最適化されたマイクロインジェクションおよびゼブラフィッシュ幼虫の画像処理用微細構造デバイス

The overall goal of this methodology is to make microinjection of zerbrafish larvae easier and more accurate. This method helps researchers using zebrafish model systems to immobilize zebrafish larvae in specific orientations in large arrays. The technique allows for easier access to specific tissues for microinjection and imaging, and is very effective when large numbers of larvae are required for applications like infection or xenograft cancer models.

My fabrication strategies in zebrafish models are highly complementary. One is man-made and simple. The other one is live and complex.

Both share the same size scale. They are both transparent. And both enable observations at single-cell resolution.

As you show here, they can be easily combined, and they could have applications to the study of virtually any major disease. After preparing a PDMS block in a Petri dish covered in a thin film of E-3, according to the text protocol, use a transfer pipet to transfer the required number of embryos onto the surface of the PDMS block. Using a hair loop or similar tool, push the embryos into the microstructure divots, particularly for dorsal and ventral orientations.

When loading larvae into the divots, it's important to have enough water covering the PDMS block to allow easy manipulation. It's also important to push them into the divots so that they remain in place during the injection. To orient zebrafish on microstructure channels previously prepared according to the text protocol, use a transfer pipet to draw E-3 off the patterned PDMS block.

The level of E-3 should drop below the edge of the block, but still be present in the reservoir and channels, and a thin layer remains on the PDMS. Using a transfer pipet, transfer 10 anesthetized embryos into the reservoir, being careful not to overflow the edges. Using a hair loop, manipulate each embryo into the funnel head, first at the entrance of the appropriate channel, ensuring that the embryo has the appropriate orientation as it enters the channel.

Then, use a hair loop to slide each embryo down the channel, until it reaches the orienting microfeatures in the channel walls that keep it in place. Following the preparation of microinjection needles and microinjection solution, use a finely tapered pipet tip to load five microliters of the solution into the needle. Mount the needle into a micromanipulator mounted on a magnetic stand, and connect it to a PICO pump microinjector.

Then, use forceps to break the tip of a needle at a 45 degree angle by pinching the tapered tip. Adjust the injection bolus size to one nanoliter by adjusting the injection time on the PICO pump controller. Adjust the angle of the microinjection needle on the micromanipulation controller, beginning with 45 degrees with the needle parallel to the length of the embryo.

A steeper angle may be used to minimize lateral movement of the embryo during the microinjection. Controlling the Petri dish with the left hand, and the need micromanipulator with the right, bring the needle as close as possible to the intended site of microinjection, such as the otic vesicle. Using the micromanipulation controller, insert the needle into the target tissue, and use the PICO pump foot switch to inject the preset volume.

If the tissue resists penetration, gently tap the micromanipulation controller knob. To release the embryos, use a transfer pipet to flow E-3 over the surface from top to bottom, or by swirling the dish such that the embryos are released into the surrounding E-3. Transfer the embryos to a recovery dish of E-3 without MS-222.

To screen the embryos using the PDMS device in the wells of a six well plate, use a 1, 000 microliter pipet or a narrow-tipped transfer pipet to prime the device by flowing E-3 through the port. Use E-3 and MS-222 to fill the well until the device is covered. Then, use a transfer pipet to deliver four previously injected embryos into each well.

Using a hair loop or similar tool, position the embryos at the entrances of the imaging channels in the desired orientation, and partially push them into the device. This will help draw them into the device evenly. Using a 1, 000 microliter pipet or transfer pipet, draw E-3 through the device from the port until the embryos are drawn into the channels in the correct orientation for imaging.

Transfer the well plate to an inverted fluorescent microscope. To image the embryos, adjust the magnification by selecting the appropriate lens, such as 10x for imaging zebrafish neturaohil migration. Adjust the light source intensity and exposure time for each channel, so that each signal is clear, but not saturated.

Finally, capture images for each fluorescent and transmitted channel. Using the microstructured channel device to stabilize embryos in the lateral orientation, the ability of neutrophils to respond to the standard chemoattractants fMLP and LTB4 microinjected into the otic vesicles, of two days post-fertilization zebrafish embryos was tested. To visualize neutrophil recruitment, injections were traced with high molecular weight rhodamine dextran, and performed in transgenic embryos with green fluorescent neutrophils.

Uninjected embryos and embryos injected with rhodamine alone were used as negative controls. Following microinjection, embryos were transferred into the imaging channel device and imaged using an EVOS fluorescent microscope at 30 minutes and five hours post injection. As expected, a small number of fluorescent neutrophils were recruited to the mock injected otic vesicle at the early time point, likely due to damaged mediated recruitment.

Interestingly, the rhodamine dextran tracer was observed to accumulate in the otoliths during the experiment. For both chemoattractants, neutrophils were recruited in higher numbers, particularly in response to LTB4. Once mastered, this technique can greatly improve the speed and accuracy of zebrafish microinjection.

Development of this technique was driven by specific challenges in our lab. We refined this approach to fit the needs of our projects on fungal infection and tumor xenografts in zebrafish models. After watching this video, you should have a good understanding of how to use microstructured surface arrays for zebrafish microinjection and for extended live imaging.