The authors would like to thank David Langenau for generously providing aquarium space; Eric Stone, John C. Moore and Qin Tang for help with zebrafish maintenance and reagents, and Anne Robertson and Elliott Hagedorn from Leonard Zon&#39;s lab for procuring the zebrafish strain used here. They would also like to thank Octavio Hurtado for advice on photolithographic techniques. FE was funded by Fellowships from Shriner&#39;s Hospital for Children and the American Australian Association. This work was funded by NIH grant GM92804.

Microinjection of larvae was approved by the Massachusetts General Hospital Subcommittee on Research Animal Care under Protocol 2011N000127.
1. Device Fabrication
NOTE: All computer assisted drawing (CAD) files used to design photolithography masks described here (Figure 1) are available for download. See Table of Materials for links.
<ol>
	<li>Fabricate the master mold wafer in a Class 1,000 clean room using standard photolithographic techniques7. For the microstructure arrays and channels, pattern the epoxy-based negative photoresist sequentially in 3 layers using 3 separate masks, according to the manufacturer&#39;s instructions: 100 &#181;m for the shallow features, 200 &#181;m for the medium features, and 400 &#181;m for the deep features. For the imaging device, pattern two layers using 400 &#181;m for the shallow features and 600 &#181;m for the deep features.</li>
	<li>Tape the completed wafer to the base of a 15 cm Petri dish for casting (Figure 2).</li>
</ol>
2. Preparation of Microstructured Surface Arrays - PDMS
<ol>
	<li>To make usable devices, cast polydimethylsiloxane (PDMS), using the master wafer as a mold. Combine 50 g of PDMS monomer with 5 g of initiator and mix thoroughly in a plastic weighing dish with a plastic fork.</li>
	<li>Pour mixture carefully over the mold.</li>
	<li>Degas in a vacuum desiccator at 16-25 inHg (54-85 kPa) for 1 h.</li>
	<li>To cure the PDMS, bake at 65 &#176;C for at least 3 h. The PDMS should cure to a firm but flexible solid.</li>
	<li>Carefully cut around the device on the master wafer using a scalpel, making sure to maintain contact between the tip of the scalpel and the wafer surface. Using forceps, peel the PDMS replica away from the wafer and transfer to a clean 10 cm Petri dish, feature side up (Figure 3).</li>
	<li>Treat the device with oxygen plasma using a plasma oven for 35 s to reduce hydrophobicity.</li>
	<li>Cover with zebrafish embryo medium (E3) containing 168 mg/L of ethyl 3-aminobenoate methanesulfonate (MS-222, pH 7.5). Remove any bubbles from deeper features by flowing water over the surface of the device using a transfer pipette.</li>
</ol>
3. Preparation of Microstructured Surface Arrays - Agarose
<ol>
	<li>To make a negative PDMS mold, first treat a PDMS device with oxygen plasma.</li>
	<li>Treat the PDMS device with 1H,1H,2H,2H-Perfluorooctyltriethoxysilane (PFDTS) silane vapor to create an inert surface8. Briefly:
	<ol>
		<li>Place the PDMS to be treated feature-side-up in a vacuum chamber within a fume hood.</li>
		<li>Next to the PDMS, place a small open glass or aluminum dish, and carefully pipette 200 &#181;L of PFDTS (98% silane) into the dish. 
		Caution: PFDTS silane is highly volatile, reacts violently with water, is flammable, and causes severe skin and eye damage on contact. Read MSDS in detail before use.</li>
		<li>Close the vacuum chamber and apply vacuum for 15-30 min, or until the PFDTS silane is completely evaporated.</li>
	</ol>
	</li>
	<li>Place the device in a Petri dish and use as a mold for fresh PDMS, prepared as described in step 2.1-2.3.</li>
	<li>Bake at 65 &#176;C for at least 3 h, then peel the entire layer of fresh PDMS from the treated device and place in a fresh Petri dish. This can then be used as a negative mold to cast agarose.</li>
	<li>To prepare agarose, boil a 2% solution of low gelling temperature agarose in E3 in a microwave until the agarose is fully dissolved.</li>
	<li>Pour the hot agarose over the PDMS negative mold and remove any bubbles in the agarose covering the device by flowing hot agarose over the features with a transfer pipette.</li>
	<li>Once bubbles are removed, refrigerate at 4 &#176;C to set the agarose.</li>
	<li>Remove the agarose block from the negative PDMS mold by deforming the PDMS from underneath by hand, transfer the agarose block to a new Petri dish, and wet with E3 containing MS-222.</li>
</ol>
4. Preparation of PDMS Imaging Devices
<ol>
	<li>To make usable devices, cast PDMS, using the master wafer as a mold. Combine 50 g of PDMS monomer with 5 g of initiator (same as used in step 2.1) and mix thoroughly in a plastic weighing dish with a plastic fork.</li>
	<li>Pour mixture carefully over the mold.</li>
	<li>Degas in a vacuum desiccator at 16-25 inHg (54-85 kPa) for 1 h.</li>
	<li>To cure the PDMS, bake at 65 &#176;C for at least 3 h. The PDMS should cure to a firm but flexible solid.</li>
	<li>Cut the devices from the Master Wafer and punch out ports using a 1.5 mm punch.</li>
	<li>Treat the devices and a 6-well glass-bottom well plate with oxygen plasma for 35 s, as described in step 2.6.</li>
	<li>Bond one device feature-side-down to each glass coverslip of the 6-well plate by placing it on an 85 &#176;C hotplate for 10 min. 
	NOTE: To confirm successful bonding, apply firm pressure to one side of the bonded device using forceps. The device should remain attached to the glass coverslip.</li>
</ol>
5. Zebrafish Culture
<ol>
	<li>Culture zebrafish embryos to 2 dpf using standard techniques3.</li>
	<li>Dechorionate embryos using forceps or 1 mg/mL protease mix (see Materials Table)3 at least 30-60 mins prior to loading onto device, to allow them to straighten.</li>
	<li>Anaesthetize embryos using (168 mg/L) MS-222 (pH 7.5) by adding 1 mL of 25X stock solution to the E3 in their Petri dish.</li>
</ol>
6. Orientation of Zebrafish on Microstructured Surface - Divot Arrays
<ol>
	<li>Prepare the PDMS block from step 2.7 (Figure 3A) in its Petri dish such that it is covered by a thin (1-2 mm) layer of E3, which allows easier manipulation of the embryos.</li>
	<li>Transfer required number of embryos (generally 10-20 per condition) onto the surface of the PDMS block using a transfer pipette.</li>
	<li>Using a micromanipulation tool (hair loop or similar), push the embryos into the microstructured divots (Figure 3A). Use of divots is particularly suited to dorsal and ventral orientations. 
	NOTE: For the divots with a tighter fit (dorsal and ventral), try positioning the embryos on the surface of the block in parallel to the divot, and then rolling them into position using a hair loop. Pushing them deeper into the divot will help to keep them stable.</li>
</ol>
7. Orientation of Zebrafish on Microstructured Surface - Microstructured Channels
<ol>
	<li>Draw E3 off the PDMS block patterned with microstructured channels (Figure 3B) using a transfer pipette until the level of E3 drops below the edge of the block, but is still present in the reservoir and channels and in a thin layer on the PDMS.</li>
	<li>Transfer 10 embryos from step 5.3 into the reservoir using a transfer pipette, being careful not to overflow the edges.</li>
	<li>Using a hair loop, manipulate each embryo into the funnel at the entrance of the appropriate channel and orient such that the head of the embryo is toward the channel and the embryo has the appropriate orientation as it enters the channel.</li>
	<li>Slide each embryo down the channel using a hair loop until it reaches the orienting microfeatures in the channel walls that help to keep it in place. Use of microstructured channels is particularly recommended for the lateral orientation. 
	NOTE: To reduce embryo sliding during microinjection, try adjusting the amount of surface tension by drawing E3 from the reservoir.</li>
</ol>
8. Microinjection of Zebrafish
<ol>
	<li>Prepare microinjection needle.
	<ol>
		<li>Make borosilicate glass microcapillary needles using a micropipette puller as previously described4. The resulting needles should be tapered to a point at one end, with a taper length of approximately 10 mm.</li>
		<li>Prepare solution to be injected, in this case 100 nM chemoattractant (N-Formylmethionine-leucyl-phenylalanine (fMLP) or Leukotriene B4 (LTB4)) and 100 nM of 70 kDa Rhodamine dextran tracer in PBS.</li>
		<li>Load 5 &#181;L of solution into the needle using a finely tapered pipette tip (see materials table).</li>
		<li>Mount the needle into a micromanipulator mounted on a magnetic stand and connected to a picopump microinjector.</li>
	</ol>
	</li>
	<li>Break the tip of needle at a 45 &#176; angle by pinching the tapered tip with forceps.</li>
	<li>Adjust injection bolus size to 1 nL by adjusting the injection time on the picopump controller.</li>
	<li>Adjust the angle of the microinjection needle. A needle angle of 45 &#176; on the micromanipulation controller is generally a good starting point, with the needle parallel to the length of the embryo. A steeper angle may be used to minimize lateral movement of the embryo during microinjection.</li>
	<li>Controlling the Petri dish with the left hand and the needle micromanipulator with the right, bring the needle as close as possible to the intended site of microinjection, in this case the otic vesicle. 
	NOTE: The otic vesicle can be identified as the oblong organ posterior to the eye containing two smaller distinct dark oblong structures (otoliths).</li>
	<li>Using the micromanipulation controller, penetrate the target tissue (in this case the otic vesicle) such that the tip of the needle is within the vesicle, and inject the preset volume using the picopump foot switch. 
	NOTE: If the tissue resists penetration, try gently tapping the micromanipulation controller knob.</li>
	<li>To release the embryos, flow E3 over the surface from top to bottom using a transfer pipette, or by swirling the dish such that the embryos are released into the surrounding E3.</li>
	<li>Transfer the embryos to a recovery dish of E3 without MS-222 using a transfer pipette.</li>
</ol>
9. Screening of Embryos Using the PDMS Imaging Device
<ol>
	<li>Prime the device from step 4.7 by flowing E3 (+ MS-222) through the port. This can be done using either a 1,000 &#181;L pipette or a narrow-tip transfer pipette.</li>
	<li>Fill the well with E3 + MS-222 until the device is covered.</li>
	<li>Deliver 4 injected embryos from step 8.7 into each well using a transfer pipette. Embryos can be pre-anesthetized if desired by incubating in E3 + MS-222 for 1-2 min prior to loading (step 5.3).</li>
	<li>Using a micromanipulation tool (hair loop or similar), orient the embryos at the entrances of the imaging channels in the desired orientation (tail-first or head-first, depending on the device used), and push them partially into the device (Figure 4A). This will help draw them into the device evenly.</li>
	<li>Draw E3 through the device from the port using either a 1,000 &#181;L pipette or transfer pipette until the embryos are drawn into the channels in the correct orientation for imaging (Figure 4B). 
	NOTE: Be careful not to suck the embryos past the trapping point, this will damage the embryos and alter their orientation.</li>
	<li>Transfer well-plate to microscope for imaging (Figure 4C).</li>
	<li>Image using an inverted fluorescent microscope.
	<ol>
		<li>Adjust magnification by selecting the appropriate lens. 10X is usually a useful magnification for imaging zebrafish neutrophil migration.</li>
		<li>Adjust light source intensity and exposure time for each channel so that each signal is clear, but not saturated.</li>
		<li>Capture images for each channel. Here, we used green fluorescent protein (GFP, Ex: 470/22, Em: 525/50), Texas Red (TxRed, Ex: 585/29, Em: 628/32) and Transmitted channels. Multichannel images can be combined during post-processing (Figure 4B-I).</li>
	</ol>
	</li>
</ol>

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The authors declare no conflicts of interest.

Here, we describe the use of devices we recently developed to facilitate 2 dpf zebrafish microinjection5, and introduce a simple agarose-free mounting device for convenient imaging of embryos. These tools highlight the utility of photolithographic techniques for fabrication of devices useful for zebrafish techniques.
We have found MSA devices particularly useful for injection of cells or particles prone to aggregation within the microinjection needle, such as fungal con...

Zebrafish have emerged as a powerful model for many fields, from studies of fundamental developmental biology to large-scale genetic and chemical screens1,2. Routine genetic manipulations, such as gene overexpression, knockdown, CRISPR/Cas9 mutagenesis, and transgenesis rely on microinjection of genetic material into the single-cell zygote, which has led to the development of simple, easy-to-use, commercially available tools for orienting and stabilizing eggs for injection3. Other approaches, such as transplantation and infection, often require microinjection into later stage embryos and larvae using larger gauge capillary needles4. However, use of larger gauge needles presents significant technical challenges, as it is more difficult to penetrate the target tissue without pushing or rolling the embryo. Under these conditions, obtaining the appropriate water tension required to stabilize the embryo while avoiding drying during the procedure is difficult, and embryos may not be ideally oriented for injection into the target tissue.
Following microinjection, it is often useful to screen injected embryos to select those that have been successfully injected, and to capture images of the initial time point. To address these challenges, we have developed a range of microstructured devices that help to stabilize 2 dpf embryos in various orientations both for microinjection5, and for rapid image-based screening post-injection.
To obtain sufficient structural resolution in these devices, we utilized photolithographic techniques. Commonly used in microelectronic industries and more recently extrapolated to microfluidic fabrication, these approaches can achieve vertical structures ranging from 1-1,000 &#181;m, a scale well suited to manipulation of zebrafish embryos and larvae. All devices were fabricated using polydimethylsiloxane (PDMS), which is cheap, physically robust, biologically inert, and transparent.
Microstructured surface arrays (MSAs) were formatted as blocks of PDMS with a patterned top surface, analogous to the simple channels in agarose blocks commonly used for egg microinjection. For post-injection screening, 6 imaging devices can be arrayed in a standard glass-bottomed 6-well plate. These devices are designed for easy loading of embryos, while the unloading procedure conveniently allows rescue of specific embryos, facilitating image-based screening approaches in a more user-friendly manner than those devices previously developed by the Beebe laboratory6.

<table>
	<tbody>
		<tr>
			<td>Dow Corning Sylgard 184&#160; Polydimethylsiloxane (PDMS)&#160;</td>
			<td>Ellsworth Adhsives</td>
			<td>184 SIL ELAST KIT 0.5KG</td>
			<td>For casting the devices. Kit includes PDMS monomer and Initiator</td>
		</tr>
		<tr>
			<td>Low gelling temperature agarose</td>
			<td>Sigma Aldrich</td>
			<td>A9414-10G</td>
			<td>For casting agarose devices</td>
		</tr>
		<tr>
			<td>PFDTS silane</td>
			<td>Sigma Aldrich</td>
			<td>448931-10G</td>
			<td>For casting of negative PDMS molds</td>
		</tr>
		<tr>
			<td>Tricaine (MS-222)</td>
			<td>Sigma Aldrich</td>
			<td>E10521-10G</td>
			<td>To anesthetize&#160; zebrafish&#160;</td>
		</tr>
		<tr>
			<td>Rhodamine Dextran 70,000 Da</td>
			<td>ThermoFisher</td>
			<td>D1818</td>
			<td>To trace microinjections</td>
		</tr>
		<tr>
			<td>Leukotriene B4 (LTB4)</td>
			<td>Cayman Chemicals</td>
			<td>20110</td>
			<td>Neutrophil chemoattractant</td>
		</tr>
		<tr>
			<td>N-Formylmethionine-leucyl-phenylalanine (fMLP)</td>
			<td>Sigma Aldrich</td>
			<td>F3506-50MG</td>
			<td>Neutrophil chemoattractant</td>
		</tr>
		<tr>
			<td>15 cm Petri dish</td>
			<td>Fisher scientific</td>
			<td>08-757-148</td>
			<td>For Casting from the master wafer</td>
		</tr>
		<tr>
			<td>Glass-bottom 6-well plates</td>
			<td>MatTek</td>
			<td>P06G-0-20-F</td>
			<td>For imaging devices</td>
		</tr>
		<tr>
			<td>Borosilicate glass microcapillaries</td>
			<td>World Scientific Instruments</td>
			<td>TW-100-4</td>
			<td>For microinjection needles</td>
		</tr>
		<tr>
			<td>Transfer pipettes</td>
			<td>Sigma Aldrich</td>
			<td>Z350796</td>
			<td>For transferring zebrafish embryos</td>
		</tr>
		<tr>
			<td>Microloader tips</td>
			<td>Fisher scientific</td>
			<td>E5242956003</td>
			<td>For loading the microinjection needles</td>
		</tr>
		<tr>
			<td>Harris Uni-Core 1.5 mm punch</td>
			<td>Ted Pella Inc.</td>
			<td>15111-15</td>
			<td>To punch ports in PDMS imaging devices</td>
		</tr>
		<tr>
			<td>No.&#160;11 Scalpel</td>
			<td>Fine Science Tools</td>
			<td>10011-00</td>
			<td>For cutting PDMS&#160;</td>
		</tr>
		<tr>
			<td>Dumont No. 5 Forceps</td>
			<td>Fine Science Tools</td>
			<td>11252-10</td>
			<td>For dechorionating embryos and breaking microinjection needle tips</td>
		</tr>
		<tr>
			<td>Marzhauser Micromanipulator</td>
			<td>ASI&#160;</td>
			<td>MM33-R</td>
			<td>For manipulating microinjection needle</td>
		</tr>
		<tr>
			<td>Magnetic stand</td>
			<td>MSC</td>
			<td>SPI - 87242624</td>
			<td>For mounting micromanipulator</td>
		</tr>
		<tr>
			<td>MPPI-3 Picopump controller</td>
			<td>ASI</td>
			<td>MPPI-3</td>
			<td>To control microinjection volume and timing</td>
		</tr>
		<tr>
			<td>EVOS inverted fluorescent microscope</td>
			<td>ThermoFisher</td>
			<td>EVOS FL</td>
			<td>To image injected embryos</td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td>Nikon</td>
			<td>SMZ745</td>
			<td>For visualizing microinjecion</td>
		</tr>
		<tr>
			<td>AutoCAD software</td>
			<td>Autodesk</td>
			<td></td>
			<td>Download AutoCAD files from: https://dx.doi.org/10.6084/m9.figshare.4282853 and on the ZFIN community protocols wiki page: https://wiki.zfin.org/display/prot/ZFIN+ Protocol+Wiki &#160;</td>
		</tr>
	</tbody>
</table>

microstructured devices for optimized microinjection and imaging of zebrafish larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

The approach described here demonstrates the design (Figure 1) and fabrication of devices for use with 2 dpf zebrafish, using photolithographic (Figure 2) and soft-lithographic (Figure 3) techniques. This method allows rapid testing of many design iterations and modifications, and alterations and optimization of microstructure dimensions for use with zebrafish at other stages of development may exten...

Watch this Scientific Journal Video about Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae at JoVE.com

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

microstructured-devices-for-optimized-microinjection-imaging

Research

JoVE Journal

Developmental Biology

9.5K Views.  Massachusetts General Hospital–Harvard Medical School–Shriners Burns Hospital. Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Video: Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Tracking Cells in GFP-transgenic Zebrafish Using the Photoconvertible PSmOrange System

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing developmental processes as they happen in the living animal. Cells of interest can be labeled by using a tissue specific promoter to drive the expression of a fluorescent protein (FP) for the generation of transgenic lines. Using fluorescent photoconvertible proteins for this purpose additionally allows to precisely follow defined structures within the expression domain. Illuminating the protein in the region of interest, changes its emission spectrum and highlights a particular cell or cell cluster leaving other transgenic cells in their original color. A major limitation is the lack of known promoters for a large number of tissues in the zebrafish. Conversely, gene- and enhancer trap screens have generated enormous transgenic resources discretely labeling literally all embryonic structures mostly with GFP or to a lesser extend red or yellow FPs. An approach to follow defined structures in such transgenic backgrounds would be to additionally introduce a ubiquitous photoconvertible protein, which could be converted in the cell(s) of interest. However, the photoconvertible proteins available involve a green and/or less frequently a red emission state1 and can therefore often not be used to track cells in the FP-background of existing transgenic lines. To circumvent this problem, we have established the PSmOrange system for the zebrafish2,3. Simple microinjection of synthetic mRNA encoding a nuclear form of this protein labels all cell nuclei with orange/red fluorescence. Upon targeted photoconversion of the protein, it switches its emission spectrum to far red. The quantum efficiency and stability of the protein makes PSmOrange a superb cell-tracking tool for zebrafish and possibly other teleost species.

We established the photoconvertible PSmOrange system as a powerful, straight-forward and cost inexpensive tool for in vivo cell tracking in GFP transgenic backgrounds. This protocol describes its application in the zebrafish model system.

The rapid development of transparent zebrafish embryos (Danio rerio) in combination with fluorescent labelings of cells and tissues allows visualizing ...

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.

Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...

In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

The zebrafish forms two kidney structures in its lifetime. The pronephros (embryonic kidney) forms during embryonic development and begins to function at 2 days post fertilization. Consisting of only two nephrons, the pronephros serves as the sole kidney during larval life until more renal function is required due to the increasing body mass. To cope with this higher demand, the mesonephros (adult kidney) begins to form during metamorphosis. The new primary nephrons fuse to the pronephros and form connected lumens. Then, secondary nephrons fuse to primary ones (and so on) to create a branching network in the mesonephros. The vast majority of research is focused on the pronephros due to the ease of using embryos. Thus, there is a need to develop techniques to study older and larger larvae and juvenile fish to better understand mesonephros development. Here, an in situ hybridization protocol for gene expression analysis is optimized for probe penetration, washing of probes and antibodies, and bleaching of pigments to better visualize the mesonephros. The Tg(lhx1a-EGFP) transgenic line is used to label progenitor cells and the distal tubules of nascent nephrons. This protocol fills a gap in mesonephros research. It is a crucial model for understanding how new kidney tissues form and integrate with existing nephrons and provide insights into regenerative therapies.

In Situ Hybridization in Zebrafish Larvae and Juveniles during Mesonephros Development

The zebrafish is an important model for understanding kidney development. Here, an in situ hybridization protocol is optimized to detect gene expression in zebrafish larvae and juveniles during mesonephros development.

The zebrafish forms two kidney structures in its lifetime. The pronephros (embryonic kidney) forms during embryonic development and begins to function at ...

Using Alizarin Red Staining to Detect Chemically Induced Bone Loss in Zebrafish Larvae

Chemically induced bone loss due to lead (Pb) exposure could trigger an array of adverse impacts on both human and animal skeletal systems. However, the specific effects and mechanisms in zebrafish remain unclear. Alizarin red has a high affinity for calcium ions and can help visualize the bone and illustrate skeletal mineral mass. In this study, we aimed to detect lead acetate (PbAc)-induced bone loss in zebrafish larvae by using alizarin red staining. Zebrafish embryos were treated with a series of PbAc concentrations (0, 5, 10, 20 mg/L) between 2 and 120 h post fertilization. Whole-mount skeletal staining was conducted on larvae at 9 days post fertilization, and the total stained area was quantified using ImageJ software. The results indicated that the mineralized tissues were stained in red, and the stained area decreased significantly in the PbAc-exposure group, with a dose-dependent change in bone mineralization. This paper presents a staining protocol for investigating skeletal changes in PbAc-induced bone defects. The method can also be used in zebrafish larvae for the detection of bone loss induced by other chemicals.

Here, we have used alizarin red staining to show that lead acetate exposure causes a bone mass change in zebrafish larvae. This staining method can be adapted to the investigation of bone loss in zebrafish larvae loss induced by other hazardous toxicants.

Chemically induced bone loss due to lead (Pb) exposure could trigger an array of adverse impacts on both human and animal skeletal systems. However, the ...

Genetic Modification of Primate Cerebral Organoids

Targeted Microinjection and Electroporation of Primate Cerebral Organoids for Genetic Modification

The cerebral cortex is the outermost brain structure and is responsible for the processing of sensory input and motor output; it is seen as the seat of higher-order cognitive abilities in mammals, in particular, primates. Studying gene functions in primate brains is challenging due to technical and ethical reasons, but the establishment of the brain organoid technology has enabled the study of brain development in traditional primate models (e.g., rhesus macaque and common marmoset), as well as in previously experimentally inaccessible primate species (e.g., great apes), in an ethically justifiable and less technically demanding system. Moreover, human brain organoids allow the advanced investigation of neurodevelopmental and neurological disorders.
As brain organoids recapitulate many processes of brain development, they also represent a powerful tool to identify differences in, and to functionally compare, the genetic determinants underlying the brain development of various species in an evolutionary context. A great advantage of using organoids is the possibility to introduce genetic modifications, which permits the testing of gene functions. However, the introduction of such modifications is laborious and expensive. This paper describes a fast and cost-efficient approach to genetically modify cell populations within the ventricle-like structures of primate cerebral organoids, a subtype of brain organoids. This method combines a modified protocol for the reliable generation of cerebral organoids from human-, chimpanzee-, rhesus macaque-, and common marmoset-derived induced pluripotent stem cells (iPSCs) with a microinjection and electroporation approach. This provides an effective tool for the study of neurodevelopmental and evolutionary processes that can also be applied for disease modeling.

Author Spotlight: Targeted Microinjection and Electroporation of Primate Cerebral Organoids for Genetic Modification  

The electroporation of primate cerebral organoids provides a precise and efficient approach to introduce transient genetic modification(s) into different progenitor types and neurons in a model system close to primate (patho)physiological neocortex development. This allows the study of neurodevelopmental and evolutionary processes and can also be applied for disease modeling.

The cerebral cortex is the outermost brain structure and is responsible for the processing of sensory input and motor output; it is seen as the seat of ...