 80 mm x 6 mm); an oxygen cylinder for inflating the collection bags at the surface; the Elbagin, which is an antimicrobial comprises 10% sodium nifurstyrenate; a polystyrene foam box in which to place the collection bags; pipettes; and, if necessary (refer to step 1.4), a rope with a length equivalent to maximum diving depth, as well as a weight of at least 2 kg.</li>
	<li>Collect the targeted fish species using a hand net while diving; while still underwater, place the captured fish in a polyethylene bag and tie the mouth of the bag with a rubberband. If it will take &#8805;1 day to transport the fish to an aquarium, also preserve water from the natural habitat in other bags, to be used for renewing the fish-keeping water. 
	NOTE: For safety, the underwater tasks must be worked by at least two people.</li>
	<li>Surface from the collecting point with the bags of fish at a speed of no more than 1 m/min: while at a depth of 10 m or more, stop ascending for 1 - 5 min every 2 m; and from a depth of 10 m to the surface stop for 1 - 5 min every 1 m. When the captured fish are unable to maintain their buoyancy in the bag (i.e., if they float up with an expanded gas bladder while still trying to swim toward the bottom), either maintain same depth for 1 - 5 min or descend to 1 - 2 m deeper. Once the fish seem to recover their buoyancy, resume ascent to the surface. 
	NOTE: If there is a possibility that the air in SCUBA tank will run out while ascending, fasten a rope to the polyethylene bag and attach a &#8805;2 kg weight; after underwater operator has safely ascended, pull this up to the surface at the same speeds stipulated above.</li>
	<li>At beach or boat, dissolve 10 ppm Elbagin in the water of each polyethylene bag after surfacing. 
	NOTE: Although some fish may not be able maintain their buoyancy just after being brought to the surface, most can be expected to recover within one day.</li>
	<li>If the density of fish in a collecting bag is high, divide the fish among more bags to prevent from them damaging each other by rubbing against each other during transport. If the fish is an aggressive species, divide them individually between bags.</li>
	<li>Inflate the collection bag with oxygen and again seal the mouth of the bag using rubberbands. When introducing the oxygen into the bag, lower the nozzle into the bag&#39;s water to increase the dissolved oxygen content; the fully expanded bag should be 1/4 full with water.</li>
	<li>Keep the collection bags with fish in a polystyrene foam box, to maintain a stable water temperature and decrease the fish&#39;s stress under dark conditions.</li>
	<li>If it will take &#8805;1 day to transport the fish to an aquarium, once per day exchange 1/4 to 1/3 of the water in each fish-keeping bag, using water retained from the habitat to which has been added 10 ppm Elbagin, and repack each with oxygen. Each day, remove dead fish and excreta on the bottom by hand net or pipette. 
	NOTE: If airplane transportation is involved, wait at least 1 day to acclimating fish to the air-pressure on surface before transportation because two steps pressure change (from underwater to surface, and surface to upper air) in the short term may be negatively affect the survival of the fish.</li>
</ol>
2. Acclimating the Fish to an Aquarium
<ol>
	<li>Float the bag containing fish in an aquarium for 30 min to equalize the water temperature.</li>
	<li>Over a 10 min period, incrementally exchange the bag water with aquarium water, to avoid shock caused by a sudden difference in water chemistry (e.g., pH, salinity). 
	NOTE: A fish that becomes shocked by too great a difference in water chemistry may display an abnormal change in body color and/or behavior. Monitor this carefully during acclimation.</li>
	<li>Dissolve 10 ppm Elbagin in the aquarium water.</li>
	<li>Thereafter, once a day for 3 days, renew 1/3 of the aquarium water, having added 10 ppm Elbagin to the replacement water.</li>
	<li>Finally, begin to eliminate the Elbagin by exchanging half the aquarium water once a day, until the color fades.</li>
</ol>
3. Injecting a Visible Implant Elastomer (VIE) Tag to Identify Individual Fish
NOTE: In this work, individual fish are identified using VIE tags; for examples, refer to Frederick10, Olsen and V&#248;llestad11, and Leblanc and Noakes12. Also, if the study species is large enough to hold in a hand, the surgical table used in step 3.2 will not be necessary.
<ol>
	<li>Determine the injection position and tag color for each individual. The safest option is to choose an injection point in the thick muscular dorsal or caudal part of the body, and to avoid injecting it into the abdomen where internal organs may be pierced. 
	NOTE: A numbering system is described in Figure 1: if the study species is large enough for injection into 8 possible positions, this system will allow the identification of 154 individuals using a single color. Ten colors of VIE tags are available commercially. Choose the distinguishable color based on the body color of fish.</li>
	<li>For fish that are smaller than can be held in the hand, prepare a surgical table as follows (refer also to Kinkel et al.13) (Figure 2).
	<ol>
		<li>Cut out a soft sponge measuring 5 cm L x 5 cm W, and which is at least 5 - 10 mm lower than the height of a Petri dish.</li>
		<li>Cut a groove in the sponge of approximately 5 - 10 mm depth, and adjust its width to that of the approximate body width of the fish. Cut the polyvinyl chloride (PVC) board (0.3 mm thickness) to 5 cm L x 5 cm W, and bend it into a valley-fold (or M-shape).</li>
		<li>Set the grooved PVC board on top of the grooved sponge and then set the sponge into the Petri dish (&#248; 160 mm, 30 mm depth). Use water from the recovery tank to fill the Petri dish until the PVC board is adequately immersed.</li>
	</ol>
	</li>
	<li>Prepare the VIE tags according to the VIE tagging manual. Mix the elastomer materials and add the mixture to the 3-mL syringe with 29-gauge needle, as contained in the kit.</li>
	<li>Prepare two water tanks to use while performing the tag injections: one for anesthesia and one for recovery; adjust the temperature and salinity concentration of these to match the water in the rearing aquarium. Use an air pump with an air stone to mildly circulate the water in the recovery tank.</li>
	<li>Prepare an anesthesia liquid by mixing 2-methylquinoline with an equal volume of 99.5% ethanol; add this to the anesthesia tank to achieve a concentration of 18 ppm. 
	NOTE: The optimum concentration of the anesthesia liquid will depend on the species and body size of the individuals. Therefore, examine the optimum concentration for the study species in advance.</li>
	<li>Transfer individual fish to the anesthesia tank and allow them to become anesthetized: that is, wait until fish do not react to being touched or to water vibrations when the outside of the tank is tapped. As the body color changes when it is anesthetized in many fish, also monitor the body color carefully and judge whether fish is anesthetized from its changes. Because there is some possibility that fish may jump out of the anesthesia tank, keep it covered using a transparent acrylic board, at least until the fish begin to move slowly. 
	NOTE: Fine-tune the time needed to anesthetize and/or adjust the concentration of anesthesia liquid, depending on the study species and its body size. If the operculum movement is ever stopped by the anesthesia, the fish will be at a high risk of death.</li>
	<li>As fish will weaken if their body is warmed, keep fingers in the relatively cold water while anesthetizing the fish.</li>
	<li>Lift a fish from the anesthesia tank and quickly measure it or record the necessary data (total length, sex, etc.). If a fish appears to recover during measurement, immediately transfer it back to the anesthesia tank.</li>
	<li>Transfer the fish to a surgical table. Set the fish ventral side down in the PVC groove. If the body size is very small, use a binocular microscope while making the injection. If the study species is large enough to hold in a hand, inject the VIE tag while it is being held.</li>
	<li>Position the beveled side of the needle toward the outside and pointed toward the head of the fish. Insert the needle subcutaneously, more or less parallel to the body, and as near to just under the skin as possible. Adjust the insertion depth depending on the fish body size and the easiness to ultimately see the tag.</li>
	<li>Inject VIE tag while withdrawing the needle, and stop the injection before the needle bevel reaches the needle entry point (this will be easy to discern if injecting the tag in a relatively wide area).</li>
	<li>On completing the tag injection, immediately transfer the fish to the recovery tank. If recovery appears too slow, gently circulate the water by hand.</li>
	<li>After recovery, return the fish to the rearing aquarium, and continue to add 10 ppm Elbagin to the water for 3 days. 
	NOTE: Under low-visibility conditions for VIE tags, UVA filtered light will facilitate recognizing the tags.</li>
</ol>
4. Counting the Demersal Adhesive Eggs
<ol>
	<li>To create an artificial spawning nest to which the fish can deposit adhesive eggs, cut an opaque PVC pipe (&#248; 5 cm, 6 cm long) in half, perpendicular to the diameter. 
	NOTE: The above pipe size is suitable for spawning the relatively small goby T. marinae. Thus, adjust the PVC pipe size and form as is appropriate for the study species.</li>
	<li>Print a 5 mm x 5 mm grid on a waterproof sheet and cut it out to fit the inside area of the PVC pipe.</li>
	<li>Fix the waterproof sheet to the inside of the PVC pipe using a rubberband. 
	NOTE: If it is difficult for females to attach eggs on the waterproof sheet, and eggs fall from the sheet, make the sheet textured with sandpaper.</li>
	<li>Cover the aquarium bottom with a layer of sand, 1 - 2 cm thick. Insert approximately one-third of the PVC pipe obliquely into the sand, with the sheet-fixed side down.</li>
	<li>After successful spawning by the fish, remove the sheet with the egg mass from the PVC pipe and place it in a Petri dish with aquarium water. Fix a new sheet inside the PVC pipe and place this back on the bottom of the aquarium. Finally, count the eggs by photographing the egg mass.
	<ol>
		<li>Alternatively, if observations of parental egg care are to be undertaken after spawning, be careful not to bring the eggs out of the water. Rather, use a Petri dish to scoop up the eggs and sheet along with aquarium water, photograph the egg mass quickly, and then carefully reaffix the sheet with the egg mass back into the same PVC pipe before returning it all to its original position on the bottom of the aquarium. Finally, count the eggs using the picture.</li>
	</ol>
	</li>
	<li>Count the eggs using ImageJ (Figure 3). These automatic and manual cell counting methods are explained in detail in the following: Basic cell counting (The University of Chicago, Integrated Light Microscopy Core Facility, http://digital.bsd.uchicago.edu/image_j.php); Particle Analysis (http://imagej.net/Particle_Analysis); Cell Counter (https://imagej.net/Cell_Counter).
	<ol>
		<li>Otherwise, if eggs are deposited densely, estimate the number of eggs by calculating the area-density ratio: count eggs in a printed 5 x 5 mm2 grid to calculate the egg density, measure the surface area that is covered with eggs by ImageJ, and estimate the total number of eggs from above density per unit and surface area. The above surface area measurement method by ImageJ is explained in the following: Set scale&#8230; (Research Services Branch, https://imagej.nih.gov/ij/docs/menus/analyze.html); it explains how to define the spatial scale of the image; Using ImageJ to Measure Surface Area (Keene State college, Academic Technology, https://dept.keene.edu/at/2011/04/15/by-matthew-ragan/). 
		NOTE: If eggs are transparent, the black waterproof sheet is more suitable for the automatic counting method because ImageJ can recognize the outline of eggs individually.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The reproductive ecology of numerous fishes has often been revealed through experimental rearing. Especially, sex change6,8,14, mate choice15,16 and intraspecific competition7,17 have been frequent topics of detailed investigations using aquarium-kept fish. Furthermore, some results observed in aquaria have been ...

Spectacular adaptive evolution is evident in the morphology, ecology and behavior of fishes1. Especially, ecological features relating to reproduction are especially diverse, and most of these can be directly influenced by individual fitness2. To gain insight into selective pressures that have led to the evolution of unique features in different fish species, direct observation of reproductive and social behaviors using live fish is often beneficial to substantiate theoretical hypotheses.
However, continuous field observations of fish may require specialized underwater equipment and facilities that are difficult to maintain. In these cases, observations of wild-caught aquarium-reared fish can be helpful3,4,5. In addition, efficient observations of fish behaviors that are otherwise rare or difficult to observe under natural conditions can become possible by manipulating experiments in aquaria6,7,8. Rearing fish under good conditions by minimizing artificial stress and physical damage is critical for accurate ecological investigations.
The pygmy goby Trimma marinae reaches 23-25 mm total length and is distributed in the western Pacific Ocean, where it is found in quiet, sheltered bays, at depths of 9-26 m9. In this work, T. marinae is used as a model to describe a series of basic techniques for the collection of fish using the self-contained underwater breathing apparatus (SCUBA), fish transport, and eventual acclimation of the fish to aquaria for direct observation of the study species&#39; reproductive behavior and ecology.

<table><tbody><tr><td>Hand net</td><td>Nisso</td><td>AQ-17</td><td>Select for the target species size.</td></tr><tr><td>Polyethylene bag</td><td>San-U Fish Farm</td><td>8194</td><td></td></tr><tr><td>Rubber band</td><td>ESCO Co. LTD.</td><td>78-0420-64</td><td>&amp;oslash; 80 mm x 6 mm</td></tr><tr><td>Oxygen cylinder</td><td>N/A</td><td>N/A</td><td>Oxgen cylinder for diving equipment suits.</td></tr><tr><td>Elbagin</td><td>Japan Pet Design Co. Ltd.</td><td>75950</td><td>Pafurazine F (provided from same company) is equivalent drug to Elbagin.</td></tr><tr><td>Polystyrene foam box</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Pipette</td><td>AS ONE</td><td>1-8625-04</td><td></td></tr><tr><td>Rope</td><td>Mizukami Kinzoku Co. LTD.</td><td>95301601</td><td></td></tr><tr><td>Weight</td><td>N/A</td><td>N/A</td><td>Weight for diving equipment suits.</td></tr><tr><td>Water tank</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Air pump</td><td>KOTOBUKI</td><td>4972814 062115</td><td></td></tr><tr><td>Air stone</td><td>KOTOBUKI</td><td>4972814 232204</td><td></td></tr><tr><td>2-Methylquinoline&amp;nbsp;</td><td>Wako</td><td>170-00376</td><td></td></tr><tr><td>Ethanol (99.5)</td><td>Wako</td><td>057-00456</td><td></td></tr><tr><td>Visible implant elastomer tag kit</td><td>Northwest Marine Technology</td><td>N/A</td><td>http://www.nmt.us/products/vie/vie.shtml</td></tr><tr><td>Soft sponge</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>PVC board</td><td>N/A</td><td>N/A</td><td>0.3 mm thickness is easy to use.</td></tr><tr><td>Petri dish</td><td>N/A</td><td>N/A</td><td>A large one, such as&amp;nbsp; &amp;oslash; 160 mm and 30 mm depth, is convenient for the injection of the VIE tag.</td></tr><tr><td>Transparent acrylic board</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>UVA filtered light&amp;nbsp;</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>PVC pipe</td><td>N/A</td><td>N/A</td><td>&amp;oslash; 5 cm</td></tr><tr><td>Waterproof sheet</td><td>SOMAR Corp.</td><td>3EKW03</td><td>The film for the plain copier.</td></tr><tr><td>Sand</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Stereo microscope</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Camera</td><td>N/A</td><td>N/A</td><td></td></tr></tbody></table>

basic methods for the study of reproductive ecology of fish in aquaria

Captive-rearing observations are valuable for revealing aspects of fish behavior and ecology when continuous field investigations are impossible. Here, a series of basic techniques are described to enable observations of the reproductive behavior of a wild-caught gobiid fish, as a model, kept in an aquarium. The method focuses on three steps: collection, transport, and observations of reproductive ecology of a substrate spawner. Essential aspects of live fish collection and transport are (1) preventing injury to the fish, and (2) careful acclimation to the aquarium. Preventing harm through injuries such as scratches or a sudden change of water pressure is imperative when collecting live fish, as any physical damage is likely to negatively affect the survival and later behavior of the fish. Careful acclimation to aquaria decreases the incidence death and mitigates the shock of transport. Observations during captive rearing include (1) the identification of individual fish and (2) monitoring spawned eggs without negative effects to the fish or eggs, thereby enabling detailed investigation of the study species' reproductive ecology. The subcutaneous injection of a visible implant elastomer (VIE) tag is a precise method for the subsequent identification of individual fish, and it can be used with a wide size range of fish, with minimal influence on their survival and behavior. If the study species is a substrate spawner that deposits adhesive eggs, an artificial nest site constructed from polyvinyl chloride (PVC) pipe with the addition of a removable waterproof sheet will facilitate counting and monitoring the eggs, lessening the investigator's influence on the nest-holding and egg-guarding behavior of the fish. Although this basic method entails techniques that are seldom mentioned in detail in research articles, they are fundamental for undertaking experiments that require the captive rearing of a wild fish.

Following the above methods, 41, 15 and 96 individuals of T. marinae were collected in April 2014, 2015 and 2016, respectively, offshore of Amami Oshima, Kagoshima Prefecture, Japan (Table 1). In each case, 25 (61%), 14 (93%) and 91 (95%) individuals lived until depositing eggs in an aquarium. As reported in Fukuda et al.3, only one fish died before the end of the observation period in 2014, and fish spawning otherwise appeared to...

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Basic Methods for the Study of Reproductive Ecology of Fish in Aquaria

A series of basic methods to enable the study of the reproductive ecology of fish kept in aquaria are described. These are useful protocols for collecting fish using SCUBA, transporting live fish, and observing the reproductive behavior of wild-caught fish kept in aquaria.

Captive-rearing observations are valuable for revealing aspects of fish behavior and ecology when continuous field investigations are impossible. Here, a ...

basic-methods-for-the-study-of-reproductive-ecology-of-fish-in-aquaria

Research

JoVE Journal

Behavior

11.5K Views.  Tokyo University of Marine Science and Technology. A series of basic methods to enable the study of the reproductive ecology of fish kept in aquaria are described. These are useful protocols for collecting fish using SCUBA, transporting live fish, and observing the reproductive behavior of wild-caught fish kept in aquaria.

Video: Basic Methods for the Study of Reproductive Ecology of Fish in Aquaria

Development of New Methods for Quantifying Fish Density Using Underwater Stereo-video Tools

The use of video camera systems in ecological studies of fish continues to gain traction as a viable, non-extractive method of measuring fish lengths and estimating fish abundance. We developed and implemented a rotating stereo-video camera tool that covers a full 360 degrees of sampling, which maximizes sampling effort compared to stationary camera tools. A variety of studies have detailed the ability of static, stereo-camera systems to obtain highly accurate and precise measurements of fish; the focus here was on the development of methodological approaches to quantify fish density using rotating camera systems. The first approach was to develop a modification of the metric MaxN, which typically is a conservative count of the minimum number of fish observed on a given camera survey. We redefine MaxN to be the maximum number of fish observed in any given rotation of the camera system. When precautions are taken to avoid double counting, this method for MaxN may more accurately reflect true abundance than that obtained from a fixed camera. Secondly, because stereo-video allows fish to be mapped in three-dimensional space, precise estimates of the distance-from-camera can be obtained for each fish. By using the 95% percentile of the observed distance from camera to establish species-specific areas surveyed, we account for differences in detectability among species while avoiding diluting density estimates by using the maximum distance a species was observed. Accounting for this range of detectability is critical to accurately estimate fish abundances. This methodology will facilitate the integration of rotating stereo-video tools in both applied science and management contexts.

We describe a new method for counting fishes, and estimating relative abundance (MaxN) and fish density using rotating stereo-video camera systems. We also demonstrate how to use distance from camera (Z distance) to estimate species-specific detectability.

The use of video camera systems in ecological studies of fish continues to gain traction as a viable, non-extractive method of measuring fish lengths and ...

Environment

Protocol for Acute and Chronic Ecotoxicity Testing of the Turquoise Killifish Nothobranchius furzeri

The killifish Nothobranchius furzeri is an emerging model organism in the field of ecotoxicology and its applicability in acute and chronic ecotoxicity testing has been demonstrated. Overall, the sensitivity of the species to toxic compounds is in the range with, or higher than, that of other model species.
This work describes protocols for acute, chronic, and multigenerational bioassays of single and combined stressor effects on N. furzeri. Due to its short maturation time and life-cycle, this vertebrate model allows the study of endpoints such as maturation time and fecundity within four months. Transgenerational full life-cycle exposure trials can be performed in as little as 8 months. Since this species produces eggs that are drought-resistant and remain viable for years, the on-site culture of the species is not needed but individuals can be recruited when required. The protocols are designed to measure life-history traits (mortality, growth, fecundity, weight) and critical thermal maximum.

Protocol for Acute and Chronic Ecotoxicity Testing of the Turquoise Killifish Nothobranchius furzeri

In this work, we describe an acute, chronic and multigenerational bioassay to study the effects of single and combined stressors on the Turquoise killifish Nothobranchius furzeri. This protocol is designed to study life-history traits (mortality, growth, fecundity, weight) and critical thermal maximum.

The killifish Nothobranchius furzeri is an emerging model organism in the field of ecotoxicology and its applicability in acute and chronic ecotoxicity ...

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.

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.

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

Genetics

Using the FishSim Animation Toolchain to Investigate Fish Behavior: A Case Study on Mate-Choice Copying In Sailfin Mollies

Over the last decade, employing computer animations for animal behavior research has increased due to its ability to non-invasively manipulate the appearance and behavior of visual stimuli, compared to manipulating live animals. Here, we present the FishSim Animation Toolchain, a software framework developed to provide researchers with an easy-to-use method for implementing 3D computer animations in behavioral experiments with fish. The toolchain offers templates to create virtual 3D stimuli of five different fish species. Stimuli are customizable in both appearance and size, based on photographs taken of live fish. Multiple stimuli can be animated by recording swimming paths in a virtual environment using a video game controller. To increase standardization of the simulated behavior, the prerecorded swimming path may be replayed with different stimuli. Multiple animations can later be organized into playlists and presented on monitors during experiments with live fish.
In a case study with sailfin mollies (Poecilia latipinna), we provide a protocol on how to conduct a mate-choice copying experiment with FishSim. We utilized this method to create and animate virtual males and virtual model females, and then presented these to live focal females in a binary choice experiment. Our results demonstrate that computer animation may be used to simulate virtual fish in a mate-choice copying experiment to investigate the role of female gravid spots as an indication of quality for a model female in mate-choice copying.
Applying this method is not limited to mate-choice copying experiments but can be used in various experimental designs. Still, its usability depends on the visual capabilities of the study species and first needs validation. Overall, computer animations offer a high degree of control and standardization in experiments and bear the potential to &#39;reduce&#39; and &#39;replace&#39; live stimulus animals as well as to &#39;refine&#39; experimental procedures.

Using the FishSim Animation Toolchain to Investigate Fish Behavior: A Case Study on Mate-Choice Copying In Sailfin Mollies

Using the novel FishSim Animation Toolchain, we present a protocol for non-invasive visual manipulation of public information in the context of mate-choice copying in sailfin mollies. FishSim Animation Toolchain provides an easy-to-use framework for the design, animation and presentation of computer-animated fish stimuli for behavioral experiments with live test fish.

Over the last decade, employing computer animations for animal behavior research has increased due to its ability to non-invasively manipulate the ...

A Standardized Protocol for Preference Testing to Assess Fish Welfare

Animal welfare assessment techniques try to take into consideration the specific needs and wants of the animal in question. Providing enrichment (the addition of physical objects or conspecifics in the housing environment) is often a way to give captive animals the opportunity to choose who or what they interact with and how they spend their time. A fundamental component of the aquatic environment that is often overlooked in captivity, however, is the ability for the animal to choose to engage in physical exercise. For many animals, including fish, exercise is an important aspect of their life history, and is known to have many health benefits, including positive changes in the brain and behavior. Here we present a method for assessing habitat preferences in captive animals. The protocol could easily be adapted to look at a variety of environmental factors (e.g., gravel versus sand as a substrate, plastic plants versus live plants, low flow versus high flow of water) in different aquatic species, or for use with terrestrial species. Statistical assessment of preference is carried out using Jacob&#39;s preference index, which ranks the habitats from -1 (avoidance) to +1 (most preferred). With this information, it can be determined what the animal wants from a welfare perspective, including their preferred location.

A fundamental aspect of assessing the welfare of animals in captivity is to ask whether the animals have what they want. Here, we present a protocol to determine housing preference in the zebrafish (Danio rerio) with respect to the presence/absence of environmental enrichment and access to flowing of water.

Animal welfare assessment techniques try to take into consideration the specific needs and wants of the animal in question. Providing enrichment (the ...

A Novel Method of Drug Administration to Multiple Zebrafish (Danio rerio) and the Quantification of Withdrawal

Anxiety testing in zebrafish is often studied in combination with the application of pharmacological substances. In these studies, fish are routinely netted and transported between home aquaria and dosing tanks. In order to enhance the ease of compound administration, a novel method for transferring fish between tanks for drug administration was developed. Inserts that are designed for spawning were used to transfer groups of fish into the drug solution, allowing accurate dosing of all fish in the group. This increases the precision and efficiency of dosing, which becomes very important in long schedules of repeated drug administration. We implemented this procedure for use in a study examining the behavior of zebrafish in the light/dark test after administering ethanol with differing 21 day schedules. In fish exposed to daily-moderate amounts of alcohol there was a significant difference in location preference after 2 days of withdrawal when compared to the control group. However, a significant difference in location preference in a group exposed to weekly-binge administration was not observed. 
This protocol can be generalized for use with all types of compounds that are water-soluble and may be used in any situation when the behavior of fish during or after long schedules of drug administration is being examined. The light/dark test is also a valuable method of assessing withdrawal-induced changes in anxiety.

A Novel Method of Drug Administration to Multiple Zebrafish Danio rerio and the Quantification of Withdrawal

A novel method for reducing variability when exposing fish to drugs is explained. Fish exposed to various patterns of ethanol exposure were found to have altered anxiety levels during withdrawal in a light/dark scototaxic assay.

Anxiety testing in zebrafish is often studied in combination with the application of pharmacological substances. In these studies, fish are routinely ...

Neuroscience

Boldness, Aggression, and Shoaling Assays for Zebrafish Behavioral Syndromes

A behavioral syndrome exists when specific behaviors interact under different contexts. Zebrafish have been test subjects in recent studies and it is important to standardize protocols to ensure proper analyses and interpretations. In our previous studies, we have measured boldness by monitoring a series of behaviors (time near surface, latency in transitions, number of transitions, and darts) in a 1.5 L trapezoidal tank. Likewise, we quantified aggression by observing bites, lateral displays, darts, and time near an inclined mirror in a rectangular 19 L tank. By dividing a 76 L tank into thirds, we also examined shoaling preferences. The shoaling assay is a highly customizable assay and can be tailored for specific hypotheses. However, protocols for this assay also must be standardized, yet flexible enough for customization. In previous studies, end chambers were either empty, contained 5 or 10 zebrafish, or 5 pearl danios (D. albolineatus). In the following manuscript, we present a detailed protocol and representative data that accompany successful applications of the protocol, which will allow for replication of behavioral syndrome experiments.

This manuscript describes the setup, implementation, and analysis of boldness, aggression, and shoaling in zebrafish and testing for the presence of a behavioral syndrome. A standardized approach for behavioral quantification will allow for easier comparison across studies. Modifications to this protocol are possible as each assay can be run individually.

A behavioral syndrome exists when specific behaviors interact under different contexts. Zebrafish have been test subjects in recent studies and it is ...

Medicine

Regular Care and Maintenance of a Zebrafish (Danio rerio) Laboratory: An Introduction

This protocol describes regular care and maintenance of a zebrafish laboratory. Zebrafish are now gaining popularity in genetics, pharmacological and behavioural research. As a vertebrate, zebrafish share considerable genetic sequence similarity with humans and are being used as an animal model for various human disease conditions. The advantages of zebrafish in comparison to other common vertebrate models include high fecundity, low maintenance cost, transparent embryos, and rapid development. Due to the spur of interest in zebrafish research, the need to establish and maintain a productive zebrafish housing facility is also increasing. Although literature is available for the maintenance of a zebrafish laboratory, a concise video protocol is lacking. This video illustrates the protocol for regular housing, feeding, breeding and raising of zebrafish larvae. This process will help researchers to understand the natural behaviour and optimal conditions of zebrafish husbandry and hence troubleshoot experimental issues that originate from the fish husbandry conditions. This protocol will be of immense help to researchers planning to establish a zebrafish laboratory, and also to graduate students who are intending to use zebrafish as an animal model.

Regular Care and Maintenance of a Zebrafish Danio rerio Laboratory: An Introduction

This protocol outlines regular maintenance and care to maintain optimal conditions for zebrafish husbandry. The video illustrates the protocol for system maintenance, regular housing, feeding, breeding, and raising of zebrafish larvae.

This protocol describes regular care and maintenance of a zebrafish laboratory. Zebrafish are now gaining popularity in genetics, pharmacological and ...

Biology

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 ...

Sperm Collection and Computer-Assisted Sperm Analysis in the Teleost Model Japanese Medaka (Oryzias latipes)

Japanese medaka (Oryzias latipes) is a teleost fish and&#160;an emerging vertebrate model for ecotoxicology, developmental, genetics, and physiology research. Medaka is also used extensively to investigate vertebrate reproduction, which is an essential biological function as it allows a species to perpetuate. Sperm quality is an important indicator of male fertility and, thus, reproduction success. Techniques for extracting sperm and sperm analysis are well documented for many species, including teleost fish. Collecting sperm is relatively simple in larger fish but can be more complicated in small model fish as they produce less sperm and are more delicate. This article, therefore, describes two methods of sperm collection in the small model fish, Japanese medaka: testes dissection and abdominal massage. This paper demonstrates that both approaches are feasible for medaka and shows that abdominal massage can be performed a repeated number of times as the fish quickly recover from the procedure. This article also describes a protocol for computer-assisted sperm analysis in medaka to objectively assess several important indicators of medaka sperm quality (motility, progressivity, duration of motility, relative concentration). These procedures, specified for this useful small teleost model, will greatly enhance understanding of the environmental, physiological, and genetic factors influencing fertility in vertebrate males.

Sperm Collection and Computer-Assisted Sperm Analysis in the Teleost Model Japanese Medaka Oryzias latipes

This article describes two quick and efficient methods for collecting sperm from the small model fish medaka (Oryzias latipes), as well as a protocol for reliably assessing sperm quality using computer-assisted sperm analysis (CASA).

Japanese medaka (Oryzias latipes) is a teleost fish and&#160;an emerging vertebrate model for ecotoxicology, developmental, genetics, and physiology ...

Basic Methods for the Study of Reproductive Ecology of Fish in Aquaria

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Rozdziały w tym wideo

Development of New Methods for Quantifying Fish Density Using Underwater Stereo-video Tools

Protocol for Acute and Chronic Ecotoxicity Testing of the Turquoise Killifish Nothobranchius furzeri

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

Using the FishSim Animation Toolchain to Investigate Fish Behavior: A Case Study on Mate-Choice Copying In Sailfin Mollies

A Standardized Protocol for Preference Testing to Assess Fish Welfare

A Novel Method of Drug Administration to Multiple Zebrafish (Danio rerio) and the Quantification of Withdrawal

Boldness, Aggression, and Shoaling Assays for Zebrafish Behavioral Syndromes

Regular Care and Maintenance of a Zebrafish (Danio rerio) Laboratory: An Introduction

Gamete Collection and In Vitro Fertilization of Astyanax mexicanus

Sperm Collection and Computer-Assisted Sperm Analysis in the Teleost Model Japanese Medaka (Oryzias latipes)