Name: light sheet-based fluorescence microscopy of living or fixed and stained tribolium castaneum embryos
Uploaded: 2017-04-28T15:00:00
Description: Watch this Scientific Journal Video about Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos at JoVE.com

We thank Sven Plath for technical support. The Glia-blue transgenic line was a kind gift from Gregor Bucher (G&#246;ttingen, Germany). The research was funded by the Cluster of Excellence Frankfurt am Main for Macromolecular Complexes (CEF-MC, EXC 115, speaker Volker D&#246;tsch) granted in part to EHKS at the Buchmann Institute for Molecular Life Sciences (BMLS, director Enrico Schleiff) at the Goethe Universit&#228;t Frankfurt am Main by the Deutsche Forschungsgemeinschaft (DFG).

1. Husbandry of Tribolium Cultures
NOTE: Standard conditions are defined as an incubation temperature of 25 &#176;C and 70% relative humidity in a 12 h bright/12 h dark cycle. For more information on Tribolium husbandry, respective guidelines are available64. This protocol requires two different flour-based media, which can be prepared in kilogram quantities and stored for several months.
<ol>
	<li>Prior to working with flour and inactive dry yeast, ...

Quality control
In live imaging assays, the preparation and recording procedure must be non-invasive, i.e. neither the mechanical and chemical handling (collection, dechorionation, mounting onto the sample holder) nor the integrated energy load during the observation should affect the viability of the specimen. For studies that characterize WT development, it is recommended to only use data from experiments in which the embryo survives the recording process, is retrie...

The red flour beetle Tribolium castaneum, which belongs to the large family of darkling beetles (Tenebrionidae), has a long history within the agricultural and life sciences and is the second best studied model insect model organism after the fruit fly Drosophila melanogaster. During the last four decades, it became a powerful and popular insect model organism in developmental genetics, in evolutionary developmental biology and, during the last twenty years, in embryonic morphogenesis for a variety of reasons:
Drosophila and Tribolium both belong to the Holometabola, but diverged approximately 300 million years ago1,2,3,4. While the embryonic development of Drosophila is commonly considered as highly derived, Tribolium shows a more ancestral mode of development that is found in a considerably larger proportion of insect species5,6,7,8,9. Firstly, Tribolium exhibits non-involuted head development, i.e. its mouthparts and antennae emerge already during embryogenesis10,11,12,13,14,15. Secondly, Tribolium follows the principles of short-germ development, i.e. abdominal segments are added sequentially from a posterior growth zone during germband elongation16,17,18,19. Thirdly, Tribolium develops and later degrades two extra-embryonic membranes i.e. the amnion, which covers the embryo only ventrally, and the serosa, which envelops the embryo completely20,21,22. Both membranes play a crucial morphogenetic23 as well as protective role against microorganisms24,25 and desiccation26. Fourthly, the embryonically developing legs are fully functional during the larval life stage and serve as the primordia for the adult legs during pupal metamorphosis27,28,29,30,31.
Due to their small size and modest demands, cultivation of Tribolium in the laboratory is fairly straightforward. Cultures of wild-type (WT) strains or transgenic lines typically consist of around 100-300 adults and can be kept within one-liter glass bottles (footprint 80 cm2) filled three to four centimeters high (about 50 g) with growth medium that consists of full grain wheat flour supplemented with inactive dry yeast. A water supply is not necessary. This allows even small laboratories to keep dozens of beetle cultures within small- or medium-sized commercially available insect incubators. Later developmental stages of Tribolium (larvae after approximately the fourth instar, pupae and adults) are easily separated from the growth medium by sieving. Synchronized embryos are obtained by incubating adults for short periods on egg-laying medium. For rapid development, beetle cultures are kept at 32 &#176;C (about four weeks per generation), while stock keeping is typically performed at 22-25 &#176;C (about ten weeks per generation).
Within the last decade, many standard techniques have been gradually adapted and optimized for Tribolium, as summarized in the Emerging Model Organisms books32. Of great importance are advanced genetic methods such as embryonic33, larval34,35 or parental36,37 RNA interference-based gene knockdown, germline transformation with either the piggyBac38,39 or the Minos40 transposase system and CRISPR/Cas9-based genome engineering41. Furthermore, the Tribolium genome has been sequenced about a decade ago42, and is now in the third round of genome assembly release43, which allows efficient and genome-wide identification and systematic analysis of genes44 or other genetic elements45,46. Additionally, the genomes of four other coleopteran species are available for comparative genetic approaches47,48,49,50. In association with the sequenced genome, two large-scale genetic analyses have been performed, i.e. an insertional mutagenesis screen51 and a systematic RNA interference-based gene knockdown screen52,53.
Fluorescence live imaging with widefield, confocal or light sheet-based microscopy (LSFM) allows to observe the embryonic morphology of Tribolium as a function of time (i.e. the morphogenesis) in a multi-dimensional context (Table 1). In widefield and confocal fluorescence microscopy, the excitation and emission light is guided through the same objective lens. In both approaches, the entire specimen is illuminated for every recorded two-dimensional plane. Hence, the specimens are subjected to very high energy levels. In LSFM, only the fluorophores in the focal plane are excited due to a decoupling of illumination and detection by using two perpendicularly arranged objective lenses (Figure 1). LSFM comes in two canonic implementations &#8211; the single plane illumination microscope (SPIM) and the digital scanned laser light sheet-based fluorescence microscope (DSLM, Figure 2) &#8211; and offers several crucial advantages over traditional approaches: (i) intrinsic optical sectioning capability, (ii) good axial resolution, (iii) strongly reduced level of photo-bleaching, (iv) very low photo-toxicity, (v) high signal-to-noise ratio, (vi) relatively high acquisition speed, (vii) imaging along multiple directions and (viii) deeper tissue penetration due to the usage of low numerical aperture illumination objective lenses54,55,56.
LSFM has already been successfully applied in Tribolium to document nearly the entire embryonic morphogenesis57 and to analyze the principles of extra-embryonic membrane rupture at the beginning of dorsal closure23. To raise the attractiveness of LSFM in the Tribolium community and for insect science in general, it is of great importance to establish standard operating procedures and to improve the methods, protocols and the pool of resources to a level where the microscope becomes an ease-of-use standard tool in developmental biology laboratories, and the biological questions stay in the center of attention.
This protocol begins with the basics of Tribolium cultivation, i.e. maintenance, reproduction and embryo collection. Next, two experimental strategies are illustrated: (i) live imaging of custom-made transgenic lines and (ii) imaging of fixed embryos that were stained with fluorescent dyes (Table 2). Subsequently, three mounting techniques with slightly different purposes are explained in detail (Figure 3 and Table 3): (i) the agarose column, (ii) the agarose hemisphere and (iii) the novel cobweb holder. The protocol then explains the data acquisition procedure with LSFM. Imaging modalities and key considerations are outlined. Finally, embryo retrieval is explained and suggestions for basic data processing are provided. In the representative results, live imaging data from two novel custom-made and the Glia-blue58 transgenic lines are shown and the respective imaging datasets are provided as a downloadable resource. Additionally, image data of fixed embryos that were stained with a variety of fluorescence dyes are presented. The discussion focuses on quality control, current limitations of the live imaging approach and the adaptation of the protocol to other species.
The protocol is written for light sheet-based fluorescence microscopes that are equipped with a sample chamber and a rotatable clamp mechanism for standardized sample holders54,59,60, which are typically cylinder-shaped elements made of metal, plastic or glass with a diameter in the millimeter range. The protocol is also suitable for both canonic implementations, i.e. SPIM and DSLM, as well as for setups with two or more illumination and detection arms61,62,63. The representative results show data in two spectral channels, green (illumination with a 488 nm laser, detection through a 525/50 bandpass filter) and red (illumination with a 561 nm laser, detection through a 607/70 bandpass filter), but the protocol can be expanded to three or four spectral channels.

<table border="0" cellpadding="0" cellspacing="0" width="573">
	<tbody>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">full grain wheat flour</td>
			<td style="width:119px;">Demeter e.V.</td>
			<td style="width:71px;">1.13E+08</td>
			<td style="width:167px;">US: whole wheat flour, UK: whole meal flour</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">405 fine wheat flour</td>
			<td style="width:119px;">Demeter e.V.</td>
			<td style="width:71px;">1.13E+08</td>
			<td style="width:167px;">US: pastry flour, UK: soft flour</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">inactive dry yeast</td>
			<td style="width:119px;">Flystuff / Genesee Scientific</td>
			<td style="width:71px;">62-106</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">phosphate-buffered saline (PBS), pH 7.4</td>
			<td style="width:119px;">Thermo Fisher Scientific</td>
			<td style="width:71px;">10010-023</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">sodium hypochlorite, ~12% active Cl</td>
			<td style="width:119px;">Sigma Aldrich</td>
			<td style="width:71px;">425044-250ML</td>
			<td style="width:167px;">Caution: sodium hypochlorite is corrosive</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">low-melt agarose</td>
			<td style="width:119px;">Carl Roth</td>
			<td style="width:71px;">6351.2</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">6-well plate</td>
			<td style="width:119px;">Orange Scientific</td>
			<td style="width:71px;">4430500</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">24-well plate</td>
			<td style="width:119px;">Orange Scientific</td>
			<td style="width:71px;">4430300</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">glass capillaries, internal &#216; 0.46 mm</td>
			<td style="width:119px;">Brand GmbH + Co KG</td>
			<td style="width:71px;">7087 09</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">SYTOX Green</td>
			<td style="width:119px;">Thermo Fisher Scientific</td>
			<td style="width:71px;">57020</td>
			<td style="width:167px;">Staining solution preparation is explained in Table 2</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">YOYO-1 Iodide</td>
			<td style="width:119px;">Thermo Fisher Scientific</td>
			<td style="width:71px;">Y3601</td>
			<td style="width:167px;">Staining solution preparation is explained in Table 2</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">BOBO-3 Iodide</td>
			<td style="width:119px;">Thermo Fisher Scientific</td>
			<td style="width:71px;">B3586</td>
			<td style="width:167px;">Staining solution preparation is explained in Table 2</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">Alexa Fluor 488 Phalloidin</td>
			<td style="width:119px;">Thermo Fisher Scientific</td>
			<td style="width:71px;">A12379</td>
			<td style="width:167px;">Staining solution preparation is explained in Table 2</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">Alexa Fluor 546 Phalloidin</td>
			<td style="width:119px;">Thermo Fisher Scientific</td>
			<td style="width:71px;">A22283</td>
			<td style="width:167px;">Staining solution preparation is explained in Table 2</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">sieve, 800 &#181;m mesh size</td>
			<td style="width:119px;">VWR International</td>
			<td style="width:71px;">200.025.222-051</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">sieve, 710 &#181;m mesh size</td>
			<td style="width:119px;">VWR International</td>
			<td style="width:71px;">200.025.222-050</td>
			<td style="width:167px;">for growth medium preparation (step 1.1)</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">sieve, 300 &#181;m mesh size</td>
			<td style="width:119px;">VWR International</td>
			<td style="width:71px;">200.025.222-040</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">sieve, 250 &#181;m mesh size</td>
			<td style="width:119px;">VWR International</td>
			<td style="width:71px;">200.025.222-038</td>
			<td style="width:167px;">for egg laying medium preparation (step 1.2)</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">glass dish, &#216; 100 mm &#215; 20 mm</td>
			<td style="width:119px;">Sigma Aldrich</td>
			<td style="width:71px;">CLS70165102</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:216px;">cell strainer, 100 &#181;m mesh size</td>
			<td style="width:119px;">BD Biosciences</td>
			<td style="width:71px;">352360</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">paint brush, head &#216; 2 mm</td>
			<td style="width:119px;">VWR International</td>
			<td style="width:71px;">149-2121</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">syringe, 1.0 ml</td>
			<td style="width:119px;">B. Braun Medical AG</td>
			<td style="width:71px;">9166017V</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">scintillation vials</td>
			<td style="width:119px;">Sigma Aldrich</td>
			<td style="width:71px;">M1152-1000EA</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:216px;">paraformaldehyde</td>
			<td style="width:119px;">Sigma Aldrich</td>
			<td style="width:71px;">158127</td>
			<td style="width:167px;">Caution: paraformaldehyde is toxic and corrosive</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">n-heptane &#8805; 99%</td>
			<td style="width:119px;">Carl Roth</td>
			<td style="width:71px;">8654.1</td>
			<td style="width:167px;">Caution: n-heptane is flammable and toxic</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:216px;">Triton X-100</td>
			<td style="width:119px;">Sigma Aldrich</td>
			<td style="width:71px;">X100-100ML</td>
			<td style="width:167px;">Caution: Trition X-100 is corrosive</td>
		</tr>
	</tbody>
</table>

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.

This protocol describes an experimental framework for fluorescence imaging of living or fixed and stained Tribolium embryos with LSFM. Due to the low levels of photo-bleaching and photo-toxicity, a direct consequence of its optical sectioning capability, LSFM is particularly well suited for long-term live imaging.
The novel AGOC{ATub&#39;H2B-mEmerald} #1 transgenic line expresses a histone2B-mEmerald fusion protein unde...

Watch this Scientific Journal Video about Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos at JoVE.com

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.

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 overall goal of this protocol is to observe the development of Tribolium castaneum embryos noninvasively over very long periods of time. Light sheet based fluorescence microscopy helps to answer morphogenesis related questions in the emerging insect model organism Tribolium castaneum addressing cell migration and proliferation patterns from the early blastoderm to the onset of muscular movement. The main advantage of this method is the critical factors such as photobleaching and phototoxicity are kept to a minimum even when the embryos are observed for several days.To mount an embryo into the sample chamber using an Agarose Column first fill a one millimeter syringe with distilled water and use a small rubber hose to attach the syringe to a glass capillary. Fill the capillary half way with distilled water then use a paint brush to transfer an embryo to the inner side of the glass capillary opening, and stick the capillary into the liquid agarose slowly drawing a few microliters of the agarose along with the embryo into the capillary. When the embryo is in place there should be a few microliters of agarose both above and below the embryo.When the agarose has solidified shorten the capillary to a length of about five millimeters. The remaining fragment should be completely filled with agarose and the embryo should be found within the second quartile. Then insert the steel cylinder with the pin into the sample chamber, and stick the capillary fragment on top of the pin, The Agarose Column should should protrude a few millimeters from the region of the capillary fragment that contains the embryo.To mount the embryo into the sample chamber using an Agarose Hemisphere first draw 200 microliters of liquid Agarose into a one millimeter syringe, and use the syringe to fill the steel pipe with the sample chamber with Agarose from the top of the pipe until a hemisphere forms at the bottom opening. When the Agarose has solidified pick up an embryo with the tip of a paint brush and rearrange the embryo so that the anterior end becomes accessible. Dip the Agarose Hemisphere into liquid Agarose to cover the hemisphere with a thin film and place the embryo with its anterior end upright on the pole of the Agarose Hemisphere.Turn the steel pipe and use the brush to correct the position of the embryo as necessary. Ideally less than half of the embryo surface should be covered in agarose and the flanks should be as steep as possible. Too much agarose on the embryo's surface leads to a mechanic constriction and a reduced gas and ion exchange, which has a dramatic effect on the embryo's survival rate.Then slowly insert the steel pipe into the sample chamber. To mount the embryo into the sample chamber using a Cobweb Holder first select an embryo with the paint brush and set the paint brush aside. Next cover the slotted hole of the cobweb holder with five to eight microliters of liquid agarose then remove most of the agarose until only a thin agarose film remains.Place the embryo on to the agarose film and adjust its position carefully moving the embryo to the x-axis center of the slotted hole and aligning its elongated anterior posterior axis with the elongated axis of the slotted hole. Make sure to place the embryo onto the agarose film within 10-15 seconds after its application that is while the agarose is still liquid or else it will not attach properly. Then slowly insert the Cobweb holder into the sample chamber positioning the holder so that it does not interfere with the excitation and emission light.For live imaging assays first configure the time-lapse, then in the transmission light mode position the embryo along the X and Y axis into the center of the field of view without exposing the embryo to the laser. Rotate the embryo 360 degrees in 90 degree steps to examine the tissue for any damage that could have occurred during the mounting process. In the florescence mode define the Z-Stack for each direction adding a 25 to 50 micrometer spacial buffer in front of and behind the embryo and define the Z spacing.For long term imaging ensure that the sample chamber is filled with sufficient imaging buffer and cover the sample chamber opening. Then start the imaging process, monitoring the correct acquisition along all of the directions in all of the channels for all of the embryos. At the end of the imaging period remove the sample holder from the sample chamber and transfer the embryo to an object slide.If an agarose column was used use a sharp blade to extract the embryos from the surrounding agarose. If an agarose hemisphere was used transfer the hemisphere with a flat side to the object slide. For the cobweb holder method use a paint brush to gently dismount the embryo from the agarose film.Then incubate the embryo loaded object slide in a small glass dish in a saturated humidity atmosphere under standard conditions until the larvae hatch. To process the data of the observed embryo open the files of interest in the appropriate imaging analysis software and rotate the Z-Stacks to align the anterior posterior axis of the embryo with the Y axis as necessary. Crop the Z-Stacks along the X and Y axis so that only a minimal background remains then calculate the Z Maximum Projection for each Z-Stack.Then depending on the transgenic line and imaging modalities make the appropriate dynamic intensity adjustments of the time-stacks to correct for any photobleaching or flora for expression fluctuations as necessary. Using a transgenic line that expresses a Histone 2B M-Emerald fusion protein under the control of the alpha-tubublin one promoter 66 hours of embryonic development at room temperature were recorded from germ band retraction to dorsal closure. This enhancer trap-line can be used to visualize neuronal clusters within the head appendages and the dynamics of cirrosa migration during dorsal closure.Imaging a similar transgenic line which carries the same trans-gene but at a different genomic location the optical sections of the transition from gastrulation to germ band elongation show the internal cell arrangement of the emerging germ band. Using the glia-blue transgenic line the acquisition of Z-Stacks along multiple directions via sample rotation allows the observation of glial cell reorganization along and around the ventral nerve cord as well as their proliferation dynamics of the dorsal side of the head lobes in the same embryo. Small florescent dyes can also be used for the specific labeling of intercellular structures to highlight certain embryonic features such as the cirrhosis gar, the posterior ventral cirrhosis cells, or the serosa amnion germ band tissue tri-layer that emerges during the serosa window closure.Dual color florescent dye staining allows the further visualization of two inter-cellular structures in the same embryo. For example the actin cytoskeleton and the nuclear envelope. Once the recording is finished it's important for the quality control of the experiment to retrieve the embryos and raise them to healthy fertile adults.This procedure can be combined with other techniques such as Parental RNA Difference to analyze knocked down phenotypes.

light-sheet-based-fluorescence-microscopy-living-or-fixed-stained

Research

JoVE Journal

Developmental Biology

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 ...

Methods for Precisely Localized Transfer of Cells or DNA into Early Postimplantation Mouse Embryos

Manipulation and culture of early mouse embryos is a powerful yet largely under-utilized technology enhancing the value of this model system. Conversely, cell culture has been widely used in developmental biology studies. However, it is important to determine whether in vitro cultured cells truly represent in vivo cell types. Grafting cells into embryos, followed by an assessment of their contribution during development is a useful method to determine the potential of in vitro cultured cells. In this study, we describe a method for grafting cells into a defined site of early postimplantation mouse embryos, followed by ex vivo culture. We also introduce an optimized electroporation method that uses glass capillaries of known diameter, allowing precise localization and adjustment of the number of cells receiving exogenous DNA with both high transfection efficiency and low cell death. These techniques, which do not require any specialized equipment, render experimental manipulations of the gastrulation and early organogenesis-stage mouse embryo possible, allowing analysis of commitment in cultured cell subpopulations and the effect of genetic manipulations in situ on cell differentiation.

We demonstrate a method for grafting cultured cells into defined sites of early mouse embryos to determine their in vivo potential. We also introduce an optimized electroporation method that uses glass capillaries of known diameter, allowing the precise delivery of exogenous DNA into a few cells in the embryos.

Manipulation and culture of early mouse embryos is a powerful yet largely under-utilized technology enhancing the value of this model system. Conversely, ...

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.

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 ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Quantitative Whole-mount Immunofluorescence Analysis of Cardiac Progenitor Populations in Mouse Embryos

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development and organogenesis are two areas of research that have benefitted greatly from these advances, due to the high quality of data that can be obtained directly from imaging pre-implantation embryos or ex vivo organs. While pre-implantation embryos have yielded data with especially high spatial resolution, later stages have been less amenable to three-dimensional reconstruction. Obtaining high-quality 3D or volumetric data for known embryonic structures in combination with fate mapping or genetic lineage tracing will allow for a more comprehensive analysis of the morphogenetic events taking place during embryogenesis.
This protocol describes a whole-mount immunofluorescence approach that allows for the labeling, visualization, and quantification of progenitor cell populations within the developing cardiac crescent, a key structure formed during heart development. The approach is designed in such a way that both cell- and tissue-level information can be obtained. Using confocal microscopy and image processing, this protocol allows for three-dimensional spatial reconstruction of the cardiac crescent, thereby providing the ability to analyze the localization and organization of specific progenitor populations during this critical phase of heart development. Importantly, the use of reference antibodies allows for successive masking of the cardiac crescent and subsequent quantitative measurements of areas within the crescent. This protocol will not only enable a detailed examination of early heart development, but with adaptations should be applicable to most organ systems in the gastrula to early somite stage mouse embryo.

Here, we describe a protocol for whole mount immunofluorescence and image-based quantitative volumetric analysis of early stage mouse embryos. We present this technique as a powerful approach to qualitatively and quantitatively assess cardiac structures during development, and propose that it may be widely adaptable to other organ systems.

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development ...

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the development of neuronal connectivity. During establishment of the retino-tectal projection, optic axons extend from the eye and navigate through distinct regions of the brain to reach their target tissue, the optic tectum. Once optic axons enter the tectum, they elaborate terminal arbors that function to increase the number of synaptic connections they can make with target interneurons in the tectum. Here, we describe a method to express DNA encoding green fluorescent protein (GFP), and gain- and loss-of-function gene constructs, in optic neurons (retinal ganglion cells) in Xenopus embryos. We explain how to microinject a combined DNA/lipofection reagent into eyebuds of one day old embryos such that exogenous genes are expressed in single or small numbers of optic neurons. By tagging genes with GFP or co-injecting with a GFP plasmid, terminal axonal arbors of individual optic neurons with altered molecular signaling can be imaged directly in brains of intact, living Xenopus tadpoles several days later, and their morphology can be quantified. This protocol allows for determination of cell-autonomous molecular mechanisms that underlie the development of optic axon arborization in vivo.

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

This protocol aims to demonstrate how to microinject a DNA/DOTAP mixture into eyebuds of one day old Xenopus laevis embryos, and how to image and reconstruct individual green fluorescent protein (GFP) expressing optic axonal arbors in tectal midbrains of intact, living Xenopus tadpoles.

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the ...

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

Light-Induced GFP Expression in Zebrafish Embryos using the Optogenetic TAEL/C120 System

Inducible gene expression systems are an invaluable tool for studying biological processes. Optogenetic expression systems can provide precise control over gene expression timing, location, and amplitude using light as the inducing agent. In this protocol, an optogenetic expression system is used to achieve light-inducible gene expression in zebrafish embryos. This system relies on an engineered transcription factor called TAEL based on a naturally occurring light-activated transcription factor from the bacterium E. litoralis. When illuminated with blue light, TAEL dimerizes, binds to its cognate regulatory element called C120, and activates transcription. This protocol uses transgenic zebrafish embryos that express the TAEL transcription factor under the control of the ubiquitous ubb promoter. At the same time, the C120 regulatory element drives the expression of a fluorescent reporter gene (GFP). Using a simple LED panel to deliver activating blue light, induction of GFP expression can first be detected after 30 min of illumination and reaches a peak of more than 130-fold induction after 3 h of light treatment. Expression induction can be assessed by quantitative real-time PCR (qRT-PCR) and by fluorescence microscopy. This method is a versatile and easy-to-use approach for optogenetic gene expression.

Optogenetics is a powerful tool with wide-ranging applications. This protocol demonstrates how to achieve light-inducible gene expression in zebrafish embryos using the blue light-responsive TAEL/C120 system.

Inducible gene expression systems are an invaluable tool for studying biological processes. Optogenetic expression systems can provide precise control ...

Multi-Photon Laser Ablation of Cytoplasmic Microtubule Organizing Centers in Mouse Oocytes

The fidelity of oocyte meiosis is critical for generating developmentally competent euploid eggs. In mammals, the oocyte undergoes a lengthy arrest at prophase I of the first meiotic division. After puberty and upon meiotic resumption, the nuclear membrane disassembles (nuclear envelope breakdown), and the spindle is assembled mainly at the oocyte center. Initial central spindle positioning is essential to protect against abnormal kinetochore-microtubule (MT) attachments and aneuploidy. The centrally positioned spindle migrates in a time-sensitive manner toward the cortex, and this is a necessary process to extrude a tiny polar body. In mitotic cells, spindle positioning relies on the interaction between centrosome-mediated astral MTs and the cell cortex. On the contrary, mouse oocytes lack classic centrosomes and, instead, contain numerous acentriolar MT organizing centers (MTOCs). At the metaphase I stage, mouse oocytes have two different sets of MTOCs: (1) MTOCs that are clustered and sorted to assemble spindle poles (polar MTOCs), and (2) metaphase cytoplasmic MTOCs (mcMTOCs) that remain in the cytoplasm and do not contribute directly to spindle formation but play a crucial role in regulating spindle positioning and timely spindle migration. Here, a multi-photon laser ablation method is described to selectively deplete endogenously labeled mcMTOCs in oocytes collected from Cep192-eGfp reporter mice. This method contributes to the understanding of the molecular mechanisms underlying spindle positioning and migration in mammalian oocytes.

An optimized protocol is presented that enables the depletion of cytoplasmic microtubule organizing centers in mouse oocytes during metaphase I using a near-infrared femtosecond laser.

The fidelity of oocyte meiosis is critical for generating developmentally competent euploid eggs. In mammals, the oocyte undergoes a lengthy arrest at ...

Investigating Development with Optogenetic Signaling Manipulation in Zebrafish

Optogenetic Signaling Activation in Zebrafish Embryos

Signaling pathways orchestrate fundamental biological processes, including development, regeneration, homeostasis, and disease. Methods to experimentally manipulate signaling are required to understand how signaling is interpreted in these wide-ranging contexts. Molecular optogenetic&#160;tools can provide reversible, tunable manipulations of signaling pathway activity with a high degree of spatiotemporal control and have been applied in vitro, ex vivo, and in vivo. These tools couple light-responsive protein domains, such as the blue light homodimerizing light-oxygen-voltage sensing (LOV) domain, with signaling effectors to confer light-dependent experimental control over signaling. This protocol provides practical guidelines for using the LOV-based bone morphogenetic protein (BMP) and Nodal signaling activators bOpto-BMP and bOpto-Nodal in the optically accessible early zebrafish embryo. It describes two control experiments: A quick phenotype assay to determine appropriate experimental conditions, and an immunofluorescence assay to directly assess signaling. Together, these control experiments can help establish a pipeline for using optogenetic tools in early zebrafish embryos. These strategies provide a powerful platform to investigate the roles of signaling in development, health, and physiology.

Author Spotlight: Manipulating Signaling in Zebrafish Embryos to Decode Cell Fate Decisions 

Optogenetic manipulation of signaling pathways can be a powerful strategy to investigate how signaling is decoded in development, regeneration, homeostasis, and disease. This protocol provides practical guidelines for using light-oxygen-voltage sensing domain-based Nodal and bone morphogenic protein (BMP) signaling activators in the early zebrafish embryo.

Signaling pathways orchestrate fundamental biological processes, including development, regeneration, homeostasis, and disease. Methods to experimentally ...