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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

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.

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.

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...

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...

Disclosures

The authors declare no competing interests.

Acknowledgements

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

NameCompanyCatalog NumberComments
Adhesive, Blu TackBostik--
Aerosol vacuum ---
AgaroseSigma-AldrichA9539-
Agarose, low gelling temperatureSigma-AldrichA9414Once melted, keep at 37oC in a block heater to remain in liquid form for repeated use.
Block heaterCole-ParmerSBH130-
BODIPY FL C5-ceramideThermo ScientificD3521To be diluted in fish water and used at 5 µM for overnight incubation.
Crocodile clips and wires---
Fiji/imageJNational 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 microscopeLeicaLeica MZ16FFluorescence microscope of other kind are also expected to be suitable.
Glass needleWorld Precision Instruments, Inc.1B100-6To be fire-polished to prevent damage of the embryos during manipulation.
Grass stimulatorGrass InstrumentsS88Stimulators of other kind are also expected to be suitable.
Kimwipes, Delicate Task WipersKimberly-Clark Professional13258179-
Laser scanning microscope (LSM) ZeissZeiss LSM 5 Exciter
Zeiss LSM 880		LSM of other kind are also expected to be suitable.
Nunc Cell-Culture Treated, 6-well plateThermo Scientific140675-
Objective, 20×/1.0W water immersionZeiss--
Pasteur Pipette, Graduated 1 mLStarlab GroupE1414-0100-
Pasteur Pipette, Micro Fine Tip 1 mLStarlab GroupE1414-1100-
Petri dish, 60 mmSigma-AldrichP5481-
Plasmid, CMV-CeruleanChristina 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-CAAXHenry 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 InstrumentsPC750-
Soldering iron---
TricaineSigma-AldrichE10521Ethyl 3-aminobenzoate methanesulfonate/ MS-222; to be dissolved in fish water and used at 0.6 mM.
VolocityPerkin Elmer/Quorum Technologies Inc--
Watchmaker forceps, No. 5---
Wire, PlatinumGoodfellowPT005142/120.40 mm in diameter; an expensive alternative of silver.
Wire, SilverAcros Organics3177300100.25 mm in diameter (a range of diameter i.e. 0.25-0.5 mm had been tested, which produced similar results).
Zebrafish, myog:H2B-mRFPDavid 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-CAAXPeter 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 softwareZeiss--

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