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

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.