We want to thank Tobias Pietzsch for providing his powerful open source software BigDataViewer. We thank the Light Microscopy Facility of the Max Planck Institute of Molecular Cell Biology and Genetics (MPI-CBG), namely Jan Peychl, Sebastian Bundschuh and Davide Accardi for technical assistance, perfect maintenance of the microscopes used in the study and for comments on the manuscript and H. Moon (MPI-CBG Scientific Computing Facility) for the BigDataServer. We thank Julia Eichhorn for assembling the Movie 1. The Norden lab members and Svea Grieb provided many helpful comments on the manuscript. Jaroslav Icha, Christopher Schmied and Jaydeep Sidhaye are members of the International Max Planck Research School for Cell, Developmental and Systems Biology and doctoral students at TU Dresden. Pavel Tomancak is supported by the ERC Starting Grant: Quantitative Analysis of the Hourglass Model of Evolution of Development and Human Frontier Science Program Young Investigator grant RGY0093/2012. Caren Norden is supported by the Human Frontier Science Program (CDA-00007/2011) and the German Research Foundation (DFG) [SFB 655, A25].

Open Access publication fees were provided by, Carl Zeiss Microscopy GmbH.

Morphogenesis is the process that shapes the embryo and together with growth and differentiation drives the ontogeny from a fertilized egg into a mature multicellular organism. The morphogenetic processes during animal development can be analyzed best by imaging of intact living specimens1-3. This is because such whole embryo imaging preserves all the components that drive and regulate development including gradients of signaling molecules, extracellular matrix, vasculature, innervation as well as mechanical properties of the surrounding tissue. To bridge the scales, at which morphogenesis occurs, the fast subcellular events have to be captured on a minute time scale in the context of development of the whole tissue over hours or days. To fulfill all these requirements, a modern implementation4 of the orthogonal plane illumination microscope5 was developed. Originally, it was named the Selective Plane Illumination Microscopy (SPIM)4; now an all-embracing term Light Sheet Fluorescence Microscopy (LSFM) is typically used. LSFM enables imaging at high time resolution, while inducing less phototoxicity than laser scanning or spinning disc confocal microscopes6,7. Nowadays, there are already many implementations of the basic light sheet illumination principle and it has been used to image a large variety of specimens and processes previously inaccessible to researchers8-11.
We would first like to highlight several key advantages of LSFM over conventional confocal microscopy approaches:
To acquire meaningful results from live imaging microscopic experiments, it is important that the observation only minimally affects the specimen. However, many organisms including zebrafish are very susceptible to laser light exposure, making it challenging to image them in a confocal microscope with high time resolution without phototoxicity effects like stalled or delayed development6,7. LSFM is currently the fluorescence imaging technique with the least disruptive effects on the sample7. Since the thin laser light sheet illuminates only the part of the specimen that is imaged at a particular time point, the light sheet microscope is using photons very efficiently. Consequently, the low light exposure allows for longer time-lapse observations of healthier specimens, e.g.12-17. Furthermore, thanks to the minimal invasiveness of LSFM, the number of acquired images is no longer dictated by how much light the sample can tolerate, but rather by how much data can be processed and stored.
Along the same lines of keeping the specimen in near physiological conditions, LSFM comes with alternative sample mounting strategies well suited for live embryos. In LSFM techniques, the embryos are typically embedded within a thin column of low percentage agarose. The mounting into agarose cylinders allows for the complete freedom of rotation, so the sample can be observed from the perfect angle (in LSFM referred to as view) and from several views simultaneously. The multiview imaging and subsequent multiview fusion is beneficial particularly for big, scattering specimens and allows capturing them with high, isotropic resolution. A summary of other possible LSFM mounting strategies can be found in the official microscope operating manual, in the chapter on sample preparation written by the lab of E. Reynaud. It is a recommended read, especially if the goal is to image different samples than described here.
The image acquisition in LSFM is wide field, camera-based, as opposed to laser scanning confocal microscope. This results in a higher signal-to noise ratio (SNR) for acquired images and can be extremely fast (tens to hundreds of frames per second). The high sensitivity of LSFM further enables imaging of weakly fluorescent samples, like transcription factors expressed at endogenous levels18 or, in the near future, endogenous proteins tagged using CRISPR/Cas9. The high SNR is also important for successful downstream image analysis. The high speed is required not only to capture rapid intracellular processes, but also to image the whole embryo from multiple views fast enough. A seamless fusion of the multiple views can only be achieved, if the observed phenomenon does not change during acquisition of these several z stacks coming from separate views.
The advantages of LSFM do not typically come at the expense of image quality. The lateral resolution of LSFM is slightly worse than the resolution of a confocal microscope. This is because the detection objectives used in LSFM have lower numerical aperture (usually 1.0 or less) compared to 1.2-1.3 of water or silicon immersion objectives on standard confocal setups. Additionally, due to the wide field detection in LSFM (absence of a pinhole), there is more out-of-focus light compared to a confocal microscope. The amount of out-of-focus light is determined by the light sheet thickness. Nevertheless, these disadvantages are compensated by the higher SNR in LSFM. In practice, this results in similar quality images compared to for example spinning disc confocal acquisition15. Consequently, this enables reliable extraction of features like cell membranes or nuclei, e.g., for cell lineage tracing15,19.
The axial resolution of LSFM is determined, in addition to the detection objective, by the light sheet thickness. The axial resolution of LSFM can in some cases surpass the resolution of confocal microscopes. First, the improvement in resolution comes when the light sheet is thinner than the axial resolution of the detection objective, which typically occurs for large specimens imaged with a low magnification objective. The second way, how the LSFM can achieve better axial resolution, is the multiview fusion, in which the high xy resolution information from different views is combined into one image stack. The resulting merged stack has an isotropic resolution approaching the values of resolution in the lateral direction20,21. The strategy for registering the multiple views onto each other described in this article is based on using fluorescent polystyrene beads as fiduciary markers embedded in the agarose around the sample20,21.
As a result of the LSFM commercialization, this technique is now available to a broad community of scientists22. Therefore, the motivation for writing this protocol is to make this technology accessible to developmental biologists lacking practical experience in LSFM and to get these scientists started using this technology with their samples. Our protocol uses the commercial light sheet microscope, which constitutes a conceptually simple microscope that is easy to operate. We would additionally like to mention other recent protocols for imaging zebrafish with home-built LSFM setups, which might be suitable to answer particular questions23-25. Another entry option to LSFM are the open platforms26,27, which use the open access principles to bring light sheet microscopy to a broader community. The documentation of both the hardware and the software aspects can be found at http://openspim.org and https://sites.google.com/site/openspinmicroscopy/.
In this protocol, we use the teleost zebrafish as a model system to study developmental processes with LSFM. Morphogenesis of the zebrafish eye is an example that underlines many of the benefits of LSFM. LSFM has already been used in the past to investigate eye development in medaka28 and in zebrafish29,30. At the early stage of eye development it is complicated to orient the embryo correctly for conventional microscopy, as the bulky yolk does not allow the embryo to lie on the side with its eye facing the objective. However, with LSFM mounting into an agarose column, the sample can be reproducibly positioned. Additionally, during the transition from optic vesicle to the optic cup stage, the eye undergoes major morphogenetic rearrangements accompanied by growth, which requires capturing a large z stack and a big field of view. Also for these challenges LSFM is superior to conventional confocal imaging. The process of optic cup formation is three-dimensional, therefore it is hard to comprehend and visualize solely by imaging from one view. This makes multiview imaging with isotropic resolution beneficial. After optic cup formation, the retina becomes increasingly sensitive to laser exposure. Thus, the low phototoxicity associated with LSFM is a major advantage for long-term imaging.
Here we present an optimized protocol for imaging of one to three days old zebrafish embryos and larvae with focus on the eye development. Our method allows recording of time-lapse movies covering up to 12-14 hr with high spatial and temporal resolution. Importantly, we also show a pipeline for data processing, which is an essential step in LSFM, as this technique invariably generates big datasets, often in the terabyte range.

<table border="0" cellpadding="0" cellspacing="0" width="908">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:387px;">Lightsheet Z.1 microscope</td>
			<td style="width:184px;">Carl Zeiss Microscopy</td>
			<td style="width:119px;"></td>
			<td style="width:219px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Low melting point agarose</td>
			<td>Roth</td>
			<td>6351.1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Low melting point agarose</td>
			<td>Sigma</td>
			<td>A4018 or A9414</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ethyl 3-aminobenzoate methanesulfonate (MESAB/MS-222/Tricaine)</td>
			<td>Sigma</td>
			<td>E10521</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">N-Phenylthiourea (PTU)</td>
			<td>Sigma</td>
			<td>P7629</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">500 nm red fluorescent beads F-Y 050</td>
			<td>Millipore (Estapor)</td>
			<td>80380495</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">20 &#956;l (1mm inner diameter, marked black) capilllaries</td>
			<td>Brand</td>
			<td>701904</td>
			<td>sold as spare part for transferpettor</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Teflon tip plungers for 20 &#956;l capillaries</td>
			<td>Brand</td>
			<td>701932</td>
			<td>sold as spare part for transferpettor</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Circular glass coverslips diameter 18 mm, selected thickness 0.17 mm</td>
			<td>Thermo scientific (Menzel-Glaser)</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">O-rings for chamber windows 17&#215;1.5 mm</td>
			<td>Carl Zeiss Microscopy</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tweezers, style 55</td>
			<td>Dumont</td>
			<td>0209-55-PO</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">50 ml Luer-Lock syringes</td>
			<td>Becton Dickinson</td>
			<td>300865</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">150 cm extension cable for infusion compatible with Luer-Lock syringes</td>
			<td>Becton Dickinson</td>
			<td>397400</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:387px;">Links</td>
			<td style="width:184px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:387px;">Link 1 Multiview reconstruction application</td>
			<td></td>
			<td></td>
			<td>https://github.com/bigdataviewer/SPIM_Registration http://fiji.sc/Multiview-Reconstruction</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:387px;">Link 2 BigDataViewer</td>
			<td></td>
			<td></td>
			<td>https://github.com/bigdataviewer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:387px;">Link 3 BigDataViewer</td>
			<td></td>
			<td></td>
			<td>http://fiji.sc/BigDataViewer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:387px;">Link 4 Multiview reconstruction application-issues</td>
			<td></td>
			<td></td>
			<td>https://github.com/bigdataviewer/SPIM_Registration/issues</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:387px;">Link 5 BigDataViewer-issues</td>
			<td></td>
			<td></td>
			<td>https://github.com/bigdataviewer/bigdataviewer_fiji/issues&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:387px;">Link 6 BigDataServer</td>
			<td></td>
			<td></td>
			<td>http://fiji.sc/BigDataServer.</td>
		</tr>
	</tbody>
</table>

using light sheet fluorescence microscopy to image zebrafish eye development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

LSFM is an ideal method for imaging developmental processes across scales. Several applications are compiled here showing both short and long-term imaging of intracellular structures, as well as cells and entire tissues. These examples also demonstrate that LSFM is a useful tool at various stages of eye development from optic cup formation to neurogenesis in the retina. Movie 1 serves as an illustration of the general LSFM approach, first showing a zoomed out view of an intact embryo in the embedding agarose cylinder in brightfield and later showing a detailed view of the retina in the fluorescence channel.
<img alt="Movie 1" src="/files/ftp_upload/53966/53966movie1.jpg" /> 
Movie 1: LSFM of the zebrafish retina. To illustrate the LSFM approach, this movie shows first in the brightfield that the embryo is imaged intact inside the agarose cylinder before switching to fluorescence. Later it is shown that the big field of view enables observation of the whole retina. Next, the movie shows a magnified small region of the retina to highlight the good subcellular resolution. An Ath5:gap-GFP37 transgenic zebrafish embryo was used for imaging. This transgene labels different neurons in the retina (mainly ganglion cells and photoreceptor precursors). The fluorescence part of the movie was captured as a single view recording with dual sided illumination using the 40X/1.0 W objective with 5 min intervals. The maximum intensity projection of a 30 &#181;m thick volume is shown. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53966/Movie1.mp4" target="_blank">Please click here to download this file.</a>
Movie 2 demonstrates, how very fast intracellular events can be captured with high resolution; in this case the growth of microtubules at their plus ends in the retinal neuronal progenitor cells. The information contained in the movie allows for tracking and quantification of microtubule plus end growth.
<img alt="Movie 2" src="/files/ftp_upload/53966/53966movie2.jpg" /> 
Movie 2: Microtubule dynamics in a single cell. This movie captures growing plus tips of microtubules labeled by the plus tip marker protein EB3-GFP38. The protein is expressed in a single retinal progenitor cell. The microtubules are growing predominantly in the direction from apical to basal (top to bottom). The average speed of the EB3 comets was measured as 0.28 &#177;0.05 &#181;m/sec. The bright spot at the apical side of the cell exhibiting high microtubule nucleation activity is the centrosome. The wild type embryo was injected with hsp70:EB3-GFP plasmid DNA. The movie was acquired 4 hr after heat shock (15 min at 37 &#176;C) at around 28 hrpost fertilization (hpf) as a single view recording with single sided illumination using the 63X/1.0 W objective and 1 sec time intervals. The single cell was cropped from a field of view covering the majority of the retina. The maximum intensity projection of two slices is shown. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53966/Movie2.mp4" target="_blank">Please click here to download this file.</a>
Figure 3 shows, how intracellular structures can be followed over many hours. Here, the centrosome within translocating retinal ganglion cells (RGCs) is captured.
<img alt="Figure 3" src="/files/ftp_upload/53966/53966fig3.jpg" /> 
Figure 3: Centrosome localization during the retinal ganglion cell translocation. This montage of a time-lapse experiment shows the position of the centrosome throughout the retinal ganglion cell (RGC) maturation. In the neuronal progenitors the centrosome is localized in the very tip of the apical process (1:00). During cell division, the two centrosomes serve as poles for the mitotic spindle (2:25). This division results in one daughter cell that differentiates into an RGC and a second daughter cell that becomes a photoreceptor cell precursor. After division, the cell body of the RGC translocates to the basal side of the retina, while the apical process remains attached at the apical side. Once the RGC reaches the basal side, its apical process detaches and the centrosome travels with it (6:15). The centrosome can be followed while gradually retracting together with the apical process (6:50, 7:20, 8:10). In the last frame (8:55) the ganglion cell is growing an axon from its basal side, while the centrosome is still localized apically. The mosaic expression was achieved by plasmid DNA injection into wild type embryo at one cell stage. The cells are visualized by the Ath5:GFP-caax (green) construct, which labels RGCs and other neurons. The centrosomes (arrowheads) are labeled by Centrin-tdTomato29 expression (magenta). The apical side of the retina is at the top of the image and the basal side at the bottom. The maximum intensity projection of a 30 &#181;m thick volume is shown. The images were cropped from a movie covering the whole retina. The movie starts at around 34 hr post fertilization (hpf). A z stack was acquired every 5 min with the 40X/1.0 W objective. Time is shown in hh:mm. Scale bar represents 10 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53966/53966fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In Figure 4 it is displayed, how single cell behavior can be extracted from data capturing the whole tissue such as in Movie 1. Translocation of an RGC can be easily tracked and its apical and basal processes followed.
<img alt="Figure 4" src="/files/ftp_upload/53966/53966fig4.jpg" /> 
Figure 4: Translocation of a single retinal ganglion cell. The RGC translocation from apical to basal side of the retina occurs after the terminal mitosis as described in Figure 3. The RGC is labeled by expression of the Ath5:gap-GFP37 transgene. The maximum intensity projection of a 30 &#181;m thick volume is shown. The images were cropped from a movie covering the whole retina. The movie starts at around 34 hpf. A z stack was acquired every 5 min with the 40X/1.0 W objective. Time is shown in hh:mm. Scale bar represents 5 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53966/53966fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Figure 5 demonstrates the ability of multiview LSFM to capture tissue scale morphogenetic processes with cellular resolution on the example of optic cup morphogenesis, during which optic vesicle transforms into an optic cup. The image quality can be vastly improved by multiview imaging, when the final image is combined from the information out of 5 different views (in this case) into one z stack with isotropic resolution. This figure illustrates the improvement in image quality after weighted-average fusion and further gain in image contrast and resolution after multiview deconvolution. The figure shows two optical slices in different orientations through the dataset. Additionally, a montage from the deconvolved dataset shows the optic cup morphogenesis over time. All the time points are then shown in Movie 3.

<img alt="Figure 5" src="/files/ftp_upload/53966/53966fig5.jpg" /> 
Figure 5: Comparison of image quality between single view and two methods of multiview fusion. (A) One optical slice of single view data shown from lateral view and (B) dorsal view. Stripe artifacts and degradation of the signal deeper inside the sample are apparent. Also a part of the image visible in the fused data (C-F) was not captured in this particular view. (C) The same optical slice, now as multiview fused data shown from lateral view and (D) dorsal view. Note that stripe artifacts are suppressed and structures deep in the sample are better resolved. (E) The same optical slice now as multiview deconvolved data shown from lateral view and (F) dorsal view. Note the increased contrast and resolution so the individual cell membranes and nuclei can be well distinguished. The resolution does not deteriorate notably deeper inside the sample. The dataset was acquired with dual sided illumination from 5 views approximately 20 degrees apart. The z stacks of about 100 &#181;m with 1.5 &#181;m step size were acquired at each view in 10 min intervals for 10 hours with the 20X/1.0 W objective. Input images for the multiview fusion and deconvolution were down sampled 2&#215; to speed up the image processing. 15 iterations of the multiview deconvolution were run. (G) The montage shows a cropped area of the dorsal view from the deconvolved data to highlight the morphogenetic events during early eye development from the optic vesicle to the optic cup stage. The two layers of the optic vesicle, which are initially similar columnar epithelia, differentiate into distinct cell populations. The distal layer close to the epidermis becomes the retinal neuroepithelium (RN) and the proximal layer close to the neural tube becomes the retinal pigment epithelium (RP). The cells in the RN elongate and invaginate to form the optic cup (1:40 to 5:00); at the same time the RP cells flatten. The surface ectoderm is induced to form a lens (1:40), which invaginates later (5:00). The movie starts at 17 hpf. Time is shown in hh:mm. All the cellular membranes are labeled by the &#946;-actin:ras-GFP transgene and all the nuclei are labeled by the hsp70:H2B-RFP transgene. Scale bar represents 30 &#181;m. FB forebrain, LE lens, OP olfactory placode, RN retinal neuroepithelium, RP retinal pigment epithelium. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53966/53966fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Movie 3" src="/files/ftp_upload/53966/53966movie3.jpg" /> 
Movie 3: Optic cup morphogenesis shown with single view and two methods of multiview fusion. The time-lapse movie illustrates the complete process of optic cup morphogenesis from the optic vesicle to the optic cup stage. It shows a single optical slice from the lateral view (top) and a single optical slice from the dorsal view (bottom). The cells of the developing optic vesicle undergo complex rearrangements to finally form the hemispherical optic cup with the inner retinal neuroepithelium and outer retinal pigment epithelium. A lens is formed from the surface ectoderm. It invaginates along with the neuroepithelium and sits in the optic cup. All cellular membranes are labeled by expression of the &#946;-actin:ras-GFP (green) transgene and the nuclei are labeled with hsp70:H2B-RFP (magenta). The movie starts at around 17 hpf. The dataset was acquired with dual sided illumination from 5 views approximately 20 degrees apart and a z stacks of about 100 &#181;m were acquired every 10 min with the 20X/1.0 W objective. Time is shown in hh:mm. Scale bar represents 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53966/Movie3-final.mp4" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development at JoVE.com

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

using-light-sheet-fluorescence-microscopy-to-image-zebrafish-eye

Research

JoVE Journal

Developmental Biology

33.8K Views.  Max Planck Institute of Molecular Cell Biology and Genetics. Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Video: Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.
This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. ...

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and scanning electron microscopy. The sub-diffraction limit resolution capabilities of super-resolution light microscopes allow improving the accuracy of the correlated data. Briefly, 110 nanometer thick cryo-sections are transferred to a silicon wafer and, after immunofluorescence staining, are imaged by super-resolution light microscopy. Subsequently, the sections are preserved in methylcellulose and platinum shadowed prior to imaging in a scanning electron microscope (SEM). The images from these two microscopy modalities are easily merged using tissue landmarks with open source software. Here we describe the adapted method for the larval zebrafish retina. However, this method is also applicable to other types of tissues and organisms. We demonstrate that the complementary information obtained by this correlation is able to resolve the expression of mitochondrial proteins in relation with the membranes and cristae of mitochondria as well as to other compartments of the cell.

This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to ...

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous system can be observed, with single cell resolution, in vivo. Here, we present an integrated protocol for the examination of neurodevelopment in C. elegans embryos. Our protocol combines imaging, lineaging and neuroanatomical tracing of single cells in developing embryos. We achieve long-term, four-dimensional (4D) imaging of living C. elegans&#160;embryos with nearly isotropic spatial resolution through the use of Dual-view Inverted Selective Plane Illumination Microscopy (diSPIM). Nuclei and neuronal structures in the nematode embryos are imaged and isotropically fused to yield images with resolution of ~330 nm in all three dimensions. These minute-by-minute high-resolution 4D data sets are then analyzed to correlate definitive cell-lineage identities with gene expression and morphological dynamics at single-cell and subcellular levels of detail. Our protocol is structured to enable modular implementation of each of the described steps and enhance studies on embryogenesis, gene expression, or neurodevelopment.

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous ...

Three-dimensional Angiogenesis Assay System using Co-culture Spheroids Formed by Endothelial Colony Forming Cells and Mesenchymal Stem Cells

Studies in the field of angiogenesis have been aggressively growing in the last few decades with the recognition that angiogenesis is a hallmark of more than 50 different pathological conditions, such as rheumatoid arthritis, oculopathy, cardiovascular diseases, and tumor metastasis. During angiogenesis drug development, it is crucial to use in vitro assay systems with appropriate cell types and proper conditions to reflect the physiologic angiogenesis process. To overcome limitations of current in vitro angiogenesis assay systems using mainly endothelial cells, we developed a 3-dimensional (3D) co-culture spheroid sprouting assay system. Co-culture spheroids were produced by two human vascular cell precursors, endothelial colony forming cells (ECFCs) and mesenchymal stem cells (MSCs) with a ratio of 5 to 1. ECFCs+MSCs spheroids were embedded into type I collagen matrix to mimic the in vivo extracellular environment. A real-time cell recorder was utilized to continuously observe the progression of angiogenic sprouting from spheroids for 24 h. Live cell fluorescent labeling technique was also applied to tract the localization of each cell type during sprout formation. Angiogenic potential was quantified by counting the number of sprouts and measuring the cumulative length of sprouts generated from the individual spheroids. Five randomly-selected spheroids were analyzed per experimental group. Comparison experiments demonstrated that ECFCs+MSCs spheroids showed greater sprout number and cumulative sprout length compared with ECFCs-only spheroids. Bevacizumab, an FDA-approved angiogenesis inhibitor, was tested with the newly-developed co-culture spheroid assay system to verify its potential to screen anti-angiogenic drugs. The IC50 value for ECFCs+MSCs spheroids compared to the ECFCs-only spheroids was closer to the effective plasma concentration of bevacizumab obtained from the xenograft tumor mouse model. The present study suggests that the 3D ECFCs+MSCs spheroid angiogenesis assay system is relevant to physiologic angiogenesis, and can predict an effective plasma concentration in advance of animal experiments.

Three-dimensional co-culture spheroid angiogenesis assay system is designed to mimic the physiologic angiogenesis. Co-culture spheroids are formed by two human vascular cell precursors, ECFCs and MSCs, and embedded in collagen gel. The new system is effective for evaluating angiogenic modulators, and provides more relevant information to the in vivo study.

Studies in the field of angiogenesis have been aggressively growing in the last few decades with the recognition that angiogenesis is a hallmark of more ...

Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval liver size in zebrafish HCC models provides a means to rapidly assess the effects of drugs and other manipulations on an oncogene-related phenotype. Here we show how to fix zebrafish larvae, dissect the tissues surrounding the liver, photograph livers using bright-field microscopy, measure liver area, and analyze results. This protocol enables rapid, precise quantification of liver size. As this method involves measuring liver area, it may underestimate differences in liver volume, and complementary methodologies are required to differentiate between changes in cell size and changes in cell number. The dissection technique described herein is an excellent tool to visualize the liver, gut, and pancreas in their natural positions for myriad downstream applications including immunofluorescence staining and in situ hybridization. The described strategy for quantifying larval liver size is applicable to many aspects of liver development and regeneration.

Here we demonstrate a method for quantifying liver size in larval zebrafish, providing a way to assess the effects of genetic and pharmacologic manipulations on liver growth and development.

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval ...

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

4D microscopy is an invaluable tool for unraveling the embryonic developmental process in different animals. Over the last decades, Caenorhabditis elegans has emerged as one of the best models for studying development. From an optical point of view, its size and transparent body make this nematode an ideal specimen for DIC (Differential Interference Contrast or Nomarski) microscopy. This article illustrates a protocol for growing C. elegans nematodes, preparing and mounting their embryos, performing 4D microscopy and cell lineage tracing. The method is based on multifocal time-lapse records of Nomarski images and analysis with specific software. This technique reveals embryonic developmental dynamics at the cellular level. Any embryonic defect in mutants, such as problems in spindle orientation, cell migration, apoptosis or cell fate specification, can be efficiently detected and scored. Virtually every single cell of the embryo can be followed up to the moment the embryo begins to move. Tracing the complete cell lineage of a C. elegans embryo by 4D DIC microscopy is laborious, but the use of specific software greatly facilitates this task. In addition, this technique is easy to implement in the lab. 4D microscopy is a versatile tool and opens the possibility of performing an unparalleled analysis of embryonic development.

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

Here, we present a protocol for preparing and mounting Caenorhabditis elegans embryos, recording development under a 4D microscope and tracing cell lineage.

4D microscopy is an invaluable tool for unraveling the embryonic developmental process in different animals. Over the last decades, Caenorhabditis elegans ...

Visualizing Ocular Morphogenesis by Lightsheet Microscopy Using rx3:GFP Transgenic Zebrafish

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&#160;protocol is&#160;provided&#160;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.

Here, a&#160;protocol is&#160;provided&#160;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.&#160;

Vertebrate eye development is a complex process that begins near the end of embryo gastrulation and requires the precise coordination of cell migration, ...