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Method Article

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

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DOI:

10.3791/70610

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August 7th, 2026

 ,  ,  ,  ,  ,  , 
1Christopher Newport University, 2Virginia Governor's School for Science and Technology, 3Bay Path University

In This Article

Summary

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.

Abstract

Understanding social behavior is a core pursuit in neurobehavioral research, often studied using social preference tasks. These tasks, originally developed for rodents and later adapted for zebrafish, are widely used to examine social dynamics and responses. The 3-chamber open-tank free-swim task (OTFST) consists of a central tank flanked by two stimulus tanks with a camera positioned to record behavior of the subject in the central tank and polarizing film and backlighting to enhance video quality and validity. This protocol uses a low-cost setup with USB webcams and readily available materials. We feature the use of DeepLabCut, an open-source markerless pose estimation tool that enables accurate video tracking of animal movements and provides positional time-series data to be analyzed for various metrics. Pilot sessions validate the system’s ability to capture robust social preference behavior. This budget-friendly approach offers an accessible and scalable platform for labs investigating social interaction in zebrafish, and the versatility and relative simplicity of the 3-Chamber OTFST make it suitable for undergraduate teaching laboratories and research projects.

Introduction

Social preference is an individual’s tendency to engage, seek, or avoid social stimuli and is a fundamental aspect of social behavior that plays a critical role in establishing bonds and hierarchies within groups of animals, including humans1,2,3,4. Understanding social preference is essential to characterizing the neural and behavioral mechanisms that underlie social interaction. Prior research indicates that deficits in social preferences can be associated with neurodevelopmental and psychiatric disorders5,6,7,8. Animal models can provide insights into the biological basis of social behavior and aid in the development of translational tools for diagnosing and treating social impairments. These animal models, ranging from rodents9,10 to aquatic species11,12, provide useful frameworks for discovery.

Rodents were first used for behavioral research in the early 19th century13, due to their physiological similarities to humans14,15. While behaviors like aggression were originally studied, systematic measures of social preference remained minimal. Researchers filled this gap by developing the three-chamber social preference task, introduced by Nadler et al.16. The apparatus consists of three interconnected chambers. Typically, one side chamber contains a novel conspecific while the other chamber remains empty. The subject is allowed to freely explore all chambers, where the time in each chamber is recorded and evaluated as a preference indicator. This apparatus and procedure, along with several variations, gained widespread popularity and have been used to study various aspects of social performance17,18,19 and social deficits in rodent models for autism spectrum disorder5,20,21. However, studies with rodents are more costly and require considerable space. As such, many researchers have instead turned toward using aquatic species for neurobehavioral research.

Aquatic species, most prominently zebrafish (Danio rerio), possess an elaborate behavioral repertoire, including complex social interactions22. Due to their utility as a biomedical research model and the importance of social interaction in their behavioral ecology, zebrafish provide a meaningful framework for social responses that can be used in comparison to humans23. Zebrafish have become increasingly popular for behavioral neuroscience research as they are inexpensive to maintain and require a relatively small holding space, making this species a cost-effective and practical model for large-scale studies of social behavior and preference24. In some studies, an adjacent tank configuration was used for studying social interactions in aquatic species25,26; however, this setup provided somewhat limited throughput for assessing social preference as only one stimulus was presented. This limitation was addressed by the adaptation of the murine three-chamber task for zebrafish in 201827. The modified apparatus for the 3-chamber open-tank free-swim task (OTFST) consisted of a single acrylamide tank divided with fixed acrylamide barriers to provide a neutral start zone and two side chambers, each side chamber containing either a familiar or novel conspecific. To measure social preference, the cumulative time spent near each side chamber was recorded, demonstrating that ethanol exposure and sam2 knockout reduced novelty preference in zebrafish27. While the rodent 3-chamber social interaction test allows complex multi-modal interactions between the subject and the test conspecific(s) through the visual, chemical, tactile, and acoustic modalities, the use of separate chambers for the aquatic 3-chamber OTFST reduces the interaction between the subject and stimulus animals to just the visual modality. While this single-modality interaction may be lower in ecological validity, it does provide more experimental control and enhances simplicity for projects conducted by undergraduate researchers. In the literature, various researchers have since explored adaptations of the 3-chamber OTFST to investigate genetic and environmental factors that influence social behavior in zebrafish28,29,30,31,32.

The present adaptation of the 3-chamber OTFST is designed to serve as a versatile and undergraduate-friendly model apparatus. The FishLab, a fully undergraduate laboratory, created and employed this setup and has demonstrated its effectiveness as a high-throughput tool for investigating social preference in zebrafish. Studies utilizing this setup have explored multiple facets of zebrafish social preference in response to manipulations of various independent variables. The first study demonstrated that zebrafish can detect live versus simulated social stimuli33, while the second investigated preference for established shoals compared to newly-formed shoals34. The most recent study examined the ability of zebrafish to differentiate between chemically-alarmed versus unalarmed shoals35. Together, these studies address different aspects of social behavior, including realism, familiarity, and threat detection, while demonstrating commonalities in shoaling preferences and assessing responses to predation risk. However, the setup used for these previous studies employed digital IP cameras mounted on professional camera rigging, a dedicated aquatic rack housing system, and proprietary software for both video acquisition and analyses, with a total cost that might be prohibitive for investigators/instructors with small operational budgets. With the advent of open-source software and high-quality USB-based webcams, a low-cost version is now feasible for investigators wishing to employ this technique with undergraduate researchers, with a total price (approximately US$2,000; excluding the computer) that is substantially lower than the expense of the high-cost system (approximately US$20,000+, excluding the computer).

The following protocol outlines the development and application of a low-cost version of the 3-chamber OTFST for zebrafish. Its affordability, simplicity, and versatility make it a useful tool for small laboratories or educational settings with limited funding. When combined with low-cost housing, such as commonly available 10-gallon aquaria, the current setup costs significantly less to deploy and maintain. Additionally, the use of open-access software such as open broadcasting system (OBS) for recording experimental sessions and various tracking applications (e.g., DeepLabCut, SLEAP, etc.) provides a low-cost, accessible framework for capturing video and generating time-series data of social interactions of zebrafish.

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Protocol

All experimental procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at Christopher Newport University (Protocol #2025-23).

1. Animals

  1. Obtain adult short-fin wild-type zebrafish23,36,37, although other strains can be used. Alternatively, obtain embryos and rear them in-house.
  2. House the fish in free-standing 10-gallon (38 L) tanks at a density of approximately two fish per gallon (3.8 L) of water. Equip tanks with active bio-filters, air stones, and submersible heaters. Alternatively, house fish on an aquatic rack system.
  3. Maintain water-quality parameters within the following ranges: pH 7.0–8.0, conductivity 400–700 µS, temperature 25–27 °C, <40 ppm nitrates, <0.2 ppm nitrites, and 0.01–0.1 ppm ammonia.
  4. Keep animals under a 12-h light:12-h dark photoperiod and feed once daily with commercial flakes.
  5. Select animals randomly from these tanks for experimental assignment.
  6. Upon completion of the experiment, repurpose the animals for other research/teaching purposes, or euthanize them by immersion in 250-500 mg/mL buffered tricaine for 30 min, followed by a physical method (e.g., decapitation) to ensure death. Use the euthanized animals for subsequent bioassays and/or neurochemical analyses.

2. 3-chamber open-tank free-swim task apparatus and software

  1. Construction of test chambers
    1. Carefully remove the plastic trim from the bottom of each 5.5-gallon tank. Use a rotary tool with a cutoff wheel to slit the plastic at the corners to facilitate removal. Once the plastic trim is slit, separate the plastic from the silicone adhesive with a common putty knife.
    2. To diffuse the backlighting, cover the back of all three tanks, the left-hand end of one tank, and the right-hand end of the other tank with the white adhesive shelf liner. Leave the ends uncovered on the tank that will be used as the center chamber.
    3. Place a black horizontal line to indicate the fill level at 4.5 cm below the top rim on the front of each chamber with an enamel marker or with black graphic layout tape.
    4. Cut one sheet of polarizing filter to fit the uncovered end side of each flanking tank. Orient the filter sheets 90° to each other. Use clear tape along all four edges to prevent water from entering the space between the aquarium glass and filter material.
      NOTE: The offset orientation of the polarizing filters will prevent stimulus animals in one flanking stimulus chamber from seeing the animals (or other stimuli) in the other flanking stimulus chamber while still allowing them to see the subject. The subject in the center tank can see the visual stimuli in both flanking chambers.
    5. Construct the polyvinyl chloride (PVC) base to align the exact placement of the tanks on the setup bench. Cut the bottom of the base from a 1/4 inch (0.635 cm) thick white PVC sheet at 21 cm x 122.5 cm and cut the end pieces from 1 x 2 PVC trim board at 21 cm long.
    6. Test fit the ends on the base with all three tanks set in place. Ensure easy insertion and removal of the divider panels (below) and confirm the flanking tanks do not abut tightly against the sides of the center tank. Once test fitting is complete, join the end pieces to the PVC base with PVC cement.
    7. To make the clear lids for each chamber, cut the polycarbonate sheet into three pieces sized to the outside dimensions of the top rim of the 5.5-gallon tank (40 cm × 21 cm). Cut a rabbet channel (6 mm × 4 mm) around the edge of each lid to provide secure positioning of the lid on top of the chamber.
    8. Cover the chambers with the lids whenever animals are held in the chambers. Place a submersible heater and an airstone connected to an aquarium air pump while animals are held in their respective chambers to provide temperature control and aeration. Use a three-way air valve to balance air flow across all three airstones.
    9. Make simple opaque dividers for placement between the center chamber and each flanking chamber to control when test subjects are provided visual exposure to social stimuli by laminating two sheets of A4 light-colored cardstock.
    10. Prepare the dividers for hanging and remote removal by punching two holes at 85 mm and 125 mm along one 210 mm edge of each laminated sheet.
    11. Cut two 75 cm lengths of high-strength twine and thread the twine through each hole and tie the leftover ends together, connecting the two sheets in a center yoke.
    12. Cut a 6 m length of twine. Tie one end of the 6 m twine to the yoke to create a central tethered line, and thread the free end of the twine through a swivel eye pulley.
    13. Mount the swivel eye pulley on the ceiling above the centermost tank and ensure the end of the twine is hanging on the outside of the experimental setup (Figure 1A).
  2. Construction of lighting setup
    1. Make a frame from 1/4 inch (0.635 cm) thick white PVC sheet for the panel light. Cut two end pieces (10 cm × 31.5 cm) and a top and a bottom (10 cm × 122 cm). Test fit the frame pieces together in such a way that the panel fits inside the framing. Once the fit is tested, join the frame pieces using PVC cement.
    2. Have a qualified electrician add a 3-conductor cord to the panel light.
    3. Have a qualified electrician wire the slider dimmer to control the output receptacle in the waterproof enclosure. Plug the panel light into this receptacle.
    4. Adjust the slider dimmer to adjust the brightness of the backlighting to 350 lux. Fill the chambers with water and move the slider dimmer to adjust the brightness at the front of the chambers to 15 lux.
      NOTE: Researchers should verify the lighting before, during, and after a study to ensure the brightness settings have been maintained.
  3. Construction of curtain and camera frame
    1. Set up the testing bench with the panel light, the tank base panel, and the three chambers. Position and adjust the two tarp poles vertically so they each fit snugly from the floor to the ceiling, at least 75 cm from the front of the tanks.
    2. Make a top pocket for each curtain by folding one end of the sheet over 60 cm and sewing a straight stitch 6 cm from the top fold to create a pocket for the frame piece to slide into.
    3. Measure the dimensions for the cross-joining and stabilizing sections for the frame setup according to the width and depth of the area to be used for the setup.
      NOTE: For the current setup in The FishLab, the vertical tarp poles were placed 146.5 cm from the back wall in the corner of the lab. The inside tarp pole is positioned 6.5 cm from the side wall, while the outside tarp pole is positioned 142.5 cm from the side wall.
    4. Cut three sections of electrical metal tubing (EMT) conduit at 143 cm to be used as stabilizing sections for the sides of the frame. Pass one section through the top pocket of the movable curtain and position horizontally 190 cm above the floor.
    5. Attach the section to the outside tarp pole with a T-clamp on one end and the custom 3D-printed socket at the other end. Attach the 3D-printed socket to the back wall using screws. Attach the other two sections to the inside tarp pole.
    6. Pass the top section through the top pocket of one of the fixed curtains and position 190 cm above the floor. Attach to the inside tarp pole with a T-clamp at one end and the other 3D-printed socket at the other end.
    7. Attach the 3D-printed socket to the side wall using screws. Place the final stabilizing section 120 cm above the floor and attach it similarly to the other two stabilizing sections.
    8. Cut another section of EMT conduit at 127 cm to be used as a stabilizing section for the front of the frame. Pass this section through the top pocket of the second fixed curtain and mount it 193 cm above the floor by attaching each end to the tarp poles using a T-clamp at each end.
    9. Cut the remaining piece of EMT conduit at 145 cm to be used as the camera-mounting bar. Place this section horizontally at 88 cm above the floor to align the cameras with the center of the tanks. Fit one end into a 3D-printed socket, which is screwed to the wall, and use the custom 3D-printed cross-clamps to hold the camera bar tightly against the tarp poles (Figure 1B).
  4. Construction of the camera setup
    1. Clamp one camera mount on the camera bar so that the horizontal and vertical centers of the camera align with the corresponding horizontal and vertical centers of the center chamber.
      NOTE: Additional cameras can be mounted in front of each flanking chamber if researchers wish to obtain digital video of the responses of live animals used as visual stimuli. The simplest setup involves placing a single camera to capture video of the center experimental chamber. For live social stimuli in the stimulus chambers, researchers may also wish to collect video of the chambers.
    2. Mount each camera on the threaded post of each camera mount. Adjust each camera so the image is centered on the observation zone of its respective chamber.
    3. Secure the USB cables for each camera on the camera bar using straps or ties to prevent dangling of the cords, which can cause cameras to misalign.
    4. Cut a section of wire rack shelving at 140 cm long and attach it vertically to the outside of the front of the frame to prevent the cameras from being bumped out of alignment. Mount the “camera cage” using T-clamps attached to the tarp poles.
      NOTE: A second camera cage can be mounted to the inside of the frame to further prevent the cameras from being bumped out of alignment; a small section of wire grid may need to be removed if the front of the cameras is “caged” to prevent occlusion of the tank images.
  5. Load and configure acquisition software - Open broadcaster studio (OBS)
    1. Download OBS and install it on a Windows-operated local computer. Launch OBS Studio and run the Auto-Configuration Wizard to optimize settings for the specific recording needs, hardware resources, and network conditions.
      NOTE: This tool is available under Tools > Auto-Configuration Wizard.
    2. Verify the basic settings before configuring the camera. Set the Base (Canvas) resolution to match the display resolution, and set the Output (Scaled) resolution to the recording resolution (e.g., 720 pixels).
    3. Create a scene by right-clicking in the Scenes box (or use the cross at the bottom). Assign a name to the scene. Use the scene to define the stream layout.
    4. Add a new source by right-clicking the Sources box (or use the plus symbol at the bottom). Select Video Capture Device to add a webcam. Assign each camera a distinct name (e.g., “camera-left”, “camera-center”, “camera-right”). Repeat this process for additional cameras or sources as needed.
    5. Position and resize sources in the preview window. Ensure the camera is visible by confirming it is listed above other sources in the Sources list. To center the camera view on the canvas, select the source in the Sources list (a red bounding box will appear), and press CTRL+D (Center to Screen).
    6. Fill the entire screen with a source by pressing CTRL+F (fit to Screen) or CTRL+S (stretch to Screen), if desired. Crop the source by holding the Alt key while dragging the bounding box. Verify cropping by checking that the edges turn green.
    7. Adjust video properties such as brightness and contrast by double-clicking the selected camera in the Sources list and selecting Configure Video. Modify settings using the properties menu.

3. Behavioral testing

  1. Acclimation
    1. Fill all 3 chambers with water from housing tanks. Place opaque barriers between each chamber and leave them in place during the entire acclimation period. Transfer one subject to the center experimental chamber and allow it to acclimate for a minimum of 1 h before testing.
      NOTE: The acclimation period for the representative results was set at 3 h. The acclimation period is necessary for animals to recover from any momentary transfer stress and adjust to the chamber environment prior to behavioral testing.
    2. At the beginning of the acclimation period, place the social stimuli in the flanking stimulus chambers. Center the artificial social stimuli in the left stimulus chamber and transfer a small shoal of 4 fish (2 males and 2 females of similar size, age, and familiarity) to the right stimulus chamber.
      NOTE: The positions of the artificial social stimuli and the live social stimuli were fixed and not counterbalanced in the present protocol, as previous studies33,34 detected no effect of stimulus position using the OTFST. Counterbalancing in this instance would require unnecessarily doubling the number of animals used for research, which is contrary to the “3Rs” of animal research. This protocol provides flexibility in the nature of the stimuli to be tested, e.g., static models, sympatric or novel species, etc.
    3. During acclimation, supply all chambers containing live animals with a submersible heater (set at 26 °C), a bubbler stone connected to the aquarium air pump, and a tank lid to maintain stable holding conditions. Remove the heaters and bubbler stones before initiating an experimental session.
  2. Experimental testing
    1. Verify the activity status of the social stimuli being tested in each of the flanking tanks and test subject in the center tank. Use the single camera on the center tank to record video for each test subject.
      NOTE: The representative results in this protocol are a replication of a single condition from Velkey et al.33 to test zebrafish preference between low-fidelity (mobile models) and high-fidelity (live shoal) social stimuli.
    2. Open OBS Studio to begin recording a session. Resize the center camera view to fill the screen by clicking and dragging the corners of the video. Select Start Recording under Controls at the bottom right of the screen.
    3. Immediately after the recording is initiated, remove the opaque barriers via the pulley-string system. Continue recording for 10 min and allow the subject to have simultaneous exposure to both social stimuli while it moves freely within the center experimental chamber.
    4. End the session by selecting Stop Recording under Controls. Access recorded sessions by selecting File in the top-left menu and choosing Show Recordings from the drop-down menu.
      NOTE: The resulting videos provide unstructured data suitable for a variety of analyses.
    5. Transfer live fish to holding tanks, remove the water from all three chambers, rinse thoroughly with water (avoid getting water between the polarized films), spray each chamber with 5% citric acid to sanitize, allow chambers to air dry overnight, and rinse at the beginning of the subsequent experimental day before being placed back on the testing bench.

4. Obtaining structured data for analyses

  1. Select a motion-analysis software application (e.g., DeepLabCut; see https://www.mackenziemathislab.org/deeplabcut). Install DeepLabCut on the computer, following the user guide at https://deeplabcut.github.io/DeepLabCut/docs/installation.html. Refer to the selected software’s information and user guide for further directions if not using DeepLabCut.
  2. Load four to five videos into DeepLabCut for model training, and extract 20–30 frames per video. Define at least three primary tracking points (e.g., head, trunk, tail) and train the network to track these points across frames.
    NOTE: For more information, consult the DeepLabCut user guide at https://deeplabcut.github.io/DeepLabCut/docs/standardDeepLabCut_UserGuide.html. The representative results use 22 frames labeled from five videos with 3 tracking points.
  3. Use a ResNet-101-based neural network to train the model. Determine the number of iterations required to complete training (between 100,000 and 200,000 is suggested). Determine a p-cutoff value between 0.5 and 1.
    NOTE: For the representative results, 150,000 iterations were used to train the model with a p-cutoff of 0.7. If the trained network falls below the 0.7 p-cutoff, increasing the video contrast or changing the labeling convention will address this problem.
  4. Deploy the fully-trained DeepLabCut model on the remaining experimental videos. Extract positional data for each trial. Follow the user guide at https://deeplabcut.github.io/DeepLabCut/docs/beginner-guides/Training-Evaluation.html.
    NOTE: For the representative results, we validated with 2 shuffles and determined the test error was 2 pixels:train 4 pixels (with an image size of 1600 x 820 pixels). The network was then used to analyze the experimental videos obtained from the present experimental setup.
  5. Use coding software, such as Python, to define and analyze movement across the three vertical arena zones of equal width to quantify preference. Measure the time in zone by at least two body points within the identified zone to reduce tracking noise. Use the formula: Total time in zone = (Total frames in zone)/(Total frame count).
  6. If a measure of thigmotaxis is desired, quantify zones by a full body length of a zebrafish from the tank walls, which will provide a measure of the passive tendency of subjects to remain near a vertical boundary. This measure is most appropriate for relatively larger experimental tanks.
  7. Use the same coding software to define and measure time spent motionless (e.g., freezing). Define the threshold of movement at 2–5 pixels per frame with a minimum consecutive frame count of 15 frames.
    NOTE: For the representative results, freezing was identified with a threshold of 3 pixels per frame for 15 frames.
  8. Compile and export measures for statistical analysis and visualizations.
    NOTE: For data analysis, refer to https://github.com/farhanaugustine/DeepLabCut-Analysis-Jupyter-Scripts. We

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Results

In order to validate the low-cost methods outlined above, one condition from the experiments conducted in Velkey et. al.33 was replicated using the present low-cost setup. Subjects (N = 23, 11 males and 12 females) were presented with the artificial moving mobile (low-fidelity) social stimuli on the left and the live shoal (high-fidelity) social stimuli on the right (Figure 2); as previous studies33,34 have no...

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Discussion

The current protocol demonstrates that the present low-cost adaptation of the 3-Chamber OTFST can be used to reliably characterize social preference in adult zebrafish as measured by time spent in zone. Using inexpensive construction materials, a USB-connected webcam, and open-source acquisition and analysis software applications, the present system successfully replicated the finding that zebrafish prefer a shoal of live conspecifics over lower-fidelity moving models. These results validate the application of a low-cost...

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Disclosures

The authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

Acknowledgements

The authors wish to thank Rommel Pagkalinawan, Robby Buck, Elizabeth Hiltz, Haley Dewitt, Maia Tankersley, Alexis Thai-Nguyen, Grady Fleming, Jacob Young, and Jack Medlin of TheFishLab team at Christopher Newport University for their contributions to animal husbandry and data collection for the validation experiment. The authors also wish to thank Samantha Commons for her assistance during her Community Captains Summer 2025 Program, including conducting the inventory of all components of the existing high-cost setup as well as the low-cost setup. The authors would also like to thank the Office of Student Research and Creative Activity at Christopher Newport University for funding provided to K.K. under the undergraduate Summer Scholars Program. Figure 2 was created in BioRender by Kinslow, K. (2025). https://BioRender.com/wbf0wph.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
10-Gallon AquariumAqueon, Inc. Franklin, WI, USA#100110010purchased at local pet supply (1 to 4)
12 Pieces 1-inch T-Pipe ClampsJoyHua Trade, ChinaASIN #B0DF19DJF1purchased from Amazon.com (1)
2-gang electrical boxSigma Engineered Solutions, Raleigh, NC, USA# 14350purchased at local hardware store (1)
3/4" Silver-Metallic electrical metallic(EMT) Conduit - 10-footWheatland Tube, Inc., Wheatland, PA, USA #0550110000purchased at local hardware store (3)
3-conductor cordUtilitech, Inc., Charlotte, NC, USAUT010710purchased at local hardware store (1)
3-prong receptacleEaton, Inc., Beachwood, OH, USA#270W-SP-LWpurchased at local hardware store (1)
3-way Air ValvePenn-Plax, Inc. 35 Marcus Blvd. Hauppauge, NY, USAModel VN3purchased at local pet supply (1)
A4 Cardstock (white), laminatedvariesvariespurchased from campus bookstore (2)
Active bio-filter Penguin 100Spectrum Brands Pet, LLC, Blacksburg, VA, USA#PF00100Bpurchased at local pet supply (1 to 4)
Air stonesPenn-Plax, Inc. 35 Marcus Blvd. Hauppauge, NY, USAModel AS6Qpurchased at local pet supply (3)
Aquarium Air TubingPenn-Plax, Inc. 35 Marcus Blvd. Hauppauge, NY, USAModel STD8purchased at local pet supply (1)
Black Fabric Twin Sheets (167.6 cm X 243.8 cm)Royale Linens, Inc., Melville, NY, USA#00011342258875purchased from Amazon.com (3)
Brio 4k WebcamLogitech International S.A., Lausanne, Switzerland960-001419purchased from Amazon.com (1 to 3)
Computer (sufficient for running Open Broadcaster Studio & 1-3 Webcams)Dell, Inc., Round Rock, TX, USAOptiPlex Micro 7010purchased from dell.com (1)
Dimmable light-emitting diode (LED) flat-panel lightLithonia Lighting-Acuity Brands, Inc. Conyers, GA, USACTCL-153P-WHpurchased from Amazon.com (1)
E-Z Up Dust Containment Pole (2 Pack)Trimaco, Inc., Cary, NC, USA# 00047034547330purchased from Amazon.com (1)
Feet Hardware TwinevariesVariespurchased at local hardware store (1)
Linear polarizing filtersB&H Foto & Electronics, Inc., New York, NY USARosco #7300purchased online (7)
Open Broadcaster SoftwareOBS Projectv32.0.0download from https://obsproject.com (1)
Polycarbonate Sheet (0.38"/9.6mm thick)Spectraglass, ltd., Perth, ScotlandUPC # 608037376817purchased from Amazon.com (1)
polyvinyl chloride (PVC) Cement, 4 oz canisterThe AZEK Co, Chicago, IL, USA#ARAD0004OZpurchased at local hardware store (1)
Rimless 5.5-Gallon AquariumAqueon, Inc. Franklin, WI, USA#100541424purchased at local pet supply (3)
Self-adhesive white plastic film CON-TACT Brand, La Miranda, CA, USA#16F-C9A952-06purchased at local hardware store (1)
Slider Dimmer switchLutron, Inc., Cooperburg, PA, USA#LECL-150H-WHpurchased at local hardware store (1)
small pulleyvariesVariespurchased at local hardware store (1)
Submersible 50W heaters (preset to 26C)Aqueon, Inc. Franklin, WI, USA#100106251purchased at local pet supply (3 to 7)
Super Clamp Camera MountJINSUI, ChinaX0040GEYLRpurchased from Amazon.com (1 to 3)
Tetra Whisper 10 Aquarium Air PumpSpectrum Pet, Home & Garden, Inc., Middleton, WI, USAUPC #046798778516purchased at local pet supply (1)
Weatherproof 2-gang electrical enclosureHubbell, Inc., Shelton, CT, USA#MM2420Cpurchased at local hardware store (1)
White PVC Sheet 1/4" x 24" 48"Royal Building Products, Bristol, TN, USA#3101purchased at local hardware store (2)
White PVC Trim Board, nominal 1 x 2 (0.75" x 1/5" x 96")Royal Building Products, Bristol, TN, USA#7832-8purchased at local hardware store (1)

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Zebrafish Social BehaviorThree Chamber TaskDeepLabCut TrackingMarkerless Pose EstimationBehavioral Video AnalysisLow Cost SetupSocial InteractionNeurobehavioral Research
Low-Cost Alternative for Examining Social Preference in Zebrafish (Danio rerio) Using the 3-Chamber Open-Tank Free-Swim Task
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