Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

The overall goal of this procedure is to generate whole genome DNA replication timing maps from zebrafish embryos in different stages of development. This method can help answer key questions in the cell cycle and chromatin biology fields, such as how chromatin changes that occur during embryonic development can affect the initiation of DNA replication forks. The main advantage of this technique is that it maps replication changes that occur in vivo at defined stages of embryonic development.

Though this method can provide insight into how DNA replication changes during zebrafish development, it also can be applied to other systems or model organisms, such as tunicates, frogs, and flies. Generally, people new to this method may struggle because it does take some practice to generate single-cell suspensions from zebrafish embryos. The night before embryos will be collected, place dozens of breeding adults into breeding tanks, using approximately equal numbers of males and females.

Depending on the experiment, use one of the following breeding strategies. For the first breeding strategy, place a few male and female zebrafish in individual breeding tanks, and use a divider to separate the males from the females. If this approach is used, set up many different breeding tanks and pool the embryos from each to ensure the biological variability is representative of the population.

For breeding strategy number two, combine dozens of male and female zebrafish in a large breeding tank. Use this strategy as long as there are a sufficiently large number of embryos. It is reasonable to assume they came from a variety of founders and are therefore genetically representative of the population.

To perform timed matings, begin as soon as the light cycles start in the morning. Allow fish to breed in shallow water for a period of 10 minutes, with a false bottom for embryos to escape through. After 10 minutes, collect embryos by pouring them through a strainer and using a wash bottle to rinse them into a 10-centimeter plate.

Pool all embryos collected at the same time point, mark them with the time of collection, and immediately place them in an incubator at 28.5 degrees Celsius. To ensure that synchronously developing embryos are collected, it is critical that batches of embryos are collected every 10 minutes. This is especially important with the earliest stages of embryonic development.

Hundreds of embryos can be expected from a single mating pair in a day. Allow adults to breed in 10-minute cycles until the number of embryos obtained is less than 20. Approximately one to 1.5 hours after collection, to sort embryos, remove them from the incubator, observe them under a dissecting microscope, and sort them to remove dead and unfertilized eggs.

Count the embryos, and place them at a density of 100 embryos per 10-centimeter plate. Add fresh E3 medium, and return the embryos to the 28.5 degree Celsius incubator. When the embryos reach the desired stage of development, verify the stage morphologically under a light microscope according to Stages of Embryonic Development of the Zebrafish.

If the embryos are 48 hpf or younger, transfer up to 1500 staged embryos to a 15-milliliter conical tube and use E3 to wash them several times. For every 100 embryos, add one milliliter of E3 and 20 microliters of Pronase solution and gently swirl. Up to 1500 embryos can be dechorionated in a single tube with 15 milliliters of E3.Lay the tube on its side, and arrange embryos in an even layer to ensure maximum surface-to-volume ratio.

Gently agitate the tube every two to three minutes until all the chorions have fallen off of the embryos. The total dechorionation time will vary depending on the Pronase activity. Remove the supernatant, and, with 10 milliliters of E3, gently rinse the embryos three times.

After placing the embryos on ice and removing the remaining E3, use one milliliter of freshly prepared, ice-cold deyolking buffer for every 500 embryos to wash the sample. Next, using a P1000, gently pipette the embryos up and down five to 10 times to disrupt yolk sack. Then, transfer one milliliter to a 1.5-milliliter microfuge tube, and centrifuge the sample at four degrees Celsius and 500 times gravity for five minutes.

The samples must be pipetted just vigorously enough to create a single-cell suspension without lysing the cells. This step will take some practice. Remove the supernatant, and use one milliliter of ice-cold 1x PBS to wash the pellet to remove the Ringer's solution.

Then, centrifuge the sample again. Carefully remove the supernatant, and resuspend the cells in one milliliter of 1x PBS. Then, place tube on ice.

Add three milliliters of ice-cold ethanol to a 15-milliliter conical tube, and gently vortex the tube of EtOH on a low setting. Using a P1000, slowly drip zebrafish embryo cell suspension into the EtOH while continuing to vortex. Add a total of one milliliter of the embryo cell suspension, for a final fixation solution of 75%EtOH.

After one milliliter of cell suspension has been added, gently swirl the tube to mix and place it on its side at negative 20 degrees Celsius for at least one hour, or store the suspension in the freezer for two to three weeks. To stain embryonic DNA, after removing EtOH-fixed embryo cells from negative 20 degrees Celsius, centrifuge the tube at 1500 times gravity and four degrees Celsius for five minutes. Place the tube on ice, and carefully remove the supernatant.

Then, using a P1000, gently resuspend cells in one milliliter of freshly prepared, cold PBS-BSA. Centrifuge the cells at 1500 times gravity and four degrees Celsius for five minutes, and place the sample on ice. Then, remove the supernatant, and, for every 1, 000 embryos, add 200 microliters of propidium iodide staining solution and resuspend the cells.

Allow the cells to incubate in PI solution for 30 minutes on ice, gently mixing every five minutes. Place a 40-micrometer nylon cell strainer on a 50-milliliter conical tube on ice. Then, gently pipette the cells to mix and filter the cell solution through the mesh.

Using the flat, inside end of a syringe plunger, gently disrupt any remaining clumps of cells on the mesh. For every 1, 000 embryos, use 500 microliters of cold PBS-BSA to rinse the cells through the filter. Place a 40-micrometer mesh over a 15-milliliter conical tube on ice, and filter the cell suspension from the 50-milliliter conical tube a second time into the 15-milliliter tube.

Finally, using an appropriate cell sorting instrument, set the laser excitation to 561 nanometers and emission detection to 610 over 20 to detect propidium iodide. Sort the cells, and carry out additional analyses according to the text protocol. To determine the read mapping quality, the sequencing data must first be aligned to the genome, using statistics of mapQ values.

In addition, the calculated read length statistics are plotted as a histogram of the distribution of insert sizes for all sequencing reads. This particular experiment resulted in a median of 170 bp. This bar graph illustrates the read statistics for the total mapped reads, the reads containing a low-quality flag, reads with pairs mapping to a different chromosome, reads with a pair beyond distance threshold, and PCR duplicates.

Representative numbers of total mapped, unmapped, and unpaired reads, coverage, and read resolution for a typical sequencing run of a zebrafish sample are listed here. After filtering, read counts are determined in variable-sized windows and the data are smoothed and normalized. Typical smoothed and normalized replication timing profiles of a representative zebrafish chromosome for biological replicates are shown here.

The profiles for biological and experimental replicates should be very similar and also display high correlation along the length of the chromosome. The profiles should also display high correlation between timing values genome-wide. The color map represents Pearson's correlation coefficient.

While attempting this procedure, it's most important to remember to ensure synchronously developing embryos are collected. Following this procedure, other methods, like morpholino injections or CRISPR-Cas9 mutagenesis, can be performed in order to answer additional questions, like what are the mechanisms by which DNA replication is coordinated with other developmental processes? After watching this video, you should have a good understanding of how to collect cells at specific stages of the cell cycle from synchronously developing zebrafish embryos.

Collection of these samples is critical for generating whole genome DNA replication timing maps.