Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Podsumowanie

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Streszczenie

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraB^hi1045, and esco2^hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Wprowadzenie

Mitosis is a critical cellular process essential for growth, differentiation, and regeneration in a living organism. Upon accurate preparation and replication of DNA in interphase, the cell is primed to divide. The first phase of mitosis, prophase, is initiated by activation of cyclin B/Cdk1. Prophase is characterized by condensation of chromatin material into chromosomes. Nuclear envelope breakdown occurs at the transition between prophase and prometaphase. In prometaphase, centrosomes, the nucleating center for spindle formation, begin to migrate to opposite poles while extending microtubules in search of kinetochore attachment. Upon attachment, conversions to end-on microtubule attachment and tension forces orient the chromosomes forming a metaphase plate¹. If all chromosomes are attached correctly, the spindle assembly checkpoint is satisfied, cohesin rings holding the sister chromatids together are cleaved, and microtubules shorten to pull sister chromatids to opposite poles during anaphase^2,3. The final phase, telophase, involves elongation of the cell and reformation of the nuclear envelope around the two new nuclei. Cytokinesis completes the division process by separating the cytoplasm of the two new daughter cells^4-6. Alteration of key mitotic pathways (i.e., spindle assembly checkpoint, centrosome duplication, sister chromatid cohesion, etc.) can result in metaphase arrest, missegregation of chromosomes, and genomic instability^7-10. Ultimately, defects in pathways controlling mitosis can cause developmental disorders and cancer, necessitating visualization of mitosis and its defects in a live, vertebrate, multi-cellular organism^10-16.

Zebrafish embryos serve as a great model organism for live imaging due to the transparent tissue, ease of microinjection, and fast development. Using zebrafish, the overall goal of this manuscript is to describe a method of live 5D (dimensions X, Y, Z, time, and wavelength) imaging of mitosis¹⁷ (Figure 1C). The use of mutant zebrafish defective in different mitotic pathways demonstrate the consequence of such defects. For this protocol, Aurora B and Esco2 mutants were chosen to illustrate these defects. Aurora B is a kinase that is part of the chromosome passenger complex (CPC) involved in spindle formation and microtubule attachment. It is also required for cleavage furrow formation in cytokinesis^18,19. In zebrafish, Aurora B deficiency leads to defects in furrow induction, cytokinesis, and chromosome segregation²⁰. Esco2, on the other hand, is an acetyltransferase that is essential for sister chromatid cohesion^21,22. It acetylates cohesin on the SMC3 portion of the ring thus stabilizing cohesin to ensure proper chromosome segregation at the metaphase-anaphase transition²³. Loss of Esco2 in zebrafish leads to chromosome missegregation, premature sister chromatid separation, genomic instability, and p53-dependent and independent apoptosis^24,25. Due to the availability, auroraB^hi1045, and esco2^hi2865 mutant zebrafish (hereafter referred to as aurB^m/m and esco2^m/m, respectively) will be used to illustrate this technique^25-27.

Coupling confocal microscopy with fluorescent-tagged cell machinery has enabled researchers to visualize chromatin and cell membrane dynamics during mitosis^25,28,29. Fluorescent-tagged histones have historically been used to visualize chromatin. Histones are nuclear proteins composed of four different pairs (H2A, H2B, H3, and H4) that are responsible for the nucleosome structure that composes chromosomes³⁰. While H2B is arguably the most used histone for fluorescent proteins in mouse and cell culture, use of Histone 2A, Family Z (H2A.F/Z) has proved well for use in zebrafish^31,32. Concanavalin A and casein kinase 1-gamma for example, localize to the cell membrane and have previously been shown effective in visualizing the cell membrane in sea urchins and drosophila^33,34. Other studies have shown that the CAAX fluorescent-tagged protein labels the cell membrane and was successful in zebrafish³¹. CAAX is a motif that is recognized by post-translational modifying enzymes such as farnesyltransferases and geranylgeranyltransferases. Modifications by these enzymes cause proteins to become membrane-associated, thus labeling the cell membrane³⁵.

Due to the prior use in zebrafish, this protocol chose to use H2A.F/Z and CAAX to label chromatin and the cell membrane. Application of this method will allow the researcher to monitor mitosis at the individual cell level to observe individual chromosome dynamics, as well as simultaneously monitor multiple cell divisions that may impact tissue differentiation and development. This article will focus on imaging the dynamics of chromosome segregation during mitosis at the individual cell level. Within this manuscript, the ability to observe several mitotic divisions, calculate division time, and decipher the mitotic phenotypes will be illustrated and discussed. By using these parameters, physiologically relevant data can be collected and applied to several disease states affected by mitotic defects.

Protokół

1. In Vitro Transcription

1. Linearize pCS2-H2A.F/Z-EGFP and/or pCS2-mCherry-CAAX vectors by NotI restriction enzyme digest³¹. Using an RNA in vitro transcription kit, generate 5' capped mRNA products from each template, according to manufacturer's protocol.
2. Purify the capped mRNA using a purification kit. Follow manufacturer's instructions. Elute with RNase-free H₂O.
3. Determine the concentration of RNA by absorbance at 260 nm using a spectrophotometer. (OD₂₆₀ x dilution x 40 µg/ml).
4. Dilute the RNA to 100 ng/µl for each H2A.F/Z-EGFP and mCherry-CAAX with RNase-free H₂O. If the RNA concentration is too low, the fluorescence will be diminished or absent. Brighter samples will diminish the concerns over phototoxicity and photobleaching. On the other hand, too much RNA can be toxic and/or cause off target effects.
   Note: Store the remaining purified mRNA in -80 °C freezer.

2. Zebrafish Breeding, Embryo Collection, and mRNA Injection ^36-38

1. Assemble breeding tanks with a barrier to separate the tank into two regions and fill each breeding tank with aquaculture system water used in the zebrafish facility.
2. In order to prevent untimely breeding, place two male fish on one side of the barrier and two female fish on the other side the night before breeding.
3. The next day, thaw the previously prepared mRNA mixture on ice. Replace the water in breeding tanks with fresh aquaculture system water and remove the barriers. Immediately after the barriers are pulled, warm an injection mold to 28.5 °C and set up the equipment for microinjection.
   Note: For information on injection molds, please refer to Gerlach³⁶.
4. Collect eggs every 10 - 15 min using a tea strainer and rinse the eggs into a clean 100 x 15 mm Petri dish with E3 Blue (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl₂, 0.33 mM MgSO₄, 10^-5% Methylene blue). E3 Blue is used to prevent fungal growth and ensure proper development of larval fish. For more information on mating and embryo collection, refer to Gerlach³⁶ and Porazinkski³⁷.
5. Embed one-cell staged embryos in a warmed injection mold and inject the RNA into the yolk in the desired amount of embryos (Figure 1A). Account for natural embryonic death and unfertilized embryos by performing embryonic injections on 15% more embryos than needed for the experiment. For additional details on microinjection of zebrafish embryos, please refer to Gerlach³⁶ and Porazinkski³⁷.
   NOTE: For first time use of mRNA, perform a dose-curve analysis to determine the optimal dose for fluorescence and viability (defined as no gross developmental defects up to 5 dpf) prior to performing 5D imaging. 150 - 200 ng/µl injected into embryos is often the optimal concentration, therefore it is a good starting point for the final concentration.
6. Carefully extract the injected embryos from the mold using a modified 9" glass Pasteur pipette. To modify the pipette, melt the end using a Bunsen burner until it forms a ball.
7. Place injected embryos in a 100 x 15 mm Petri dish in E3 Blue and house in a 28.5 °C incubator.
8. Six hr post-injection, remove any dead or unfertilized embryos from the plates and add clean E3 Blue. House the embryos at 28.5 °C.

3. Preparation and Embedding of Live Zebrafish Embryos for Imaging (Figure 1B)

1. Two hr before imaging, screen the injected embryos for GFP using a fluorescent dissecting microscope. Place bright green GFP-expressing embryos in a new 100 x 15 mm Petri dish with E3 Blue.
2. Boil a stock solution of 1% low melt agarose by adding 1 g of low melt agarose to 100 ml of E3 Blue. After using the agarose, cover the flask with aluminum foil. The stock solution remains useful for up to one month.
3. Aliquot 3 ml of the melted agar into a 17 x 100 mm culture tube. Keep the agarose warm by placing the culture tube in a 42 °C water bath until ready for use. Prepare a 15 mM Tricaine solution in deionized water to anesthetize the zebrafish embryos³⁶.
   NOTE: If imaging at earlier time points is desired, the concentration of agar can be decreased as low as 0.3%³⁹.
4. Bring the 15 mM Tricaine, screened embryos, low melt agarose, E3 Blue, and a 35 mm glass coverslip bottom culture dish to a dissection light microscope. Carefully remove the embryo's chorion with #5 tweezers. Do this for three embryos.
5. Place the dechorionated embryos in a separate container to be anesthetized. The lid of the coverslip bottom dish is often used for this purpose. Using a transfer pipette, add three drops (approximately 150 µl) of 15 mM Tricaine to the dish of 5 ml E3 blue (if using the lid of the coverslip bottom dish) or until embryos have been sufficiently anesthetized. In addition, add 3-4 drops (approximately 150 - 200 µl) of 15 mM Tricaine solution to the 1% melted agarose tube.
6. Using a p200 pipette with 1 cm of the pipette tip cut off; transfer the anesthetized embryos to the coverslip-bottomed dish. Remove any excess E3 Blue:Tricaine solution.
7. Slowly add 5 - 10 µl of low-melt agarose:Tricaine solution over the embryos, keeping each drop separate to ensure the embryos do not accidentally drift close to one another.
   Warning: If the agarose is too warm, it will damage the embryo. A good temperature to maintain the agar at is 42 °C.
8. Using a 21 G 1 ½ needle, gently orient the embryo in the agarose to the desired position. When using an inverted microscope for time-lapse imaging, orient the region of interest (ROI) as close to the coverslip as possible.
   Note: For general purposes, the tail region offers ease of orientation and clarity due to the relatively thin tissue (Figure 1A). Other tissues, such as the epithelial layer surrounding the yolk and fin folds, can be used^28,29. These tissues offer great clarity, however these regions are only a few cell layers thick. For the purpose of this protocol, it is beneficial to image the tail region to acquire as many cell divisions as possible.
9. Allow a few min for partial solidification of the agar. Use the needle to break apart a small piece of agar to test its solidification. When a piece of agar can be pulled away from the drop, proceed to the next step.
10. Cover the entire coverslip with low melt agar forming a dome over the embedded embryos. Allow the agar to solidify before moving the dish for confocal imaging (Figure 1A).
11. During the agar solidification process (takes approximately 10 min), prepare 3 ml of E3 Blue solution with five drops of (approximately 250 µl) 15 mM Tricaine to be placed over the embedded embryos during imaging.

4. 5D Confocal Imaging of Live Zebrafish Embryos^40,41

NOTE: See Ariga⁴⁰ and O'Brien⁴¹ for details on how to perform 5D imaging using other confocal systems. For Z-interval, Z-stack, Z-depth, time interval, and 5D definitions see Figure 1C.

1. Open the imaging software and set the microscope to 60X NA 1.4 objective lens. Apply immersion oil to the objective 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 bring the objective lens upward to meet the culture dish.
2. Click on the eye piece icon and switch to the GFP filter on the microscope. Focus on the ROI. Focusing on the tissue closest to the coverslip will offer the best imaging results.
3. Remove the interlock. Select the GFP and mCherry channels (pre-set wavelengths in software) and set the line averaging option to normal.
4. Use "View/Acquisition Controls/A1 Scan Area" command to open the A1 Scan Area tool.
5. Begin scanning. Using the axis-controller, position the embryo so that the scan area is filled with as much of the zebrafish as possible. The laser power does not need to be optimal at this point. Lower the laser power to avoid unnecessary photobleaching.
6. Use the "View/Acquisition Controls/ND acquisition" command to open the ND acquisition control panel.
7. Begin scanning to set the Z-stack parameters. Set the Z-stack upper limit to where the cells are not in focus and lower limit to where cells are no longer visible. Allow for a 3 µm space above the sample for growth and cell movement, which may expand into the imaging field. The Z-stack for the figures shown in this protocol covered a Z-depth of 40 µm in the embryo tail.
8. Set the Z-interval step size to 2 µm. On average, a cell is 10 µm in diameter; therefore 2 µm will produce five intervals of imaging data to be analyzed for each cell.
   NOTE: The depth of the image that can be acquired depends on the Z interval. Z resolution is sacrificed to gain overall depth in the Z dimension in order to image as many cells undergoing mitosis as possible (large Z-depth). The inverse is true, in that, by decreasing the step size, Z resolution is gained, while Z depth is sacrificed (small Z-depth).
9. Adjust the image laser power, HV, and offset levels. For the experiments demonstrated in this protocol, use the following laser power, HV, and offset levels set at the corresponding levels, respectively, for the GFP channel; 2 - 5, 120 - 140, and -9 to -11. For the mCherry channel, use the following laser power, HV, and offset levels, respectively; 3 - 6, 120 - 140, and -3 to -8. Once the parameters are set, shut off the scan to prevent unnecessary laser exposure that may cause phototoxicity and photobleaching of the sample.
10. Select the 2x line averaging icon. "No averaging" produces a grainy image while "4x line averaging" drastically increases the scan time. Use of 2x line averaging provides the best image quality and fastest scan time.
11. Select the appropriate time interval and time duration necessary for the experiment. For wild-type divisions, two-minute time intervals for 2 hr is best for determining mitotic duration (used in Figures 1, 2, 3A, and 3B). Divisions that activate the spindle assembly checkpoint for longer than 30 min are more suitable for five minute intervals for four hr in order to preserve fluorescence as demonstrated in Figure 3C.
12. Check the "save to file" box and name the file to automatically save the file as it is being acquired. Double check all parameters are set correctly and hit "start run".
13. After acquisition is complete, to view the file in a three dimensional format, click on the volume threshold icon.

Wyniki

Figure 2 demonstrates the ability to observe many cell divisions using a wide field view of an AB wild-type zebrafish tail. Over seven mitotic cells are imaged in a 14 min time frame (Movie 1). Within the two hr time-course, over 40 mitotic events were captured. On average, 50 dividing cells were observed in the AB and 30 dividing cells in aurB^m/m embryos (Figure 2B). To account for the number of cells imaged,...

Dyskusje

Use of this method allows one to infer nuclear envelope breakdown, formation of a metaphase plate by microtubule-kinetochore attachments, and segregation of sister chromatids to form two new cells in vivo and in a time-dependent manner. The ability to observe mitosis in zebrafish is advantageous over fixed samples and cell culture systems because the cells are being imaged in the natural physiology, the tissue is transparent which allows for fluorescent proteins to be used, they develop relatively fast, and time...

Materiały

Name	Company	Catalog Number	Comments
pCS2 vectors	Gift from K. Kwan		For plasmid of interest
NotI-HF restriction enzyme	New England BioLabs	R3189S	For restriction digest of plasmid
mMessage SP6 kit	Life Technologies	AM1340	For in vitro transcription
RNeasy Mini kit	Qiagen	74104	For purifying mRNA
100 x 15 mm Petri dishes	Fisher Scientific	FB0875712	For housing embryos
microinjection mold	homemade		For holding embryos during microinjection
Agarose II	Amresco	0815-25G	For embedding embryos
Tricaine	Sigma-Aldrich	E10521-10G	For anesthetizing embryos
Sodium Chloride	Sigma-Aldrich	S9888	For embryo water (E3 Blue), dissolved in UltraPure H2O
Potassium Chloride	Sigma-Aldrich	P3911	For embryo water (E3 Blue), dissolved in UltraPure H2O
Calcium Chloride Dihydrate	Sigma-Aldrich	C8106	For embryo water (E3 Blue), dissolved in UltraPure H2O
Magnesium Sulfate	Fisher Scientific	M7506	For embryo water (E3 Blue), dissolved in UltraPure H2O
Methylene Blue Hydrate	Sigma-Aldrich	MB1	For embryo water (E3 Blue), dissolved in UltraPure H2O
100 mm culture tube	Fisher Scientific	50-819-812	For melted agar
35 mm glass coverslip bottom culture dish	MatTek Corp	P35G-0-20-C	No. 0, 20 mm glass, For embedding embryos
#5 tweezers	Dumont	72701-D	For dechorionating embryos
21 G 1 1/2 gauge needle	Becton Dickinson	305167	For positioning embryos in agar
Dissecting microscope	Nikon AZ100		For screening and embedding embryos, any dissecting scope will do
Confocal microscope	Nikon A1+		For time-lapse imaging
Confocal software	NIS Elements AR 4.13.00		For image acquisition and processing