Imaging Subcellular Structures in the Living Zebrafish Embryo

Summary

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

Abstract

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Introduction

In vivo imaging provides direct visualization of cellular behaviors in the most physiological context. The transparency of zebrafish embryos, their rapid and external development and a rich array of genetic tools that permit fluorescent labeling have all contributed to the growing use of in vivo microscopy to elucidate the dynamics of key developmental events. Imaging studies of nervous system development in zebrafish have for example greatly expanded our knowledge of the behavior of neural progenitor cells and the fate of their progeny including their subsequent migration, differentiation and circuit integration^1-8.

The stage is now set to investigate the subcellular dynamics underlying these cellular behaviors. Indeed, zebrafish are already being exploited as tools for in vivo cell biology. It is now possible to visualize mitochondria^9-11, centrosomes^2,8,12-14, Golgi¹⁵, the microtubule⁴ and actin¹⁶ cytoskeleton, endosomes¹⁷ and components of the apical membrane complex^1,18, among other subcellular structures in zebrafish embryos in vivo. So far, much of what is known about the function of these organelles comes from studying their behavior in cultured cells. While in vitro studies have yielded tremendous insight into cell biology, cells in culture do not fully represent the complexity of the in vivo situation and therefore do not necessarily reflect the function and dynamics of subcellular organelles in vivo. Zebrafish embryos offer a viable in vivo alternative to examining subcellular dynamics.

As vertebrates, zebrafish possess many organ systems (e.g., neural retina) that are homologous to those found in mammalian species. Additionally, zebrafish embryos are increasingly being used to model human diseases^19,20, including those related to centrosomal function (e.g., microcephaly²¹ and Leber's congenital amaurosis²²) and to mitochondrial function (e.g., Parkinson's disease²³, tauopathies^10,24 and Barth syndrome²⁵). In vivo imaging at the cellular and subcellular level in these instances will permit a better understanding of the cell biology underlying these pathological states.

The overall goal of the methods described here is to provide a comprehensive guide to investigate organelles and other subcellular structures in zebrafish embryos using in vivo light microscopy. The entire work-flow involved in visualizing and tracking subcellular structures in vivo is described - from genetic labeling approaches, to generating transiently expressing and stable transgenic fish, and finally to imaging using wide-field and confocal microscopy. While each of these procedures is used by numerous zebrafish laboratories, the protocols described are optimized and streamlined for investigating the dynamics of subcellular structures. Two specific aspects of the work described here warrant mention: First, the use of the Gal4-UAS expression system in multiple configurations to genetically label organelles in specific cell-types. Second, a direct comparison of wide-field and confocal microscopy to image subcellular structures in vivo.

Current strategies to genetically label organelles and other subcellular structures in zebrafish either make use of capped mRNA^1,4,8 or DNA based constructs where promoter elements directly drive the expression of fusion proteins^9,14,15. In vitro transcribed capped RNA results in rapid and broad expression, that is not tissue-specific however. Additionally, expression levels diminish over time as the capped RNA is diluted or degraded. Thus the use of RNA based constructs to examine organelle dynamics at later stages in development is limited (usually up to 3 days post-fertilization).

These limitations can be overcome by using DNA constructs, where spatial and temporal control of expression is determined by specific promoter elements. When DNA based constructs are used in the context of the Gal4-UAS system significant improvements to transgene expression levels are observed^26,27. In this bipartite expression system, cell-type specific promoter elements drive the expression of a transcriptional activator Gal4, while reporter genes are cloned downstream of the Gal4-binding upstream activating sequence (UAS). By combining UAS reporters with appropriate Gal4 drivers, expression can be restricted to specific cell-types, circumventing the need to clone reporter genes behind different promoters every time a specific expression pattern is desired. Furthermore, the expression of multiple UAS reporter genes can be driven by a single Gal4 activator. The Gal4-UAS system thus provides a versatile and flexible genetic approach for subcellular labeling.

Wide-field and confocal microscopes are the workhorses of most laboratories. Wide-field systems typically use an arc lamp as a light source and detect the emitted light with a sensitive camera that is placed at the end of the light path. This imaging modality is typically restricted to thin samples as out-of focus light obscures in-focus information in thicker samples. Confocal microscopes differ from wide-field systems in that they are built to favor signals that originate from the focal plane over those that originate out of focus (i.e., "optical sectioning")²⁸. To achieve optical sectioning a pinhole is placed in the emission path in a conjugate position to the point light source. Lasers are used as light sources and signals are detected with photomultiplier tubes (PMTs). Practically, a laser beam is swiped over the sample point-by-point and the fluorescence emission at each spot (pixel) is detected by the PMT.

Here we image the very same subcellular structures in living zebrafish embryos using both wide-field and confocal microscopy to provide a direct comparison of both microscopy modalities. The underlying aim of providing such comparisons is to offer guidelines for choosing the most appropriate microscopy technique for the specific question at hand.

Using the approaches described here we demonstrate Gal4-UAS based genetic labeling of mitochondria and centrosomes. These organelles are imaged in different cell-types of the nervous system and in muscle cells using wide-field and confocal microscopy to demonstrate the suitability of each imaging modality. The methods described here can easily be adapted for investigating other organelles and subcellular structures in the living zebrafish embryo.

Protocol

All animal experiments were performed in accordance with local regulations of the government of Upper Bavaria (Munich, Germany). 

1. Labeling Organelles and Other Subcellular Structures

NOTE: Here genetic reporter constructs that fluorescently tag centrosomes, mitochondria and cell membranes are described.

1. Use conventional cloning methods²⁹ to generate fusion proteins that fluorescently label centrosomes and mitochondria. Clone the coding sequence of zebrafish centrin4 (cetn4) in-frame with a fluorescent protein² (FP) such as Yellow fluorescent protein to generate cetn4-YFP. Clone the mitochondrial targeting sequence of subunit VIII of cytochrome c oxidase in-frame with an FP such as Cyan fluorescent protein to generate mitoCFP^9-11.
2. To restrict expression of the fusion proteins (cetn4-YFP or mitoCFP) to specific cells of the zebrafish embryo (e.g. neurons or muscle cells) use the bipartite Gal4-UAS expression system^26,27. First generate a UAS reporter construct. Use conventional methods to clone the fusion protein in a UAS expression vector, downstream of the Gal4-binding UAS cassette (variants range from 1x to 14x UAS, see Discussion)^26,27,30 and a minimal promoter (E1b basal promoter from the carp βactin gene), and generate UAS:cetn4-YFP and UAS:mitoCFP.
   NOTE: To prepare UAS reporter constructs for stable transgenic line generation additional elements need to be cloned (see 3 below)
3. To visualize the cellular context in which centrosomes or mitochondria are labeled, generate UAS reporter constructs in which cell membranes are labeled by FPs. Clone the first twenty amino acids of zebrafish Gap43 (containing palmitoylation sites) in-frame with an FP to target that FP to the cell membrane (memFP)^31,32.
4. Obtain driver constructs or transgenic lines in which cell-type specific promoter elements drive the expression of the transcriptional activator Gal4-VP16²⁷, Gal4FF³³ or KalTA4³⁰.
5. Combine the UAS reporter constructs with an appropriate driver construct to restrict expression to the desired cell-type of interest (e.g., in neurons or muscle cells). To do this co-inject Gal4 driver and UAS reporter constructs at the one-cell stage of fertilized eggs (for detailed instructions see 2 below) to generate transiently expressing fish. Alternatively, generate a UAS reporter stable transgenic line (for detailed instructions see 3 below) and cross to a Gal4 driver stable transgenic line to generate offspring bearing both transgenes.
   NOTE: It is also possible to inject UAS reporter construct/s into fertilized eggs from a Gal4 driver stable transgenic line or a Gal4 driver construct into fertilized eggs from a UAS reporter stable transgenic line.
6. To co-express multiple distinct fusion proteins using the Gal4-UAS system, use a Gal4 driver and UAS reporter/s in one of the following configurations: A. Multiple, separate UAS reporter constructs (e.g., UAS:mitoCFP and UAS:memYFP co-injected with a Gal4 driver construct)¹⁰, B. Multiple UAS cassettes on a single reporter (e.g., UAS:cetn4-YFP, UAS:memCerulean)¹² C. Bi-directional UAS reporter (e.g. UAS:memYFP,mitoCFP)^2,10,24 (Figure 1).

Figure 1. Strategies to co-express fusion proteins using the Gal4-UAS system.
Co-expression of multiple fusion proteins can be achieved by using UAS reporter cassettes in various configurations: (A) multiple, separate UAS-driven constructs, (B) multiple UAS cassettes on a single construct or (C) a bidirectional UAS cassette on a single construct. Please click here to view a larger version of this figure.

2. Generate Transiently Expressing Fish

NOTE: The following is an adaptation of previously published protocols^34,35. A basic injection setup should include a stereomicroscope with a magnification range up to 12x, a micromanipulator with a micropipette holder and a source of air pressure. Prepare 2.1-2.5 ahead of time:

1. To prepare egg microinjection chambers, prepare a solution of 1.5 % (w/v) agarose in water. Add agarose powder to water and microwave until the agarose is completely dissolved. Fill a petri dish (100 x 15 mm) with this solution.
   1. Allow the agarose solution to cool to approximately 45 ^oC before placing a plastic microinjection mold (40 mm x 66 mm, with 6x 50 mm long ridges that are 1.5 mm wide, 1 mm deep and spaced 3 mm apart) on the surface of the molten agarose, taking care not to introduce bubbles at the interface.
   2. After the agarose sets, store at 4 ^oC O/N. Carefully remove the mold using a spatula. Prepare multiple microinjection chambers at a time, store at 4 ^oC and use repeatedly over several weeks.
2. Use a micropipette puller to generate glass injection capillaries with a long shank³⁶.
   NOTE: The specific settings used to generate injection capillaries should be determined empirically. Injection capillaries can be pulled ahead of time and stored prior to use.
3. Measure the concentration of plasmid DNA (Gal4 driver and UAS reporter constructs) for microinjection using a spectrophotometer according to the manufacturer's manual. Measure the ratio of absorbance at 260 nm and 280 nm (A260/280). Dilute the DNA to a final concentration of 100 ng/µl in nuclease-free water.
   NOTE: A A260/280 ratio of ~1.8 signifies pure DNA. Consider using a commercial kit (based on silica-based binding matrix) to purify the plasmid DNA to be used for injection. Plasmid DNA should be of high quality to prevent toxicity.
4. Prepare 10 µl injection-mix by combining DNA (0.5 to 2.5 µl of a 100 ng/µl stock for a final concentration of 5 to 20 ng/μl), Danieau's solution (0.33 µl of 30x stock), Phenol Red (0.25 µl, optional) and nuclease-free water to a 10 μl total volume.
   NOTE: Empirically determine the concentration of DNA that yields suitable expression. Gal4 driver and UAS reporter plasmids need to be co-injected. No expression will ensue when only the Gal4 driver or UAS reporter plasmid is injected into wild-type fertilized eggs. If however the fertilized eggs are from a Gal4 driver line then injection of one or several UAS reporter plasmids suffices. Conversely, if microinjecting into a UAS reporter line, only the Gal4 driver plasmid needs to be injected. Finally, while Phenol Red can greatly help to visualize injections, it is also a fluorescent compound itself. Therefore, Phenol Red could obscure visualizing FPs if imaging is planned before 24 hr post-fertilization (hpf) as it may not be sufficiently diluted by this time.
5. The evening before injections are planned, set up several pairs (typically 5 to 10) of male and female zebrafish for breeding. Use breeding tanks with an insert containing a gridded bottom and a removable divider to separate the male and female fish.
   NOTE: Male and female zebrafish can generally be distinguished by their body shape. Males tend to be slender while females tend to exhibit a ventrally protruding abdomen. Males additionally tend to have a yellowish-red coloring on their ventral abdominal area.
6. Immediately prior to DNA injections, remove the dividers to allow the male and female to mate. Approximately 15 to 30 min after removing the divider, check for eggs at the bottom of the tank. Transfer the adult fish using a fishing-net to a new breeding tank.
7. Pour the water, containing the newly-laid fertilized eggs, out of the breeding tank into a sieve (e.g., a plastic tea-strainer). Wash the eggs in the sieve using a spray bottle containing Danieau's solution. Invert the sieve onto a petri dish and use a spray bottle with Danieau's solution to recover all the eggs (typically 50 to 200 eggs per breeding pair of adult fish).
8. Using a microloader pipette, back-fill a pulled injection capillary with the DNA injection-mix. Mount the injection capillary into the holder of a micromanipulator and trim the tip, under the visual control of a stereomicroscope, using forceps to create a micropipette that can penetrate the chorion and cell cytoplasm while being able to deliver a suitable volume of the DNA.
   NOTE: The degree to which the tip of the injection capillary is trimmed should be determined empirically and can best be judged when injecting eggs.
9. To determine the volume of DNA for injection, discharge a drop of the injection solution into mineral oil. Measure the radius (r) of the drop using a calibration micrometer and calculate its volume (4/3 π r³). Modulate the volume by adjusting the injection pressure and/or the duration of each injection pulse.
10. Using a plastic pipette, transfer all the collected eggs (typically 50-200, from 2.7 above) into the trenches of a microinjection chamber. Under the visual control of a stereomicroscope, use forceps to orient the eggs so that the cell cytoplasm (transparent) and yolk (dense, non-transparent) can be readily identified.
11. Under the visual control of a stereomicroscope, inject DNA into the cell cytoplasm of the fertilized egg, taking care to ensure that the injected volume (1 to 2 nl) corresponds to approximately 10% of the cell's volume.
    NOTE: Phenol red in the DNA solution aids in the visualization of the volume injected.
12. After injection, loosen the eggs from the trenches of the injection mold by flushing them with a spray bottle containing Danieau's solution. Subsequently transfer eggs into a fresh petri dish using a plastic transfer pipette and maintain in a 28.5 ^oC incubator.
13. Maintain un-injected eggs as controls to determine whether injections contribute to high mortality rates, either as a result of physical damage sustained during micropipette penetration or the DNA mix.
    NOTE: High mortality rates following injections could be due to numerous factors, including excessively high DNA concentration or injection volumes and/or plasmids with contaminants. To prevent toxicity from low quality DNA preps, commercial kits (based on silica-based binding matrix) can be used.
14. At regular intervals following injections, use a stereomicroscope to screen for unfertilized eggs and dead or malformed embryos and discard these.
    NOTE: Deem the eggs that appear to be at the one-cell stage several hours after fertilization, as unfertilized. Identify malformed embryos by gross anatomical defects such as a compromised anterior-posterior body axes. Amorphous material within a chorion suggests that the embryo did not proceed through developmental stages and died. To ascertain if embryos that were microinjected undergo a normal course of development consult anatomical atlases³⁷.
15. Transfer the embryos using a plastic transfer pipette to Danieau's solution containing 1x 1-phenyl 2-thiourea (1xPTU) between 10 and 24 hpf to prevent pigment formation. Maintain the embryos in Danieau's solution containing PTU for the duration of the experiment.
    NOTE: In addition to melanophores, iridophores on the surface of the skin can be problematic for imaging. While PTU does not inhibit iridophore formation, mutant lines with reduced numbers of iridophores such as roy orbison (roy) exist and can be used³⁸.
16. After embryos hatch (usually on the second or third day post-fertilization, 2 or 3 dpf), discard the chorions and exchange the Danieau's solution containing PTU.
    NOTE: These steps are important to ensure the quality of the embryo medium and the viability of the healthy embryos.

3. Generate Stable Transgenic Lines

NOTE: Stable transgenic lines can be efficiently generated using the Tol2 transposon system. A Tol2 transposon vector containing the transgene of interest is co-injected along with mRNA encoding the transposase enzyme into fertilized eggs at the one-cell stage. Transposase protein derived from the injected mRNA catalyzes the excision of the transgene cassette from the transposon vector and its integration into the genome³⁹.

1. Clone the transgene reporter cassette (e.g., UAS:mitoCFP) between Tol2 inverted terminal repeats in one of the Tol2 vectors currently used in the zebrafish community as described⁴⁰.
   NOTE: Tol2 vectors containing a selectable cassette in which promoter elements drive FP expression in organ systems unrelated to the area of interest (e.g., heart⁴¹ or lens⁴²) are useful for screening UAS reporter transgenic lines in the absence of Gal4 transactivation.
2. Using a commercial kit (based on silica-based binding matrix), purify the plasmid DNA to be used for injection. Ensure that the plasmid DNA is of high quality to prevent toxicity.
3. Transcribe Tol2 transposase-encoding mRNA using an appropriate transcription vector such as PCS-TP⁴³. Briefly, linearize PCS-TP and use a commercial kit to generate capped mRNA and follow the manufacturer's protocol. Take care to avoid contamination with RNases (e.g., use RNAse-free pipette tips and tubes). Aliquot the RNA, at a concentration of 100 ng/µl, for single-use and store at -80 ^oC.
4. On the morning of the injections, thaw an aliquot of transposase mRNA on ice. Mix the transposon DNA vector (2 µl of 100 ng/µl) and transposase mRNA (2 µl of 100 ng/µl) in a 1:1 ratio, and add 6 µl of RNAse-free water to bring the total volume to 10µl.
   NOTE: A concentration of 20 ng/µl is recommended for each but the optimal concentration should be determined empirically. Use RNAse-free water for dilution. Use gloves when handling the DNA-RNA injection mix and keep it on ice for the entire period of injections to prevent degradation of the mRNA.
5. Using the detailed instructions in section 2 above, inject the DNA-RNA injection mix into fertilized eggs at the one-cell stage. Under visual control of a stereomicroscope target the cell cytoplasm at the one-cell stage for the highest rates of transgenesis efficiency. Maintain the injected embryos at 28.5 ^oC until ready to screen for transgene expression.
   NOTE: PCR can be done to check for efficiency of the Tol2 transposase as previously described⁴⁴.
6. Screen embryos in a petri dish for transgenesis using the oculars of a fluorescence dissecting microscope.
   NOTE: Use the appropriate filter sets of the microscope to visualize FP expression of the selectable cassette (i.e., fluorescence in the heart or lens) at 2 or 3 dpf. Identify potential transgenic embryos by expression (in heart or lens) and raise these embryos to adulthood (F₀ generation) using standard protocols⁴⁵. Alternatively, identify potential transgenic embryos by PCR as described in ⁴⁶.
7. Once the UAS reporter F₀ fish are 2.5 - 3 months of age, cross them (see 2.5 for details) to non-transgenic wild-type fish to acquire eggs to establish an F₁ generation. Alternatively, cross the UAS reporter F₀ fish to a Gal4 driver line to establish an F₁ generation. In the latter case, the UAS-driven transgene can be directly monitored in the embryos obtained from the cross.
8. Use a fluorescence dissecting microscope to screen F₁ embryos for expression of the transgene or selectable cassette.
   NOTE: When expression is confirmed then the transgene can be said to be germline transmitted. Transgenesis is non-Mendelian at the F₀ stage. The number of embryos expressing the transgene may vary from a small number to the vast majority in individual clutches. Transposon integration in different F₀ fish may vary greatly, resulting in different expression patterns. Therefore maintain F₁ fish from different F₀ founders as separate sub-lines. Transposon integrations in F₁ fish are stable and are passed onto subsequent generations in a Mendelian fashion.
9. Maintain each F₀ fish in separate tanks to re-identify transgene carriers.

4. Prepare Embryos for Imaging on an Upright Microscope

NOTE: The procedures described here have been optimized for imaging on upright microscopes with long-working distance water-dipping-cone objectives.

1. Use a fluorescence dissecting microscope to screen embryos for transgene expression.
   NOTE: Expression of a UAS reporter transgene will depend on the specific Gal4 driver line used. Select embryos for imaging experiments based on desired expression pattern.
2. Transfer the selected embryos using a plastic pipette to a separate petri dish containing Danieau's buffer with 1x PTU and 1x Tricaine.
   NOTE: When labeling is sparse (only a few cells in an organ system) mount the embryos in agarose (see 4.3-4.7 below) and screen for transgene expression using a wide-field microscope with high magnification objectives (see 5.1 below). Tricaine anesthetizes the fish and should be effective within seconds. If fish are not immobilized it is likely that the Tricaine has degraded. Possible reasons for this degradation include poor storage of Tricaine, which is light-sensitive⁴⁷.
3. Prepare the agarose to embed the fish by dissolving low-melting agarose powder in Danieau's buffer to a final concentration of 0.7%. Aliquot (1 ml) and keep in a heat-block at 40 °C until ready to use.
   NOTE: Higher concentrations of agarose (upto 1.5%) can also be used to embed fish.
4. Add 50 μl of a 20x stock of Tricaine and 20 μl of a 50x stock solution of PTU to the 1 ml aliquot of low-melting agarose and mix well by tapping the side of the tube. Return the tube to the heat-block. Using a plastic transfer pipette, gently pipette a few (1-10) anesthetized embryos into the agarose, transferring as little liquid as possible to avoid diluting the agarose.
5. Using a plastic pipette transfer all the embryos with a small amount of agarose to a glass bottom petri dish.
6. Working relatively fast, use forceps to position the embryos in the desired orientation, depending on the structure to be imaged.
   NOTE: Orient embryos on their side if the retina or Rohon-Beard (RB) sensory neurons should be imaged.
7. Allow the agarose to solidify for at least 15 mins. Add Danieau's buffer containing 1xPTU and 1xTricaine to cover the embryos embedded in agarose. Place the dish containing agarose-embedded embryos in an incubator at 28.5 ^oC until ready to image.

5. Imaging Cellular and Subcellular Structures using Wide-field or Confocal Microscopy

NOTE: Wide-field microscopy and point-scanning confocal microscopy are the most widely used modalities to image fluorescently labeled zebrafish embryos. Table 1 summarizes the main advantages and disadvantages of both systems. For both forms of microscopy, embryos are mounted in agarose as described in 4 above, and maintained at 28.5 ^oC throughout the imaging experiment by using a heating chamber on the stage of the microscope. Importantly, the dish with agarose-embedded embryos is allowed to equilibrate to 28.5 ^oC before imaging commences as temperature fluctuations cause drift in the z-dimension. When conducting long-term imaging experiments (over several hours), place a Plexiglas cover over the petri-dish to reduce evaporation of the buffer.

	Wide-field	Point-scanning confocal
Cost	Relatively inexpensive	Expensive (approx. 5x more)
Photo-bleaching	Low	High. The sample is exposed to laser light above and below the focal plane.
Photo-toxicity	Low	High (see Photo-bleaching above)
Acquisition speed	Fast	Slow
Resolution in xy	Abbe’s law	Abbe’s law (can be improved by app. 40% using a very small pinhole; however, in most settings this has little practical application)
Optical sectioning	Poor	Yes (can be adjusted by pinhole size)
Tissue penetration	Limited to superficial structures (e.g., Rohon-Beard cells or muscle fibers)	Limited to < 100 mm from the surface of the embryo

Table 1. General comparison of wide-field and point-scanning confocal microscopy

1. Wide-field microscopy
   NOTE: Here guidelines are provided for imaging zebrafish embryos using an upright wide-field microscope with long-working distance water-dipping-cone objectives. The microscope is equipped with a cooled CCD camera, and filter sets for visualizing different fluorophores mounted on an automated filter wheel for rapid acquisition of multiple channels. Image acquisition is controlled by μManager, an open source microscopy software package⁴⁸. A specific example for imaging mitochondria in RB sensory neurons is provided. RB cells are genetically labeled using membrane targeted YFP and their mitochondria are CFP-tagged (see 1.1 above).
   1. Mount embryos on their side in low-melting agarose as described in 4 above.
   2. Allow the dish with mounted fish to equilibrate to 28.5 ^oC in a heating chamber on the stage of the microscope before commencing imaging.
   3. Use a low magnification water-dipping-cone objective and look through the oculars of the microscope to choose an area on the surface of the embryo to image.
      NOTE: If the stable transgenic MitoFish reporter line, Tg(UAS-E1b:mYFP,mitoCFP)^mde6, is used in conjunction with a driver line such as HuC:Gal4 then most RB neurons are labeled, and appear as a dense mesh-work of neurite arbors on the surface of the embryo. If microinjection of DNA constructs is used to achieve sparse labeling (e.g., Sensory:Gal4-VP16 with UAS:mitoCFP and UAS:MA-YFP), screen embryos to identify double-labeled cells.
   4. Open the microscope software (μManager 1.4) and click on "Illumination'' under "Configuration settings" to define the correct filter sets for the wavelength of interest (YFP or CFP for the current example). Under "Camera settings" enter the exposure time required to acquire a suitable image.
      NOTE: The "Illumination Settings" is pre-set by the user upon installation of the software and is loaded when opening μManager. If the software is not configured to control the filter wheel, manually move the filter wheel in the turret to the appropriate position.
   5. Change to a long-working distance water-dipping-cone objective ranging from 40-100x magnification. Choose the objective that has the highest numerical aperture (NA) and is chromatically corrected (apochromat).
   6. Click on "Live" to choose the field of view: In the case of the RB neuron, image mitochondrial transport at the stem axon, emanating from the cell body, or in the peripheral arbor.
      NOTE: The peripheral arbors of RB neurons in the caudal fin fold of the embryo provide an ideal opportunity to image a large field of view that is flat and can therefore be captured on a single frame with a wide-field microscope. It is therefore important to mount the embryo as flatly as possible on its side, so that the caudal fin fold is parallel to the bottom of the petri dish.
   7. After choosing a field of view, click on the "rectangular selection" button in the ImageJ menu, define a region of interest (ROI) and click on "ROI" in the µManager window. After selecting the ROI for imaging, click "Stop" and "Save" to take an image of the YFP channel to record the local morphology of the neurite/s.
   8. To image mitochondrial transport click on the "Multi-D Acq." button. Observe an additional window ("Multi-dimensional Acquisition") and select the number of time-points as well as the interval between time-points on this window. Use frame rates of 0.3-1 Hz. Carry out recordings for at least 10 min to collect as many data-points as possible.
   9. In the "Multi-dimensional Acquisition" window, click on "Channels", add and define the wavelength for imaging (CFP for mitochondria in this example) and the time of exposure. Keep exposure times to below 400 msec for imaging mitochondrial transport.
   10. Click on the option "Save Images" in the "Multi-dimensional Acquisition" window, to automatically save the files in a specific folder outlined in the 'Directory root'. Click on "Acquire" at the upper right corner to start time-lapse imaging.
   11. Manually readjust the focus while time-lapse recording to compensate for any drift in the z dimension.
       NOTE: Using these parameters mitochondrial transport in RB neurons can be imaged for as long as 4 hr.
2. Confocal microscopy
   NOTE: Here guidelines are provided for imaging with an upright Olympus FV1000 confocal microscope and long-working distance water-dipping-cone objectives. The microscope is equipped with multiple laser lines and several detectors (conventional PMTs and more sensitive gallium arsenide phosphide detectors) that allow for multichannel imaging. Specific reference is made to Olympus Fluoview software. However, the imaging parameters described here should be easily transferrable to other confocal systems.
   1. Mount embryos in low-melting agarose in an orientation appropriate for the organ/structure to be imaged, as described in 4 above.
   2. Allow the dish with mounted fish to equilibrate to 28.5 ^oC in a heating chamber on the stage of the microscope before commencing imaging.
   3. Use long-working distance water-dipping-cone objectives for confocal microscopes in an upright configuration. Choose objectives with the highest possible NA to maximize the amount of fluorescent signals that can be collected and have the best resolving power. When collecting images from multiple channels choose chromatically corrected objectives (apochromat).
   4. Open the microscope software and click on "Trans Lamp" or "Epi Lamp" to use either transmitted or fluorescence light respectively to identify the region of interest via the oculars of the microscope.
   5. Use the software to set up the following scanning parameters to acquire image stacks:
      1. In the "Acquisition Setting" window verify that the objective chosen for imaging matches the objective that appears on the drop-down menu of available objectives.
         NOTE: This is to ensure that any pre-calculated parameters for a particular objective (e.g., lateral and axial resolution, size of the confocal aperture etc.) hold true and can be reliably used.
      2. Select the appropriate laser line/s, and excitation and emission dichroic filters to image specific fluorescent protein(s). Do this by clicking on the "Dye list" button in the "Image Acquisition Control" window and choose the appropriate fluorophore/s. Alternatively, click on the "Light path and dyes" button to set these parameters manually.
         NOTE: When the "Dye List" button option is chosen, the software automatically selects the appropriate laser lines, excitation and emission dichroic filters and adjusts the size of the confocal aperture appropriately.
      3. In the "Acquisition Setting" window, adjust the "Zoom" factor and "Size aspect ratio" (i.e., 512x512, 1024x1024 etc.) of the scanned image to obtain the pixel size necessary to best resolve the structures being imaged. Set the pixel size to be about half the theoretical resolution of the objective, thereby following Nyquist sampling criteria. To determine the pixel size of the acquired image click on the button with the symbol for information ("i") in the "Image Acquisition Control" window.
      4. Set the scan speed to the fastest possible (2 μs/pixel) in the "Acquisition Setting" window.
      5. In the "Image Acquisition Control" window click on Kalman line averaging to reduce noise.A factor of 2 to 3 usually suffices.
      6. Select a sequential scanning mode when imaging fluorophores with overlapping spectra. To do this, click on "Sequential" and "Line" in the "Image Acquisition Control" window.
      7. Adjust the power output of the relevant laser line/s (typically below 5%). The specific values need to be determined empirically.
      8. Click on "XY Repeat" or the "Focus x2" or "Focus x4" button to continuously scan the selected region while adjusting the detector settings for each channel. Adjust "HV", "Gain" and "Offset" for each channel to acquire images that have the highest dynamic range of grey values.
         NOTE: The specific values for these settings need to be determined empirically. "HV" adjusts the voltage on the detector to change its sensitivity, "Offset" adjusts the output signal of the detector and "Gain' multiplies the output signal of the detector by a constant factor.
      9. Press Ctrl + H to visualize the acquired images via a look-up table (HiLo) that identifies under-saturated (appear blue) and over-saturated pixels (appear red), both of which should generally be avoided. Re-evaluate the power output of the relevant laser line/s and the detector settings.
         NOTE: Using high laser power and high levels of the "HV" of the detector will lead to more signal. Higher laser power will however lead to increased bleaching and photo-toxicity. Increasing the "HV" will lead to increased noise. Therefore, a compromise needs to be found when setting laser power and "HV" to acquire images with acceptable levels of noise while preventing photo-damage (see also Table 1).
      10. Collect images from a defined volume by focusing on the upper and lower limits of the area of interest. In the "Acquisition Setting" window click on the "End Set" and "Start Set" set buttons of the z-stack window at the upper and lower limits of the volume to be imaged. Select a step size that is half of the z-resolution for the given objective (e.g., if z resolution is 2 µm choose 1 µm). To determine the z-resolution of the objective click on the button with the symbol for information ("i") in the "Image Acquisition Control" window.
      11. Set up a time series to collect z-stacks at a temporal frequency that is appropriate for the dynamics of the subcellular structure being imaged. In the "TimeScan" sub-window enter the frequency at which z-stacks should be acquired ("Interval"') and the number of times the images should be acquired ("Num").
      12. Click on the "Depth" and "Time" buttons in the "Image Acquisition Control" window to confirm that a z-stack and time-series will be acquired. Finally click on the "XYZT" button to begin acquiring time-lapse images.
      13. After completion of image acquisition, observe "Series Done" on the software interface. Click on it and save the images in "oib" format to record the images and metadata associated with them.
          NOTE: Images obtained on the wide-field microscope and confocal microscope can be visualized and analyzed using software such as FIJI, a free public domain image processing program.

Results

Here the use of wide-field and confocal microscopy to image mitochondria and centrosomes in living zebrafish embryos is directly compared and contrasted. Depending on the location of the cells in which organelle dynamics are to be examined and the inherent frequency of the specific subcellular events, generally either wide-field or confocal microscopy is the better choice. We imaged organelles in RB neurons located on the surface of the embryo and in retinal cells located deeper. The supe...

Discussion

Here, we demonstrate the versatility of the Gal4-UAS expression system to fluorescently tag mitochondria, centrosomes and the cellular membranes of specific cell-types in vivo in zebrafish embryos. Many fluorescent fusion proteins that label other organelles or subcellular structures can be found in the published literature and can be obtained from the respective laboratory, commercial sources or non-commercial plasmid depositories (e.g., Addgene). To design a new fluorescent fusion protein, several par...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Agarose (2-hydroxyethylagarose)	Sigma-Aldrich	A4018-10G	Low-gelling temperature Type VII
Block heater	Eppendorf		Thermomixer compact
Ca(NO3)2 Calcium nitrate hydrate, 99.996%	Aldrich	202967-50g	To prepare 30x Danieau's
CCD camera	Qimaging	Retiga Exi Fast 1394
Ceramic Coated Dumont #5 Forceps	Dumont - Fine Science Tools	11252-50	#5 Forceps
Confocal laser-scanning microscope	Olympus	FV1000 Fluoview
Culture dish heater	Warner Instrument Corporation	DH-35	Heating ring
Ethyl 3-aminobenzoate methanesulfate salt	PharmaQ	Tricaine PharmaQ-25g	Tricaine (anesthetic)
Fluorescence dissecting microscope	Leica	M205 FA
GeneClean kit	MP Biomedicals	111001200
Glass Bottom Culture Dishes	MatTek Corporation	P35G-0-14-C	35 mm petri dish, 14 mm microwell, No. 0 coverglass
Glass needles	World Precision Instruments Inc.	TW100F-4	For microinjections
HEPES	Sigma	H3375-250g	To prepare 30x Danieau's
High vacuum grease	Dow Corning	DCC000001242 150g	Silicon dioxide grease
KCl 99%	Sigma-Aldrich	S7643-5kg	To prepare 30x Danieau's
MgSO4.7H2O   Magnesium sulfate heptahydrate 98+% A.C.S reagent	Sigma-Aldrich	230291-500g	To prepare 30x Danieau's
Microinjector	Eppendorf		FemtoJet
Microloader tips	Eppendorf	930001007	0.5-20 ul
Micromanipulator	Maerzhaeuser Wetzlar	MM33 Rechts/00-42-101-0000/M3301R
Micropipette holder	Intracel	P/N 50-00XX-130-1
mMESSAGE mMACHINE SP6 Transcription Kit	Ambion	AM1340	To transcribe PCS-Transposase
NaCl BioXtra >99.5%	Sigma-Aldrich	P9541-1kg	To prepare 30x Danieau's
Nanophotometer			To measure DNA/RNA concentration
Needle puller	Sutter Instrument	P-1000	Flaming/Brown
NIR Apo 40x/0.80W	Nikon		Water-dipping-cone objective
N-Phenylthiourea Grade I, approx. 98%	Sigma	P7629-10G	PTU (prevents pigmentation)
Petri dishes	Sarstedt AG	821472	92 x 16mm
Plastic molds	Adaptive Science Tools	TU-1	For microinjections
Plexiglas cover-with a hole	Custom-made		The hole in the Plexiglas cover should be 3 mm larger than the diameter of the water-dipping-cone objective
Tea-strainer (Plastic)			To collect zebrafish eggs
Temperature controller	Warner Instrument Corporation	TC-344B	Dual Automatic Temperature Controller
Transfer pipettes	Sarstedt AG	86.1171	3.5mL plastic transfer pipettes
UMPlanFI 100x/1.00W	Olympus		Water-dipping-cone objective
UMPlanFLN 20x/0.50W	Olympus		Water-dipping-cone objective
Widefield microscope	Olympus	BX51WI
PTU (50x Stock)			Dissolve 76 mg PTU in 50 ml distilled water Stir vigorously at room temperature  Store at -20 oC in 1 ml aliquots Use at 1x working solution
Tricaine (20x Stock)			Dissolve 200 mg Tricaine in 48 ml distilled water Add 2 ml 1M Tris base (pH9) Adjust to pH 7  Store at -20 oC in 1 ml aliquots Use at 1x working solution
Danieau's Solution (30x Stock)			1,740 mM  NaCl  21 mM      KCl  12 mM      MgSO4.7H2O  18 mM      Ca (NO3)2 150 mM    HEPES buffer  Distilled water upto 1 L  Store at 4 oC  Use at 0.3x working solution