Name: microstructured devices for optimized microinjection and imaging of zebrafish larvae
Uploaded: 2017-12-08T13:00:00.000+00:00
Duration: 7 min 40 s
Description: Watch this Scientific Journal Video about Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae at JoVE.com

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 conidia or human cancer cells, which require the use of a larger bore needle for delivery. Injection into the otic vesicle (Figure 4) presents a particular challenge, as embryos often roll upon contact with the needle. We find that use of microstructured channels that orient and stabilize the zebrafish in a lateral orientation greatly improved both the throughput and accuracy of this technique in our hands.
For rapid imaging of laterally oriented zebrafish at different time points, such as assessment of neutrophil recruitment or xenograft metastasis, avoiding agarose increased mounting consistency between embryos and reduced chances of damage during post-imaging rescue. These devices also show promise for extended time lapse imaging, and have been used to successfully capture multichannel, multiplane images with minimal fish movement over 12 h. Because they are not constrained by agarose, active elongation of the larvae during this period (~300 &#181;m from 54-78 hpf)11 is unrestricted and can proceed normally.
The most critical steps in the preparation and use of these tools are common to the use of most PDMS based microfluidic devices: devices must be thoroughly wetted and bubbles avoided. To avoid such issues, wet the devices in E3 immediately after they are fabricated, while the surface hydrophilicity conferred by oxygen plasma treatment is still present. Re-treatment with oxygen plasma can also be used to transiently restore hydrophilicity to aged PDMS devices.
The precise geometries used here take advantage of the high resolution of photolithographic techniques, but the current designs limit use to zebrafish during the second day post-fertilization. Re-design of geometries based on the current platform would allow successful orientation of 1 dpf and 3 dpf for microinjection. Orientation of more buoyant 4 dpf larvae with inflated swim bladders would likely be more challenging, and require snug geometries in combination with higher water tension. Complex devices integrating flow or vacuum might provide an alternative approach applicable to a wider range of developmental stages, but would also require use of pumps and tubing, increasing cost and risk of device failure.
PDMS is a cost-effective, useful material for fabricating devices like those described here, providing biocompatibility, transparency, and reusability. One drawback of using PDMS devices for zebrafish is hydrophobicity, which can make maintenance of shallow layers of E3 on the surface of the device challenging. This problem can be avoided by casting the microinjection devices in agarose, although this limits the reusability and increases the fragility of devices. A preferred alternative would be identification of a biocompatible surface treatment that increases the hydrophilicity of PDMS12, or use of an appropriate plastic substrate.
Current methods for microinjection of zebrafish at 2 dpf use either water tension4 or simple channels cast in agarose using commercial molds to immobilize and position the larvae. These approaches provide limited control of orientation and can be complicated by rapid drying of embryos. The microstructures integrated into the divots and channels used here are designed to orient and immobilize the larvae without depending entirely on water tension, thus reducing the risk of drying. Divots made using 3D printed molds have previously been used for orientation of embryos for imaging purposes13, although the depth of these divots made them inaccessible for microinjection.
Use of microfluidic systems for zebrafish imaging is becoming increasingly common14. 
Multiple flow-through systems have been developed to allow observation of zebrafish development over time with the ability to introduce compounds on demand15,16,17. More complex devices have been designed to array embryos while still within their chorion, which provides a simple spherical geometry that can be easily manipulated with relatively high throughput in these systems18,19. Several groups have developed platforms for arraying and imaging larval stage zebrafish in various orientations6,20, the most user-friendly being the passive micropump driven &#34;ZEBRA&#34; system developed by the Beebe lab6. Unlike the ZEBRA system, the device that we describe here uses suction, generated using a standard transfer pipette, to draw the larvae into the channels, while parallel rather than sequential loading of larvae allows rescue of individual larvae post-imaging by applying positive flow through the same port. Furthermore, our approach requires a smaller footprint, so PDMS devices can be bonded to conventional glass-bottom 6-well plates for imaging.
The current experimental design utilizes separate devices for microinjection and imaging primarily so that they can be used with conventional microinjection equipment and standard inverted microscopes. Microfluidic approaches for automated microinjection of zebrafish eggs already exist21, and in combination with &#34;fish-trap&#34; type methods for highly accurate arraying and orientation of larvae20, it is foreseeable that devices integrating automated larval injection and imaging may also one day become a reality.

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 Polydimethylsiloxane (PDMS)</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 zebrafish</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. 11 Scalpel</td><td>Fine Science Tools</td><td>10011-00</td><td>For cutting PDMS</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</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.remotexs.ntu.edu.sg/10.6084/m9.figshare.4282853 and on the ZFIN community protocols wiki page: https://wiki.zfin.org/display/prot/ZFIN+ Protocol+Wiki &amp;nbsp;</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 extend their application.
We demonstrate use of these devices for challenging microinjection techniques and convenient imaging. The otic vesicle (Figure 4D, white dashed outline) provides a useful, immune-privileged and isolated site for testing neutrophil recruitment in zebrafish9. To test the ability of zebrafish neutrophils to respond to standard chemoattractants, 100 nM of fMLP and LTB4 were microinjected into the otic vesicles of 2 dpf zebrafish embryos (Figure 4F, J), using the microstructured channel device (Figure 1G) to stabilize embryos in the lateral orientation. Injections were traced with high molecular weight Rhodamine dextran, and performed in Tg(mpx:EGFP) embryos to facilitate visualization of neutrophil recruitment. Uninjected embryos (Figure 4D, H) and embryos injected with Rhodamine alone (Figure 4E, I) were used as negative controls. Following microinjection, embryos were transferred into the imaging channel device (Figure 1A, B) and imaged using an EVOS fluorescent microscope (Figure 4C) at 30 min and 5 h post injection (Figure 4D-G and Figure 4H-K, respectively). As expected, a small number of fluorescent neutrophils were recruited to the mock-injected otic vesicle at the early timepoint (Figure 4I), likely due to damage-mediated recruitment. Interestingly, the Rhodamine dextran tracer was observed to accumulate in the otoliths during the experiment. For both chemoattractants (Figure 4F, J), neutrophils were recruited in higher numbers, particularly in response to LTB4 (Figure 4F).
The representative results shown here demonstrate the successful use of this method for the microinjection and imaging of zebrafish. In the context of zebrafish neutrophil recruitment assays, application of these tools in combination with detailed live imaging techniques and sophisticated cell migration analysis10 may provide a useful platform for future experiments in this field.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56498/56498fig1.jpg" /> 
Figure 1: Computer assisted drawing (CAD) design of photolithography masks. (A-C) CAD design for microstructured surface divots for positioning in lateral (A), ventral (B), and dorsal (C) orientations. Different colored lines represent CAD mask designs for different features, with green 100 &#181;m, blue 200 &#181;m, and red 400 &#181;m feature heights. Scale bar = 500 &#181;m. (D-F) CAD design for microstructured channels for positioning in lateral (D), ventral (E), and dorsal (F) orientations. Different colored lines represent mask designs for different features, with green 100 &#181;m, blue 200 &#181;m, and red 400 &#181;m feature heights. Scale bar: 500 &#181;m. (G-I) CAD design for imaging devices designed for loading head-first (G) or tail-first (H and I). Blue lines represent 400 &#181;m features, while red lines represent 600 &#181;m features. Arrows show the direction in which larvae are loaded. Scale bar = 200 &#181;m. This figure has been modified from Ellett et al.5 <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56498/56498fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56498/56498fig2.jpg" /> 
Figure 2: Silicone master wafer. (A) Top section of master wafer shows features for orienting individual larvae into divots. (B) Bottom section of master wafer showing design for microstructured channels. This figure has been modified from Ellett et al.5 <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56498/56498fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56498/56498fig3.jpg" /> 
Figure 3: Microstructured PDMS devices. (A) PDMS block cast from master wafer showing divots for orienting individual larvae. (B) PDMS block cast from master wafer showing microstructured channels. This figure has been modified from Ellett et al.5 <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56498/56498fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56498/56498fig4.jpg" /> 
Figure 4: Representative results: Imaging recruitment of zebrafish neutrophils following microinjection delivery of chemoattractants into the otic vesicle. (A-B) Loading and mounting embryos for imaging. (A) The head-first device with embryos ready for loading, with the port indicated (black arrow). (B) The loaded embryos after they have been drawn into position (yellow arrow). (C) 6-well glass-bottom plate with imaging devices on microscope. This format allows imaging of 4 laterally oriented embryos per well (24 embryos total), using a standard inverted microscope. (D-G) Neutrophils (EGFP-green) in an uninjected (D) embryo and following otic vesicle microinjection of Rhodamine dextran (red, open white arrowhead) alone (E), 100 nM LTB4 (F), and 100 nM fMLP (G) 30 min post injection (mpi). (H-K) Neutrophils (EGFP-green) in an uninjected (H) embryo and 5 h following otic vesicle microinjection of Rhodamine dextran (red, open white arrowhead) alone (I), 100 nM LTB4 (J), and 100 nM fMLP (K) 5 h post injection (hpi). Scale bar = 200 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56498/56498fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

9.5K Views.  Massachusetts General Hospital–Harvard Medical School–Shriners Burns Hospital. Microinjection van zebravis embryo's en larven is een essentieel maar uitdagende techniek gebruikt in vele zebrafish modellen. Hier presenteren we een scala van microscale instrumenten om te helpen bij de stabilisatie en de oriëntatie van de zebravis voor zowel microinjection en beeldbewerking.

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

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

Bioengineering

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 ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression studies. In zebrafish (Danio rerio), microinjection of RNA, DNA, proteins, antisense oligonucleotides and other small molecules into the developing embryo provides researchers a quick and robust assay for exploring gene function in vivo. In this video-article, we will demonstrate how to prepare and microinject in vitro synthesized EGFP mRNA and a translational-blocking morpholino oligo against pkd2, a gene associated with autosomal dominant polycystic kidney disease (ADPKD), into 1-cell stage zebrafish embryos. We will then analyze the success of the mRNA and morpholino microinjections by verifying GFP expression and phenotype analysis. Broad applications of this technique include generating transgenic animals and germ-line chimeras, cell-fate mapping and gene screening. Herein we describe a protocol for overexpression of EGFP and knockdown of pkd2 by mRNA and morpholino oligonucleotide injection.

Microinjection is a well-established and effective method for introducing foreign substances into fertilized zebrafish embryos. Here, we demonstrate a robust microinjection technique for performing mRNA overexpression, and morpholino oligonucleotide gene knockdown studies in zebrafish.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression ...

Biology

Microinjection of Zebrafish Embryos to Analyze Gene Function

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic oligonucleotides with antisense complementarity to target RNAs, can be added to the embryo to reduce the expression of a particular gene product.  Conversely, processed mRNA can be added to the embryo to increase levels of a gene product.  The vehicle for adding either mRNA or morpholino to an embryo is microinjection.  Microinjection is efficient and rapid, allowing for the injection of hundreds of embryos per hour.  This video shows all the steps involved in microinjection.  Briefly, eggs are collected immediately after being laid and lined up against a microscope slide in a Petri dish.  Next, a fine-tipped needle loaded with injection material is connected to a microinjector and an air source, and the microinjector controls are adjusted to produce a desirable injection volume.  Finally, the needle is plunged into the embryo's yolk and the morpholino or mRNA is expelled.

This video shows how morpholino or mRNA can be injected into zebrafish embryos at the one-cell stage to decrease or increase the level of specific gene products during subsequent development.

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Micromanipulation of Gene Expression in the Adult Zebrafish Brain Using Cerebroventricular Microinjection of Morpholino Oligonucleotides

Manipulation of gene expression in tissues is required to perform functional studies. In this paper, we demonstrate the cerebroventricular microinjection (CVMI) technique as a means to modulate gene expression in the adult zebrafish brain. By using CVMI, substances can be administered into the cerebroventricular fluid and be thoroughly distributed along the rostrocaudal axis of the brain. We particularly focus on the use of antisense morpholino oligonucleotides, which are potent tools for knocking down gene expression in vivo. In our method, when applied, morpholino molecules are taken up by the cells lining the ventricular surface. These cells include the radial glial cells, which act as neurogenic progenitors. Therefore, knocking down gene expression in the radial glial cells is of utmost importance to analyze the widespread neurogenesis response in zebrafish, and also would provide insight into how vertebrates could sustain adult neurogenesis response. Such an understanding would also help the efforts for clinical applications in human neurodegenerative disorders and central nervous system regeneration. Thus, we present the cerebroventricular microinjection method as a quick and efficient way to alter gene expression and neurogenesis response in the adult zebrafish forebrain. We also provide troubleshooting tips and other useful information on how to carry out the CVMI procedure.

In this article, we demonstrate a method for manipulation of gene expression in the ventricular cells of the adult zebrafish telencephalon using antisense morpholino oligonucleotides. We present this method as an efficient and quick protocol that can be used for functional studies in the adult vertebrate brain.

Manipulation of gene expression in tissues is  required to perform functional studies. In this paper, we demonstrate the  cerebroventricular ...

Neuroscience

Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for live high-resolution microscopic imaging of dynamic biological phenomena in space and time with cellular resolution. However, the traditional method of agarose encapsulation for live imaging can impede larval development and tissue regrowth. Therefore, this manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which was designed and fabricated as a functionally compartmentalized device to orient larvae for high-resolution microscopy while permitting caudal fin transection within the device and subsequent unrestrained tail development and re-growth. This device allows for wounding and long-term imaging while maintaining viability. Given that the zWEDGI mold is 3D printed, the customizability of its geometries make it easily modified for diverse zebrafish imaging applications. Furthermore, the zWEDGI offers numerous benefits, such as access to the larva during experimentation for wounding or for the application of reagents, paralleled orientation of multiple larvae for streamlined imaging, and reusability of the device.

This manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which is a compartmentalized device designed to orient and restrain zebrafish larvae. The design permits tail transection and long-term collection of high-resolution fluorescent microscopy images of wound healing and regeneration.

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for ...

4-Dimensional Imaging of Zebrafish Optic Cup Morphogenesis

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of visual impairment, yet mechanisms of eye morphogenesis are still poorly understood. The basic organization of the embryonic eye is conserved throughout vertebrates, thus live imaging of zebrafish embryos has become a powerful approach to directly observe eye development at real time under normal and pathological conditions. Dynamic cell processes including movements, morphologies, interactions, division, and death can be visualized in the embryo. We have developed methods for uniform labeling of subcellular structures and timelapse confocal microscopy of early eye development in zebrafish. This protocol outlines the method of generating capped mRNA for injection into the 1-cell zebrafish embryo, mounting embryos at optic vesicle stage (~12 hours post fertilization, hpf), and performing multi-dimensional timelapse imaging of optic cup morphogenesis on a laser scanning confocal microscope, such that multiple datasets are acquired sequentially in the same imaging session. Such an approach yields data that can be used for a variety of purposes, including cell tracking, volume measurements, three-dimensional (3D) rendering, and visualization. Our approaches allow us to pinpoint the cellular and molecular mechanisms driving optic cup development, in both wild type and genetic mutant conditions. These methods can be employed directly by other groups or adapted to visualize many additional aspects of zebrafish eye development.

This protocol describes an approach for in toto labeling and multidimensional imaging of zebrafish early eye development. We describe labeling, embedding, and four dimensional (4D) imaging using laser scanning confocal microscopy, and considerations for optimizing acquisition of datasets for dissecting mechanisms of optic cup morphogenesis.

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of ...

Low-cost Polyethylene Terephthalate Lamination Microfluidics Designs for Multiplexed Zebrafish Imaging

Zebrafish embryos are transparent and thus uniquely suited for noninvasive intravital imaging of fundamental processes, such as wound healing and immune cell migration. Microfluidic devices are used for entrapment to support long-term imaging of multicellular organisms, including zebrafish. However, the fabrication of these devices using soft lithography requires specialized facilities and competency in 3D printing, which may not be accessible to every lab. Our adaptation of a previously developed low-cost polyethylene terephthalate lamination method for constructing microfluidic devices increases accessibility by enabling design fabrication and iteration for a fraction of the technical investment of conventional techniques. We use a device made with this method, the Rotational Assistant for Danio Imaging of Subsequent Healing (RADISH), to accommodate drug treatment, manual wounding, and long-term imaging of up to four embryos in the same field of view. With this new design, we successfully capture gross morphological characteristics of the calcium&#160;signal around laser ablation and manual transection wounds for multiple embryos in the 2 h immediately following injury, as well as neutrophil recruitment to the wound edge for 24 h.

An innovative method for fabricating microfluidic devices using polyethylene terephthalate (PET) lamination significantly reduces the cost and complexity of entrapping and imaging multiple live zebrafish embryos.

Zebrafish embryos are transparent and thus uniquely suited for noninvasive intravital imaging of fundamental processes, such as wound healing and immune ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

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