Published: April 5th, 2021
Here, a protocol is provided for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software.
Vertebrate eye development is a complex process that begins near the end of embryo gastrulation and requires the precise coordination of cell migration, proliferation, and differentiation. Time-lapse imagining offers unique insight to the behavior of cells during eye development because it allows us to visualize oculogenesis in vivo. Zebrafish are an excellent model to visualize this process due to their highly conserved vertebrate eye and their ability to develop rapidly and externally while remaining optically transparent. Time-lapse imaging studies of zebrafish eye development are greatly facilitated by use of the transgenic zebrafish line Tg(rx3:GFP). In the developing forebrain, rx3:GFP expression marks the cells of the single eye field, and GFP continues to be expressed as the eye field evaginates to form an optic vesicle, which then invaginates to form an optic cup. High resolution time lapse imaging of rx3:GFP expression, therefore, allows us to track the eye primordium through time as it develops into the retina. Lightsheet microscopy is an ideal method to image ocular morphogenesis over time due to its ability to penetrate thicker samples for fluorescent imaging, minimize photobleaching and phototoxicity, and image at a high speed. Here, a protocol is provided for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software. The resulting dataset can provide valuable insights into the process of ocular morphogenesis, as well as perturbations to this process as a result of genetic mutation, exposure to pharmacological agents, or other experimental manipulations.
Embryonic development is a complex process that requires the precise coordination of many different events. The formation of the vertebrate eye begins in the developing forebrain, where a portion of the cells are specified as the eye field. These cells will evaginate toward the surface ectoderm, giving rise to two bilateral optic vesicles1,2,3,4,5,6,7,8,9,10. Contact with the surface ectoderm then induces an invagination of the optic vesicle into an optic cup. The surface ectoderm will give rise to the anterior structures of the eye, such as the lens and cornea, while the optic cup will give rise to the neural retina and retinal pigmented epithelium1,2,3,5,6,7,8,9,10,11,12. Disruptions in this process can lead to congenital defects such as microphthalmia, anophthalmia, and coloboma (MAC). At this time, there are no options to correct these defects3,5,6,7,9,10,11,12. Further studies of the mechanisms of ocular morphogenesis and the problems that can lead to MAC will provide a foundation of knowledge that will potentially lead to treatments. One powerful tool to investigate the dynamic behaviors of cells during eye development is time-lapse imagining, which allows this process to be visualized and characterized in vivo and in real time.
Zebrafish (Danio rerio) are an excellent model to visualize early ocular development using time-lapse imaging. They have a highly conserved vertebrate eye and possess the ability to develop rapidly and externally while remaining optically transparent1. Zebrafish provide a great resource for time-lapse imaging due to these characteristics that mammalian models lack. Time-lapse imaging studies of zebrafish eye development are greatly facilitated by use of the transgenic zebrafish line Tg(rx3:GFP)13. Rx3 (Retinal homeobox protein 3) is a transcription factor essential for eye development14. Rx3 is the first of the three rx genes in the zebrafish to be expressed, starting its expression mid-gastrulation, approximately 8 h post fertilization (hpf)15,16. The rx3:GFP transgene can be visualized in the developing forebrain starting at the 1 somite stage (ss), approximately 10 hpf15,17,18,19,20. In the developing forebrain, rx3:GFP expression marks the cells of the single eye field, and GFP (green fluorescent protein) continues to be expressed through the remainder of ocular morphogenesis. High resolution time lapse imaging of rx3:GFP expression, therefore, allows us to track the single eye field through time as it develops into the retina17,18,20.
Time-lapse imaging studies of zebrafish development have primarily been performed using confocal or Lightsheet microscopy. Confocal microscopy is advantageous in that it allows for the precise imaging of samples along the z-axis and reduces fluorescent background signal. However, it is limited by the amount of time it takes to acquire an image, sample position rigidity, and its propensity toward photobleaching and phototoxicity of live samples. Lightsheet microscopy is an ideal method to image ocular morphogenesis over time due to its ability to penetrate thicker samples for fluorescent imaging, increased flexibility in sample orientation, minimized photobleaching and phototoxicity, and imaging at a high speed21,22,23,24,25,26,27,28,29,30,31. The spatial resolution achievable with current light sheet microcopy systems is approximately 250-500 nanometers (nm). Although this is not significantly different from what can be obtained with confocal microscopy, the sample can be freely rotated and imaged from multiple angles, improving both imaging depth and resolution, and offering much greater flexibility for in vivo time lapse imaging experiments than the confocal platform27,32. For these reasons, Lightsheet microscopy is quickly becoming the favored method for time-lapse imaging studies of zebrafish development. This protocol describes the steps of quantifying oculogenesis through the imaging of Rx3:GFP transgenic zebrafish using a commercially available Lightsheet microscope33 and details a pipeline for image analysis using the arivis software platform.
All experiments involving the use of zebrafish were carried out in accordance with protocols established by the University of Kentucky Institutional Animal Care and Use Committee (IACUC).
1. Sample preparation
2. Zeiss Lightsheet Z.1 setup
3. Image analysis
The dataset displayed here was imaged using the protocol described above. A Tg(Rx3:GFP) embryo was imaged starting at the 1 somite stage (ss) through 24 hpf, a total time period of 14 h, with the images acquired at 5 min intervals. Time-lapse imaging allows for easy selection and comparison of any time-point that shows a phenotype of interest. Figure 5 demonstrates a set of high-resolution images that were rendered from the dorsal vantage point at select developmental time points. The pipeline run in arivis Vision4D builds a mask that represents the developing eye as identified by fluorescent signal. In Figure 5 and Videos 1-6, the mask can be visualized in comparison to the fluorescent rendering of the developing eye. Additionally, Table 2 displays the volume data from the developing eye at every imaging point. This dataset includes the segment name, id, volume in both µm3 and voxel count, the mean fluorescence intensity of the object, the time point the object was identified, and the object's surface area in µm2. It is important to note that when the eye field separates into two optic vesicles (starting at Timepoint 64), there remains a third region that is Rx3:GFP positive in the forebrain, which will contribute to the hypothalamus38,39,40 (Figure 5D-M). This shows up in the volume data represented in Table 2 (highlighted in yellow starting at Timepoint 71) and can easily be separated out from the optic vesicles, since it is much smaller in volume than either optic vesicle.
Figure 1: Sample preparation. (A) Positioning of embryos in a glass capillary. The arrow points to an embryo in the capillary. (B) Glass capillary and capillary holder parts. (C) Partially assembled capillary holder. (D) Fully assembled capillary holder. (E) Lightsheet mounting chamber. The arrow indicated the white line used to orient the capillary holder. (F) Capillary holder properly mounted in the Lightsheet. The arrow shows the matching white lines, indicating proper orientation of the capillary holder. Please click here to view a larger version of this figure.
Figure 2: Lightsheet imaging set-up. (A) Switchboard to turn on the Lightsheet, computer, and incubation unit. The numbers indicate the order of operations. (B) Lightsheet objective chamber. (C) Imaging chamber. (D) Syringe and tubing that will be connected to the imaging chamber. (E) The imaging chamber properly positioned within the objective chamber with all of the tubes connected to the appropriate ports to the right. Please click here to view a larger version of this figure.
Figure 3: Sample positioning. (A) Locate Capillary and Locate Sample buttons in Zen Software as indicated by the arrows. (B) The positioning on the glass capillary. The arrow indicates the edge of the glass capillary positioned just above the lens of the objective. (C) The embryo suspension beyond the glass capillary. The arrow indicated the embryo suspended in agarose beneath the glass capillary in front of the objective's lens. (D) View of the embryo through the objective. (E) The ErgoDrive control panel. Please click here to view a larger version of this figure.
Figure 4: Important icons for navigating arivis Vison4D. Each panel has the icon function identified from left to right. (A) Open, Save, Close. (B) Analysis Panel, Show Objects Table, Open Track Editor. (C) Copy current viewer content as an image into clipboard, Toggle Bookmarks, Create a high-resolution image for the current view, Toggle Storyboard. (D) Show Measure Box, Show Orientation Cross, Show Legend, Show Scale Bar. (E) Show as 2D Viewer, Show as Gallery Viewer, Show as 4D Viewer, Show as Info Viewer, Show as Projection Viewer. F) Refresh all Keyframes, Add Keyframe, Add Keyframe sequence, Insert Keyframe, Remove all Keyframes, Export Movie, Load Storyboard, Save Storyboard, Adjust the target time of the entire movie, First Keyframe, Play, Pause, Stop, Last Keyframe. Please click here to view a larger version of this figure.
Figure 5: High-resolution images and eye field masks. (A-M) A set of high-resolution images that were rendered from the dorsal vantage points; (A'-M') the eye field masks for each corresponding timepoint. Each image set is notated by the time it was acquired from the start of imaging and the corresponding developmental stage in either somite stage (ss) or hours post fertilization (hpf). Please click here to view a larger version of this figure.
Video 1: Time-lapse video of Tg(rx3:GFP) zebrafish embryo from 1 ss-24 hpf. Please right click here to download the video (save link as).
Video 2: Time-lapse video of Tg(rx3:GFP) zebrafish embryo from 1 ss-24 hpf with the eyefield as identified by the arivis Vision4D pipeline. Please right click here to download the video (save link as).
Video 3: 360° rotation of a Tg(rx3:GFP) zebrafish embryo at 1 ss. Please right click here to download the video (save link as).
Video 4: 360° rotation of a Tg(rx3:GFP) zebrafish embryo eye field mask at 1 ss. Please right click here to download the video (save link as).
Video 5: 360° rotation of a Tg(rx3:GFP) zebrafish embryo at 24 hpf. Please right click here to download the video (save link as).
Video 6: 360° rotation of a Tg(rx3:GFP) zebrafish embryo eye field mask at 24 hpf. Please right click here to download the video (save link as).
Table 1: arivis Vision4D pipeline for volume analysis of the developing eye field. Please click here to download this Table.
Table 2: Volume and surface area of the developing eye field acquired by arivis Vision4D. The rows pertaining to the presumptive hypothalamus are highlighted in yellow to distinguish them from the optic vesicles. Please click here to download this Table.
In this protocol, the Lightsheet microscope was used to perform time-lapse imaging of eye development and the resulting data were analyzed. The resulting dataset can provide valuable insights into the process of ocular morphogenesis, as well as perturbations to this process as a result of genetic mutation, exposure to pharmacological agents, or other experimental parameters. Here the protocol demonstrated how this dataset can be obtained and provided an example of how to analyze the volume of the eye field through early development. This data was found to be reproducible and consistent (less than 10% variation in volume) across biological replicates, bearing in mind that slight differences in embryo staging prior to the start of the run can lead to some variation in final volume measurements.
Care should be taken in the initial positioning of the embryo in the capillary and in positioning the embedded embryo in front of the objective. Orientation plays an important role in preventing the embryo from growing and moving out of the view of the objective. The embryos have a round shape at 10 hpf, which makes it challenging to guarantee a specific orientation in the capillary. Ideally, the body of the embryo will be positioned laterally in the capillary. Loading multiple embryos in the capillary will increase the likelihood of having a well-positioned embryo.
In this procedure, the embryo is embedded in agarose in order to suspend it in front of the imaging and illumination objectives. Choosing the correct concentration of the low melting temperature agarose is critical. Too high of a concentration will constrict the embryo and prevent it from properly developing; too low of a concentration will result in the agarose falling apart and not holding the embryo. The concentration optimal for this protocol is a final concentration of 1% low melting temperature agarose30,31.
Another element that should be taken into consideration is the level of saturation. As the eyefield grows and differentiates, the strength of the Rx3:GFP signal intensifies. Therefore, when setting the initial imaging parameters, the exposure and laser power should be reduced to undersaturate the image. This will prevent the image from becoming oversaturated as the Rx3:GFP gets brighter over time. Modifications can be made to correct for undersaturation in image processing but oversaturation cannot be corrected after the images have been acquired.
There are a few additional modifications that can be made to this protocol that may be advantageous to some projects that are not described in this paper. For example, it is possible to set up Multiview imaging in the image acquisition set up. This parameter would allow multiple embryos at different positions along the y-axis to be sequentially imaged at each time interval. While adding complexity to the data set, it would increase the rate of data collection. Additionally, in terms of image processing, it is possible to quantify the eye field by other parameters. Here, we described how to quantify the data in terms of the eye field volume. Alternatively, a pipeline could be made to quantify and track individual cells or determine the rate of optic vesicle evagination.
As previously mentioned, both confocal and Lightsheet microscopy have been used to perform time-lapse imaging studies of zebrafish. Lightsheet was specifically chosen for this project due to its superior ability to image through a thick (>1 mm) sample, because it is equipped with an incubation unit to maintain an ideal temperature environment for the zebrafish embryo, and because its ability to image at a faster rate than confocal microscopy allows for image acquisition at the numerous time intervals required for this protocol no accompanying damage or photobleaching of the embryo21,22,23,24,25,26,27,28,29,30,31. It is also important to note that the Lightsheet microscope is equipped to image the signal from multiple fluorophores. The Lightsheet microscope used in this study has solid state laser excitation lines at 405, 445, 488, 515, 561, and 638 nm, which could be useful for imaging transgenic embryos expressing more than one fluorescent reporter transgene.
While this protocol details instructions for image acquisition analysis specifically using the Lightsheet Z.1 Dual Illumination Microscope System and arivis Vision4D analysis software, there are other commercially available Lightsheet microscopes made by Leica, Olympus, and Luxendo, as well as image analysis software by Imaris, that could be used to achieve similar results. The selection of equipment and software for this protocol was determined by the availability at our institution.
In summary, it is anticipated this protocol will provide a solid starting point for conducting time-lapse imaging using Lightsheet microscopy, and for image quantification of early eye development in zebrafish.
The authors declare no competing financial interests.
Research reported in this publication was supported by the Office of The Director of the National Institutes of Health (NIH) under Award Number S10OD020067 and by NIH award R01EY021769 (to A.C.M.). We are grateful for the assistance of Doug Harrison and Jim Begley in the Arts & Sciences Imaging Center at the University of Kentucky, and to Lucas Vieira Francisco and Evelyn M. Turnbaugh for expert zebrafish care.
|60mL Syringe Luer-Lok Tip
|Agarose, Low Melting Temperature
|arivis Vision4D Software
|Dumoxel N3C Forceps
|E3 fish buffer (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4)
|Glass Capillary – Size 2 (~1mm)
|Heidelberger Extension Line 100cm
|Light & Filter Set – Royal Blue
|Petri Dish (100 x 15 mm)
|Stereomicroscope Fluorescence Adapter
|Teflon Tipped Plunger – Size 2
|This line was established by Rembold et al.13
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