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Developmental Biology

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

Published: June 16, 2022 doi: 10.3791/64063
* These authors contributed equally

Summary

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.

Abstract

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.

Introduction

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.

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Protocol

All research described was performed in compliance with institutional guidelines and under suitable licenses from UK Home Office in accordance with the Animal (Scientific Procedures) Act 1986 and subsequent modifications. Embryos/larvae should be reared at 28.5 °C until completion of gastrulation but may then be kept at 22-31 °C to control the rate of development. Fish may be scanned or stimulated at room temperature.

1. Anesthetize zebrafish larvae

  1. Cross suitable fluorescent reporter adult fish such as Tg(Ola.Actb:Hsa.HRAS-EGFP)vu119Tg reference36 or Tg(α-actin:mCherry-CAAX)pc22Tg reference37 and collect embryos as described38.
  2. At the time of choice, such as 2 days post-fertilization (dpf), anesthetize embryos briefly using Tricaine-containing fish medium (either fish water or E3 medium), and screen for EGFP or mCherry under a fluorescence microscope, such as a Leica MZ16F. If one has many embryos, select those with the brightest signal. Return embryos to normal fish medium immediately after screening.

2. Mounting fish for confocal scanning

  1. Turn on the confocal laser scanning system and lasers, to let the system stabilize for 30-60 min.
    NOTE: Here, we used a Zeiss LSM 5 Exciter microscope with upright materials stand (which enhances working distance) equipped with a 20x/1.0 W water-immersion objective.
  2. Prepare 1% low melting agarose (LMA) and keep in a 37 °C heat block for repeated use in a 1.5 mL tube. To avoid heat-shock, remove the LMA aliquot from the heat block and let it cool to just above setting before applying to the larva, testing against one's skin to judge the appropriate temperature, as when assessing the temperature of baby formula milk.
  3. Select fish to be mounted and transiently anesthetize each fish in turn with Tricaine (0.6 mM in fish medium).
  4. Take a 60 mm diameter Petri dish that has been coated with a layer of 1% agarose and place on stage of a dissecting microscope.
  5. Transfer the larva with a 1 mL plastic Pasteur pipette onto the 60 mm coated Petri dish and remove as much transferred medium as possible. Then, still using the Pasteur pipette, place 5 to 10 drops of LMA onto the fish and rapidly position horizontally in lateral view with forceps (or a fire-polished fine glass needle) before the LMA sets. For optimal imaging, it is desirable to position the larva close to the upper surface of the LMA.
    1. Alternatively, using a 1 mL plastic Pasteur pipette, collect the larva with as little fish medium as possible, and transfer the larva into the aliquot of cooled LMA. Allow the larva to sink for 5 s to become fully surrounded by LMA. Then, retrieve the larva and transfer it in a drop of LMA onto the agarose-coated Petri dish. Quickly orientate and position the larva as described above.
  6. Orient the larva with both its anteroposterior and dorsoventral axes within 10° of the horizontal (see the note 'On error and its correction', at point 4.6 below).
    1. If larva is not correctly mounted horizontally near the surface of the agarose drop, remove and re-embed. Larvae can be easily retrieved by gentle suction using a microfine 1 mL plastic Pasteur pipette, and LMA can be gently removed using Kimwipes. Practice really does make perfect in the mounting procedure; spend an afternoon embedding some unimportant larvae before trying this on a real experiment.
      ​NOTE: On microscope design: Many labs use inverted confocal microscopes for imaging through a coverslip. We have found that the repeated embedded and removal of fish held in agarose under a coverslip for observation in an inverted microscope leads to greater loss of samples during repeated scanning than in the described procedure with an upright microscope. For this reason, the use of an upright system is recommended, if available. Nevertheless, a key to high quality data is the proper selection and use of objective and scan parameters, a subject too large for discussion here.

3. Confocal scanning

  1. When LMA has set, flood the dish with around 10 mL of Tricaine-containing fish medium. If planning to capture confocal stacks, let the mounted fish rest for at least 10 min before proceeding to scanning, as some agarose swelling occurs.
  2. Load the sample dish to the stage of the confocal system, locate the larva and focus on the desired somite. Somite 17 may be chosen because of its ease of localization near the anal vent and ease of imaging. Check by counting somites from anterior.
    NOTE: The first somite is fused behind the ear and has no anterior border, but can be readily observed to have striated muscle fibers.
  3. Set up as if to capture a Z-stack by defining top (i.e., just above the skin) and bottom (i.e., just below the notochord, so as to include the entire somite, even if the fish is mounted slightly skewed). Both left and right sides can be captured as desired. This will ensure that all the rapid YZ scans capture the desired region(s).
  4. Capture an XY image as follows. Orient the scan area with respect to the fish, as the confocal software permits. Position the fish with the anteroposterior axis parallel to the imaged X axis and dorsoventral axis parallel to the Y axis with somite 17 in the center of the field as shown in Supplementary File 1. Focus on a mid-level plane in the uppermost myotome in which the whole epaxial and hypaxial somite halves together with the vertical and horizontal myosepta are visible and capture a high resolution XY image. Remember to name and save the image.
  5. Capture one or more YZ images as follows. In the Representative Results (below) the accuracy of 2-slice and 4-slice methods is compared. In the 2-slice approach, single XY and YZ scans are employed. In the 4-slice approach, three YZ scans are averaged to give a more accurate estimate of myotome volume. If required by the confocal software, re-orient the scan field.
    1. Draw a precisely dorsal to ventral line across the chosen somite perpendicular to the anteroposterior axis of the fish at a selected anteroposterior position. Perform a Z-stack line scan.
    2. Repeat the YZ line scan three times at defined anteroposterior positions along the selected myotome to capture YZa, YZm, and YZp. Representative results are shown in Figure 2A. Name and save these images together with the related XY image.
      ​NOTE: On selection of YZ planes: The myotome is V-shaped and its form changes during growth. To obtain the most accurate assessment of myotome 17 volume with the 4-slice method, position YZa on the anterior tip of the myotome, YZp on the posterior tips of the myotome at dorsal and ventral extremes, and YZm halfway between YZa and YZp. Assuming the somite tapers uniformly, the mean of measurements from each YZ section will represent the myotome as a whole. For the 2-slice method, the single YZ scan should be positioned at the posterior end of the horizontal myoseptum, which roughly corresponds to the anteroposterior center of the myotome of interest (such as YZm). Alternatively, a set of three YZ sections may be taken at anterior, middle, and posterior of the horizontal myoseptum but, as shown below, such measurement will slightly over- or under-estimate myotome volume (for rostral and caudal somites, respectively, due to myotome tapering). Fundamentally, consistency in positioning of YZ slice plane(s) between fish and experiments is key to reproducibility.

4. Analysis

  1. As the myotome changes size along the fish in a graded manner, always work with the same somite in comparative studies.
  2. To measure and calculate the myotome volume, use the confocal software (such as the .lsm files created using Zeiss ZEN microscope software) or open-source universal image analysis software, such as Fiji/ImageJ.
    NOTE: If changing file formats, make sure the Z-step size is correctly transferred, as not all software can read proprietary confocal file formats correctly. For example, to import a ZEN line scan image into Fiji, first use the File/Export command to export as .tif in the Full resolution image window - single plane format, and then import into Fiji. Although YZ scan.lsm can be opened directly in Fiji, the resulting YZ images are generally compressed in the Z dimension due to incorrect evaluation of the Z step size.
  3. Analysis using ZEN
    1. First, open the XY scan.lsm files in ZEN. Go to the Graphics tab and select the Line tool. Draw a line between the two vertical myosepta of somite 17 spanning the entire the myotome length (parallel to the anteroposterior axis of the fish). Check the M box to reveal the values of the measurement (Length = 89.71 µm, see Supplemental File 2).
    2. Open the YZ scan.lsm files. Under the Graphics tab, select the Closed Bezier tool. Draw around the perimeter of the myotome. Once completed, check the M box, this would reveal the value of the measurement (Area = 11980.01 µm2, see Supplemental File 3).
    3. Record the values of each measurement manually in a spreadsheet. Average the CSA measurements as required. Volume of the myotome can be calculated as Volume = Myotome length x CSA, i.e., 89.71 µm x 11980.01 µm2 = 1.075 x 106 µm3.
  4. Analysis using Fiji/ImageJ
    1. Open the XY scan.lsm files in Fiji/ImageJ. Check whether XY images directly opened in Fiji are correctly calibrated in scale, as they should be.
    2. Select the Straight Line tool from the icons. Draw a line along the length of somite 17 as described in step 4.3.1. Set measurement parameters by going to Analyze, then select Set Measurements…, and check the following boxes Area and Display Label. To measure, simply press the hot key M, or go to Analyze menu and select Measure. A resulting pop-up window lists all measurement values (i.e., Length = 90.023 µm; see Supplemental File 4). The results can be saved in form of .csv and opened in Microsoft Excel or similar for subsequent analysis.
    3. To measure CSA on the YZ images, open YZ images in .tif format as described in step 4.2.
    4. Calibrate the YZ .tif images as they are uncalibrated when exported. Parameters for the calibration can be obtained in ZEN by going to the Info of the selected images: record the Scaling X (0.489 µm) and Scaling Z values (0.890 µm; see Supplemental File 5). Next, while the images are open in Fiji, go to Image and select Properties…. Input 0.489 µm for the Pixel Width and Pixel Height, and 0.890 µm for the Voxel Depth. Check the Global box to apply the calibration universally if repeated measurement of YZ images is anticipated (see Supplemental File 6).
      NOTE: Make sure all YZ images are captured using the same scanning parameters; restart Fiji/ImageJ or modify the calibration values if a new set of calibration is required.
    5. To measure the CSA of the calibrated YZ images, select the Polygon Selections tool from the icons. Draw around the perimeter of the somite, and press M to reveal the values of the measurement (Area = 11980.395 µm2; see Supplemental File 7). Volume of the myotome can be calculated as Volume = Myotome length x CSA, i.e., 90.023 µm x 11980.395 µm2 = 1.079 x 106 µm3.
    6. Repeat the measurements on the other XY and YZ images. It is recommended to use the same software for all measurements within an experimental series for consistency. The volume estimate from each software is similar but not identical due to the distinct drawing tools, i.e., ZEN = 1.074 × 106 µm3 and Fiji/ImageJ = 1.079 × 106 µm3. Growth of the myotome between two time points (i.e., 3 to 4 dpf) can be calculated as: (Volume 4 dpf - Volume 3 dpf)/ Volume 3 dpf × 100%.
      NOTE: On error and its correction. During mounting, the fish should be orientated with its sagittal plane (i.e., the anteroposterior and dorsoventral axes) as close as possible to horizontal, to avoid yaw and roll, respectively. This is because both the myotome length L measured from the XY scan and the CSA measured from a YZ scan will be over-estimated if the fish shows yaw (rotation around the dorsoventral axis) due to oblique anteroposterior mounting. Neither pitch nor roll during mounting should affect measurements after scanning as described in section 3. Nevertheless, dorsoventral rotation (roll) degrades image quality. Simple trigonometry shows that up to 10° of yaw will give 3% error in volume measurement, as measured L and CSA each increase in proportion to (cosq)-1, where q is the angle away from anteroposterior horizontal (yaw). 15° and 20° off will give 7% and 13% over-estimates of volume, respectively.
      As the notochord is cylindrical, inclusion of the whole notochord in the YZ scan can be used to calculate the angle and extent of obliquity from the orientation and magnitude of the major and minor axes and thereby correct the measured L and CSA to maximize accuracy. Corrected CSA = Measured CSA x Notochord minor axis/Notochord major axis. Corrected L = Measured L x Notochord minor axis/Notochord axis in microscope Z direction.
      A further consideration permits additional correction of L. As the myotome grows, it skews in the coronal plane (normal to the dorsoventral axis) such that the medial myotome is slightly anterior to the lateral myotome. Viewed from dorsal, the vertical myosepta on left and right sides form a broad chevron pointing anterior. If yaw is low, this morphology does not affect measurement of L. But if yaw is significant, trigonometrical correction becomes challenging and a better approach is to measure True L directly by estimating the XYZ coordinates of the two points where the anterior and posterior vertical myosepta meet the notochord at the horizontal myoseptum. Simple trigonometry permits calculation of True L from these coordinates as L = SQRT[(X2 - X1)2 + (Y2 - Y1)2 + (Z2 - Z1)2]. Weaknesses of this last approach are that a) selection of the points can vary with operator and b) no visual record of the points chosen is retained. This consideration does not affect CSA correction.

5. Optional method: Remove and re-introduce muscle electrical activity

  1. Create a stimulation chamber.
    1. Take a 6 x 35 mm well plate, create two small openings (<5 mm in diameter, 1 cm apart) on each side of each well (see Figure 1) using a narrow soldering iron.
      NOTE: Handle the hot soldering iron with care and work in a fume hood if desired to avoid inhaling vapor.
    2. Thread a pair of silver or platinum wires (~20 cm long) through the openings of each well (see Figure 1). Reusable adhesive material (e.g., BluTack) can be applied near the openings to keep the wires in place, and ensure a 1 cm separation between the wires (see Figure 1).
  2. At 3 dpf, split fish into three conditions: fish medium Control, Inactive, and Inactive+Stim.
    1. For Inactive and Inactive+Stim groups, anesthetize larvae at 72 hours post-fertilization (hpf) with Tricaine (0.6 mM).
      NOTE: Following reference38, frozen aliquots of tricaine stock are thawed and diluted (40 µL/mL fish medium, to a final concentration of 0.6 mM) before adding to fish. Do not add tricaine straight into the water containing fish, as some fish could receive high doses. Tricaine stock should be used within a month and never be re-frozen.
    2. For the fish medium Control fish, leave them un-anesthetized.
  3. At selected time(s) after the onset of tricaine exposure (i.e., at 80 hpf), prepare the Inactive+Stim group for stimulation.
    1. Prepare 60 mL of 2% agarose (1.2 g of agarose powder in 60 mL of fish medium), and melt fully using microwave, cool, add tricaine and pour 4 mL into each well of the stimulation chamber (Figure 1).
    2. Immediately add custom-made 4-well combs in between the electrodes (created by cutting out plastics (e.g., polypropylene) of desired dimensions and sticking together using Superglue; see Figure 1). Allow 10 min for gel to set. Remove combs carefully to create four rectangular wells.
    3. Fill each well with tricaine water and place a single anesthetized Inactive+Stim larva in each well using a micropipette, with their anteroposterior axis perpendicular to the electrodes (see Figure 1).
    4. Check under the dissecting fluorescent microscope whether each fish is fully anesthetized within each well of the chamber.
  4. Connect an adjustable electrophysiological pattern-generating stimulator to the chamber via a Polarity Controller, using crocodile clips connected to each of the electrodes on one side of the chamber (see Figure 1).
    NOTE: The polarity controller is used to reverse the polarity every 5 s, so as to prevent electrolysis and corrosion of the electrodes.
  5. Stimulate fish. For example, 1s with a train of 200, 20 V pulses, with 0.5 ms pulse duration and 4.5 ms pulse separation, once every 5 s gives an effective repeated-tetanic contraction resistance regime.
  6. Regularly check under the microscope to confirm the fish are being stimulated; the example electrical stimulus should induce a visible bilateral contraction and slight movement, once every 5 s.
  7. For a resistance/high force regime, stimulate the fish at a high frequency for a bout of 5 min, three times, with each bout separated by 5 min of rest.
    NOTE: While fish on one side of the chamber are resting, the crocodile clips can be connected to the electrode pair on the other side of the chamber, and those additional fish stimulated.
  8. After stimulation, carefully remove fish from each well by gently flushing them out with a plastic pipette and return to the incubator in fresh tricaine-containing fish medium.
  9. Pour away the tricaine water from within the chamber and use forceps to cut around and remove the agarose from each well. Rinse the wells with tap water and allow to dry.
    NOTE: If using silver wire electrodes, occasionally silver oxide may accumulate on the surface of the wire after a stimulation experiment. As silver oxide is less conductive than silver, to maintain reproducibility, carefully rub silver oxide off the wire using Kimwipes before re-using the setup.

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Representative Results

A rapid and precise measure of somite volume
A method of sample preparation, data acquisition, and volumetric analysis that allows the rapid measurement of muscle growth in zebrafish larvae is described. Muscle size can be measured in live animals using fish labeled on their plasma membranes with a membrane-targeted GFP (β-actin:HRAS-EGFP) or mCherry (α-actin:mCherry-CAAX). Larvae were transiently anesthetized using tricaine, mounted in low-melting-point agarose and imaged using confocal fluorescence microscopy. Somite 17 was chosen for analysis of muscle size given its accessibility at the trunk-tail interface31. In practice, as in theory, myotome volume can be calculated as the product of myotome length (L) and the average of three cross-sectional area measures (CSA) (Figure 2A, which is referred to as the 4-slice method).

To validate this method, whole myotome confocal Z-stacks of live zebrafish larvae (Figure 2B) were obtained, and somite volume was calculated by multiplying the sum of somite 17 profile areas in each slice (80-100 slices) by the inter-slice distance (1-1.2 µm). A strong correlation was observed between the 4-slice and full stack methods of calculation (Figure 2C). However, the 4-slice method generally gave a slightly larger volume estimate (~2% larger on average) (Figure 2D). This difference could be due either to a) obliquity of the samples giving erroneously large volume measurement or b) the observation that zebrafish myotomes taper along the anteroposterior axis, being smaller toward the tail of the animal. As the YZa, YZm, and YZp CSA measurements are toward the anterior portion of the somite 17 myotome chevron (Figure 2A), the latter interpretation was tested by volumetric analysis of the myotomes of somites 16 and 18. Each somite was about 7% larger than the one behind (Figure 2E). Further analysis revealed that a 2-slice method requiring only a single YZ measurement, located in the middle of the somite where the epaxial and hypaxial halves meet at the horizontal myoseptum (YZm), and the XY slice, gives a reasonably accurate estimate of myotome volume (Figure 2F,G). The 2-slice approach enables more rapid data acquisition when time constraints limit the number of fish that can be scanned. In summary, these data show that myotome volume can be measured rapidly and accurately in live zebrafish larvae. Single larvae were successfully, repeatedly, measured over a 6-day period with this method.

For comparative study of growth of an identified tissue unit, the described method provides reliable and precise volume estimates. To obtain accurate absolute volumes, however, a number of modifications can be applied. First, correction can be made for errors caused by oblique mounting either by tilting the dish or microscope stage to obtain better transverse YZ and parasagittal XY slices during imaging or through using the notochord profile in YZ slices; the cross-sectional minor axis reveals the true diameter of the cylindrical notochord, whereas its major axis reveals the angle and magnitude of obliquity (see Note in point 4.6 above). Second, the location of XY and YZ sectioning during scanning must be selected to reflect, accurately, the desired myotome(s). (See Note in point 4.6 above). Note, however, that the form of the myotome chevrons changes depending on developmental stage and this must be taken into account when selecting YZ scans. Lastly, to determine whether changes in myotome 17 reflect muscle growth throughout the axis, myotomes further into the trunk or tail regions may be measured by the described method.

Repeated measurements reveal somite growth
An advantage of the described method is the ease of repeated analysis on single fish. Individual embryos and larvae can be repeatedly embedded, measured, and released without suffering obvious long-term effects (Figure 2H). The myotome grows detectably both in L and CSA between 2 and 5 dpf, leading to steady increase in volume (Figure 2H). Growth was imaged in this way between 1 and 8 dpf and larvae were also released after imaging and grown to adulthood. Analyses further into the late larval period are expected to be possible, although effects of repeated imaging on feeding behavior would need to be monitored carefully on comparison with siblings that are not imaged. Importantly, development of pigmentation can obscure imaging. Whereas pigmentation is not a problem in properly orientated fish, as the melanophore stripes do not prevent the required measurements, pigment can be more problematic in obliquely mounted samples. The use of pigmentation mutant lines, such as roy;mitfa39, is anticipated, which would extend the time window of growth measurements until the limits of practicable confocal scan depth are reached.

A further advantage of the described procedure is the ease of detailed analysis of growth of single fibers in comparison with their whole myotome over short periods. By mosaic labeling of fibers through DNA injection, a method was developed to detect nuclear acquisition and growth in individual identified fibers over 4 h, and then permit re-analysing at later times (Figure 2I). Moreover, growth of the whole myotome can be measured over 12 h or less26 (and data not shown). By repeatedly measuring the same fish, inter-individual variability is eliminated and a small number of animals can yield statistically robust results26.

Manipulations which change somite volume can readily be detected
The current protocol allows one to interrogate changes in muscle growth under various biological and physiological conditions, such as altered physical inactivity. Anesthetic tricaine, which blocks nerve action potentials by inhibiting voltage-gated Na+ channels40, was used to induce muscle inactivity in β-actin:HRAS-EGFP larvae. As shown previously26, inactivity for 24 h between the third and fourth dpf greatly reduced myotome volume, indicating that larval muscle growth is activity-dependent (Figure 3A). The effect of inactivity can also be investigated using other detection methods which reveal the structure of the somite, such as mCherry-CAAX (either in a transgenic line or by mRNA injection) or by overnight immersion of larvae in BODIPY dye (Figure 3A). The latter approach, while removing the need to cross fish onto transgenic backgrounds or inject embryos cannot be used for repeated measurements due to toxicity of BODIPY. Thus, the current method allows one reproducibly to measure changes in the volume of muscle tissue.

As described above, genetic marking methods can be used to make repeated measurements of myotome volume, permitting tracking of change in muscle size over successive days in individual fish. As individual fish and entire lays at the same development stage differ in absolute myotome volume (Figure 3A; perhaps due the size or health of eggs), the ability to measure growth of each individual reduces the effects of individual variation by permitting paired sample statistical analyses. Repeatedly measuring the same fish reduces the number of fish needed to detect effects robustly. To illustrate this effect, results from analyzing the growth of each individual from 3 to 4 dpf in populations of active and inactive larvae were compared with analysis of the same two populations using only the single measure of myotome size made at 4 dpf. From lay to lay, larger variation in apparent reduction of myotome size was observed when measuring myotome volume at 4 dpf only compared to measuring 4/3 dpf volume for each individual (Figure 3B). Note the greater range of reduction (68%-91%) in the 4 dpf only measurements, compared to the 4/3 dpf method (78%-89%) and the weak correlation between the two measures. Although, as expected, the mean reduction across all 13 biological replicates was similar in each assessment (Figure 3C) being 82.62 ± 1.01% (mean ± SEM, n = 13) for the 4/3 dpf method and 82.31 ± 1.92% for the 4 dpf only method, the estimated error with the 4 dpf only method was almost double that with the 4/3 dpf method. Thus, quantifying individual growth through repeated measurement is the more accurate method, by eliminating size variability between fish within the same lay, as demonstrated previously26. Nonetheless, as no significant difference in the reduction in myotome volume caused by inactivity was observed when measuring myotome volume at 4 dpf only (Figure 3C), the data suggest that ~6-8 fish are sufficient to average out inter-individual size difference within a lay. Clearly, when size changes are small the 4/3 dpf method is preferred.

Analysis of the cellular basis of growth
Myotome CSA is determined by muscle fiber number and fiber size. Fiber number can be estimated by counting the number of fast and slow fibers on three YZ sections (YZa, YZm, YZp). Although regions of two adjacent myotomes are contained in such YZ sections, as shown by the presence of vertical myosepta (VM) in most sections (Figure 2A), these counts accurately reflect fiber number. Over-counting occurs at VMs due to the tapering of pairs of fibers from each adjacent segment at the VM (Figure 4A). Such overcounting can be accounted for using the following equation: Fiber number = Total fiber count - (fibers in contact with VM) / 2 reference33. Furthermore, average fiber volume can be determined by dividing myotome volume by the number of fibers. We have used these analyses to reveal that activity controls both cellular aspects of growth (Figure 4B,C).

It is evident from Figure 2A that fibers vary in size across the myotome, a reality that is not reflected in calculated average fiber volume measurements. By drawing around each fiber of somite 17 from two fish, measured fiber cross-sectional area was shown to range from 28 µm2 to 217 µm2 (Figure 4D). In reality, however, many fibers are angled obliquely within the myotome, so such CSA measures do not reflect the true CSA of a fiber perpendicular to its long axis. Conversely, due to the varying angles of fibers within the myotome, all fibers running at orientations that are not aligned anteroposteriorly have lengths that differ from the myotome length. Despite these caveats, which can only be circumvented either by complete segmentation of the myotome into single fiber volumes or by calculation after measuring the angle of obliquity of each fiber, the measured CSAs provide an estimate of fiber size diversity in each fish. For example, individual fibers decreased in CSA with inactivity, resulting in a shift to the left in the cumulative frequency curve with respect to active (un-anesthetized) control larvae (Figure 4E). As inactive fish have ~10 fewer fibers than active fish (Figure 4B), either the 10 smallest fibers from active un-anesthetized fish (on the assumption that they are the new ones) or the 10 largest (as the alternative extreme) were omitted from the comparison, which showed that most loss of the myotome volume is due to lack of fiber growth, rather than failure of new fiber formation (Figure 4F). Taken together, these data show that the described method allows detailed investigation of the role of physical activity on the formation and growth of muscle tissue.

Reimposition of activity by electrical stimulation
Physical activity is required for muscle growth (Figure 3A). A method to re-impose muscle contractions by electrical stimulation in otherwise inactive larvae, evoking a strong contractile response is described (Figure 5A, Supplemental File 8). Although precise stimulation parameters are described here (Figure 5B) to maximally activate the musculature, the protocol can be altered (by changing current amplitude, frequency, pulse duration, etc.) to control muscle activation and exercise dosage. Thus, the current method provides a controlled activity stimulus with standardized behavior between activity bouts, overcoming an important limitation of current animal models of exercise18.

The described methods demonstrate the potential of using zebrafish larvae to study various aspects of muscle growth (e.g., hyperplasia and hypertrophy). In particular, myogenesis in zebrafish larvae is shown to be amenable to analysis through pharmacologically-induced inactivity and electrically-induced contractility. The approach allows the study of the molecular mechanisms by which physical activity leads to muscle growth in vivo.

Figure 1
Figure 1: Design of a stimulation chamber for re-introduction of muscle electrical activity to zebrafish larvae. A 6-well cell culture plate fitted with electrodes permits maintenance of larvae in wells made within 2% agarose gel. Custom 4-well combs are made to create rectangle wells (dimensions as indicated) in agarose to maintain position and orientation of individual fish larvae, and to prevent them from contacting the silver wires during vigorous twitching upon electrical stimulation. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Measuring muscle volume in live zebrafish α-actin:mCherry-CAAX (A,B,E) or β-actin:HRAS-EGFP transgenic larvae (H,I). Larvae imaged from lateral view and shown dorsal to top, anterior to left in upper panels (A,B,E). (A) In the 4-slice method, myotome volume was calculated by multiplying the length of somite 17 (L) measured between vertical myosepta (VM) on an XY plane (blue arrow) by the average of three cross-sectional areas (CSA) measured on YZ planes (YZm, YZa, and YZp; dashed yellow lines). (B) In the Full Stack method, the area of somite 17 myotome was measured (examples outlined in yellow) in each slice of a whole Z-stack of a live zebrafish larva and the sum of myotomal areas was multiplied by the inter-slice distance. (C) High concordance between the 4-slice and Full Stack methods. Colors denote individual β-actin:HRAS-EGFP (squares) or α-actin:mCherry-CAAX (circles) fish. (D) Myotome volume is slightly, but significantly, larger when calculated using the 4-slice method. (E) In the tapering zebrafish larvae, more anterior somites are larger than posterior somites, as measured by the 4-slice method. (F,G) Strong correlation between volume measurements made using the average of three CSA sections (4-slice method) or a single YZm CSA (2-slice method). p-values show results of two tailed t-tests with equal variance (D,G) or one way ANOVA with Bonferroni post hoc tests (E). ns, not significant. (H) Measurement of myotome 16 volume, length (L) and cross-sectional area (CSA) from 2 to 5 dpf in three individual fish (colors) expressing β-actin:HRAS-EGFP and myog:H2B-mRFP. YZm images of the green individual at each timepoint are shown above. (I) Single fiber growth measured by automated constant-threshold segmentation. A β-actin:HRAS-EGFP;myog:H2B-mRFP fish mosaically labeled by injection of a CMV:Cerulean plasmid at the 1-2 cell stage. A single Cerulean marked fiber in somite 10 was scanned with high resolution full XYZ stacks repeatedly on a Zeiss LSM880. Images shown are representative single slices (left) and the projection of the three dimensional segmented volume (right). Each datapoint on the graph represents a single scan of the same fiber, at 3 dpf (0 h), after 4 h, and at 5 dpf (54 h). Triplicate scans were made and segmented per time-point to show reproducibility. White arrowheads point to fiber nuclei; note that two nuclei are added between 3 and 5 dpf. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Activity-dependent muscle growth in somite 17. (A) Inhibiting activity for 24 h by application of tricaine (pink) between 3 and 4 dpf reduces myotome volume both in transgenic lines and in non-transgenic fish stained with BODIPY compared to vehicle controls on siblings (blue). Volume quantified by 4-slice method. Symbol shape indicates replicate experiments from distinct lays (biological replicates). Large symbols denote mean ± SEM values. Small faint symbols show the volume of individual replicate larvae. (B) Comparison of myotome volume reduction in inactive fish compared to control siblings determined by single measurement of myotome volume at 4 dpf (4 dpf only, upper schematic) or change in myotome volume between 3 and 4 dpf (4/3 dpf, lower schematic). Each symbol represents the mean volume of myotome 17 in ~5 inactive fish from a single lay divided by the mean myotome 17 volume of ~5 active control siblings. (C) No difference was observed in the mean reduction in muscle growth in inactive fish, when measuring by the 4 dpf only or 4/3 dpf methods. Numbers within bars represent total number of fish analyzed. p-values show results of two way ANOVA with Bonferroni post hoc tests (A) or two tailed t-test with unequal variance (C). Please click here to view a larger version of this figure.

Figure 4
Figure 4: Cellular level changes in muscle growth caused by inactivity. (A) Schematic showing how overcounting occurs at VMs due to double counting of tapering fibers where the blue and red myotomes meet. Note that the average fiber count is 6 but the true mean value is 5.5. The corrected count gives a better approximation. (B,C) Fiber number (B) and average fiber volume (C) are reduced in inactive (pink) compared to active (bue) larvae. Symbol shape indicates replicate experiments from distinct lays (biological replicates). Large symbols denote mean ± SEM values. Smaller faint symbols show the value of individual replicate larvae. Numbers within bars represent total number of fish analyzed. (D) In single larvae, each fiber profile in myotome 17 was outlined and CSA determined. Boxes show mean ± SEM. p-values show results of two-way ANOVA with Bonferroni post hoc tests (B,C) or two tailed t-test with equal variance (D). (E,F) Cumulative frequency curves showing fiber size distribution in active control (blue) and inactive (pink) larvae. Comparison of all fibers (E), or after omission of presumed-nascent small or, at the alternative extreme, large fibers (F) shows that fiber size increase, not increase in fiber number, primarily accounts for activity-driven growth of the myotome. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Brief electrical stimulation evokes tetanic muscle contractions in anesthetised zebrafish larvae. (A) Sequential images (captured from video in Supplemental File 8) showing how electrical stimulation triggers maximal contractions of an anaesthetized larva at 3 dpf. Time scale in seconds. Red boxes indicate movements at the start of three successive 1 s stimulation trains. Each image is a 40 ms exposure. (B) Schematic showing the electrical stimulation regime, in which a 1 s train of 200 high frequency, 20 V electrical impulses is given every 5 s. Please click here to view a larger version of this figure.

Supplemental File 1: Schematics of captured images of perfectly (top left) and imperfectly mounted larvae with respect to the microscope XYZ reference frame (black axes). Myotome (green), notochord (yellow), neural tube (tan), measured myotomal parameters (red), measured notochord parameters (black arrows), and possible or actual fish rotations (blue). Please click here to download this File.

Supplemental File 2: Screenshot showing somite length measurement from XY images in ZEN. Please click here to download this File.

Supplemental File 3: Screenshot showing CSA measurement from YZ images in ZEN. Please click here to download this File.

Supplemental File 4: Screenshot showing somite length measurement from XY images in Fiji/ImageJ. Please click here to download this File.

Supplemental File: Screenshot showing extraction of calibration parameters for YZ images from ZEN. Please click here to download this File.

Supplemental File 6: Screenshot showing calibration of YZ images in Fiji/ImageJ. Please click here to download this File.

Supplemental File 7: Screenshot showing CSA measurement from YZ images in Fiji/ImageJ. Please click here to download this File.

Supplemental File 8: Representative video showing muscle contraction evoked by direct electrical stimulation (length 1 s) of a tricaine-anaesthetized 3 dpf larva. Please click here to download this File.

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Discussion

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 β-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 'slice' 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.

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Disclosures

The authors declare no competing interests.

Acknowledgments

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 from King's College London. This work benefited from the trigonometrical input of David M. Robinson, scholar, mentor, and friend.

Materials

Name Company Catalog Number Comments
Adhesive, Blu Tack Bostik - -
Aerosol vacuum  - - -
Agarose Sigma-Aldrich A9539 -
Agarose, low gelling temperature Sigma-Aldrich A9414 Once melted, keep at 37oC in a block heater to remain in liquid form for repeated use.
Block heater Cole-Parmer SBH130 -
BODIPY FL C5-ceramide Thermo Scientific D3521 To be diluted in fish water and used at 5 µM for overnight incubation.
Crocodile clips and wires - - -
Fiji/imageJ National Institutes of Health, NIH - -
Fish medium, Fish water - - Circulating system water collected from the fish facility.
Fish medium, E3 medium - - E3 is described in The Zebrafish Book. http://zfin.org (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, and 0.33 mM MgSO4 in distilled water).
Fluorescence microscope Leica Leica MZ16F Fluorescence microscope of other kind are also expected to be suitable.
Glass needle World Precision Instruments, Inc. 1B100-6 To be fire-polished to prevent damage of the embryos during manipulation.
Grass stimulator Grass Instruments S88 Stimulators of other kind are also expected to be suitable.
Kimwipes, Delicate Task Wipers Kimberly-Clark Professional 13258179 -
Laser scanning microscope (LSM)  Zeiss Zeiss LSM 5 Exciter
Zeiss LSM 880
LSM of other kind are also expected to be suitable.
Nunc Cell-Culture Treated, 6-well plate Thermo Scientific 140675 -
Objective, 20×/1.0W water immersion Zeiss - -
Pasteur Pipette, Graduated 1 mL Starlab Group E1414-0100 -
Pasteur Pipette, Micro Fine Tip 1 mL Starlab Group E1414-1100 -
Petri dish, 60 mm Sigma-Aldrich P5481 -
Plasmid, CMV-Cerulean Christina L. Hammond (University of Bristol) pCS2+_cerulean_kanR plasmid injected at 25-75 pg at one-cell stage.  Citation: Bussman J, and Schulte-Merker S. (2011) Development 138:4327-4332. doi: 10.1242/dev.068080.
Plasmid, pCS-mCherry-CAAX Henry Roehl (University of Sheffield) - For in vitro transcription using the SP6 promoter (plasmids containing other membrane labelling markers can be used);
synthesised capped mRNA to be injected at 100-200 pg at one-cell stage.
Pulse Controller  Hoefer Scientific Instruments PC750 -
Soldering iron - - -
Tricaine Sigma-Aldrich E10521 Ethyl 3-aminobenzoate methanesulfonate/ MS-222; to be dissolved in fish water and used at 0.6 mM.
Volocity Perkin Elmer/Quorum Technologies Inc - -
Watchmaker forceps, No. 5 - - -
Wire, Platinum Goodfellow PT005142/12 0.40 mm in diameter; an expensive alternative of silver.
Wire, Silver Acros Organics 317730010 0.25 mm in diameter (a range of diameter i.e. 0.25-0.5 mm had been tested, which produced similar results).
Zebrafish, myog:H2B-mRFP David M. Langenau (Massachusetts General Hospital; Harvard Stem Cell Institute) - ZFIN official name: Tg(myog:Hsa.HIST1H2BJ-mRFP), fb121Tg.  http://zfin.org/ZDB-ALT-160803-2  Citation: Tang Q, Moore JC, Ignatius MS, Tenente IM, Hayes MN, Garcia EG, Torres Yordán 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.
Zebrafish, α-actin:mCherry-CAAX Peter D. Currrie (ARMI, Monash University) - ZFIN official name: Tg(actc1b:mCherry-CAAX), pc22Tg.  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 träge 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.
Zebrafish, β-actin:HRAS-EGFP - - ZFIN official name: Tg(Ola.Actb:Hsa.HRAS-EGFP), vu119Tg. http://zfin.org/ZDB-ALT-061107-2  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.
ZEN software Zeiss - -

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Cite this Article

Attwaters, M., Kelu, J. J., Pipalia, T. G., Hughes, S. M. Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity. J. Vis. Exp. (184), e64063, doi:10.3791/64063 (2022).More

Attwaters, M., Kelu, J. J., Pipalia, T. G., Hughes, S. M. Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity. J. Vis. Exp. (184), e64063, doi:10.3791/64063 (2022).

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