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.
Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many myopathies affecting skeletal muscle function can be quickly and easily assessed in zebrafish over the first few days of embryogenesis. By 24 hr post-fertilization (hpf), wildtype zebrafish spontaneously contract their tail muscles and by 48 hpf, zebrafish exhibit controlled swimming behaviors. Reduction in the frequency of, or other alterations in, these movements may indicate a skeletal muscle dysfunction. To analyze swimming behavior and assess muscle performance in early zebrafish development, we utilize both touch-evoked escape response and locomotion assays.
Touch-evoked escape response assays can be used to assess muscle performance during short burst movements resulting from contraction of fast-twitch muscle fibers. In response to an external stimulus, which in this case is a tap on the head, wildtype zebrafish at 2 days post-fertilization (dpf) typically exhibit a powerful burst swim, accompanied by sharp turns. Our method quantifies skeletal muscle function by measuring the maximum acceleration during a burst swimming motion, the acceleration being directly proportional to the force produced by muscle contraction.
In contrast, locomotion assays during early zebrafish larval development are used to assess muscle performance during sustained periods of muscle activity. Using a tracking system to monitor swimming behavior, we obtain an automated calculation of the frequency of activity and distance in 6-day old zebrafish, reflective of their skeletal muscle function. Measurements of swimming performance are valuable for phenotypic assessment of disease models and high-throughput screening of mutations or chemical treatments affecting skeletal muscle function.
Over the past decade zebrafish have been increasingly used to study muscle cell biology and disease. The rapid external development of the zebrafish embryo, coupled with its optical clarity, allows the direct visualization of muscle formation, growth, and function. The process of muscle development is highly conserved in zebrafish and this has allowed the successful modeling of a range of muscle diseases including muscular dystrophies and congenital myopathies1-8. Detailed examination of zebrafish models has not only provided novel insights into the pathobiology of these conditions but also provided a platform for the testing of suitable therapies6,9-13.
The analysis of zebrafish models of muscle diseases relies on reliable and reproducible assays to measure muscle performance. Previous studies have successfully measured the force generating capability of the zebrafish trunk muscle in fish between 3 and 7 dpf by electrically stimulating contraction of an immobilized fish attached to a force transduction system14. This can provide detailed measurements of force but are not ideally suited to higher throughput experiments and there are advantages to measuring muscle performance during swimming. At 2 dpf zebrafish muscle is fully functional and the fish can elicit burst swimming movements in response to stimuli. The touch-evoke escape response assay is used to measure acceleration during a burst swimming motion, which can be used as a measure of contractile force.
One of the most utilized measures of muscle function in myopathy patients is the 6 min walk test, which records the total distance walked on a hard flat surface15,16. We have applied a comparable test to measure muscle function in 6 dpf zebrafish larvae, whereby we monitor the total distance swum, and the total number of movements made by each larva over a 10 min period. This is performed using an automated tracking system, which provides reliable and high-throughput measurements of muscle performance. Both muscle tests are highly reproducible and have been used to quantify differences in muscle performance in zebrafish myopathy models8.
1. Touch-evoked Response Assay
2. Locomotion Assay — 10-min Swim Test
Touch evoked response assay can be used to determine the speed and acceleration of swimming movements which is a proportional measure of muscle force. In response to a mechanical stimulus, such as a small tap on the head 2 dpf wild type zebrafish exhibit a fast swimming action. Videos were captured and analyzed for two different zebrafish myopathy models: Tg(ACTA1D286G-eGFP), a model of nemaline myopathy that has been shown to have significant muscle weakness, and a model of Duchenne muscular dystrophy in which severe muscle defects have been described at 5 dpf19,20. Images from a video of a typical touch evoked assay are represented in Figure 1A. Acceleration of the zebrafish was examined and found to peak within the first 0.2 sec of the burst swimming escape response (Figure 1B). This peak maximum acceleration provides a measure that is proportional to the force generating capacity of the skeletal muscle. The maximum acceleration values were averaged to obtain a mean maximum acceleration value (± standard error of the mean) for each strain: Tg(ACTA1D286G-eGFP): mean = 276.0 ± 28.8 m/sec2, n = 3 independent replicate experiments comprising 15 individual fish; wildtype control: mean = 500.8 ± 50.28 m/sec2, n = 3 independent replicate experiments comprising 15 individual fish; dmdpc2-/- mutant: mean = 249.9 ± 19.1 m/sec2, n=3 independent replicate experiments comprising 12-19 individual fish; dmdpc2+/- heterozygotes: mean = 235.9 ± 8.7 m/sec2, n = 3 independent replicate experiments comprising 16-27 individual fish; dmdpc2+/+ wildtype homozygotes: mean = 230.9 ± 8.7 m/sec2, n = 3 independent replicate experiments comprising 8-27 individual fish (Figure 1C). As expected, the Tg(ACTA1D286G-eGFP) fish were found to have a significant decrease in maximum acceleration indicating reduced muscle function, which is consistent with mouse models and patient data8,21,22. The dmdpc2-/- mutant fish however, showed no difference in maximum acceleration, at 2 dpf, consistent with the detection of muscle defects from 3 dpf20 (Figure 1D).
Locomotion assays were performed at 6 dpf to determine the activity and distance swum by zebrafish strains as an indication of muscle performance. Following testing, a diagrammatic representation of the swimming movements over the ten-minute testing period was generated, with red and green lines representing periods of slow and fast movement respectively and black lines representing periods of inactivity (Figure 2). Individual wildtype zebrafish show high activity with relatively no periods of inactivity as opposed to Tg(ACTA1D286G-eGFP) fish, which are less active over the testing period (Figure 2B).
The swimming behavior was quantified by averaging the individual values of the number of movements and the distance swum by each fish (Figure 3). Both, Tg(ACTA1D286G-eGFP) fish (Figure 3A and 3B) and dmdpc2-/- mutant fish (Figure 3C and 3D) were found to have a significant decrease in the mean number of movements and distance swum compared to their respective controls: Tg(ACTA1D286G-eGFP) fish: mean number of movements = 94.3 ± 13.6, mean distance swum = 112.9 ± 18.4 mm, n = 3 independent replicate experiments comprising 45 fish; wild type controls: mean number of movements = 177.4 ± 14.0, mean distance swum = 300.2 ± 22.8 mm, n = 3 independent replicate experiments comprising 45 fish; dmdpc2-/- mutant: mean number of movements = 163.3 ± 30.0, mean distance swum: 298.4 ± 60.37 mm, n = 3 independent replicate experiments comprising 12-20 fish; dmdpc2+/- heterozygotes: mean number of movements = 362.3 ± 38.8, mean distance swum: 660.3 ± 86.1mm n = 3 independent replicate experiments comprising 17-27 fish; dmdpc2+/+ wildtype homozygotes: mean number of movements = 341.9 ± 91.6, mean distance swum = 574.3 ± 170.9mm n = 3 independent replicate experiments comprising 8-25 fish.
Figure 1: Quantification of touch-evoke response assay for 2 dpf zebrafish embryos. (A) Snapshot images of a control zebrafish during touch-evoke assays at 2 dpf. (B) Acceleration profile for the first 0.2 sec of a single Tg(ACTA1D286G-eGFP) (red) and single control (blue) zebrafish following application of the touch stimulus. The maximum acceleration is represented by the dotted lines. (C, D) Quantification of the maximum acceleration (m/sec2) recorded from touch-evoked response assays of (C) Tg(ACTA1D286G-eGFP) zebrafish and (D) dmdpc2-/- mutant fish compared to control zebrafish at 2 dpf. Error bars represent ± SEM for 3 replicate experiments, *p <0.05. Please click here to view a larger version of this figure.
Figure 2: Representation of locomotion assays for zebrafish embryos. (A) Zebrafish embryos are placed in 48-well plates and locomotion is recorded from above using an infrared digital camera. (B) Schematic of zebrafish movement during the testing period with red lines depicting fast movements, green lines depicting slow movements and black lines depicting inactivity (as determined by the detection thresholds entered in the software). Please click here to view a larger version of this figure.
Figure 3: Quantification of locomotion assays for 6 dpf zebrafish larvae. Quantification of the (A) number of movements and (B) distance travelled by Tg(ACTA1D286G-eGFP) zebrafish compared to control zebrafish at 6 dpf. Quantification of the (C) number of movements and (D) distance travelled by dmdpc2-/- mutant fish compared to control zebrafish at 6 dpf. Error bars represent ± SEM for 3 replicate experiments, *p <0.05, **p <0.01. Please click here to view a larger version of this figure.
Many different animal models including mice, dogs, zebrafish, flies and worms have contributed towards our understanding of the genetic and molecular basis of muscle diseases, and assisted in the development of therapeutic approaches to combat them. The zebrafish boasts several advantages for the study of muscle disease. The zebrafish provides a genetically manipulable system to assess complex muscle patterning in a suitable physiological environment, which is not possible in in vitro culture systems. Unlike other vertebrate animal models, the large number of fish produced, together with its optical clarity, facilitates rapid, high-throughput in vivo chemical and genetic screening.
Here we describe the development of zebrafish movement assays to provide a high-throughput and automated method to assess muscle performance during zebrafish embryogenesis. For both assays it must be acknowledged that circadian rhythms and external environmental stimuli will significantly affect zebrafish swimming behavior17,18. Repeated testing of the same zebrafish will also lead to habituation causing a decrease in response to the tactile stimulus23. Therefore, in order to achieve reproducible results between experiments each zebrafish embryo should only be tested once, the time of the day and lighting conditions should be standardized, and water temperature needs to be tightly regulated.
Using the touch evoked analysis at 2 dpf we can directly measure the maximum acceleration of a burst swimming action, which is proportional to muscle force. Previous techniques in zebrafish have examined muscle force by tying both ends of the embryos to experimental equipment following which muscle contraction is stimulated using an electric field and the force-generating capability of the muscle14 is measured. Whilst this method measures the force generating capacity of the larval muscle, it does not measure the actual force generated by the larval muscle during swimming. We therefore developed a method to indirectly assess the force generated during the normal larval swimming motion to provide an overall measure of muscle health. The high-speed video system, capable of recording individual zebrafish movements at a frame rate of 1,000 frames/sec can be used to identify small but significant differences in muscle function, which are not directly distinguishable by eye. It will be of interest to see how previously reported changes in electrically stimulated force-generation correlate with changes in swimming performance.
In addition the touch evoked response assays can also be used to assess swimming kinematics, such as the shape and speed of the body wave during the swimming motion24, to give a quantitative measurement of the locomotory behavior.
Due to the spontaneous movement of zebrafish larvae after 3 dpf, we were not able to perform the touch-evoke assays to measure muscle function. Conversely, we measured muscle performance over a longer period by determining the distance swum by zebrafish larvae at 6 dpf. This test, although an indirect measure of muscle function, can be used to identify fish displaying impaired muscle performance8 or neurodegeneration25,26. This test not only provides a measurement analogous to the 6 min walk test but is also suitable for automated high-throughput in vivo drug or mutagenesis screens.
The authors have nothing to disclose.
We thank Viewpoint for their kind sponsorship of this manuscript. This work was funded by an Australian National Health and Medical Research Council (NHMRC) Project Grant (APP1010110).
21G X 1' Blunt Needle | Terumo/Admiral Medical Supplies | TE2125 | |
48-well plates | Sigma | M8937 | |
90mm Petri Dishes | Pacific Laboratory Products PT | S90001 | |
High Speed Camera | Baumer | HXC20 | |
http://www.randomization.com | N/A | Steps 1.1.2, 2.1.3 | |
Incubator | Thermoline Scientific | TEI-43L | |
Plastic Pipette | VWR | 16001-188 | |
StreamPix5 | NorPix | Step 1.2.3 | |
Temperature Control Unit | Viewpoint | ||
Tweezers, style 8 | ProSciTech | T04-821 | |
Zebrabox System | Viewpoint | ||
Zebralab | Viewpoint | Steps 1.3.1, 2.2.1 |