ライブゼブラフィッシュ胚における有糸分裂とダイナミクスを観察します

The overall goal of this procedure is to monitor the fidelity of chromosome segregation during mitosis in a live zebrafish embryo by using flourescently labeled chromatin and cell-membrane proteins that will be captured using high-resolution confocal microscopy. This method can help answer key questions in the mitosis field, such as how do mitotic defects result in a developmental defect, or tumor formation in a live vertebrate organism? The main advantage of this technique is that we are observing mitosis in a live vertebrate organism that allows phsyiologically relevant data to be collected.

After breeding adult zebrafish and injecting eggs with H2A. F/Z-EGFP and/or pCS2 mCherry-CAAX RNA, approximately two hours before imaging, use a fluorescent dissecting microscope to identify GFP-positive embryos. Transfer bright green, GFP-expressing embryos into a new, 100 by 15 millimeter Petri dish with E3 blue medium.

Boil a stock solution of 1%low-melt agarose in E3 blue. Aliquot three milliliters of the melted agarose into a 17 by 100 millimeter culture tube. Keep the agarose warm by placing the culture tube in a 42-degree Celsius water bath until ready for use.

Gather 15-millimolar tricaine, screened embryos, low-melt agarose, E3 blue, and a 35-millimeter glass coverslip-bottom culture dish, and a dissecting light microscope. Next, with number five tweezers, carefully remove the chorions from three embryos, and place the embryos in a separate container to be anesthetized. Then, using a transfer pipette, add three drops of 15-millimolar tricaine into the dish containing the embryos and five milliliters of medium.

Add three to four drops of 15-millimolar tricaine solution to the tube of 1%melted agarose. When the embryos are sufficiently anesthetized, using a P200 Pipetman, with one centimeter of the pipette tip cut off, transfer the anesthetized embryos to the coverslip-bottomed dish. Remove any excess E3 blue tricaine solution.

Slowly add five to 10 microliters of low-melt-agarose-tricaine solution over the embryos, keeping each drop separate, to ensure that the embryos do not drift close to one another. Then, using a 21-gauge one-and-a-half needle, gently orient the embryos in the agarose to the desired position, orienting the region of interest as close to the coverslip as possible if using an inverted microscope. Due to the shifting that may occur, it may be necessary to return to each embryo to make sure they stay in the required position.

This may have to be done multiple times, til the ideal position is achieved as the agar solidifies. After a few minutes, to test if it is set, use a needle to break apart a small piece of agarose. When it can be pulled away from the drop, use additional low-melt agarose to form a dome over the embedded embryos, covering the entire coverslip.

While the agarose solidifies, prepare three milliliters of E3 blue medium with five drops of 15-millimolar tricaine, to be placed over the embedded embryos during imaging. To carry out 5D confocal imaging of live zebrafish embryos, open the imaging software, and set the microscope to a 60X numerical aperture 1.4 lens. Apply immersion oil to the lens, and place the culture dish in the slide holder on the microscope stage.

Using the axis controller, center the embryo of interest above the objective lens, and raise the lens to meet the culture dish. Pour E3 blue and tricaine solution over the agar-embedded embryos. Next, click on the Eye Port icon to view embryos through the eyepiece.

Disable the Eye Port selection, and choose View, Acquisition Controls, A1 Scan Area, to open the A1 Scan Area tool. Select the GFP channel, restart the scan, and focus on a region of interest close to the coverslip. Stop the scan, select the GFP and mCherry channels, and set the line-averaging option to normal.

Now, choose View, Acquisition Controls, ND Acquisition, to open the A1 ND Acquisition tool. Begin scanning. Then, using the axis controller, position the embryos so that the scan area is filled with as much of the zebrafish as possible.

Set the Z-stack upper limit to where the cells are out of focus, allowing for an extra three micrometers of space above the sample, for growth and cell movement. Then, set the lower limit to where the cells are no longer visible, and set the Z-interval step size to two micrometers. For the experiments demonstrated here, adjust the imaging laser power, HV or gain, and offset levels, respectively, as listed below.

Once the parameters are set, shut off the scan to prevent unnecessary laser exposure that may cause phototoxicity and photobleaching of the sample. Now, select the 2X line averaging icon, which provides the best image quality and fastest scan time. Select the appropriate time interval and time duration necessary for the experiment.

For wild-type cell divisions, two minute time intervals for a duration of two hours is adequate for determining mitotic duration. Divisions that activate the spindle assembly checkpoint for longer than thirty minutes are more suitable for four-to-five-minute intervals for four hours, in order to preserve flourescense, as demonstrated here. To automatically save the file as it is being acquired, check the Save to File box, and name the file.

Finally, double-check that all parameters are set correctly, and hit Run Now. After acquisition is complete, to view the file in a three-dimensional format, click on the Volume Threshold icon. This figure demonstrates the ability to observe many cell divisions using a wide field of view of an AB wild-type zebrafish tail.

The cell divisions are denoted by asterisks. The corresponding movie is shown here. On average, 50 dividing cells in AB, and 30 dividing cells in the aurora B mutant were observed.

To account for the reduced number of mitotic cells imaged in the aurora B mutant embryo, the ratio of mitotic cells to the number of cells imaged was calculated, suggesting that there is not a lower number of cells going through mitosis in the aurora B mutant embryos, but, rather, a fewer number of total cells being imaged. To quantify mitotic duration, the number of time intervals that take up one cell division are multiplied by the interval of time between each Z-stack. As revealed in this graph, the average AB wild-type division time is 25 minutes, and 58 minutes for the aurora B mutant.

As seen in wild-type embryos, each mitotic phase can be distinguished. However, mitotic defects are observed in the aurora B mutant embryo, which include mitotic arrest and filed cytokinesis, resulting in a binucleated cell, and subsequent formation of micronuclei, which support the increased division time measured in this mutant. Once mastered, this technique can be done in one hour, from screening embryos to setting up the confocal microscope parameters.

While attempting this procedure, it's important to remember the context of the experiment at hand, as it pertains to the region of interest, time duration, time interval, and Z-depth. This will help maximize efficiency, as well as prevent photobleaching and phototoxicity. Following this procedure, other methods, like chromosome spreads, can be performed to answer additional questions about the cause of mitotic defects observed, or use of a drug treatment to determine the effects on mitotic progression, duration, or cell fate.

After its development, this technique paved the way for researchers in the field of biology to explore mitosis in the context of developmental and cancer biology using zebrafish. After watching this video, you should have a good understanding of how to embed zebrafish embryos, capture live imaging of mitosis, and analyze mitotic frequency, duration, and cell fate. Don't forget that working with high-powered lasers and fluorescent light can be detrimental to the eyes, and precautions, such as avoiding direct eye contact with the lasers, should always be taken while performing this procedure.