The authors are deeply indebted to the efforts of Hughes lab members Drs Seetharamaiah Attili, Jana Koth, Fernanda Bajanca, Victoria C. Williams, Yaniv Hinits, Giorgia Bergamin, and Vladimir Snetkov for development of the described protocols, and to Henry Roehl, Christina Hammond, David Langenau and Peter Currie for sharing plasmids or zebrafish lines. SMH is a Medical Research Council (MRC) Scientist with Programme Grant G1001029, MR/N021231/1, and MR/W001381/1support. MA held a MRC Doctoral Training Programme PhD Studentship&#160;from King&#39;s College London. This work benefited from the trigonometrical input of David M. Robinson, scholar, mentor, and friend.

The authors declare no competing interests.

Here we report a method for accurate and efficient estimation of the muscle volume of live zebrafish larvae at stages or in genetic variants in which pigmentation is not a big hinderance to imaging and when transient anesthesia and/or immobilization is well tolerated. Whereas we have employed laser scanning confocal microscopy, the approaches described are applicable to spinning disk confocal or light sheet microscopy and to any other method that creates stacks of images at distinct focal planes. A series of increasingly sophisticated approaches to tissue size and cell content estimation is described. Each method has advantages and limitations, which we show can be quantified. A major limitation in studying tissue growth is the difficulty of analyzing growth changes in real time as growth rates alter in response to a series of molecular or sub-cellular events catalyzed by acutely administered stimuli. Moreover, individual variation can create problems when separate individuals are compared. The current approach allows measurement of tissue growth over periods of less than a day in single live individuals. Application of the approach can be envisaged on the minute timescale.
The described methods permit analyses that were hitherto impracticable. Using the 4-slice method, the imaging portion of the muscle growth assay can be completed on a sample size of around 20 fish within an hour by a trained operator. This is in stark contrast to the conventional Full Stack method, which takes at least 3 h for the same number of fish (i.e., three times longer). If high resolution images are required for subsequent sub-cellular analyses, the Full Stack method can easily require 30 min per fish, making growth assays of cohorts of animals at a similar developmental stage impossible. In contrast, the 2- or 4-slice methods permit rapid high quality image capture. Moving to consideration of image analysis, the advantages of the 2- or 4-slice method in saving of operator time (in the absence of automated image segmentation) over the Full Stack method are enormous. Each fish requires about 20 min for Full Stack analysis, but only 3-4 min for 4-slice analysis. Operator time can be further conserved using the almost-as-accurate 2-slice method. Thus, the described method is efficient and thereby increases flexibility in experimental design.
The major limitation of the 4- or 2-slice methods is that both methods are estimates, due to the overlapping chevron shape of somites (e.g., the volume of regions of two neighboring somites (e.g., myotome 16 and 17). It is shown that this can over-estimate the actual myotome 17 volume by around 2%, depending on precisely where YZ slices were selected. Moreover, manual tracing of the somite borders during measurements might contribute to variations in estimates, although little inter-experimenter difference was found (data not shown). Measurement errors might be addressed using thresholding, filtering, and segmentation algorithms to acquire surface area in a more objective and reproducible manner. However, customizations will still be required to account for variations in the background fluorescence (such as due to embedding and/or thickness of the LMA) and the expression level of fluorescence proteins of individual larva over time. Note that such automated measurements will be even more challenging if a non-muscle-specific reporter is used, such as the &#946;-actin:HRAS-EGFP line. Nonetheless, under many circumstances, for example, when comparing effects of manipulations between fish subjected to different treatments that are expected to affect all muscle tissue, the inaccuracy may be immaterial. However, if maximal accuracy is required for comparison to fiber or nuclear numbers counted solely from myotome 17, for example, the &#39;slice&#39; methods can be improved. This can be achieved either by using the arduous Full Stack method, by mathematical correction by multiplying the measured 4-slice volumes by 0.98, or by moving the location of the YZ CSA scans posteriorly to reflect more accurately the true CSA of myotome 17.
A second limitation of the method is its sensitivity to the mounting orientation of the fish. In practice, skilled operators can orient fish within reasonable limits most of the time, even when working quickly to embed many samples. Modifications to equipment on the microscope stage can be envisaged that would allow correction of yaw and roll prior to scanning. Without such apparatus, a method to quantify misorientation that can then be used to correct the measured volumes is described. Moreover, even if misorientation increases variability in measured volume, and thus reduces the chance of observing small effect sizes, in many situations such variation will affect control and experimental samples similarly. So false positive results are unlikely, if operators are aware of the issue.
The described methods have initially been applied to analyze the role of electrical activity in muscle growth, a subject with a long history of analysis in a wide range of species (reviewed in18). To this end, simple methods to block endogenously triggered activity are described in detail and replaced with controlled patterned electrical stimulation in the zebrafish larva. Advantages of this approach are the removal of neural feedback controls40, the elimination of the effects of altered nutrition and the ability to analyze circadian effects on growth itself, rather than on growth proxies, such as protein turnover26. As different patterns of electrical activity trigger distinct muscle responses, regulating fiber type, size, and metabolism41,42,43,44,45,46,47,48, the current methods open the zebrafish to such analyses.
The methods described offer a suite of techniques with which many aspects of muscle physiology, cell biology, and pathology can be analyzed in unprecedented temporal and spatial resolution by taking advantage of the relatively unexplored zebrafish. The current approaches could clearly be applied to other species, regions of the body, and developmental stages. The rapid early growth of zebrafish larvae makes detection of acute effects of manipulations on tissue growth and morphogenesis particularly attractive areas of study. Moreover, zebrafish muscle is shown to share various growth mechanisms and controls with mammals. While zebrafish are vertebrates that conserve many aspects of muscle molecular genetics, cell, and developmental biology with humans, there are also significant differences in the control of muscle growth. For example, slow and fast fiber types are more clearly spatially segregated in fish and the innervation of muscle shows differences. Furthermore, it must be borne in mind that, so far, we have only been able to analyze the early stages of development. Similar analyses at later stages are envisaged.

Regulated tissue growth, comprising increase in cell number (hyperplasia) and/or cell size (hypertrophy), is a crucial factor in development, regeneration, and ecological and evolutionary adaptation. Despite huge advances in molecular genetic understanding of both cell and developmental biology over recent decades, mechanistic understanding of the regulation of tissue and organ size is still in its infancy. One reason for this lacuna in knowledge is the difficulty of quantifying tissue growth in living organisms with the necessary spatial and temporal accuracy.
Various aspects of growth of whole organisms can be measured repeatedly over time, revealing growth curves for each individual1,2,3,4,5. Increasingly sophisticated scanning methods, such as dual X-ray absorptiometry (DXA), computerized tomography (CT), and magnetic resonance imaging (MRI), permit the tracking of growth of whole organs and other body regions (for example, individual identified skeletal muscles) in single individuals, both human and in model organisms6,7,8,9,10. However, these methods do not yet have the resolution to reveal individual cells and thus the links between cellular behaviors and tissue level growth have been hard to discern. To make such links, traditional studies have often relied upon cohorts of similar individual animals, a few of which are sacrificed at successive timepoints and then analyzed in cytological detail. Such approaches require averaging the observed changes across groups of (preferably similar, but nevertheless variable) individuals and thus suffer from a lack of temporal and spatial resolution, making it hard to find correlated events at the cellular level suggestive of cause and effect.
Studies on invertebrate model organisms, initially in C. elegans and D. melanogaster, have circumvented these problems by developing optical microscopy to achieve cellular resolution and accurately measure growth over time in single individuals. Such studies have revealed strikingly invariant cell lineage behaviors in the growth of these small model organisms11,12,13,14,15,16,17. However, many animals, including all vertebrates, have indeterminate cell lineages, and control tissue growth by mysterious feedback processes that serve to turn the genetically-encoded growth program into a functional three dimensional organism with all its constituent tissues and organs suitably matched in size. To understand these complex growth processes, it is desirable to image whole tissues or organs over time in single individuals that can be experimentally manipulated by genetic, pharmacological or other interventions at a time of choice and the effect subsequently analyzed.
Each vertebrate skeletal muscle has a defined size, shape and function, and well-characterized interactions with adjacent tissues, such as bone, tendon, and nerves. Some muscles are small, lie just under the skin and are therefore good candidates for high-resolution imaging studies. Similar to most organs, each muscle grows throughout embryonic, postnatal, and juvenile life, before reaching a stable adult size. Muscle, however, also has a unique ability to change size during adult life, dependent upon use and nutrition18, and this property has a major impact on organismal fitness, sporting performance, and independent living. Loss of muscle mass and function in old age, sarcopenia, is an issue of increasing concern for societies faced with an ageing population19,20,21.
We and others have focused on the growth of defined blocks of skeletal muscle tissue in the segmentally-repeating body of zebrafish larvae, as an apparently closed system containing several hundred cells in which tissue growth, maintenance, and repair can be observed and manipulated22,23,24,25,26. While some quantitative work has previously been reported25,26,27,28,29,30,31,32,33,34,35, no detailed and validated method of measuring muscle growth in cellular detail in individual vertebrate organisms over time is available. Here an efficient protocol for how to perform such repeated measurements is described, along with validation, and an example of its use to analyze changes in both hypertrophic and hyperplastic growth in response to altered electrical activity is provided.

<table><tbody><tr><td>Adhesive, Blu Tack</td><td>Bostik</td><td>-</td><td>-</td></tr><tr><td>Aerosol vacuum&amp;nbsp;</td><td>-</td><td>-</td><td>-</td></tr><tr><td>Agarose</td><td>Sigma-Aldrich</td><td>A9539</td><td>-</td></tr><tr><td>Agarose, low gelling temperature</td><td>Sigma-Aldrich</td><td>A9414</td><td>Once melted, keep at 37&lt;sup&gt;o&lt;/sup&gt;C in a block heater to remain in liquid form for repeated use.</td></tr><tr><td>Block heater</td><td>Cole-Parmer</td><td>SBH130</td><td>-</td></tr><tr><td>BODIPY FL C5-ceramide</td><td>Thermo Scientific</td><td>D3521</td><td>To be diluted in fish water and used at 5 &amp;micro;M for overnight incubation.</td></tr><tr><td>Crocodile clips and wires</td><td>-</td><td>-</td><td>-</td></tr><tr><td>Fiji/imageJ</td><td>National Institutes of Health, NIH</td><td>-</td><td>-</td></tr><tr><td>Fish medium, Fish water</td><td>-</td><td>-</td><td>Circulating system water collected from the fish facility.</td></tr><tr><td>Fish medium, E3 medium</td><td>-</td><td>-</td><td>E3 is described in The Zebrafish Book. http://zfin.org (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl&lt;sub&gt;2&lt;/sub&gt;, and 0.33 mM MgSO&lt;sub&gt;4 &lt;/sub&gt;in distilled water).</td></tr><tr><td>Fluorescence microscope</td><td>Leica</td><td>Leica MZ16F</td><td>Fluorescence microscope of other kind are also expected to be suitable.</td></tr><tr><td>Glass needle</td><td>World Precision Instruments, Inc.</td><td>1B100-6</td><td>To be fire-polished to prevent damage of the embryos during manipulation.</td></tr><tr><td>Grass stimulator</td><td>Grass Instruments</td><td>S88</td><td>Stimulators of other kind are also expected to be suitable.</td></tr><tr><td>Kimwipes, Delicate Task Wipers</td><td>Kimberly-Clark Professional</td><td>13258179</td><td>-</td></tr><tr><td>Laser scanning microscope (LSM)&amp;nbsp;</td><td>Zeiss</td><td>Zeiss LSM 5 Exciter&lt;br/&gt; Zeiss LSM 880</td><td>LSM of other kind are also expected to be suitable.</td></tr><tr><td>Nunc Cell-Culture Treated, 6-well plate</td><td>Thermo Scientific</td><td>140675</td><td>-</td></tr><tr><td>Objective, 20&amp;times;/1.0W water immersion</td><td>Zeiss</td><td>-</td><td>-</td></tr><tr><td>Pasteur Pipette, Graduated 1 mL</td><td>Starlab Group</td><td>E1414-0100</td><td>-</td></tr><tr><td>Pasteur Pipette, Micro Fine Tip 1 mL</td><td>Starlab Group</td><td>E1414-1100</td><td>-</td></tr><tr><td>Petri dish, 60 mm</td><td>Sigma-Aldrich</td><td>P5481</td><td>-</td></tr><tr><td>Plasmid, CMV-Cerulean</td><td>Christina L. Hammond (University of Bristol)</td><td></td><td>pCS2+_cerulean_kanR plasmid injected at 25-75 pg at one-cell stage.&amp;nbsp; Citation: Bussman J, and Schulte-Merker S. (2011) Development 138:4327-4332. doi: 10.1242/dev.068080.</td></tr><tr><td>Plasmid, pCS-mCherry-CAAX</td><td>Henry Roehl (University of Sheffield)</td><td>-</td><td>For in vitro transcription using the SP6 promoter (plasmids containing other membrane labelling markers can be used);&lt;br/&gt; synthesised capped mRNA to be injected at 100-200 pg at one-cell stage.</td></tr><tr><td>Pulse Controller&amp;nbsp;</td><td>Hoefer Scientific Instruments</td><td>PC750</td><td>-</td></tr><tr><td>Soldering iron</td><td>-</td><td>-</td><td>-</td></tr><tr><td>Tricaine</td><td>Sigma-Aldrich</td><td>E10521</td><td>Ethyl 3-aminobenzoate methanesulfonate/ MS-222; to be dissolved in fish water and used at 0.6 mM.</td></tr><tr><td>Volocity</td><td>Perkin Elmer/Quorum Technologies Inc</td><td>-</td><td>-</td></tr><tr><td>Watchmaker forceps, No. 5</td><td>-</td><td>-</td><td>-</td></tr><tr><td>Wire, Platinum</td><td>Goodfellow</td><td>PT005142/12</td><td>0.40 mm in diameter; an expensive alternative of silver.</td></tr><tr><td>Wire, Silver</td><td>Acros Organics</td><td>317730010</td><td>0.25 mm in diameter (a range of diameter i.e. 0.25-0.5 mm had been tested, which produced similar results).</td></tr><tr><td>Zebrafish, myog:H2B-mRFP</td><td>David M. Langenau (Massachusetts General Hospital; Harvard Stem Cell Institute)</td><td>-</td><td>ZFIN official name: Tg(myog:Hsa.HIST1H2BJ-mRFP), fb121Tg.&amp;nbsp; http://zfin.org/ZDB-ALT-160803-2&amp;nbsp; Citation: Tang Q, Moore JC, Ignatius MS, Tenente IM, Hayes MN, Garcia EG, Torres Yord&amp;aacute;n N, Bourque C, He S, Blackburn JS, Look AT, Houvras Y, Langenau DM. Imaging tumour cell heterogeneity following cell transplantation into optically clear immune-deficient zebrafish. Nat Commun. 2016 Jan 21;7:10358. doi: 10.1038/ncomms10358.</td></tr><tr><td>Zebrafish, &amp;alpha;-actin:mCherry-CAAX</td><td>Peter D. Currrie (ARMI, Monash University)</td><td>-</td><td>ZFIN official name: Tg(actc1b:mCherry-CAAX), pc22Tg.&amp;nbsp; http://zfin.org/ZDB-ALT-150224-2 Citation: Berger J, Tarakci H, Berger S, Li M, Hall TE, Arner A, and Currie PD. Loss of Tropomodulin4 in the zebrafish mutant tr&amp;auml;ge causes cytoplasmic rod formation and muscle weakness reminiscent of nemaline myopathy. Dis Model Mech. 2014 Dec;7(12):1407-15. doi: 10.1242/dmm.017376.</td></tr><tr><td>Zebrafish, &amp;beta;-actin:HRAS-EGFP</td><td>-</td><td>-</td><td>ZFIN official name: Tg(Ola.Actb:Hsa.HRAS-EGFP), vu119Tg. http://zfin.org/ZDB-ALT-061107-2&amp;nbsp; Citation: Cooper MS, Szeto DP, Sommers-Herivel G, Topczewski J, Solnica-Krezel L, Kang HC, Johnson I, and Kimelman D. Visualizing morphogenesis in transgenic zebrafish embryos using BODIPY TR methyl ester dye as a vital counterstain for GFP. Dev Dyn. 2005 Feb;232(2):359-68. doi: 10.1002/dvdy.20252.</td></tr><tr><td>ZEN software</td><td>Zeiss</td><td>-</td><td>-</td></tr></tbody></table>

real time and repeated measurement of skeletal muscle growth in individual live zebrafish subjected to altered electrical activity

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish with fluorescently-marked plasma membranes can be scanned in a confocal laser scanning microscope in order to determine the volume of muscle tissue and the number of muscle fibers present. Efficient approaches for the measurement of cell number and size in live animals over time are described and validated against more arduous segmentation methods. Methods are described that permit the control of muscle electrical, and thus contractile, activity. Loss of skeletal muscle contractile activity greatly reduced muscle growth. In larvae, a protocol is described that allows reintroduction of patterned electrical-evoked contractile activity. The described methods minimize the effect of inter-individual variability and will permit analysis of the effect of electrical, genetic, drug, or environmental stimuli on a variety of cellular and physiological growth parameters in the context of the living organism. Long-term follow-up of the measured effects of a defined early-life intervention on individuals can subsequently be performed.

Watch this Scientific Journal Video about Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity at JoVE.com

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish ...

real-time-repeated-measurement-skeletal-muscle-growth-individual-live

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

1.9K Views.  King’s College London. Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

Video: Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

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

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.

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 ...

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the cells are located. Specifically, they have been developed to analyze the electrical activity of excitable cells, such as spinal neurons. In such a scenario, it is crucial to preserve the integrity of the spinal cell, but it is also important to preserve the anatomy and physiological shape of the systems involved. Indeed, the comprehension of the manner in which the nervous system-and other complex systems-works must be based on a systemic approach. For this reason, the live zebrafish embryo was chosen as a model system, and the spinal neuron membrane voltage changes were evaluated without interfering with the physiological conditions of the embryos. 
Here, an approach combining the employment of zebrafish embryos with a FRET-based biosensor is described. Zebrafish embryos are characterized by a very simplified nervous system and are particularly suited for imaging applications thanks to their transparency, allowing for the employment of fluorescence-based voltage indicators at the plasma membrane during zebrafish development. The synergy between these two components makes it possible to analyze the electrical activity of the cells in intact living organisms, without perturbing the physiological state. Finally, this non-invasive approach can co-exist with other analyses (e.g., spontaneous movement recordings, as shown here).

Protocols described here allow for the study of the electrical properties of excitable cells in the most non-invasive physiological conditions by employing zebrafish embryos in an in vivo system together with a fluorescence resonance energy transfer (FRET)-based genetically encoded voltage indicator (GEVI) selectively expressed in the cell type of interest.

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Neuroscience

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, the muscle stem cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. In many muscle wasting conditions, including muscular dystrophy and ageing, this process is impaired resulting in the inability of muscle to regenerate. The process of muscle regeneration in zebrafish is highly conserved with mammalian systems providing an excellent system to study muscle stem cell function and regeneration, in muscle wasting conditions such as muscular dystrophy. Here, we present a method to examine muscle regeneration in zebrafish models of muscle disease. The first step involves the use of a genotyping platform that allows the determination of the genotype of the larvae prior to eliciting an injury. Having determined the genotype, the muscle is injured using a needle stab, following which polarizing light microscopy is used to determine the extent of muscle regeneration. We therefore provide a high throughput pipeline which allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle regeneration is driven by tissue resident muscle stem cells, which are impaired in many muscle diseases such as muscular dystrophy, and this results in the inability of muscle to regenerate. Here, we describe a protocol that allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Biology

Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional measures of muscle health. In this video article, we demonstrate a method to measure force generation during contraction of zebrafish larval trunk muscle. Force measurements are accomplished by placing an anesthetized larva into a chamber filled with a salt solution. The anterior end of the larva is tied to a force transducer and the posterior end of the larva is tied to a length controller. An isometric twitch contraction is elicited by electric field stimulation and the force response is recorded for analysis. Force generation during contraction provides a measure of overall muscle health and specifically provides a measure of muscle function. Although we describe this technique for use with wild-type larvae, this method can be used with genetically modified larvae or with larvae treated with drugs or toxicants, to characterize muscle disease models and evaluate treatments, or to study muscle development, injury, or chemical toxicity.

Force measurements can be used to demonstrate changes in muscle function due to development, injury, disease, treatment or chemical toxicity. In this video, we demonstrate a method to measure force during a maximal contraction of zebrafish larval trunk muscle.

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional ...

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular dystrophies1-3, congenital myopathies4,5, and disruptions in sarcomeric assembly6,7, due to high genomic and structural conservation with mammals8. Muscular disorganization and locomotive impairment can be quickly assessed in the zebrafish over the first few days post-fertilization. Two assays to help characterize skeletal muscle defects in zebrafish are birefringence (structural) and touch-evoked escape response (behavioral).
Birefringence is a physical property in which light is rotated as it passes through ordered matter, such as the pseudo-crystalline array of muscle sarcomeres9. It is a simple, noninvasive approach to assess muscle integrity in translucent zebrafish larvae early in development. Wild-type zebrafish with highly organized skeletal muscle appear very bright amidst a dark background when visualized between two polarized light filters, whereas muscle mutants have birefringence patterns specific to the primary muscular disorder they model. Zebrafish modeling muscular dystrophies, diseases characterized by myofiber degeneration followed by repeated rounds of regeneration, exhibit degenerative dark patches in skeletal muscle under polarized light. Nondystrophic myopathies are not associated with necrosis or regenerative changes, but result in disorganized myofibers and skeletal muscle weakness. Myopathic zebrafish typically show an overall reduction in birefringence, reflecting the disorganization of sarcomeres.
The touch-evoked escape assay involves observing an embryo&#39;s swimming behavior in response to tactile stimulation10-12. In comparison to wild-type larvae, mutant larvae frequently display a weak escape contraction, followed by slow swimming or other type of impaired motion that fails to propel the larvae more than a short distance12. The advantage of these assays is that disease progression in the same fish type can be monitored in vivo for several days, and that large numbers of fish can be analyzed in a short time relative to higher vertebrates.

The zebrafish is now an established and powerful tool for modeling muscular dystrophies, congenital myopathies, and related neuromuscular diseases. Birefringence and touch-evoked escape behavior are two common noninvasive assays used to determine the degree of muscular disorganization and locomotive impairment of zebrafish embryos during early development.

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular ...

Live Imaging of Synaptic Vesicle Recycling in the Neuromuscular Junction of Dissected Larval Zebrafish

Neuronal communication is mediated by synaptic transmission, which depends primarily on the release of neurotransmitters stored in synaptic vesicles (SVs) in response to an action potential (AP). Since SVs are recycled locally at the presynaptic terminal, coordination of SV exocytosis and endocytosis is important for sustained synaptic transmission. A pH-sensitive green fluorescent protein, called pHluorin, provides a powerful tool to monitor SV exo/endocytosis by targeting it to the SV lumen. However, tracking AP-driven SV recycling with the pHluorin-based probes is still largely limited to in vitro culture preparations because the introduction of genetically encoded probes and subsequent optical imaging is technically challenging in general for in vivo animal models or tissue preparations. Zebrafish is a model system offering valuable features, including ease of genetic manipulation, optical clarity, and rapid external development. We recently generated a transgenic zebrafish that highly expresses a pHluorin-labeled probe at motor neuron terminals and developed a protocol to monitor AP-driven SV exo/endocytosis at the neuromuscular junction (NMJ), a well-established synapse model that forms in vivo. In this article, we show how to prepare larval zebrafish NMJ preparation suitable for pHluorin imaging. We also show that the preparation allows time-lapse imaging under conventional upright epifluorescence microscope, providing a cost-effective platform for analyzing NMJ function.

The zebrafish larval neuromuscular junction is an attractive model for studying synaptic physiology. It is amenable to many experimental techniques, including electrophysiology and optical imaging. Here, we describe a protocol for imaging synaptic transmission using a pHluorin-based probe under an upright epifluorescence microscope.

Neuronal communication is mediated by synaptic transmission, which depends primarily on the release of neurotransmitters stored in synaptic vesicles (SVs) ...

In Vivo Confocal Fluorescence Imaging of Neural Activity Induced by Sensory Stimulation in Partially Restrained Larval Zebrafish

Zebrafish larvae are a promising vertebrate model system for studying the neural mechanisms of behavior. Their translucence and relatively simple neural circuitry facilitate the use of optogenetic techniques in cellular analyses of behavior. Fluorescent indicators of in vivo neural activity, such as GCaMP6s, have been widely used to study the neural activity associated with simple behaviors in larval zebrafish. Here, we present a protocol for detecting sensory-induced activity in semi-restrained zebrafish larvae using the transgenic line Tg(elav3:GCaMP6s). In particular, we use the chemical agent allyl isothiocyanate to induce a robust, reproducible fluorescent response in a brain region at the border of the hindbrain and spinal cord. We discuss the potential uses of GCaMP6s for optical monitoring of neural activity during a range of behavioral paradigms and the limitations of this technique. Our protocol outlines an accessible approach for monitoring dynamic, behavior-related in vivo neural activity in the larval zebrafish brain.

Here, we present a detailed protocol to examine neural activity in brain regions of transgenic zebrafish that express GCaMP calcium indicators using confocal microscopy.

Zebrafish larvae are a promising vertebrate model system for studying the neural mechanisms of behavior. Their translucence and relatively simple neural ...