Name: Using an Automated 3D-tracking System to Record Individual and Shoals of Adult Zebrafish (Video) | JoVE Journal
Uploaded: 2013-12-05T16:00:00.000+00:00
Duration: 14 min 3 s
Description: Explore automated 3D-tracking of zebrafish shoals using advanced algorithms and refraction-corrected imaging. This method enables precise analysis of kinematic, spatial, and social behaviors for neuroscience research.

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The authors have no acknowledgements.

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<li>Wong, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Analyzing+habituation+responses+to+novelty+in+zebrafish+%28Danio+rerio%29%5BTitle%5D)+AND+%22Behav.+Brain+Res%22%5BJournal%5D)">Analyzing habituation responses to novelty in zebrafish (Danio rerio).</a> Behav. Brain Res. 208, 450-457 (2010).</li>
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</ol>

Hans Maaswinkel, Liqun Zhu and Wei Weng are employees of xyZfish, Ronkonkoma NY, that produces the tracking system used in this article.

The experimental procedure shown here allows recording individual zebrafish, pairs of zebrafish (e.g. to study mating or aggression) and shoals of zebrafish (as shown in the representative example). The system is especially suited for simultaneously obtaining information about 3D (vertical and horizontal) distribution, kinematic characteristics (e.g. velocity, acceleration, turning angle) and social parameters (e.g. social cohesion). In the following, we list a few important issues to consider when planning experiments.
Depending on the experimental procedure, different behavioral systems can be studied. For instance, to explore neophobia (i.e. fear for new environments) the recording should be started immediately after introducing the zebrafish to the observation tank (after waiting just for a few seconds for the water to settle down to diminish noise), without prior habituation11,17,37. On the other hand, some research topics might require the zebrafish to be well habituated, for instance when studying the effects of alarm pheromones16. When studying groups of zebrafish, the experimenter should be aware that although the software minimizes switching of tags, it is at this moment not possible to follow individual zebrafish throughout. This is also not important for shoaling studies where the emphasis is on social cohesion, but would be crucial when exploring aggression or mating. Manual reassignment of tags is an option, but can be very time-consuming.
The color of the walls of the observation container and of the observation chamber can have great influence on the behavior of the zebrafish. For instance, white walls have been described to induce aversion responses36, 37. When trying different colors, keep two things in mind: 1) it is important that the front wall (i.e. the wall closest to the camera) remains transparent and that a lid (if one is used) also has to be transparent; 2) when selecting a color never used before, first determine whether the contrast between zebrafish and background is appropriate for recording. Do a test run. Changing the camera settings for recording (Gain, Shutter, Red and Blue, as explained in steps 2.5 and 2.6) and the settings for trajectory reconstruction (e.g. color and size thresholds, as explained in steps 3.3, 3.5, 3.11, and 3.12) can make a big difference. Again, experience with the system will go a long way to decide what is possible.
Depending on the purpose of the experiment, a light source with different characteristics (e.g. spectrum, intensity) might be desirable. The LED bars can easily be replaced. Make sure that the light reaches every part of the container.
If the experiment requires that the zebrafish remains in the observation container for several hours, make sure to provide aeration. An air tube can be attached to the back right corner by using a clip. Check on the computer screen that the clip does not interfere with the recording, i.e. that the zebrafish is at no time hidden from view. In Figures 2A and 2B the black clip can be seen in the top view. Ideally, aeration should be provided before or between recordings, but not during recording as to prevent noise. Also note that switching strong aeration on or off might startle the zebrafish.
If the experiment requires keeping the zebrafish in the container overnight, it is useful to cover the observation chamber with a transparent lid to prevent the zebrafish from leaping out (which is a very common occurrence). If the cover is not removed before recording (as not to disturb the zebrafish), consider the following two points: 1) make sure that the mirror images of the light sources do not obstruct the recording (e.g. by shifting the light sources or by choosing a lid that does reflect light only minimally); 2) consider that the lid might become foggy when used overnight, especially since in this case aeration has also to be provided; one way to reduce this is to choose a lid with small holes in it.
For some experiments it might be essential to determine the baseline behavior of the zebrafish before administering a drug. In this case, the drug can be added between sessions by way of a tube (similar to the aeration tube discussed above).
Although the focus here was on recording zebrafish (Danio rerio), other fish species can, of course, also be recorded with the system. It has been used to study behaviors in (juvenile) goldfish (Carassius auratus), tiger barb (Barbus tetrazona), and green swordtail (Xiphophorus helleri).
Finally it should be noted that other dimensions of the observation tanks are principally possible. However, it would require reconfiguring the system and finding a good balance between spatial and temporal resolution when using much larger observation containers. Furthermore, the shape of the container has to be cuboid. The number of zebrafish or other experimental subjects is theoretically unlimited; however tag swapping increases with number of subjects.

Zebrafish has become an important model for biological and pharmacological research1-3. Tracking of movement and spatial distribution patterns of individual zebrafish and of shoals of zebrafish is invaluable. Older studies apply manual quantification based on recorded images4. Now, 2D automatic video tracking systems are commonly used5. However, since zebrafish and many other aquatic and nonaquatic animals move in a three-dimensional space, the interest in 3D systems is growing6. The described 3D system was first published in 20077. For technical details and comparisons with other systems see8. The basic advantage of this system as compared to other systems is that it acquires two (optionally three) synchronous and independent views using a mirroring system and that is uses a two-step calibration protocol. In the first step, the 3D-geometry of the stereo system (formed by the camera and the mirrors) is calibrated. In the second step, the nonlinear errors introduced by light refraction are corrected. When studying aquatic animals, the impact of light refraction can be considerable when left uncorrected. Other so far published systems (even newer developments9) omit the two-step calibration procedure. For technical details about object identification, occlusion with multi-object tracking, jigging noise etc. the reader is referred to the original publications8,9. The system has been validated for the study of anxiety-like behavioral responses10,11, shoaling8,11,12 and, in a preliminary study, for antagonistic behavior in zebrafish13. It has also been tested for other fish species (goldfish8, swordtail, tiger barb) and for mice9.
In the following the protocols for calibration and for using the system are described and the results of an experiment testing the effects of the NMDA-receptor noncompetitive antagonist MK-801 on shoaling zebrafish are presented as a representative example to demonstrate the type of data that can be obtained with the system.
Both in the natural environment and in aquariums, zebrafish usually swims in shoals14. Shoaling preferences seems to depend on several factors, such as rearing, sex, and food availability14,15. Interestingly, the distance between individual zebrafish in a shoal becomes smaller when signals of potential danger are present, such as alarm pheromones16 or novelty11. The shoaling paradigm is therefore also being used to test anxiolytic drugs17. Dominance hierarchies18 and social learning19 are fascinating aspect of zebrafish behavior. Because of its highly developed social behavioral character, zebrafish have also become of interest for autism research20,21.
MK-801 has been reported to alter social behavior in zebrafish5,22,23, which is reminiscent of its effects on mammals24. However, MK-801 has also many other effects on other behaviors (such as swimming behavior, inhibitory avoidance and place preference25-27). Therefore it is crucial, in initial experiments to obtain as much information as possible from basic observations. This will provide the basis for designing more targeted experiments.
Survey of the recording apparatus and software
In its current configuration, the observation container (25 cm x 25 cm x 18 cm, L x W x H), holding the zebrafish, is positioned in the observation chamber (91 cm x 46 cm x 56 cm, L x W x H). The chamber also contains a camera, a mirror which is suspended at approximately &#62;45&#176; angle above the container, and LED-light bars. Figure 1 shows a simplified diagram. The camera records both the front view and the top view (mirror) to construct the 3D trajectories of the zebrafish by determining its x-, y-, and z-coordinates. For orientation purposes, the coordinate vectors are presented on the left-front corner of the observation container (position relative to camera), which corresponds with the vectors presented on the 3D views (e.g. Figure 15) generated by the Path Reconstruction Module (see further below). The sampling rate depends on the camera and the computer. In our experiments it is approximately 40 frames/sec&#160;(fps). The spatial resolution of the recorded frames is 640 x 480 pixels (this configuration was chosen as optimal balance between spatial and temporal resolution). Windows XP or higher is required.
Before the first recording, the system has to be calibrated to determine locations of the walls of the container and the water surface, to determine the scaling factor of the recorded frames and to adjust for the sizable error generated by refraction of light when passing from water to air. In principle, calibration has only to be performed during the initial installation and after parts of the setup, such as camera or mirror, are moved or replaced. Nevertheless, it is good practice to check from time to time whether the calibration is still correct and recalibrate, if necessary.
The software consists of three modules. The Recordings Module records the frames (two example frames are shown in Figure 2). The Trajectory Module reconstructs the paths for every zebrafish. The Data Processing Module calculates a range of standard kinematic and spatial parameters. For the purpose of further analysis, the data can be imported into spreadsheet or statistical software.

<table><tbody><tr><td>AABT-3D Tracking Hardware</td><td>xyZfish</td><td>AABT II</td><td>Includes camera, video card, observation compartment with mirror and lighting</td></tr><tr><td>AABT-3D Calibration equipment</td><td>xyZfish</td><td>calib</td><td>Contains calibration panel and side mirror</td></tr><tr><td>AABT-3D Observation containers</td><td>xyZfish</td><td>cubes</td><td>&amp;nbsp;</td></tr><tr><td>AABT-3D Tracking software</td><td>xyZfish</td><td>v.1.0</td><td>&amp;nbsp;</td></tr><tr><td>Computer</td><td>optional</td><td>N/A</td><td>Windows XP or higher</td></tr><tr><td>(+)-MK-801 hydrogen maleate</td><td>Sigma</td><td>M107</td><td>&amp;nbsp;</td></tr><tr><td>Air pump</td><td>Fusion</td><td>500</td><td>&amp;nbsp;</td></tr><tr><td>Air line tubing 3/16&amp;rdquo;</td><td>Lee&amp;rsquo;s Aquarium and Pet Products</td><td>14508</td><td>&amp;nbsp;</td></tr><tr><td>Medium Binder Clips</td><td>OfficeDepot</td><td>561339</td><td>To attach airline to observation container</td></tr></tbody></table>

using an automated 3d-tracking system to record individual and shoals of adult zebrafish

Using an Automated 3D-tracking System to Record Individual and Shoals of Adult Zebrafish

Like many aquatic animals, zebrafish (Danio rerio) moves in a 3D space. It is thus preferable to use a 3D recording system to study its behavior. The presented automatic video tracking system accomplishes this by using a mirror system and a calibration procedure that corrects for the considerable error introduced by the transition of light from water to air. With this system it is possible to record both single and groups of adult zebrafish. Before use, the system has to be calibrated. The system consists of three modules: Recording, Path Reconstruction, and Data Processing. The step-by-step protocols for calibration and using the three modules are presented. Depending on the experimental setup, the system can be used for testing neophobia, white aversion, social cohesion, motor impairments, novel object exploration etc. It is especially promising as a first-step tool to study the effects of drugs or mutations on basic behavioral patterns. The system provides information about vertical and horizontal distribution of the zebrafish, about the xyz-components of kinematic parameters (such as locomotion, velocity, acceleration, and turning angle) and it provides the data necessary to calculate parameters for social cohesions when testing shoals.

The use of a 3D automatic video system that can track individual and groups of zebrafish is described. As application example we explore the effects of the NMDA-receptor antagonist MK-801 on shoals of zebrafish.

Like many aquatic animals, zebrafish (Danio rerio) moves in a 3D space. It is thus preferable to use a 3D recording system to study its behavior. The ...

The overall goal of this procedure is to study the three dimensional movement patterns of zebra fish or other aquatic animals and to analyze their kinematic, spatial, and social characteristics. This allows detection of altered behaviors due to genetic mutations or chemical and environmental factors. This is accomplished by first calibrating the system, which also corrects for the sizable image distortion generated by refraction.The second step is to record video frames at relatively high temporal resolution from at least two different angles simultaneously. Next, the software reconstructs the trajectories of the fish. Advanced algorithms are applied to reduce the effect of noise and to infer the position of a fish when occlusion or clustering occurs.The final step is to analyze the data and to perform statistical analysis for velocity acceleration, turning angle, horizontal and vertical distribution in the observation container, inter individual distances and other parameters of interest. Ultimately, the high speed calibrated 3D tracking is useful to study the behavioral deficits in mutant zebrafish and the effects of drugs on a variety of behavioral systems. For the purpose of drug discovery, One advantage of the system is that it allows studying behavior of fish in 3D.This is important because fish naturally move in all three dimensions and when recording multiple fish simultaneously, 2D recording can be very misleading when investigating the spatial configurations of the group. Another advantage of the system is that the corrects for light refraction, which when left uncorrected for, can result in significant measurement errors In its current configuration. The observation container holding the zebra fish is positioned in the observation chamber.The chamber also contains a camera, a mirror, which is suspended at approximately 45 degrees above the container and LED light bars. Before recording, calibrate the system to determine the container wall and water surface locations and the scaling factor of the recorded frames also adjust for the error generated by refraction of light when passing from water to air. Once calibration is complete, a test run should be performed with a dummy fish.Recalibration should be performed if the positions of the mirror or camera are off or if there are problems with tracking. To begin a recording session, place the observation container filled up to the marks with water into the observation chamber. The feet of the container should be inserted into the alignment holes.Orient the containers such that the semitransparent or opaque walls face the far side and to the right as seen from the camera. Next, switch on the lights in the compartment, then put the fish into the container and close the curtains. Open the record module and image of the front and top view of the container will appear on screen.Initially, the acquired image may not be ideal and hence might need some adjustment to obtain sufficient contrast between the fish and the background. To adjust the image, select camera control, and then in the general settings window that opens, deselect the boxes shown here, and then adjust shutter and gain in such a way that the image is not overexposed. Next, on the left side of the camera control window, select white balance color, and in the new window, deselect the auto box and adjust the red and blue values to obtain the best color contrast in both the front and top view.Multiple adjustments may be necessary after the adjustments are complete. From the main menu of the record module, select data location to create or select a folder. Each recording should have its own folder, which should be a sub folder of the experiment folder.Now press start recording and enter a whole number for the duration of the recording in minutes, then press okay and the experimental recording session will commence. Once the recordings are completed, copy and paste the animal container data and the correspondence data sub folders from the calibration folder to the folders previously created During recording to begin path reconstruction, open the software's trajectory module. Then press data location to select the folder that was created with the record module.Next, press the system setup button to open the parameters window first, adjust the top end bottom view color thresholds. A good starting value is 1000. Next, select top end, bottom view sized thresholds.A good starting size for a mid-sized zebra fish is 10 or use 20 for a big fish and five for a small fish. The higher these thresholds, the less likely the software is to pick up noise. However, with too high a value, it might not be able to detect the fish.It may become necessary to readjust these thresholds later on in the experiment. For upper size threshold, a value of 3000 is usually sufficient. And for specific length, keep the 220 preset value.The number of fish refers to the number of fish in the observation tank. Max numb of points refers to the total number of frames, record or remember this number and press.Okay. Then select acquire background.The next window will ask to fill in from and two fields. After loading the background, adjust the three red boxes using the mouse, the boxes mark the areas in which the software is looking for the fish. Minimize the box on the left.Since the side view is used only for calibration, now press the start generating trajectories button and path reconstruction will start. Ideally, the tag should be placed on the image of the fish if it is often placed incorrectly, open the info window. Here we see an ideal case where only the outlines of the fish are seen.In the case of excessive noise, press stop in the main menu and choose new color and size thresholds. Also, check if the red boxes can be reduced slightly. For example, at the top, to exclude the noise generated by water movements, watch the trajectory on the 3D screen.By right clicking with the mouse, the researcher can select between rotation and translations of the 3D configuration. To begin data processing, open the data processing module, select project on the toolbar, and then from the pull down menu, select new and enter an experiment name. Next, select edit project.Then add to project followed by new group and fill in a name for the experimental group. Now select the experimental group in the right window. Note that the toolbar will change.Select edit group, then add to group, and then new measurements. This will open a browser window. Next, open the folder created during recording and select the sub folder trajectories and then the point smoothened file.In the example presented here for group A, six files were loaded and for group B four files. To begin the analysis, select the point smoothened file in the right window. Then select data processing and single value endpoints from the pull down menu.By way of illustration, select duration freezing. A window will open to specify freezing in terms of speed threshold and duration threshold to familiarize yourself with the program. Try other examples such as travel distance depth distribution, which allows one to determine the number of depth levels and burst frequency.When selecting the option, batch processing using default threshold about 200 endpoints will be calculated in this context. Default means that delimiters have been set. For example, currently depth distribution is calculated for three depth levels and angular distribution is calculated for four sections.After generating the parameters, a new window will open asking for the location and name of the file to be created. These will be saved in CSV format and can be imported into other software. Here, figure A shows a typical example frame for control.Zebra fish, while figure B shows a typical example frame for zebrafish treated with MK 801 both recorded during a morning session. Notice that the controlled zebra fish were close to the bottom and close together. In contrast, MK 801 zebra fish were close to the top and swam at considerable distance from one another.Here, two representative trajectories for controlled zebra fish and zebra fish treated with 10 micromolar. MK 801 are presented for a morning session and for an afternoon session. In this figure, average social distances for controls and fish treated with 10 micromolar MK eight one are presented for both sessions, and here the timelines of the average inter-individual distances are presented for both groups.Here in figure A, we see the distribution of the zebra fish over 10 equal depth levels for the morning session, and in B, the distribution for the afternoon session. The temporal distribution over the four radial zones of the experimental groups is seen here. The inset shows the locations of these zones from center to periphery, and here we see the temporal distribution over the four quadrants of the experimental groups.The inset shows the location of the quadrants. Double lines represent white walls, single lines represent transparent walls. The camera is located left of the inset IE, closest to quadrants one and four.Black asterisks denote comparisons between the groups from the morning session and gray asterisks for those from the afternoon sessions. Finally, the X, Y, Z components of travel distance for the zebra fish during the morning and the afternoon sessions are shown here. Mean and standard error mean are presented.When working with the system, remember that the data are only as good as the recording, so make sure that the system is well calibrated and that the camera settings are optimal. The rest is easy, and it's just a matter of becoming familiar with the different steps.

using-an-automated-3d-tracking-system-to-record-individual-shoals

Research

JoVE Journal

Biology

The VisioTracker is an automated system for the quantitative analysis of visual performance of larval and small adult fish based on the recording of eye movements. It features full control over visual stimulus properties and real-time analysis, enabling high-throughput research in fields such as visual system development and function, pharmacology, neural circuit studies and sensorimotor integration.

VisioTracker, an Innovative Automated Approach to Oculomotor Analysis

The video describes the avoidance and thigmotaxis assay of zebrafish larvae. The video examines the behavior of zebrafish larvae in response to visual aversive stimulus.

Zebrafish Avoidance and Thigmotaxis Assay: A High-Throughput Method to Examine Larval Behavior in Response to Aversive Visual Stimulus

Our laboratory developed a novel high-throughput automated imaging system that is useful for the detection of several different behaviors in 7-day-old zebrafish larvae. The system can be used for detecting subtle changes in behavior after the larvae have been exposed to environmental toxicants or pharmaceuticals.

Automated High-throughput Behavioral Analyses in Zebrafish Larvae

Zebrafish are an excellent model to study muscle function and disease. During early embryogenesis zebrafish begin regular muscle contractions producing rhythmic swimming behavior, which is altered when the muscle is disrupted. Here we describe a touch-evoked response and locomotion assay to examine swimming performance as a measure of muscle function.

Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish

Here, we present a protocol to design and fabricate a zebrafish embryo arraying template, followed by a detailed procedure on the use of such template for high throughput zebrafish embryo arraying into a 96-well plate.

A 3-dimensional (3D)-printed Template for High Throughput Zebrafish Embryo Arraying

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.

Behavioral Tracking and Neuromast Imaging of Mexican Cavefish

We describe a behavioral protocol designed to assess how zebrafish’s personalities influence their response to water currents and weak magnetic fields. Fishes with the same personalities are separated based on their explorative behavior. Then, their rheotactic orientation behavior in a swimming tunnel with a low flow rate and under different magnetic conditions is observed.

Assessing the Influence of Personality on Sensitivity to Magnetic Fields in Zebrafish

Capable of functional recovery after spinal cord injury, adult zebrafish is a premier model system to elucidate innate mechanisms of neural regeneration. Here, we describe swim endurance and swim behavior assays as functional readouts of spinal cord regeneration.

Assessment of Swim Endurance and Swim Behavior in Adult Zebrafish

This protocol utilizes light-sheet imaging to investigate cardiac contractile function in zebrafish larvae and gain insights into cardiac mechanics through cell tracking and interactive analysis.

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Multi-chamber social preference tasks have been adapted from rodents to zebrafish, facilitating the study of social behavior in model organisms. This protocol validates a low-cost 3-chamber open-tank task using webcams and open-source software to measure zebrafish social preference, making it accessible to undergraduate research and teaching.

Low-Cost Alternative for Examining Social Preference in Zebrafish (Danio rerio) Using the 3-Chamber Open-Tank Free-Swim Task

Explore automated 3D-tracking of zebrafish shoals using advanced algorithms and refraction-corrected imaging. This method enables precise analysis of kinematic, spatial, and social behaviors for neuroscience research.

Using an Automated 3D-tracking System to Record Individual and Shoals of Adult Zebrafish

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