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

All animals were handled in strict accordance with protocols approved by the Oklahoma Medical Research Foundation Institutional Animal Care and Use Committee.

1. Setting up adult zebrafish for breeding

1. Use a large cohort of adult male and female zebrafish of a single strain for breeding. There are small differences in the genetic makeup of zebrafish of even a single strain, use a large cohort to ensure results are representative of the genetic variability of the population and not restricted to a small subset of the population.
2. The night before embryos will be collected, place dozens of breeding adults into breeding tanks, use approximately equal numbers of males and females. Depending on the experiment, use one of the following breeding strategies.
   1. Breeding strategy 1: Place a few male and female zebrafish in individual breeding tanks, separate the males from the females with a divider. If this approach is used, setup many different breeding tanks and pool the embryos from each to ensure the biological variability is representative of the population.
   2. Breeding strategy 2: 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.

2. Timed matings – collecting, sorting, and housing zebrafish embryos for experiments

1. Perform timed matings, beginning as soon as the light cycles being in the morning. Discard any embryos generated overnight.
2. Allow fish to breed in shallow water (3-5 cm) for a period of 10 min, with a false bottom for embryos to escape through.
3. After 10 min, collect embryos by pouring through a strainer and rinsing into a 10-cm plate with a wash bottle. Pool all embryos collected at the same time point, mark them with time of collection, and place immediately in an incubator at 28.5 °C.
4. Hundreds of embryos can be expected from a single mating pair in a day. Allow adults to breed in 10 min cycles until the number of embryos obtained is less than twenty.
5. Approximately 1-1.5 h after collection, sort embryos to remove dead and unfertilized embryos. To sort embryos, remove from incubator and observe under a dissecting microscope.
   1. Count embryos and place at a density of 100 embryos per 10 cm plate.
   2. Add fresh E3 medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl₂, 0.33 mM MgSO₄) and return embryos to the 28.5 °C incubator.

3. Dechorionate, deyolk, and fix zebrafish embryos

NOTE: This section of the protocol is designed for embryos prior to 48 h post fertilization (hpf). There is no need to remove the chorions of embryos at later stages of development (after 48 hpf), as they often naturally fall off. There is no need to deyolk or remove the chorions of fish older the 5 days post fertilization (dpf).

1. When embryos reach desired developmental age, verify appropriate stage of development morphologically under a light microscope according to "Stages of Embryonic Development of the Zebrafish"¹⁷.
2. Transfer up to 1500 staged embryos to a 15-mL conical tube and wash several times with E3.
   1. Add 1 mL of E3 and 20 µL of Pronase solution (30 mg/mL) for every 100 embryos and gently swirl. Up to 1500 embryos can be dechorionated in a single tube with 15 mL of E3.
   2. Lay tube on its side and arrange embryos in an even layer to ensure maximum surface to volume ratio. Gently agitate tube every 2-3 min until all chorions have fallen off of the embryos. The total dechorionation time will vary depending on the Pronase activity.
3. Remove supernatant and gently rinse embryos 3 times with 10 mL of E3. Place embryos on ice for the remainder of this section of the protocol.
4. Remove E3 and wash embryos with freshly prepared ice-cold deyolking buffer (0.5x Ginzburg Fish Ringer's solution, without calcium: 55 mM NaCl, 1.8 mM KCl, 1.25 mM NaHCO₃).
   1. Add 1 mL of ice-cold deyolking buffer for up to 500 embryos.
   2. Using a P1000, gently pipette embryos up and down 5-10 times to disrupt yolk sack.
   3. Transfer 1 mL to a 1.5 mL microfuge tube and centrifuge at 4 °C and 500 x g for 5 min.
5. Remove supernatant and wash pellet with 1 mL of ice-cold 1x PBS (phosphate buffered saline, pH 7.0) to remove Ringer's solution. Centrifuge at 4 °C and 500 x g for 5 min.
6. Carefully remove supernatant and resuspend cells in 1 mL of 1x PBS. Place tube on ice.
7. Add 3 mL of ice-cold ethanol (EtOH) to a 15-mL conical tube. Vortex tube of EtOH gently on low setting.
   1. Using a P1000, slowly (1 drop per second) drip zebrafish embryo cell suspension into EtOH while vortexing gently.
   2. Add a total of 1 mL of the embryo cell suspension, for a final fixation solution of 75% EtOH.
8. After 1 mL of cell suspension has been added, gently swirl tube to mix and place at -20 °C for at least 1 h. It is recommended to place the tubes on their side to ensure maximum surface to volume ratio and reduce the number of embryos and cells that stick together. Store embryo cells at -20 °C for 2-3 weeks.

4. Staining DNA and FACS sorting embryos

NOTE: This section of the protocol is designed for embryos at 1 dpf.

1. Remove EtOH-fixed embryo cells from -20 °C and centrifuge at 4 °C 1500 x g for 5 min.
   1. Place tube on ice and carefully remove supernatant. Using a P1000, gently resuspend cells in 1 mL of freshly prepared cold PBS-BSA (1x PBS, 1% bovine serum albumin).
   2. Centrifuge cells at 4 °C and 1500 x g for 5 min and place on ice.
2. Remove supernatant and resuspend cells in propidium iodide (PI) staining solution (50 µg/mL PI, 100 µg/mL RNase A, PBS, pH 7.0), using 200 µL for every 1000 embryos.
   1. Allow cells to incubate in PI solution for 30 min on ice, gently mixing every 5 min.
   2. Place a 40 µm nylon cell strainer on a 50-mL conical tube on ice. Gently pipette cells to mix and filter through the mesh.
      NOTE: If multiple tubes of embryos from the same timepoint were collected in section 3 of the protocol, combine these embryos at this point so that they are all sorted together.
   3. Using the flat inside end of a syringe plunger, gently disrupt any remaining clumps of cells on the mesh.
   4. Rinse cells through the filter with 500 µL of cold PBS-BSA for every 1000 embryos.
   5. Place a 40-µm mesh over a 15-mL conical tube on ice and filter the cell suspension from the 50-mL conical tube a second time into the 15-mL tube.
3. Using an appropriate cell sorting instrument, set the laser excitation to 561-nm and emission detection to 610/20 to detect propidium iodide.
   1. Mix 15 mL tube of stained cells briefly by vortexing and place in instrument at 4 °C.
   2. Run cells at a slow flow rate, ideally 1.0 mL/min, in order to obtain a clean cell cycle profile and avoid mixing G1 or G2/M cells with the S phase population.
   3. First, gate for FSC-A x SSC-A (forward scatter area by side scatter area) to remove debris (Figure 1A).
      NOTE: Cells from the embryos can be varying sizes depending on the stage and cell type. The neutral density (ND) filter may need to be switched between 1.0 and 1.5 depending on the size of cells present. Gate a large area to collect most cells and remove debris.
   4. Second, gate for SSC-area (SSC-A) by PI-area (PI-A), to remove debris and cell doublets (Figure 1B).
   5. Third, gate for SSC-A by SSC-W (width) to remove any remaining cell doublets (Figure 1C).
4. Display the gated populations on a histogram with cell number (count) on the y-axis and propidium iodide area (PI-A) on the x-axis. For 24 hpf embryos, this should give a stereotypical cell cycle profile as in Figure 1D.
   1. To sort the S phase population, set the gate from the right side of the G1 peak to the left side of the G2 peak, including only a minimal amount of the G1 and G2 populations. For 24 hpf embryos, this will typically comprise about 10-15% of the gated population (see Figure 1D).
   2. To sort the G1 population, set the gate on the left side of the G1 peak, not to pass the top of the peak. Adjust the width of the G1 gate as narrow as possible (see Figure 1D).
   3. Back gate the G1 and S phase populations to ensure proper initial gating (Figure 1E).
5. Sort the G1 and S phase populations into 15 mL conical tubes with 3 mL of PBS-BSA. The goal should be to obtain between 500,000 and 1 million or more sorted cells per fraction (G1 and S phase).
6. After sorting is complete, centrifuge tubes at 1500 x g and 4 °C for 10 min.
   NOTE: The small cell pellet will be difficult to see and it is important to avoid disrupting it, so use preferably a swinging bucket rotor over a fixed-angle rotor.
7. Carefully remove supernatant and resuspend pellet with 1 mL of 1x PBS. Transfer cells to a 1.5 mL microfuge tube and centrifuge at 6,000 x g and 4 °C for 5 min.
8. Carefully remove all liquid from tube and freeze dry pellet at -20 °C. Store the pellet at -20 °C for 2-3 weeks.

5. DNA isolation, RNase treatment, and DNA purification

1. Remove cell pellet from -20 °C and place on ice.
   1. Add 400 µL of SDS buffer (50 mM Tris-HCl (pH 8.0), 10 mM EDTA, 1 M NaCl, 0.5% SDS) to pellet and resuspend.
   2. Add proteinase K to a final concentration of 0.2 mg/mL and mix by vortexing briefly.
   3. Incubate samples at 56 °C for 2 h, vortex every 15 min.
2. After 2 h, perform phenol-chloroform extraction by adding 400 µL of phenol-chloroform (Phenol:Chloroform:Isoamyl Alcohol 25:24:1, saturated with 10 mM Tris, pH 8.0, 1mM EDTA).
   1. Vortex for 20 s and centrifuge sample at 16,000 x g for 5 min to separate phases.
   2. Carefully remove the upper aqueous phase (400 µL) and transfer to a 1.5 mL tube.
3. Perform EtOH precipitation by adding 40 µL of 3M sodium acetate (pH 5.2), 2 µL of glycogen, and 1 mL of EtOH.
   1. Vortex to mix thoroughly and place at -20 °C overnight to precipitate DNA. Sample can also be precipitated at -80 °C or on dry ice for 1 h.
   2. Centrifuge sample at 4 °C and 16,000 x g for 30 min to pellet DNA.
   3. Discard supernatant and wash pellet with 1 mL 70% EtOH. Centrifuge at 4 °C and 16,000 x g for 5 min.
   4. Discard supernatant and air-dry pellet at ambient temperature for 2-3 min.
4. Dissolve pellet in 96 µL of autoclaved water, and add 4 µL of RNase A (2 mg/mL).
   1. Mix well by vortexing and incubate at 37 °C for 1 h.
5. Perform phenol-chloroform extraction by adding 100 µL of phenol-chloroform and following procedure in steps 5.2.1 – 5.2.2.
6. Perform EtOH precipitation by adding 10 µL of 3M sodium acetate, 1 µL glycogen, and 250 µL of EtOH and following the procedure in steps 5.3.1 – 5.3.5.
7. Resuspend pellet in 60 µL of TE buffer (10 mM Tris-HCl (pH 8.0), 10 mM EDTA (pH8.0)) and determine DNA concentration using a quantitative fluorescence-based method. Store DNA at -20 °C.
   NOTE: It is important to quantify DNA using fluorescence-based DNA binding dyes, or other means more reliable than UV spectrometer-based methods.

6. Preparation of DNA libraries and next generation sequencing

NOTE: A G1 copy number reference sample for each biological source is required for each sequencing run (i.e. WT, mutant, transgenic, cell line, etc). Compare all S phase samples from the same biological source in the same sequencing run to the same G1 reference. Run at least two biological replicates of each sample to ensure consistency between samples.

1. Dilute 1 µg of DNA to a final volume of 50 µL and transfer to a specialized tube for sonication.
2. Shear genomic DNA for a target of 250-300 bp in a focused ultrasonicator, according to manufacturer's specifications and protocol.
3. Verify quality and size of sheared genomic DNA using an automated electrophoresis machine, according to manufacturer's specifications and protocol. Ensure there is a large peak of ~250-300 bp for the sheared genomic DNA.
4. Prepare sequencing libraries using a DNA library preparation kit with multiplex oligos for sequencing on Illumina platform, according to manufacturer's protocol.
5. Verify quality of library prep using automated electrophoresis machine, according to manufacturer's protocol. Ensure there is a large peak of ~400-450 bp, and there are minimal peaks for primer dimers (80-85 bp) and adapter dimers (~120 bp).
6. Quantify libraries using a DNA library quantification kit for next-generation DNA sequencing, according to manufacturer's protocol.
7. Dilute 15 µL of library to a concentration of 5 nM and combine multiplexed libraries as required to achieve desired uniquely mapping read counts per sample.
   NOTE: The number of uniquely mapping reads per sample will determine the resolution. Use as few as 5 million mapped reads to assess replication timing for the zebrafish genome. It is recommended to start with 10-20 million reads.
8. Perform paired-end 100 bp whole genome DNA sequencing of samples on a next generation sequencing platform, according to manufacturer's instructions.

7. Analysis of sequencing data

NOTE: The instructions in this section are intended as a guideline for analysis. Use additional methods for sequencing alignment, filtering, processing, etc. This section of the protocol will deal with the preferred method of analysis in this work. If additional methods are used, adjust the parameters and functions to suit those purposes. The commands below are entered in Ubutnu or Mac terminal, with the appropriate packages installed.

1. Obtain the most recent versions of the zebrafish genome (danRer10, as of when this protocol was written) from UCSC genome browser using the following commands:
   wget ftp://hgdownload.cse.ucsc.edu/goldenPath/danRer10/bigZips/danRer10.fa.gz-O danRer10.fa.gz
   1. Uncompress the fa.gz file with the following command:
      gunzip danRer10.fa.gz
   2. Align sequencing data to genome using BWA-mem using the following commands:
      bwa mem -t 8 danRer10.fa read_1.fastq read_2.fastq > output.sam
2. Convert .sam file to .bam file using samtools with the following command:
   samtools view -bS input.sam > output.bam
   1. Sort the aligned sequencing files using samtools with the following command:
      samtools sort output.bam > output_sorted.bam
   2. Index the .bam file using samtools with the following command:
      samtools index output.bam
3. Mark PCR duplicates with Picard using the following commands:
   java -Xmx2g -jar picard.jar MarkDuplicates
   INPUT=input.bam
   OUTPUT=output.bam
   REMOVE_DUPLICATES=false
   METRICS_FILE=output_metrics.txt
4. Generate text (.txt) files of reads for each chromosome using the following command:
   for i in {1.25}; do CHROM="chr"$i; echo $CHROM; samtools view input.bam $CHROM | cut -f 2,4,5,7,8,9 > readsChr${i}.txt; done
   NOTE: The columns in the resulting .txt file are Flag, Location, mapQ, pair chr, pair location, and fragment size, respectively.
5. Read .txt files into Matlab (or any other similar software) using the textscan function.
   1. Filter out low mapping quality reads by setting a mapQ threshold of 30.
   2. Remove reads with flags for not primary alignment, PCR duplicates, or pair mapping to a different chromosome.
   3. Filter out any reads with their pair beyond a distance threshold of 2000 bp.
   4. Evaluate statistics of resulting reads to determine number of read statistics, coverage, insert size, and quality (Figure 2).
6. Determine read coverage in 100 bp fixed windows for each sample by counting the number of reads in 100 bp intervals across the length of each chromosome.
   1. Calculate the median read count for all windows.
   2. Filter out regions of high coverage by removing windows greater than 5 standard deviations from the median.
7. Define regions for analysis in the G1 reference samples by finding segments along each chromosome with 200 reads using sliding windows.
   1. Determine read depth in each S phase sample by counting the number of S phase reads in the 200-read count windows defined for the accompanying G1 reference.
8. Smooth the raw data using the csaps function in the software, with an interpolation factor (p) of 10^-16.
9. Normalize smoothed data to z-score with a mean of 0 and standard deviation of 1.