Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.
Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.
Posterior body elongation is an essential process in embryonic development by which the embryo extends to form a large part of the body axis. It is an example of a complex morphogenetic process by which multiple cell behaviors act coordinately to generate the morphogenesis at the level of individual tissues. These differential tissue deformations then act together to generate the elongation of the posterior body at the whole structure level. To understand how these processes are controlled and coordinated during development, we must be able to follow these processes at multiple scales (i.e. at the level of molecules, cells, cell populations and tissues) and to relate this directly to the morphogenesis of the entire structure.
Zebrafish embryos are ideal for imaging posterior body elongation as their optical transparency and small size allows for the application of minimally invasive light imaging approaches well suited to live imaging.1 This has been evidenced by a series of recent publications that have shed light on posterior body development at the level of molecules,2 single cells,3 and inter-tissue behaviors,4 as well as at the level of cell population and whole organ.5
Advanced imaging techniques such as confocal, multi-photon microscopy and selective plane illumination microscopy (SPIM) are enabling the long term imaging of developmental processes with decreased effect of light toxicity and photo-bleaching. Robust techniques for the mounting of live samples are required to achieve three goals: 1) sufficient support to maintain samples in the correct orientation during transfer to the microscope and during acquisition, 2) sufficient freedom of movement of the sample to allow for the outgrowth of the posterior body region without affecting its normal development, and finally 3) a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups.
This protocol introduces a mounting technique for imaging the development of the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving the posterior body region to elongate and develop normally. As such, it is also an appropriate method for the long-term imaging of other regions of the developing body as the agarose enables imaging by standard light imaging techniques. This protocol demonstrates mounting of embryos in a lateral orientation, although it is also possible to mount embryos in alterative orientations. It will further show how to adapt the method for upright, inverted and vertical light-sheet microscopy setups.
1. Preparation of Solutions and Pulled Glass Needle
2. Embedding of Embryos for Inverted or Upright Microscopy
3. Removal of Excess Agarose Around the Posterior Body
NOTE: This section describes the procedure by which agarose is removed from the region surrounding the posterior body. In the case of posterior body elongation, it is important to ensure that the tail can grow-out normally. By removing the agarose after the embryo has been completely included by agarose, the embryo is left enclosed by the head region and approximately half of the yolk sac.
Figure 1: Diagrams of Mounting Set-ups. (A) The diagram shows the position of the mounted embryo within the center glass ring of a petri dish. On the right is a zoom of the embryo with each successive cut through the surrounding agarose shown with dottted red lines. (B) The mounted embryos is diagrammed in lateral view displaying the ease of access for both inverted and upright objectives. (C) A similar digram showing how embryos can be mounted for vertical light sheet imaging set-ups. Please click here to view a larger version of this figure.
4. Mounting of Embryos for Vertical Light-sheet Microscopy
NOTE: This is a variation on the method outlined above that allows for the access of multiple objectives for imaging of samples by vertically-orientated SPIM. The idea behind this variation is to lift the sample slightly higher than the bottom of the dish, to allow for easy access of two imaging objectives.
The protocol outlined above details a versatile technique for the mounting of zebrafish embryos for long-term time lapse imaging. An example of this is shown in Figure 2A and in animated/video Figure 1. Embryos were injected at the 1 cell stage with mRNA encoding the photoconvertible fluorescent protein kikumeGR. At the 15 somite stage they were mounted as described above and imaged for 12 hr on an inverted confocal microscope with a 10X objective. The resulting confocal stacks were maximally projected to be displayed as shown. This movie clearly shows that the posterior body is allowed to move freely throughout the duration of the movie and shows similar changes in morphology as seen in embryos that are allowed to develop freely of their chorion in normal culture conditions.
An additional example is shown in Figure 2B and animated/video Figure 2. Here, embryos were injected at the 16 cell stage with a combination of mRNAs encoding histone 2B-mCherry protein (that labels the nuclei) and eGFP:CAAX box (that labels membranes; for details see reference5). They were then imaged on an upright multiphoton microscope with a water immersion 25X objective as diagrammed in Figure 1B. Images were taken for 3 hr from the 10 somite stage in order to visualize cellular behaviors during tailbud formation. Cells can be seen to be generating active protrusions and directional movements as the tailbud forms normally.
Figure 2: Examples of Time-lapse Imaging of Posterior Body Elongation at 10X Magnification. (A) Consecutive frames of a representative movie covering the process of axial elongation. Embryos are shown in lateral view with posterior to the right side of the image. (B) Consecutive frames of a higher magnification movie showing individual cell behaviors during tailbud formation. Embryos are shown in lateral view with posterior to the right. Dotted lines outline the forming tailbud. Small colored lines show tracks of nuclei to follow individual cell movements. Please click here to view a larger version of this figure.
Animated/Video Figure 1: Time-lapse Movie of Posterior Body Elongation of a KikumeGR mRNA Injected Zebrafish Embryo by Confocal Microscopy. Embryo is shown with anterior to the left and posterior to the right. Images were taken at 10 min intervals at 10X magnification. Please click here to view a larger version of this figure.
Animated/Video Figure 2: Time-lapse Movie of Posterior Body Elongation of a Nuclear-mCherry and Membrane-GFP Labelled Zebrafish Embryo Imaged by Two-photon Microscopy. Embryo is shown with anterior to the right and posterior to the left. Images were taken at 2 min intervals at 25X magnification. Colored lines display tracks of cells for the last ten time-steps for each tracked cell. Please click here to view a larger version of this figure.
This mounting technique enables embryos to be kept still during transfer to the microscope and over long-term time-lapse imaging experiments aimed at following posterior body elongation at multiple length scales. Furthermore, it is versatile in that it allows for imaging on both upright and inverted microscopy set-ups, and a suggestion is made for how this can be further adapted to vertically orientated SPIM.
A critical step in this protocol is the careful removal of excess agarose surrounding the posterior body that is important for allowing the normal development of this structure. It is important to take care here in not damaging the embryo, particularly when removing the agarose around the top of the yolk. In addition, care must be taken when removing the cut agarose block from around the embryo, as sometimes it is possible to lose the embryo from the mold during this process. For these reasons, this is a low-throughput method that is not suitable for the mounting of many zebrafish embryos simultaneously, in contrast to already published methods utilizing 3D printed plastic molds.8,9 However, embryos are highly stable by this mounting method and therefore can be transported to the microscope much easier, and retain their 3D orientation throughout posterior body elongation.
Another critical step is the accuracy of the initial lateral orientation of the embryo, avoiding any antero-posterior or dorso-ventral tilt. Either tilt will preclude the lateral vision of the posterior body extension, which is the most convenient angle of view for further analysis. The antero-posterior tilt will in addition result in the posterior body growing up or down with respect to the horizontal plane, which will mean the acquisition of higher stacks in z and thus time-lapses with a lower time resolution.
This method is versatile and has allowed for imaging of cell division rates with the use of the Fucci line5,10 as well as multi-scalar morphometric analyses.5 However, given the dramatic overall displacement of the tailbud during the elongation of the posterior body axis, only relatively low magnification objectives can be used to capture the entire process such as shown in Figure 2. This is because higher magnification results in the region of interest leaving the field of view within 3 – 4 hr of acquisition (Video Figure 1). Therefore, one way to further improve the imaging of the tailbud is to have an automated tracking algorithm that can track the overall movement of the tailbud and adjust the x,y,z position of the microscope stage during acquisition. Given the increased stability of the embryo in all axial dimensions with the mounting method described here, it should be amenable to such an approach.
The authors have nothing to disclose.
Estelle Hirsinger: Core funding from the Institut Pasteur and Agence Nationale de la Recherche (ANR-10-BLAN-121801 DEVPROCESS). Estelle Hirsinger is from the Centre National de la Recherche Scientifique (CNRS). Benjamin Steventon was funded by the Agence Nationale de la Recherche (ANR- 10-BLAN-121801 DEVPROCESS), then a Roux fellowship (Institut Pasteur) then an AFM-Téléthon fellowship (number 16829). He is now supported by a Wellcome Trust/Royal Society Sir Henry Dale Fellowship.
CONSUMABLES | |||
Glass-bottomed dishes | Mattek | P35-1.5-10-C | 35mm petri dish, 10mm microwell. No. 1.5 cover glass |
Capillaries for injection needles | Sutter | BF 120-94-10 | We use orosilicate glass with filament, OD 1.20 mm, ID 0.94 mm, length 10 cm. However, filament needles are not necessary and most injection standard needles should work. |
Micro-scalpel | Feather | P-715 | Micro Feather disposable opthalmic scalpel with plastic handle |
Pasteur Pipettes | 230 mm long | ||
REAGENTS | |||
Tricaine | Sigma-Aldrich | A5040 | |
Low-melting point agarose | Sigma-Aldrich | A9414 | |
EQUIPMENT | |||
Fine forceps | FINE SCIENCE TOOLS GMBH | 11252-30 | Dumont #5 |
Needle puller | Sutter | P97 | Heating-filament needle puller |
Binocular dissecting microscope | Leica | S8 Apo |