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.
Due to their renowned regenerative capacity, adult zebrafish are a premier vertebrate model to interrogate mechanisms of innate spinal cord regeneration. Following complete transection of their spinal cord, zebrafish extend glial and axonal bridges across severed tissue, regenerate neurons proximal to the lesion, and regain their swim capacities within 8 weeks of injury. Recovery of swim function is thus a central readout for functional spinal cord repair. Here, we describe a set of behavioral assays to quantify zebrafish motor capacity inside an enclosed swim tunnel. The goal of these methods is to provide quantifiable measurements of swim endurance and swim behavior in adult zebrafish. For swim endurance, zebrafish are subjected to a constantly increasing water current velocity until exhaustion, and time at exhaustion is reported. For swim behavior assessment, zebrafish are subjected to low current velocities and swim videos are captured with a dorsal view of the fish. Percent activity, burst frequency, and time spent against the water current provide quantifiable readouts of swim behavior. We quantified swim endurance and swim behavior in wild-type zebrafish before injury and after spinal cord transection. We found that zebrafish lose swim function after spinal cord transection and gradually regain that capacity between 2 and 6 weeks post-injury. The methods described in this study could be applied to neurobehavioral, musculoskeletal, skeletal muscle regeneration, and neural regeneration studies in adult zebrafish.
Adult zebrafish are eminently used to investigate mechanisms of neuromuscular and musculoskeletal development and disease modeling1,2,3. Zebrafish are capable of efficient, spontaneous repair of multiple tissues, including the brain, spinal cord, and skeletal muscle4,5,6,7. The remarkable capacity to regenerate neuromuscular tissues and model diseases is attracting a growing scientific community into adult zebrafish research1,2,3. However, while assays of locomotion and swim behavior are available and standardized for larval zebrafish, there is a growing need to develop analogous protocols in adult fish8,9,10,11. The goal of this study is to describe protocols to quantify swim endurance and swim behavior in adult zebrafish. We present these protocols in the context of spinal cord regeneration research. However, the behavioral protocols described here are equally applicable to studies of neural and muscle regeneration, neuromuscular and musculoskeletal development, as well as neuromuscular and musculoskeletal disease modeling.
Zebrafish reverse paralysis within 8 weeks of complete spinal cord transection. Unlike poorly regenerative mammals, zebrafish display pro-regenerative immune, neuronal, and glial injury responses that are required for functional spinal cord repair12,13,14. An ultimate readout of functional spinal cord repair is the ability of the lesioned tissue to regain its function after injury. A suite of standardized methods to assess functional regeneration in rodents include locomotor, motor, sensory, and sensorimotor tests15,16,17. Widely used tests in mouse spinal cord injury include the locomotor Basso Mouse Scale (BMS), forelimb motor tests, tactile sensory tests, and grid walking sensorimotor tests15,17. In contrast with mammalian or larval zebrafish systems, behavioral tests in adult zebrafish are less developed, yet much needed to accommodate the growing needs of the tissue regeneration and disease modeling communities.
Complete spinal cord transections result in complete paralysis caudal to the injury site. Shortly after the injury, paralyzed animals are less active and avoid swimming as much as possible. To compensate for lost swim capacity, paralyzed animals display short, frequent bursts by overusing their pectoral fins, which lie rostral to the lesion. This compensatory swim strategy results in rapid exhaustion and lower swim capacity. As the zebrafish spinal cord regenerates, animals regain a smooth oscillatory swim function caudal to the lesion, allowing for increased swim endurance and improved swim behavior parameters. Here, we describe methods to quantify zebrafish swim endurance at increasing water current velocities and swim behavior at low current velocities.
Adult zebrafish of the Ekkwill and AB strains were maintained at the Washington University Zebrafish Core Facility. All animal experiments were performed in compliance with IACUC institutional animal protocols.
NOTE: An example of the experimental setup is shown in Figure 1A. The calibration lid (customized), swim endurance lid (customized), and swim behavior lid (standard, enclosed tunnel lid) are shown in Figure 1B. The experimental workflow is presented in Figure 2.
1. Swim tunnel preparation and calibration
2. Assessment of swim endurance
NOTE: Experimental groups are divided into groups of 10 or fewer animals for swim endurance.
3. Capturing videos for swim behavior assay
NOTE: Only up to five animals can be tracked at a time. If experimental groups are larger than five animals, multiple videos can be taken for each group, where the first video tracks five or fewer animals and the second video tracks the other five or fewer animals. For longitudinal studies that aim to track individual animals over time, fish can be individually housed and tracked across multiple time points. All scripts for tracking and analyzing are available via GitHub (see Table of Materials).
4. Analyzing movies for swim behavior assessment
NOTE: Movie recording and analysis can be completed on separate days.
We set up the swim tunnel as described in section 1 of this protocol (Figure 1). We assessed the swim endurance (section 2 of this protocol) as well as swim behavior (sections 3 and 4 of this protocol) of adult zebrafish at baseline and after spinal cord injury (Figure 2).
For establishing baseline motor function, we examined the swim endurance of wild-type zebrafish under increasing water current velocities (Figure 3A). In this assay, wild-type zebrafish swam for 41 min before getting exhausted. Fish were then subjected to complete spinal cord transections as previously described and swim endurance assays were performed6. After anesthetizing zebrafish using MS-222, a small incision is made with fine scissors to transect the spinal cord 4 mm caudal to the brainstem region. A complete transection was confirmed visually. To confirm the loss of swim capacity after spinal cord surgery, injured animals were assessed at 2 or 3 days post-injury (dpi). At this time point, zebrafish are completely paralyzed caudal to the lesion site. Swim endurance was assessed at 2, 4, and 6 weeks post-injury (wpi). At 2 wpi, lesioned fish lost 60% of their swim endurance capacity (Figure 3A). Regenerating fish gradually regained swim endurance at 4 and 6 wpi. These results indicated that wild-type zebrafish are capable of regaining swim endurance capacity after spinal cord injury.
To examine zebrafish swim behavior during spinal cord regeneration, we tracked the swim behavior of wild-type animals at 0 cm/s water current velocity or under constant, low current velocities of 10 and 20 cm/s (Figure 3B). Average tracks of fish position in the swim tunnel chamber were used for visual assessment of swim behavior at low current velocities (Figure 3B). In this assay, uninjured controls swam steadily in the front part of the swim tunnel chamber (closer to the source of water current), which corresponds to an elevated Y position (Figure 3B). In contrast, at 2 wpi, injured fish were not able to maintain steady swim capacity against the current. Consequently, their swim tracks are more irregular with an overall decrease in Y position (Figure 3B). Y position increased at 4 and 6 wpi, indicating that regenerating animals gradually regained their ability to swim in the front of the swim tunnel chamber. To quantify swim behavior parameters, we calculated the percent activity, position in the swim tunnel (Y position), and time swam against the current (Figure 3C–E). Relative to uninjured controls, lesioned animals at 2 wpi were markedly less active (Figure 3C), stalled in the rear quadrant of the swim tunnel (Figure 3D), and lost their ability to swim against low current velocities (Figure 3E). Consistent with their innate ability to achieve functional recovery, lesioned animals gradually normalized swim behavior parameters at 4 and 6 wpi (Figure 3C–E). The swim endurance and swim behavior parameters together offered quantifiable readouts of swim function and functional spinal cord repair in zebrafish.
Figure 1: Swim tunnel set up and customized lids. (A) Representative images of the swim tunnel set up including zoomed top and side views of the swim tunnel chamber. (B) Images of the swim tunnel lids used for the various applications described in this protocol. A standard, fully enclosed swim tunnel lid is used for swim behavior assays (section 3 of this protocol). A modified swim tunnel lid that accommodates a handheld digital flow meter is used for calibration (section 1 of this protocol). A modified swim endurance lid, containing a removable lid at the posterior end of the swim tunnel chamber, allows for the removal of exhausted fish during swim endurance testing (section 2 of this protocol). Please click here to view a larger version of this figure.
Figure 2: Experimental pipeline to assay for swim endurance and swim behavior in adult zebrafish. For swim endurance, fish swam against an increasing water current until exhaustion. For swim behavior, swim parameters are assessed in the absence of and at low current velocities. Please click here to view a larger version of this figure.
Figure 3: Functional recovery in wild-type zebrafish after spinal cord injury. (A) Motor function determined by swim endurance assays for wild-type zebrafish at baseline and 2, 4, and 6 wpi. Dots denote individual animals from two independent experiments. (B) Swim behavior assays tracked wild-type zebrafish under low water current velocities. The average Y position is shown at each time point throughout tracking (0 cm/s for 5 min, 10 cm/s for 5 min, and 20 cm/s for 5 min). (C–E) Percent activity (C), average Y position in the tunnel (D), and time swam against the flow (E) were quantified at 20 cm/s. For all quantifications, two independent experiments are shown. n = 30 in the pre-injury condition; n = 23 at 2 wpi, n = 20 at 4 wpi, n = 18 at 6 wpi. One-way ANOVA was used for statistical analyses. Error bars represent the Standard Error of the Mean (SEM). *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001. Please click here to view a larger version of this figure.
Supplementary File 1: Please click here to download this File.
Adult zebrafish are a popular vertebrate system for modeling human diseases and studying mechanisms of tissue regeneration. CRISPR/Cas9 genome editing has revolutionized reverse genetic studies for modeling disease in zebrafish; however, large-scale genetics in adult zebrafish has been hindered by biological and technical challenges, including the unavailability of adult zebrafish tissues to high-throughput phenotyping. Given the complex anatomy of adult zebrafish, prolonged histological processing is required to obtain and analyze tissue architecture. The swim endurance and swim behavior assays described in this study can be used to pre-screen for neural, muscular, or skeletal phenotypes at a medium throughput before histology. Moreover, as studies of tissue regeneration aim to improve functional tissue repair, the protocols described in this study will be widely applicable, if not essential, for studies of neural, muscular, and skeletal regeneration research.
Functional locomotion assays have been integral to our understanding of neural development and regeneration. Standard locomotor assays are widely available in vertebrate species, including rats, mice, and larval zebrafish. Mouse and rat model systems have behavioral and functional locomotion assays such as the BMS16 and BBB17,18, respectively. Similarly, a host of protocols have been described to measure locomotion, startle response, and behavior in larval zebrafish. These protocols are efficient to reveal behavioral differences among experimental groups in light and dark environments, predator evasion, and activity19,20,21. Here, we describe quantifiable, reproducible methods to measure functional recovery after spinal cord injury in adult zebrafish.
Note that this study presents several limitations. First, behavioral studies are highly dependent on genetic and environmental factors. To control for genetic variability, we used siblings to control for age, sex, and the genetic background across experimental groups22,23. To control for environmental factors, we made sure experiments are performed at the same time of the day, under controlled temperature and lighting conditions20. Second, while the swim behavior assay is less sensitive to biased analysis, the researcher running the swim endurance assay determines when a fish reaches exhaustion and proceeds to remove exhausted fish from the swim tunnel chamber without interrupting the rest of the fish cohort. Thus, our swim endurance experiments were performed by a single researcher who is blinded to experimental conditions. It is particularly important to avoid researcher-to-researcher variability for longitudinal studies of experimental groups over time. Finally, collisions between fish can complicate the tracking analysis in swim behavior assays. We thus recommend performing swim behavior assays for groups of five or fewer fish to minimize the chances of collisions between fish.
In consideration of critical steps, we note that injured fish can be fragile especially in the early days after injury. We thus recommend handling fish with extreme care. For swim endurance assays, collecting fish with the PVC tube as quickly as possible, either head or tail first, reduces the chance of secondary injuries during the collection process. For swim behavior assays, the flush pump can occasionally create waves, distorting the movies and causing analysis errors. In this case, the flush pump can be briefly turned off. However, we do not recommend turning off the flush pump for extended times to ensure water is constantly circulating between the swim tunnel chamber and the buffer tank. Monitoring movie recording allows for immediate termination and restarting of the movie if a frame has been dropped or a fish strays from the recording area. In additional considerations, while performing the tracking analysis, if the R code gives an error during file processing, the most likely problem is in the naming of the files. The program is made to function under a very specific naming strategy: Timepoint_Group_Subgroup_Stock number_Anything else (for example, 0_A_1_00001_WildtypeGroupA.avi). This naming allows for multiple time points, groups, and subgroups to be plotted and aligned together. Finally, while checkpoints have been built into the analysis script to ensure proper tracking, it is important to carefully check the analysis output. The program will automatically ask whether the fish number is correct, and prompt movies for re-analysis if the fish number is incorrect. Straight-line artifacts may appear in the Average Y position plot, indicating that an extra object has been recognized as a fish. In this instance, the best course of action is to carefully watch the movie to exclude extra artifacts that tend to appear at a higher flow speed.
The authors have nothing to disclose.
We thank the Washington University Zebrafish Shared Resource for animal care. This research was supported by the NIH (R01 NS113915 to M.H.M.).
AutoSwim software | Loligo Systems | MI10000 | Optional – for Automatic control of current velocity |
Customized lid | Loligo Systems | MI10001 | This customized lid is used for swim endurance |
DAQ-BT | Loligo Systems | SW10600 | Optional – for Automatic control of current velocity |
Eheim pump | Loligo Systems | PU10160 | 20 L/min. This pump is placed in theflow-through tank. |
Fiji | Fiji | Freely available through Image J (Fiji) | Specific script available at https://github.com/MokalledLab/SwimBehavior |
Flowtherm | Loligo Systems | AC10000 | Handheld digital flow meter – for calibration |
High Speed Camera | Loligo Systems | VE10380 | USB 3.0 color video camera (4MP) |
IR light panel | Loligo Systems | VE10775 | 450 x 210 mm, placed under the swim tunnel chamber |
Monofocal lens | Loligo Systems | VE10388 | 25mm manual lens |
PVC Tubing | VWR | 60985-534 | 5/16 x 7/16" Wall thickness: 1/16" |
R Studio | R Studio | Freely available. Version 3.6 with extra packages. | Specific script available at https://github.com/MokalledLab/SwimBehavior |
Swim tunnel respirometer | Loligo Systems | SW10060 | 5L (120V/60Hz). The system includes the swim chamber, motor, manual control of water current velocity, 1 pump placed inside the chamber, standard swim tunnel lid for swim behavior, and modified swim tunnel lid for calibration |
uEye Cockpit | IDS | Freely available software to control camera parameters | Alternative cameras and accompanying softwares could be used |
Vane wheel flow probe | Loligo Systems | AC10002 | Digital flow probe – for calibration |