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In This Article

Summary

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Abstract

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

Introduction

In the early phases of development, pools of specialized progenitor cells divide repeatedly in proliferative zones, producing diverse arrays of daughter cells. The cells born during this developmental period will then differentiate and travel to form the nascent organs. In the nervous system, progenitors such as radial glia give rise to immature neurons in ventricular zones. As neurons migrate away from ventricles and mature, the expanding tissue eventually forms the highly complex structures of the brain1,2,3,4,5,6. The coordination between division of progenitors and differentiation and migration of neurons will determine the eventual size, shape, and thus function of the brain, directly impacting behavior7,8,9,10. While tight control over these processes is clearly crucial for normal brain development, the global mechanisms that regulate these dynamics are not well understood. Here we describe a tool to study nervous system development at a cellular resolution, allowing researchers to visualize progenitor cells and neurons in vivo in the developing zebrafish brain with Brainbow and track their behavior over time via time-lapse confocal microscopy11. The approach can also be adapted to visualize other parts of the developing embryo.

To observe and distinguish among cells in the developing zebrafish brain, we have adapted the Brainbow cell-labeling technique11. Brainbow utilizes the randomly determined, combinatorial expression of three distinct fluorescent proteins (FPs) to label a population of cells. While the default expression for Brainbow expression is the red FP dTomato, recombination by the enzyme Cre recombinase results in expression of mCerulean (cyan fluorescent protein, CFP) or yellow fluorescent protein (YFP)12,13. The combined amount of each FP expressed in a cell gives it a unique hue, allowing clear visual distinction from neighboring cells. Additionally, when a progenitor cell divides, each daughter cell will inherit the color from its mother cell, producing color-coded clones and allowing researchers to trace cell lineage11,14. While originally used to analyze neuronal circuitry in mice12, Brainbow has since been expressed in a wide variety of model organisms, including zebrafish15.

Our technique builds on previous multicolor labeling and imaging methods to directly image multiple color-coded clones over time in living zebrafish. Due to their optical transparency as embryos, zebrafish are well suited to imaging experiments16, and previous studies have utilized Brainbow in zebrafish to study a variety of tissues, including the nervous system11,15,17,18,19,20,21,22,23,24,25,26,27. The ability to directly image into the living organism, along with their rapid ex utero development, make zebrafish a valuable model of vertebrate development. In contrast to the mammalian brain, the entire proliferative zone of the zebrafish hindbrain is readily available for imaging without disruption to its endogenous environment6. This allows experiments to be conducted in the living organism, rather than in in vitro or fixed tissue preparations. In contrast to fixed imaging experiments, in vivo studies allow for a longitudinal design, producing hours of data that can be analyzed for patterns, thus increasing the likelihood of observing relatively rare events. Depending upon the speed and length of the events of interest, researchers may choose to perform short (1–2 h) or long (up to ~16 h) time-lapse imaging experiments. By using the zebrafish heat shock promoter 70 (hsp70, hsp), Brainbow expression can be temporally controlled28,29. Additionally, the mosaic expression induced by this promoter is well suited for labeling and tracking many clones11.

The ability to visually identify multiple clones within the living brain is an advantage of this method. Important previous studies that investigated the role of clones within development of the nervous system utilized retroviral vectors to label a single progenitor cell and its progeny using a single FP or other readily visualized protein. Such labeling allows researchers to observe a single clone over time, either in vitro or in vivo2,30,31,32,33,34,35,36,37,38. In contrast to methods to track the behavior of cells within one clone, the distinct colors of Brainbow allow researchers to observe dynamics among clones. Additionally, by using Brainbow to label many clones within the brain, additional data on clonal behavior is collected relative to techniques that label a single clone11. Importantly, the approaches described here can be expanded to generate developmental comparisons between fish that have undergone different genetic or pharmacological manipulations18. Overall, these advantages make time-lapse in vivo confocal imaging of Brainbow-expressing zebrafish ideal for researchers exploring development of the vertebrate nervous system, particularly those interested in the role of clones.

Protocol

Procedures involving animal subjects have been approved by the Institutional Animal Care and Use Committee (IACUC) at Lewis & Clark College.

1. Microinjection of Zebrafish Embryos

  1. Set up wild type, adult zebrafish in sex-segregated mating tanks the afternoon prior to performing microinjections39,40.
  2. Prepare the DNA solution in the morning of the microinjections. Dilute hsp:Zebrabow11 plasmid DNA to a concentration of ~10 ng/μL in 0.1 mM KCl, along with 2.5% phenol red and 3.75U Cre recombinase enzyme.
  3. Perform microinjections of DNA solution into one-cell zebrafish embryos within 45 min of fertilization39,41. Inject approximately 4.2 nL of this solution into each embryo, equivalent to ~42 pg of plasmid DNA.
    NOTE: The microinjection step can be omitted if a transgenic, Brainbow-expressing zebrafish line is mated instead, such as Tg(ubi:Zebrabow)15 and Tg(neurod:Zebrabow)18. Performing microinjections can be advantageous because it allows researchers to titrate the copy number and labeling density. Furthermore, a second DNA construct that tags or manipulates a specific gene product can be injected alongside Brainbow if desired (e.g., a far-red fluorescent protein that complements Brainbow18).
  4. Maintain injected embryos in Petri dishes of E3 medium39 in a 28 °C incubator for 24 h.

2. Heat Shock to Induce Brainbow Expression

NOTE: If the plasmid DNA injected or the transgene expressed does not utilize the hsp70 promoter to drive the expression, the heat shock step can be skipped, and healthy embryos should be immediately transferred to phenylthiourea (PTU) at 24 h post fertilization (hpf).

  1. At 24 hpf, cull dead and deformed embryos from the group of injected embryos. Then, transfer the healthy embryos to a 50 mL tube, with up to 20 embryos/tube.
  2. Fill the tubes with 10 mL of E3. Place a cap on top of each tube but do not close tightly.
  3. Place the tube rack containing the 50 mL tubes upright in a 37 °C water bath. Ensure that the water level in the bath is higher than the level of the E3 in the tubes and leave them for 80 to 90 min.
  4. Remove the tube rack with the embryos from the water bath and return to the 28 °C incubator upright. Allow up to 1 h for the E3 to cool and the embryos to gradually re-acclimate to the temperature. Then, transfer the embryos to Petri dishes of 0.2 mM PTU in E3 warmed in the incubator to prevent pigmentation of embryos.
    CAUTION: PTU is a toxic chemical that should be handled with care and disposed of appropriately. Wearing gloves is suggested.

3. Screening Embryos for Brainbow Expression

  1. At 2 to 4 h after the heat shock, examine the embryos under a standard fluorescence dissection microscope for expression of CFP or YFP, which indicates successful Brainbow recombination (from default expression of dTomato).
    NOTE: If multiple fluorescence filter options are available, screening for CFP is the most efficient way to identify well-recombined fish. If CFP and YFP filters are not available, YFP can also be dimly visualized under green fluorescent protein (GFP) filters due to the overlap in their spectral profiles.
  2. Select embryos with robust FP expression throughout and transfer to a separate dish with PTU. Image embryos at 1 day post fertilization (dpf; 28 hpf or later) or maintain embryos in PTU and image at 2 or 3 dpf.

4. Mounting Embryos for In Vivo Imaging

  1. Prior to the day of the experiment, prepare the imaging chamber and embryo manipulation tool.
    1. Prepare the imaging chamber by carefully supergluing a plastic ring to the center of a 60 mm Petri dish.
    2. Optionally, prepare a custom-made embryo manipulator to move embryos within the dish and orient embryos while mounting. Construct the manipulator by supergluing a small length of nylon fishing line (~1/2 in, ~6 lb) to the end of a wooden cotton swab stick that has been cut to a ~4 in length.
  2. If necessary (i.e., imaging at 2 dpf or earlier), before mounting the embryos dechorionate using syringes under a dissection microscope.
  3. Anesthetize zebrafish embryos.
    1. Fill a 60 mm Petri dish halfway with E3 and add 5–6 drops of 4 mg/mL MS-222 Tricaine-S to a final concentration of ~0.2 mM. Swirl to mix.
      CAUTION: Tricaine should be handled with care and disposed of appropriately.
    2. Transfer embryos from the PTU to the tricaine solution, ensuring as little PTU is transferred as possible. While fish reflexes should cease after 1–2 min, leave the embryos in tricaine for up to 10 min to ensure thorough anesthetization for time-lapse imaging experiments.
    3. Confirm that the embryos are properly anesthetized. Assess the startle reflex by touching the embryo tails gently with the embryo manipulator. If a reflex movement is observed, add an additional drop of tricaine to the dish as far from the embryos as possible and swirl to mix. Observe the embryo heart rate under a dissection scope. If it is abnormally slow, add additional drops of E3 to the dish to reduce the tricaine concentration.
  4. Mount anesthetized embryos in agarose.
    1. Transfer the fish to the center of plastic ring within the imaging chamber and remove as much excess E3 as possible using a fine-tipped transfer pipette.
    2. Using a clean transfer pipette, cover the fish with 1.0% low-melt agarose (LMA) in E3 stored at 42 °C and fill the entire plastic ring with a thin layer of agarose. Use a transfer pipette to gently pull the embryos up into the pipette tip and back into the agarose without introducing air bubbles.
    3. Before the agarose hardens, quickly use an embryo manipulator to orient the fish appropriately. If imaging with an upright microscope, position the embryos as close to the upper surface of the agarose as possible. Position the embryos parallel to the bottom of the imaging chamber with the tail straight.
      NOTE: For hindbrain imaging, the embryos can be positioned in a dorsal or sagittal view. Care should be taken to ensure that the heads of the embryos do not sink while the agarose is still liquid. Other orientations may be appropriate to target other developing tissues for imaging. For inverted microscopes, mounting would be done differently, typically positioning the fish close to the bottom of a glass-bottomed Petri dish.
  5. Wait for the agarose to harden, then fill the imaging chamber with E3. Add as much E3 as possible to account for the evaporation over the course of imaging.

5. Time-lapse Confocal Imaging of Developing Zebrafish Hindbrain

  1. Place the imaging chamber on the confocal microscope in preparation for the imaging.
    1. Place the imaging chamber with the fish and E3 on the confocal microscope.
    2. Select an objective with a high numerical aperture and a long working distance, such as a 20x water immersion objective (1.0 NA).
    3. Find a region with appropriately dense and bright labeling of the cell type(s) of interest.
      NOTE: A temperature-controlled stage/apparatus on the microscope may also be used to regulate temperature and humidity during long imaging sessions. This can decrease evaporation.
  2. Set up the acquisition parameters for Brainbow imaging.
    1. Image each FP channel sequentially. If using a commercial software (e.g., Zen software), this is done by preparing three “tracks”. Use an Argon laser to excite CFP at 458 nm and YFP at 514 nm. Use a DPSS 561 nm laser to excite dTomato. Collect emissions between 463–509 nm for CFP, 519–555 nm for YFP, and 566–691 nm for dTomato.
    2. For on-screen display, code CFP as blue, YFP as yellow, and dTomato as red.
      NOTE: These settings will vary depending upon which laser lines and other features are available on the confocal microscope. To increase the speed of imaging, CFP and dTomato channels can be imaged simultaneously without appreciable bleed-through. If a separate FP is also being imaged, such as a far-red FP, this can be imaged sequentially or simultaneously with YFP.
    3. Set up the general imaging parameters to take 16-bit images with a resolution of at least 1024 x 1024 and averaging twice. Adjust the zoom depending on the region and cell type of interest (i.e., for hindbrain imaging with a 20x objective the zoom will range from 1.0–2.5). Maximize the field of view to allow for growth of the fish over time.
    4. Viewing one track at a time (and turning off other lasers to prevent photobleaching), optimize the acquisition settings for each FP individually (e.g., laser power, pinhole size, photomultiplier tube gain, etc.).
    5. Select Z-stack range to image.
      NOTE: For long imaging sessions (>2 h), take into consideration that the fish will continue to grow throughout the course of imaging, thus both the XY field and the Z-stack range should account for this with extra room. If imaging the hindbrain, include space in the field for the tissue to move rostrally during the imaging period. Space also needs to be included for growth in the Z dimension. Include in the image approximately 10–20 μm above the region of interest, whether the fish is positioned for sagittal or dorsal view.
  3. Set up parameters for time-lapse imaging.
    1. Select a time interval. Interval length ranges between 10–30 min to track mitotic and apoptotic events in the developing hindbrain. Interval length will vary depending on the speed of the events of interest and the total time it takes to capture a single Z-stack.
    2. Select the length of the imaging session. Time-lapse imaging sessions generally occur overnight and can last at least 16 h.
  4. Run the experiment.
    NOTE:     The fish can be euthanized immediately following imaging or carefully dissected from the agarose using syringes and returned to E3 to recover. Recovered fish can then be imaged again at a later time point, though cell position and depth will shift during this period due to overall growth.

6. Time-lapse File Management

  1. Import the image file into the analysis software.
    1. If using Zen software, save the file in .czi format once image acquisition is completed. Make sure to save raw data in a format compatible with Fiji42 and/or other software.
    2. Import into Fiji software using BioFormats Importer.
      NOTE: Time-lapse files will be large and can require significant amounts of time and memory to open for analysis. Thus, it is helpful to be strategic in how they are saved and opened. It can be useful to save a lower-quality version that can be opened more easily to search for events of interest by eye.
  2. If using Fiji, create smaller, more manageable subsets of time-lapse files. Generate subsets either by time (e.g., viewing full Z-stack at the first time point) or by depth (e.g., viewing the first 50 Z-sections at each time point) by clicking Image|Stacks|Tools|Make Substack.

7. Quantitative Clonal Color Analysis of Brainbow Images

  1. Measure the mean red, green, and blue (RGB) intensity values for the cells of interest in Fiji.
    NOTE:     These instructions are specific to image analysis in Fiji. However, researchers may prefer to obtain mean RGB intensity values in an alternative software program.
    1. Ensure that the file is correctly cropped so that there is no black space around the edges of the image. Black space will artificially lower the minimal intensity measurements needed for the analysis. If the file does need to be cropped, select the Rectangle Selection Button and draw a region of interest (ROI) around the field of view, excluding all black space. Click Image|Crop.
    2. Set the measurement tool to take mean, minimum, and maximum gray value measurements. Click Analyze|Set Measurements. Ensure the checkboxes for Min & Max Gray Value and Mean Gray Value are both selected.
    3. Viewing the raw data files or subsetted Z-stacks in Fiji, find cells of interest for clonal color analysis. In the hindbrain, putative clones can be visually identified by their shared hue as well as their radial orientation. Do not select cells that are in close contact with, and thus difficult to resolve from, neighboring cells of another color. It may be difficult to quantify their hue.
    4. Find the center Z-plane of the cell of interest using the Z-scrollbar. Right-click the Oval Selection Button and select the Elliptical Selection Button. Other selection tools will be appropriate for different cell types. Draw an elliptical ROI around the central ~2/3 of the cell body.
    5. Using the C-scrollbar, select the red (dTomato) channel. Take the mean intensity measurement for this channel by selecting Analyze|Measure. Repeat with the same ROI and focal plane for the green (YFP) and blue (CFP) channels and save all mean intensity values.
    6. Click anywhere on the image to remove the elliptical ROI in the cell of interest.
    7. To measure background levels for normalization, use the C-scrollbar and select the red (dTomato) channel. Take the minimum intensity measurement for the whole field in this channel by selecting Analyze|Measure. Repeat with the same focal plane for the green (YFP) and blue (CFP) channels and save all minimum intensity values.
  2. Calculate relative RGB channel weights for the cells of interest.
    NOTE:     Calculation of relative RGB channel weights from these intensity values can be performed in a variety of software programs, such as Microsoft Excel or R43.
    1. Normalize the mean intensity measurement of the cell ROI for each channel by subtracting the minimal intensity from the entire field in that channel and focal plane.
    2. Sum the normalized mean RGB intensity values from each channel to find the total normalized RGB intensity for the cell.
    3. Calculate the relative channel weight for each channel by dividing the normalized mean intensity value for that channel by the total normalized RGB intensity.
  3. Display relative RGB channel weights as ternary plots using the Ternary package for R43,44. Ternary plots are useful to visualize the clustering of similar colors in 3D RGB space.
  4. Calculate the color spread coefficient to quantify the difference between cell colors.
    1. Calculate the Euclidean distance between the two colors in 3D RGB space using the following equation:
      figure-protocol-16111
      where R, G, and B are the relative RGB channel weights calculated in 7.2.
    2. For ease of comprehension, normalize the color distance by dividing by √2, the maximum possible distance between colors, to obtain a color spread coefficient between 0 and 1, where 0 indicates exactly the same color and 1 indicates completely different colors (e.g., pure red and pure blue).

Results

This section illustrates examples of results that can be obtained using the in vivo multicolor time-lapse imaging approach described here. We show that Brainbow color-coded clones of cells in the proliferative ventricular zone of the developing zebrafish hindbrain14 (Figure 1).

Typically, when Brainbow-labeled cells were arranged along a particular radial fiber, they shared the same color (Figure 1D), which cou...

Discussion

This protocol describes a method to visualize clones of progenitor cells and neurons in the developing zebrafish hindbrain and follow them in vivo using Brainbow and time-lapse confocal microscopy11. The major advantage of this protocol in comparison to in vitro or ex vivo studies is the ability to directly observe the proliferative zone of the vertebrate brain in its natural milieu over time. This technique builds on previous studies that labeled a single clone using retroviral vectors. In c...

Disclosures

The authors have nothing to disclose.

Acknowledgements

We thank Y. A. Pan, J. Livet and Z. Tobias for technical and intellectual contributions. This work was supported by the National Science Foundation (Award 1553764) and the M.J. Murdock Charitable Trust.

Materials

NameCompanyCatalog NumberComments
1.5mL transfer pipetGlobe Scientific, Inc.134020
1-phenyl-2-thiourea (PTU)Alfa AesarL06690Diluted to 0.2 mM in E3 to prevent embryo pigmentation
50ml conical tubesCorning352070For heat shocking embryos
6 lb nylon fishing lineSecureLineNMT250For making embryo manipulators
7.5mL transfer pipetGlobe Scientific, Inc.135010
CaCl2SigmaC3881For E3
Cotton swabsPuritan867-WC NO GLUEFor making embryo manipulators
Cre recombinaseNew England BiolabsM0298M
Digital dry bathGenemate490016-616Used to store LMA at 40°C
Epifluorescence dissection scope
Glass capillary tubesWorld Precision InstrumentsTW100F-4
IncubatorForma Scientific3158To maintain embryos at 28°C
Injection plate moldsAdaptive Science ToolsTU-1
Isotemp water bathFisher Scientific2320For heat shocking embryos
KClAMRESCO0395For E3 and for DNA solution for injections
Laser-scanning confocal microscopeZeissLSM710
LE agaroseGenemateE3120To create agarose injection plates
Low-melt agarose (LMA)AMRESCOJ234
Mating tanksAquaneering, Inc.ZHCT100
Methylene blueSigmaM9140For E3
MgSO4Sigma9397For E3
MicromanipulatorWorld Precision InstrumentsM3301
Micropipette PullerSutter Instrument Co.P-97
MS-222 Tricaine-SWestern Chemical, Inc.Stock made at 4 mg/mL in reverse osmosis (RO) water, then added dropwise to E3 to final concentration of 0.2 mM to anesthetize embryos
NaClJ.T. Baker4058-01For E3
Petri dishes (90 mm, 60 mm)Genesee Scientific32-107GTo house embryos and create imaging chamber (60 mm)
Phenol redSigmaP0290
Soft stitch ring markersClover Needlecraft, Inc.354For creating imaging chamber with Petri dish
Super glue (Ultra gel control)Loctite1363589For making embryo manipulators
Syringe needlesBeckton DickinsonBD329412For dechorionating embryos

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