This work was supported by National Institute of General Medical Sciences of the National Institutes of Health through grants 5P20GM103636-02 (including Flow Cytometry core support) and 1R01GM121703, as well as awards from the Oklahoma Center for Adult Stem Cell Research.

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
<ol>
	<li>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.</li>
	<li>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.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
2. Timed matings - collecting, sorting, and housing zebrafish embryos for experiments
<ol>
	<li>Perform timed matings, beginning as soon as the light cycles being in the morning. Discard any embryos generated overnight.</li>
	<li>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.</li>
	<li>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 &#176;C.</li>
	<li>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.</li>
	<li>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.
	<ol>
		<li>Count embryos and place at a density of 100 embryos per 10 cm plate.</li>
		<li>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 &#176;C incubator.</li>
	</ol>
	</li>
</ol>
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).
<ol>
	<li>When embryos reach desired developmental age, verify appropriate stage of development morphologically under a light microscope according to &#34;Stages of Embryonic Development of the Zebrafish&#34;17.</li>
	<li>Transfer up to 1500 staged embryos to a 15-mL conical tube and wash several times with E3.
	<ol>
		<li>Add 1 mL of E3 and 20 &#181;L 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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Remove E3 and wash embryos with freshly prepared ice-cold deyolking buffer (0.5x Ginzburg Fish Ringer&#39;s solution, without calcium: 55 mM NaCl, 1.8 mM KCl, 1.25 mM NaHCO3).
	<ol>
		<li>Add 1 mL of ice-cold deyolking buffer for up to 500 embryos.</li>
		<li>Using a P1000, gently pipette embryos up and down 5-10 times to disrupt yolk sack.</li>
		<li>Transfer 1 mL to a 1.5 mL microfuge tube and centrifuge at 4 &#176;C and 500 x g for 5 min.</li>
	</ol>
	</li>
	<li>Remove supernatant and wash pellet with 1 mL of ice-cold 1x PBS (phosphate buffered saline, pH 7.0) to remove Ringer&#39;s solution. Centrifuge at 4 &#176;C and 500 x g for 5 min.</li>
	<li>Carefully remove supernatant and resuspend cells in 1 mL of 1x PBS. Place tube on ice.</li>
	<li>Add 3 mL of ice-cold ethanol (EtOH) to a 15-mL conical tube. Vortex tube of EtOH gently on low setting.
	<ol>
		<li>Using a P1000, slowly (1 drop per second) drip zebrafish embryo cell suspension into EtOH while vortexing gently.</li>
		<li>Add a total of 1 mL of the embryo cell suspension, for a final fixation solution of 75% EtOH.</li>
	</ol>
	</li>
	<li>After 1 mL of cell suspension has been added, gently swirl tube to mix and place at -20 &#176;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 &#176;C for 2-3 weeks.</li>
</ol>
4. Staining DNA and FACS sorting embryos
NOTE: This section of the protocol is designed for embryos at 1 dpf.
<ol>
	<li>Remove EtOH-fixed embryo cells from -20 &#176;C and centrifuge at 4 &#176;C 1500 x g for 5 min.
	<ol>
		<li>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).</li>
		<li>Centrifuge cells at 4 &#176;C and 1500 x g for 5 min and place on ice.</li>
	</ol>
	</li>
	<li>Remove supernatant and resuspend cells in propidium iodide (PI) staining solution (50 &#181;g/mL PI, 100 &#181;g/mL RNase A, PBS, pH 7.0), using 200 &#181;L for every 1000 embryos.
	<ol>
		<li>Allow cells to incubate in PI solution for 30 min on ice, gently mixing every 5 min.</li>
		<li>Place a 40 &#181;m 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.</li>
		<li>Using the flat inside end of a syringe plunger, gently disrupt any remaining clumps of cells on the mesh.</li>
		<li>Rinse cells through the filter with 500 &#181;L of cold PBS-BSA for every 1000 embryos.</li>
		<li>Place a 40-&#181;m 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.</li>
	</ol>
	</li>
	<li>Using an appropriate cell sorting instrument, set the laser excitation to 561-nm and emission detection to 610/20 to detect propidium iodide.
	<ol>
		<li>Mix 15 mL tube of stained cells briefly by vortexing and place in instrument at 4 &#176;C.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Second, gate for SSC-area (SSC-A) by PI-area (PI-A), to remove debris and cell doublets (Figure 1B).</li>
		<li>Third, gate for SSC-A by SSC-W (width) to remove any remaining cell doublets (Figure 1C).</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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).</li>
		<li>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).</li>
		<li>Back gate the G1 and S phase populations to ensure proper initial gating (Figure 1E).</li>
	</ol>
	</li>
	<li>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).</li>
	<li>After sorting is complete, centrifuge tubes at 1500 x g and 4 &#176;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.</li>
	<li>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 &#176;C for 5 min.</li>
	<li>Carefully remove all liquid from tube and freeze dry pellet at -20 &#176;C. Store the pellet at -20 &#176;C for 2-3 weeks.</li>
</ol>
5. DNA isolation, RNase treatment, and DNA purification
<ol>
	<li>Remove cell pellet from -20 &#176;C and place on ice.
	<ol>
		<li>Add 400 &#181;L of SDS buffer (50 mM Tris-HCl (pH 8.0), 10 mM EDTA, 1 M NaCl, 0.5% SDS) to pellet and resuspend.</li>
		<li>Add proteinase K to a final concentration of 0.2 mg/mL and mix by vortexing briefly.</li>
		<li>Incubate samples at 56 &#176;C for 2 h, vortex every 15 min.</li>
	</ol>
	</li>
	<li>After 2 h, perform phenol-chloroform extraction by adding 400 &#181;L of phenol-chloroform (Phenol:Chloroform:Isoamyl Alcohol 25:24:1, saturated with 10 mM Tris, pH 8.0, 1mM EDTA).
	<ol>
		<li>Vortex for 20 s and centrifuge sample at 16,000 x g for 5 min to separate phases.</li>
		<li>Carefully remove the upper aqueous phase (400 &#181;L) and transfer to a 1.5 mL tube.</li>
	</ol>
	</li>
	<li>Perform EtOH precipitation by adding 40 &#181;L of 3M sodium acetate (pH 5.2), 2 &#181;L of glycogen, and 1 mL of EtOH.
	<ol>
		<li>Vortex to mix thoroughly and place at -20 &#176;C overnight to precipitate DNA. Sample can also be precipitated at -80 &#176;C or on dry ice for 1&#160;h.</li>
		<li>Centrifuge sample at 4 &#176;C and 16,000 x g for 30 min to pellet DNA.</li>
		<li>Discard supernatant and wash pellet with 1 mL 70% EtOH. Centrifuge at 4 &#176;C and 16,000 x g for 5 min.</li>
		<li>Discard supernatant and air-dry pellet at ambient temperature for 2-3 min.</li>
	</ol>
	</li>
	<li>Dissolve pellet in 96 &#181;L of autoclaved water, and add 4 &#181;L of RNase A (2 mg/mL).
	<ol>
		<li>Mix well by vortexing and incubate at 37 &#176;C for 1&#160;h.</li>
	</ol>
	</li>
	<li>Perform phenol-chloroform extraction by adding 100 &#181;L of phenol-chloroform and following procedure in steps 5.2.1 - 5.2.2.</li>
	<li>Perform EtOH precipitation by adding 10 &#181;L of 3M sodium acetate, 1 &#181;L glycogen, and 250 &#181;L of EtOH and following the procedure in steps 5.3.1 - 5.3.5.</li>
	<li>Resuspend pellet in 60 &#181;L 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 &#176;C. 
	NOTE: It is important to quantify DNA using fluorescence-based DNA binding dyes, or other means more reliable than UV spectrometer-based methods.</li>
</ol>
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.
<ol>
	<li>Dilute 1 &#181;g of DNA to a final volume of 50 &#181;L and transfer to a specialized tube for sonication.</li>
	<li>Shear genomic DNA for a target of 250-300 bp in a focused ultrasonicator, according to manufacturer&#39;s specifications and protocol.</li>
	<li>Verify quality and size of sheared genomic DNA using an automated electrophoresis machine, according to manufacturer&#39;s specifications and protocol. Ensure there is a large peak of ~250-300 bp for the sheared genomic DNA.</li>
	<li>Prepare sequencing libraries using a DNA library preparation kit with multiplex oligos for sequencing on Illumina platform, according to manufacturer&#39;s protocol.</li>
	<li>Verify quality of library prep using automated electrophoresis machine, according to manufacturer&#39;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).</li>
	<li>Quantify libraries using a DNA library quantification kit for next-generation DNA sequencing, according to manufacturer&#39;s protocol.</li>
	<li>Dilute 15 &#181;L 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.</li>
	<li>Perform paired-end 100 bp whole genome DNA sequencing of samples on a next generation sequencing platform, according to manufacturer&#39;s instructions.</li>
</ol>
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.
<ol>
	<li>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
	<ol>
		<li>Uncompress the fa.gz file with the following command: 
		gunzip danRer10.fa.gz</li>
		<li>Align sequencing data to genome using BWA-mem using the following commands: 
		bwa mem -t 8 danRer10.fa read_1.fastq read_2.fastq &#62; output.sam</li>
	</ol>
	</li>
	<li>Convert .sam file to .bam file using samtools with the following command: 
	samtools view -bS input.sam &#62; output.bam
	<ol>
		<li>Sort the aligned sequencing files using samtools with the following command: 
		samtools sort output.bam &#62; output_sorted.bam</li>
		<li>Index the .bam file using samtools with the following command: 
		samtools index output.bam</li>
	</ol>
	</li>
	<li>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</li>
	<li>Generate text (.txt) files of reads for each chromosome using the following command: 
	for i in {1.25}; do CHROM=&#34;chr&#34;$i; echo $CHROM; samtools view input.bam $CHROM | cut -f 2,4,5,7,8,9 &#62; readsChr${i}.txt; done 
	NOTE: The columns in the resulting .txt file are Flag, Location, mapQ, pair chr, pair location, and fragment size, respectively.</li>
	<li>Read .txt files into Matlab (or any other similar software) using the textscan function.
	<ol>
		<li>Filter out low mapping quality reads by setting a mapQ threshold of 30.</li>
		<li>Remove reads with flags for not primary alignment, PCR duplicates, or pair mapping to a different chromosome.</li>
		<li>Filter out any reads with their pair beyond a distance threshold of 2000 bp.</li>
		<li>Evaluate statistics of resulting reads to determine number of read statistics, coverage, insert size, and quality (Figure 2).</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>Calculate the median read count for all windows.</li>
		<li>Filter out regions of high coverage by removing windows greater than 5 standard deviations from the median.</li>
	</ol>
	</li>
	<li>Define regions for analysis in the G1 reference samples by finding segments along each chromosome with 200 reads using sliding windows.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Smooth the raw data using the csaps function in the software, with an interpolation factor (p) of 10-16.</li>
	<li>Normalize smoothed data to z-score with a mean of 0 and standard deviation of 1.</li>
</ol>

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	<li>Rhind, N., Gilbert, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+replication+timing.">DNA replication timing.</a> Cold Spring Harb Perspect Biol. 5 (8), a010132 (2013).</li><li>Pope, B. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Topologically+associating+domains+are+stable+units+of+replication-timing+regulation.">Topologically associating domains are stable units of replication-timing regulation.</a> Nature. 515 (7527), 402-405 (2014).</li><li>Rivera-Mulia, J. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+changes+in+replication+timing+and+gene+expression+during+lineage+specification+of+human+pluripotent+stem+cells.">Dynamic changes in replication timing and gene expression during lineage specification of human pluripotent stem cells.</a> Genome Res. 25 (8), 1091-1103 (2015).</li><li>Hiratani, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+reorganization+of+replication+domains+during+embryonic+stem+cell+differentiation.">Global reorganization of replication domains during embryonic stem cell differentiation.</a> PLoS Biol. 6 (10), e245 (2008).</li><li>Hiratani, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genome-wide+dynamics+of+replication+timing+revealed+by+in+vitro+models+of+mouse+embryogenesis.">Genome-wide dynamics of replication timing revealed by in vitro models of mouse embryogenesis.</a> Genome Res. 20 (2), 155-169 (2010).</li><li>Koren, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+relationship+of+DNA+replication+timing+to+different+forms+of+human+mutation+and+variation.">Differential relationship of DNA replication timing to different forms of human mutation and variation.</a> Am J Hum Genet. 91 (6), 1033-1040 (2012).</li><li>Ryba, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Abnormal+developmental+control+of+replication-timing+domains+in+pediatric+acute+lymphoblastic+leukemia.">Abnormal developmental control of replication-timing domains in pediatric acute lymphoblastic leukemia.</a> Genome Res. 22 (10), 1833-1844 (2012).</li><li>Sima, J., Gilbert, D. 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The authors have nothing to disclose.

Zebrafish provide a new and unique in vivo model system to study DNA replication timing. When timed matings are performed as detailed in this experimental protocol, thousands of embryos can be collected in a single day for experiments. These embryos develop synchronously through precisely timed and distinctly characterized stages of development. Zebrafish can be easily and accurately staged by morphology using a stereomicroscope, as zebrafish embryos develop externally and are optically clear. This protocol details the use of zebrafish embryos to study replication timing throughout vertebrate development.
Areas of the protocol that may present problems include ensuring synchronous development of zebrafish embryos, loss of cells during preparation, FACS sorting, and library preparation. The timing of zebrafish development is temperature dependent, therefore use zebrafish adapted to light/dark cycles and housed at 28.5 &#176;C for experiments. To ensure synchronous development, perform precisely timed matings, and collect embryos in very small time-windows (10 min or less). It is critical to maintain embryos at 28.5 &#176;C and ensure they spend minimal time at ambient temperature. The timing of zebrafish development is also highly dependent on the density of embryos in a given area. It is critical to not house embryos at a density higher than 100 embryos per 10 cm plate, or they will not develop properly. The optimal time to remove dead and unfertilized embryos is when they have progressed to the 4-16 cell stage and can easily be identified as fertilized (use &#34;Stages of Embryonic Development of the Zebrafish&#34;17 as a guide). Dead embryos typically appear as black balls and unfertilized embryos appear as 1-cell. Work as quickly as possible to reduce the time at ambient temperature.
When dechorionating embryos, it is important to ensure all chorions are removed from the embryos. Keep embryos in pronase&#160;solution until most chorions have come off. Discard any remaining embryos with chorions intact or remove their chorions with forceps. Pipette embryos using a slow and smooth motion. Do not pipette rapidly as it will subject the cells to undue stress and result in loss of cells. It is also important not to use a P200 as this can cause increased shear stress that results in breakage of cells. Conversely, a plastic transfer pipette may not have a small enough bore or provide enough shear stress to adequately disrupt the yolk sack and disaggregate the cells. Alternative pipettes could be used provided the bore and shear would be equivalent to the optimal P1000.
When FACS sorting 28 hpf, if a stereotypical cell cycle profile is not obtained, adjust the voltage on the PI laser so that the G1 peak is centered around 50K. The gains for the FSC and SSC may also need to be adjusted such that the gating and cell populations appear similar to Figure 1. If there is still difficulty, the ND filter should be switched, and the FSC and SSC voltages adjusted to ensure the majority of cells are within the gate. It should be obvious whether there is PI staining, as there will be a range that approximates doubles for the PI-A. The histogram should show two clear peaks, a large peak at 2N DNA for the G1 cells, and a smaller peak at double the intensity for the 4N DNA content of G2/M cells. If this is still not obtained there may either be low numbers of intact cells, problems with the PI staining, or problems with fixation and the protocol will need to be optimized accordingly.
Cells from early embryos (prior to 10 hpf) will not give typical cell cycle profiles, as a high percentage of these cells are in S phase. These samples can be treated as S phase samples, and compared to a G1 reference from 24 hpf embryos. Embryos between 10 hpf and 24 hpf may give intermediate cell cycle profiles and can be sorted if desired. The S-phase population can also be identified and sorted by incorporating thymidine analogs such as EDU (5-Ethynyl-2&#39;-deoxyuridine) or BrdU (Bromodeoxyuridine) in brief pulses prior to fixation, however, this is not necessary for accurate determination of replication timing.
Ideally the library preparation should be started with 1 &#181;g of DNA. Theoretically ~300,000 sorted cells would yield about 1 &#181;g of DNA (estimating ~3.3 pg/nucleus), however, there is always loss throughout the protocol. 500,000 to 1 million sorted cells should give more than 1 &#181;g of DNA. A typical 24 hpf embryo will have approximately 18,000 cells, 10-15% of which will be in S-phase. Embryos at earlier stages of development will have considerably less cells, but will have higher percentages of cells in S-phase.
Performing accurate purification and size selection using magnetic beads is critical to eliminating unwanted elements from the library preparation. Follow the manufacturer&#39;s instructions carefully and additional optimization may be required. A small amount of PCR dimers is typically not a major problem, but adapter dimers will sequence very efficiently so they should be minimized. This can be achieved by adjusting the initial adapter to DNA ratio, and performing accurate size selection during library preparation.
Single-end sequencing may also be appropriate for different experimental needs. Single-end is cheaper and provides most of the required information since this approach relies on counting reads; paired-end provides a higher ability to remove PCR and optical duplicates and to resolve areas of repetitive sequences and other structural variations (which are common in the zebrafish genome). The analysis portion below will deal only with paired-end sequencing reads. Shorter sequencing reads may also be appropriate. This will provide a higher number of reads at a lower cost, but the reads may be more difficult to map. The analysis portion below is optimized for 100 bp reads, and adjustments may be necessary if shorter read length is used.
This protocol can also easily be adapted for additional uses of the zebrafish model. It has already been used for analysis of zebrafish tailfin fibroblasts (ZTF cells), a primary cell culture line isolated from adult zebrafish tailfin. To study isolated cell types from zebrafish, transgenic markers can be used to identify and isolate individual cell types before staining and sorting for DNA content. One potential strategy would be to use a transgenic line expressing a fluorescent marker driven by a tissue-specific promoter, and to first isolated cells by FACS sorting, followed fixation and sorting for DNA content. Additionally, the use of cell permeable DNA dyes may allow for simultaneous isolation of individual cell types as well as G1 and S phase populations. These adaptations to this protocol could also enable replication timing studies in other in vivo models.
An exciting possibility for the future use of this protocol is to study the role of replication timing in disease. There are several cancer models in zebrafish, some spontaneous and other inducible, that can be used to determine when changes in replication timing occur during oncogenesis. This will allow assessment of the role replication timing plays in tumorigenesis and disease progression. Furthermore, zebrafish are an excellent model for drug screening and can be used to identify drugs that affect replication timing regulation, which have the potential for use in cancer treatment.
The zebrafish is a promising new model to study replication timing. This protocol will afford many others the opportunity to utilize this model in studying the role of replication timing in development and disease. This protocol can be adapted for use with other in vivo developmental model systems, such as Drosophila melanogaster and Xenopus laevis, and look forward to future findings from these and other organisms.

For cells to successfully divide, they must first accurately and faithfully replicate their entire genome. Genome duplication occurs in a reproducible pattern, known as the DNA replication timing program1. DNA replication timing is correlated with chromatin organization, epigenetic marks, and gene expression2,3. Changes in replication timing occur throughout development, and are significantly related to transcriptional programs and alterations to chromatin marks and organization4,5. Furthermore, replication timing is correlated with mutational frequencies, and changes in timing are observed in various types of cancer6,7,8. Despite these observations, the mechanisms and determinants of replication timing establishment and regulation are still largely unknown, and the role it plays in development and disease is undetermined. In addition, until recently the genome-wide replication timing changes that occur throughout vertebrate development had only been examined in cell culture models.
Zebrafish, Danio rerio, are well suited to study replication timing in vivo during development, as a single mating pair can yield of hundreds of embryos that develop rapidly with many similarities to mammalian development9,10. Furthermore, throughout zebrafish development, there are changes to the cell cycle, chromatin organization, and transcriptional programs that share relationships with DNA replication timing11. Zebrafish are also an excellent genetic model, as they are particularly amenable to manipulation by transgenesis, mutagenesis, and targeted mutations, and genetic screens have identified many genes required for vertebrate development12. Therefore, zebrafish can be used to identify genes involved in replication timing establishment and maintenance and to observe the effects of deregulating replication timing on vertebrate development. Transgenic lines can also be used to assess replication timing from individual cell types isolated at different developmental timepoints or in disease conditions. Importantly, there are various zebrafish models of human disease that can be used to investigate the role of replication timing in disease formation and progression9,13,14.
Recently, the first replication timing profiles were generated from zebrafish, establishing it as a model system to study replication timing in vivo15. To accomplish this, cells were collected from zebrafish embryos at multiple stages of development and in a cell type isolated from adult zebrafish. Cells were then sorted by FACS (fluorescence-activated cell sorting) based on DNA content to isolate G1 and S phase populations. Using the G1 sample as a copy number control, copy number variations in S phase populations were determined and used to infer relative replication timing16. Changes in replication timing can then be directly compared between different developmental samples and cell types and this was used to determine changes in replication timing that occur in vivo throughout vertebrate development. This method offers several advantages over other genomic methods, chiefly that it does not require labeling with thymidine analogs or immunoprecipitation of DNA4,6.
Here is detailed the protocols to profile genome-wide DNA replication timing at high-resolution in zebrafish. These protocols have been used to determine relationships with genomic and epigenetic features in the zebrafish genome, as well as profiling changes in these relationships that occur throughout development. These protocols are also easily adapted to study changes in replication timing in mutant strains of zebrafish and in disease models. Additionally, these methods provide a foundation that can be expanded upon to study replication timing in specific cell types, by first sorting out the individual cell types from the zebrafish. The zebrafish can serve as an excellent in vivo model system to study replication timing and to ultimately reveal the biological functions of this important epigenetic trait.

<table><tbody><tr><td>NaCl</td><td>Fisher Scientific</td><td>BP358-10</td><td></td></tr><tr><td>KCl</td><td>Fisher Scientific</td><td>P217-500</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Fisher Scientific</td><td>C79-500</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>EMD Millipore</td><td>MMX00701</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Fisher Scientific</td><td>BP328-500</td><td></td></tr><tr><td>Pronase</td><td>Sigma</td><td>10165921001</td><td>protease solution</td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>Sigma</td><td>D1408</td><td></td></tr><tr><td>Ethanol (EtOH)</td><td>KOPTEC</td><td>V1016</td><td></td></tr><tr><td>Bovine serum albumin (BSA)</td><td>Sigma</td><td>A9647-100G</td><td></td></tr><tr><td>Propidium Iodide (PI)</td><td>Invitrogen</td><td>P3566</td><td></td></tr><tr><td>Tris-HCl</td><td>Fisher Scientific</td><td>BP153-500</td><td></td></tr><tr><td>EDTA</td><td>Sigma</td><td>E9844</td><td></td></tr><tr><td>SDS</td><td>Santa Cruz</td><td>sc-24950</td><td></td></tr><tr><td>Proteinase K</td><td>NEB</td><td>P8107S</td><td></td></tr><tr><td>Phenol:Chloroform</td><td>Sigma</td><td>P3803-100ML</td><td></td></tr><tr><td>Sodium acetate</td><td>J.T.Baker</td><td>3470</td><td></td></tr><tr><td>Glycogen</td><td>Ambion</td><td>AM9510</td><td></td></tr><tr><td>RNase A</td><td>Thermo Scientific</td><td>EN0531</td><td></td></tr><tr><td>Quanit-iT</td><td>Invitrogen</td><td>Q33130</td><td>Reagents for fluorescence-based DNA quantification</td></tr><tr><td>Covaris AFA microTUBE</td><td>Covaris</td><td>520045</td><td>specialized tube for sonication</td></tr><tr><td>Covaris E220 Sonicator</td><td>Covaris</td><td>E220</td><td>focused ultrasonicator</td></tr><tr><td>Agilent 4200 Tapestation</td><td>Agilent</td><td>G2991AA</td><td>automated electrophoresis machine</td></tr><tr><td>D1000 ScreenTape</td><td>Agilent</td><td>5067-5582</td><td>Reagents for automated electrophoresis machine</td></tr><tr><td>NEBNext Ultra DNA Library Prep Kit for Illumina</td><td>NEB</td><td>Cat#E7370L</td><td>DNA library preparation kit</td></tr><tr><td>NEBNext Multiplex Oligos Kit for Illumina (Index Primers Set 1)</td><td>NEB</td><td>Cat#E7335S</td><td>multiplex oligos for DNA library preparation kit</td></tr><tr><td>NEBNext Multiplex Oligos Kit for Illumina (Index Primers Set 2)</td><td>NEB</td><td>Cat#E7500S</td><td>additional multiplex oligos for DNA library preparation kit</td></tr><tr><td>NEBNext Library Quant Kit for Illumina</td><td>NEB</td><td>E7630L</td><td>quantification kit for library preparation</td></tr><tr><td>Agencourt AMPure XP beads</td><td>Beckman Coulter</td><td>A63882</td><td>magnetic beads</td></tr><tr><td>Illumina HiSeq 2500</td><td>Illumina</td><td>SY&amp;ndash;401&amp;ndash;2501</td><td>next generation DNA sequencing platform</td></tr><tr><td>40 &amp;micro;m Falcon Nylon Cell Strainer</td><td>Fisher Scientific</td><td>08-771-1</td><td></td></tr><tr><td>VWR Disposable Petri Dish 100 x 25 mm</td><td>VWR</td><td>89107-632</td><td></td></tr><tr><td>6.0 mL Syringe for Nichiryo Model 8100</td><td>VWR</td><td>89078-446</td><td></td></tr><tr><td>Posi-Click Tubes, 1.7 mL, Natural Color</td><td>Denville Scientific</td><td>C2170 (1001002)</td><td>Dnase/Rnase free</td></tr><tr><td>Vortex Genie 2</td><td>Scientific Industries</td><td>SI-0236</td><td></td></tr><tr><td>Wash Bottles</td><td>VWR</td><td>16650-022</td><td>Low-Density Polyethylene, Wide Mouth</td></tr><tr><td>Strainer</td><td>VWR</td><td>470092-440</td><td>6.9 cm, fine mesh</td></tr><tr><td>Corssing tank</td><td>Aquaneering</td><td>ZHCT100</td><td>individual breeding tank</td></tr><tr><td>iSpawn</td><td>Techniplast</td><td>N/A</td><td>large breeding tank</td></tr><tr><td>FACSAria II</td><td>BD biosciences</td><td>N/A</td><td>cell sorting machine</td></tr><tr><td>Wild M5a steromicroscope</td><td>Wild Heerbrugg</td><td>N/A</td><td>dissecting microscope</td></tr><tr><td>Qubit 3 Fluorometer</td><td>Thermo Scientific</td><td>Q33216</td><td>quantitative fluorescence-based method for determining DNA concentration</td></tr><tr><td>Matlab</td><td>Mathworks</td><td>version 2017a</td><td></td></tr><tr><td>Matlab Statistics Toolbox</td><td>Mathworks</td><td>version 11.1</td><td></td></tr><tr><td>Matlab Curve Fitting Toolbox</td><td>Mathworks</td><td>version 3.5.5</td><td></td></tr></tbody></table>

profiling dna replication timing using zebrafish as an in vivo model system

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Using published replication timing data, representative replication timing profiles and quality control measures are provided15. The initial steps of processing involve aligning the sequencing data to the genome, calculating read length and genome coverage statistics, and filtering low quality, unpaired, and PCR duplicate reads. Read statistics for a typical zebrafish sequencing sample are shown in Figure 2. After filtering, read counts are determined in variable-sized windows and the data are smoothed and normalized. Typical smoothed/normalized replication timing profiles of a representative zebrafish chromosome for biological replicates are shown in Figure 3A. The profiles for biological and experimental replicates should be visually very similar (see Figure 3A) and also display high correlation along the length of the chromosome (Figure 3B). They should also display high correlation between timing values genome-wide (Figure 3C). Autocorrelation, a method to assess pattern continuity, should be used to evaluate the degree of structure in the data set (Figure 3D). The autocorrelation for structured mature timing programs should be high at short distances and gradually decrease as the distance increases. Samples that show high reproducibility between biological and experimental replicates and a strong autocorrelation signal should be considered as high quality samples and can be combined to obtain a higher coverage sample.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57146/57146fig1.jpg" /> 
Figure 1: Sorting zebrafish embryos for replication timing analysis. (A) Gated population of all cells sorted for forward scatter area (FSC-A) by side scatter area (SSC-A). The colors represent the density of bins in the dot plot with high-to-low density represented by colors ranging from red-to-green-to-blue. (B) Gated population of the FSC x SSC gated cells sorted for SSC-A by propidium iodide area (PI-A). (C) Gated population of the SSC x PI gated cells sorted for SSC-A by side scatter width (SSC-W). (D) Histogram of all gated populations displaying a stereotypical cell cycle profile. Sort gates for cells in G1 and S phase are displayed by the grey shaded boxed with black lines. (E) Backgated G1 and S phase populations show that the initial gating captured the majority of cells. <a href="https://cloudflare.jove.com/files/ftp_upload/57146/57146fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57146/57146fig2.jpg" /> 
Figure 2: Sequencing read statistics for a typical zebrafish sample. (A) Reap mapping quality from statistics of mapQ values. (B) Histogram of the distribution of insert sizes for all sequencing reads. (C) Read statistics for total mapped reads, reads containing low quality flag, reads with pairs mapping to a different chromosome (pair diff chr), reads with pair beyond distance threshold (distant pair), and PCR duplicates. (D) Representative numbers of total mapped, unmapped, and unpaired reads, coverage, and read resolution for a typical sequencing run of a zebrafish sample. <a href="https://cloudflare.jove.com/files/ftp_upload/57146/57146fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57146/57146fig3.jpg" /> 
Figure 3: Representative zebrafish replication timing results and quality control measures. (A) Replication timing along the length of a representative chromosome for two biological replicates of 28 hpf embryos (Siefert, 2017). (B) Correlation between replication timing values in 2 Mb windows along the length of a representative chromosome for the biological replicates of 28 hpf embryos shown in Figure 3A. (C) Genome-wide correlation between replication timing values for biological (rep1 and rep2) and experimental (rep3) replicates of 28 hpf embryos. The color map represents Pearson&#39;s correlation coefficient. (D) Autocorrelation for a structured mature timing program displays high autocorrelation at close distances that decreases gradually over increasingly longer distances. <a href="https://cloudflare.jove.com/files/ftp_upload/57146/57146fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System at JoVE.com

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.

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

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

profiling-dna-replication-timing-using-zebrafish-as-an-vivo-model

Nanyang Technological University

Research

JoVE Journal

Genetics

7.8K Views.  Oklahoma Medical Research Foundation. Zebravis onlangs gewend waren als een modelsysteem in vivo DNA replicatie timing tijdens ontwikkeling bestuderen. Hier is gedetailleerde de protocollen voor het gebruik van embryo's de zebravis profiel replicatie timing. Dit protocol kan gemakkelijk worden aangepast om te bestuderen van replicatie timing mutanten, individuele celtypes, ziekte modellen en andere soorten.

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

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

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, auroraBhi1045, and esco2hi2865 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.

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.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Developmental Biology

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short generation time, are easy to handle, display a high reproductive rate, low cost, and are easily amenable to genetic manipulations by microinjection of DNA in embryos. Recent advances in gene editing tools are enabling precise introduction of mutations and transgenes in zebrafish. Disease modeling in zebrafish often leads to larval phenotypes and early death which can be challenging to interpret if genotypes are unknown. This early identification of genotypes is also needed in experiments requiring sample pooling, such as in gene expression or mass spectrometry studies. However, extensive genotypic screening is limited by traditional methods, which in most labs are performed only on adult zebrafish or in postmortem larvae. We addressed this problem by adapting a method for the isolation of PCR-ready genomic DNA from live zebrafish larvae that can be achieved as early as 72 h post-fertilization (hpf). This time and cost-effective technique, improved from a previously published genotyping protocol, allows the identification of genotypes from microscopic fin biopsies. The fins quickly regenerate as the larvae develop. Researchers are then able to select and raise the desired genotypes to adulthood by utilizing this high-throughput PCR-based genotyping procedure.

Zebrafish have been used as reliable genetic model organisms in biomedical research, especially with the advent of gene-editing technologies. When larval phenotypes are expected, DNA extraction and genotype identification can be challenging. Here, we describe an efficient genotyping procedure for zebrafish larvae, by tail clipping, as early as 72-h post-fertilization.

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short ...

Preparation of Meiotic Chromosome Spreads from Zebrafish Spermatocytes

Meiosis is the key cellular process required to create haploid gametes for sexual reproduction. Model organisms have been instrumental in understanding the chromosome events that take place during meiotic prophase, including the pairing, synapsis, and recombination events that ensure proper chromosome segregation. While the mouse has been an important model for understanding the molecular mechanisms underlying these processes, not all meiotic events in this system are analogous to human meiosis. We recently demonstrated the exciting potential of the zebrafish as a model of human spermatogenesis. Here we describe, in detail, our methods to visualize meiotic chromosomes and associated proteins in chromosome spread preparations. These preparations have the advantage of allowing high resolution analysis of chromosome structures. First, we describe the procedure for dissecting testes from adult zebrafish, followed by cell dissociation, lysis, and spreading of the chromosomes. Next, we describe the procedure for detecting the localization of meiotic chromosome proteins, by immunofluorescence detection, and nucleic acid sequences, by fluorescence in situ hybridization (FISH). These techniques comprise a useful set of tools for the cytological analysis of meiotic chromatin architecture in the zebrafish system. Researchers in the zebrafish community should be able to quickly master these techniques and incorporate them into their standard analyses of reproductive function.

Nuclear surface spreads are an indispensable tool for studying chromosome events during meiosis. Here we demonstrate a method to prepare and visualize meiotic chromosomes during prophase I from zebrafish spermatocytes.

Meiosis is the key cellular process required to create haploid gametes for sexual reproduction. Model organisms have been instrumental in understanding ...

Fluorescent Protein Tagging in Zebrafish Larvae Using CRISPR/Cas9 Knock-In

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in vivo imaging due to its transparency during the early developmental stage. Here, we provide a detailed protocol for performing high-efficiency fluorescence gene KI, rapid screening for KI founders, and low-abundance protein tracing in zebrafish larvae, which will lay a critical foundation for subsequent physio-pathological studies in zebrafish. The current protocol includes complete steps for the sgRNA design for the gene of interest, sgRNA in vitro transcription, Cas9 mRNA in vitro transcription, in vivo sgRNA screen for the one with the highest efficiency, donor plasmid design and construction, microinjection in zebrafish larvae, KI founder screen and zebrafish live imaging. Critical steps, troubleshooting tips, quality control methods, and advantages and applications of this protocol are included and discussed. This protocol assures quick and accurate results at a low cost and has been validated by multiple trials.

Here, we present a protocol to label proteins with a fluorescence protein tag in zebrafish larvae, a newly developed and modified in vivo system especially useful for visualizing low-abundance proteins in zebrafish.

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in ...

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, the muscle stem cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. In many muscle wasting conditions, including muscular dystrophy and ageing, this process is impaired resulting in the inability of muscle to regenerate. The process of muscle regeneration in zebrafish is highly conserved with mammalian systems providing an excellent system to study muscle stem cell function and regeneration, in muscle wasting conditions such as muscular dystrophy. Here, we present a method to examine muscle regeneration in zebrafish models of muscle disease. The first step involves the use of a genotyping platform that allows the determination of the genotype of the larvae prior to eliciting an injury. Having determined the genotype, the muscle is injured using a needle stab, following which polarizing light microscopy is used to determine the extent of muscle regeneration. We therefore provide a high throughput pipeline which allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle regeneration is driven by tissue resident muscle stem cells, which are impaired in many muscle diseases such as muscular dystrophy, and this results in the inability of muscle to regenerate. Here, we describe a protocol that allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, ...

Rapid and Efficient Spatiotemporal Monitoring of Normal and Aberrant Cytosine Methylation within Intact Zebrafish Embryos

Cytosine methylation is highly conserved across vertebrate species and, as a key driver of epigenetic programming and chromatin state, plays a critical role in early embryonic development. Enzymatic modifications drive active methylation and demethylation of cytosine into 5-methylcytosine (5-mC) and subsequent oxidation of 5-mC into 5-hydroxymethylcytosine, 5-formylcytosine, and 5-carboxylcytosine. Epigenetic reprogramming is a critical period during in utero development, and maternal exposure to chemicals has the potential to reprogram the epigenome within offspring. This can potentially cause adverse outcomes such as immediate phenotypic consequences, long-term effects on adult disease susceptibility, and transgenerational effects of inherited epigenetic marks. Although bisulfite-based sequencing enables investigators to interrogate cytosine methylation at base-pair resolution, sequencing-based approaches are cost-prohibitive and, as such, preclude the ability to monitor cytosine methylation across developmental stages, multiple concentrations per chemical, and replicate embryos per treatment. Due to the ease of automated in vivo imaging, genetic manipulations, rapid ex utero development time, and husbandry during embryogenesis, zebrafish embryos continue to be used as a physiologically intact model for uncovering xenobiotic-mediated pathways that contribute to adverse outcomes during early embryonic development. Therefore, using commercially available 5-mC-specific antibodies, we describe a cost-effective strategy for rapid and efficient spatiotemporal monitoring of cytosine methylation within individual, intact zebrafish embryos by leveraging whole-mount immunohistochemistry, automated high-content imaging, and efficient data processing using programming language prior to statistical analysis. To current knowledge, this method is the first to successfully detect and quantify 5-mC levels in situ within zebrafish embryos during early development. The method enables the detection of DNA methylation within the cell mass and also has the ability to detect cytosine methylation of yolk-localized maternal mRNAs during the maternal-to-zygotic transition. Overall, this method will be useful for the rapid identification of chemicals that have the potential to disrupt cytosine methylation in situ during epigenetic reprogramming.

This paper describes a protocol for the rapid and efficient spatiotemporal monitoring of normal and aberrant cytosine methylation within intact zebrafish embryos.

Cytosine methylation is highly conserved across vertebrate species and, as a key driver of epigenetic programming and chromatin state, plays a critical ...

Metabolic Profile Analysis of Zebrafish Embryos

A growing goal in the field of metabolism is to determine the impact of genetics on different aspects of mitochondrial function. Understanding these relationships will help to understand the underlying etiology for a range of diseases linked with mitochondrial dysfunction, such as diabetes and obesity. Recent advances in instrumentation, has enabled the monitoring of distinct parameters of mitochondrial function in cell lines or tissue explants. Here we present a method for a rapid and sensitive analysis of mitochondrial function parameters in vivo during zebrafish embryonic development using the Seahorse bioscience XF 24 extracellular flux analyser. This protocol utilizes the Islet Capture microplates where a single embryo is placed in each well, allowing measurement of bioenergetics, including: (i) basal respiration; (ii) basal mitochondrial respiration (iii) mitochondrial respiration due to ATP turnover; (iv) mitochondrial uncoupled respiration or proton leak and (iv) maximum respiration. Using this approach embryonic zebrafish respiration parameters can be compared between wild type and genetically altered embryos (mutant, gene over-expression or gene knockdown) or those manipulated pharmacologically. It is anticipated that dissemination of this protocol will provide researchers with new tools to analyse the genetic basis of metabolic disorders in vivo in this relevant vertebrate animal model.

Zebrafish represent a powerful vertebrate model that has been under-utilised for metabolic studies. Here we describe a rapid way to measure the in vivo metabolic profile of developing zebrafish that allows the comparison of different mitochondrial function parameters between genetically or pharmacologically manipulated embryos, thereby increasing the applicability of this organism.

A growing goal in the field of metabolism is to determine the impact of genetics on different aspects of mitochondrial function. Understanding these ...

Biology

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The specification of replication timing is thought to be a dynamic process regulated by tissue-specific and developmental cues that are responsive to epigenetic modifications. However, the mechanisms regulating where and when DNA replication initiates along chromosomes remains poorly understood. Homologous chromosomes usually replicate synchronously, however there are notable exceptions to this rule. For example, in female mammalian cells one of the two X chromosomes becomes late replicating through a process known as X inactivation1. Along with this delay in replication timing, estimated to be 2-3 hr, the majority of genes become transcriptionally silenced on one X chromosome. In addition, a discrete cis-acting locus, known as the X inactivation center, regulates this X inactivation process, including the induction of delayed replication timing on the entire inactive X chromosome. In addition, certain chromosome rearrangements found in cancer cells and in cells exposed to ionizing radiation display a significant delay in replication timing of &gt;3 hours that affects the entire chromosome2,3. Recent work from our lab indicates that disruption of discrete cis-acting autosomal loci result in an extremely late replicating phenotype that affects the entire chromosome4. Additional 'chromosome engineering' studies indicate that certain chromosome rearrangements affecting many different chromosomes result in this abnormal replication-timing phenotype, suggesting that all mammalian chromosomes contain discrete cis-acting loci that control proper replication timing of individual chromosomes5.
Here, we present a method for the quantitative analysis of chromosome replication timing combined with fluorescent in situ hybridization. This method allows for a direct comparison of replication timing between homologous chromosomes within the same cell, and was adapted from6. In addition, this method allows for the unambiguous identification of chromosomal rearrangements that correlate with changes in replication timing that affect the entire chromosome. This method has advantages over recently developed high throughput micro-array or sequencing protocols that cannot distinguish between homologous alleles present on rearranged and un-rearranged chromosomes. In addition, because the method described here evaluates single cells, it can detect changes in chromosome replication timing on chromosomal rearrangements that are present in only a fraction of the cells in a population.

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

A quantitative method for the analysis of chromosome replication timing is described. The method utilizes BrdU incorporation in combination with fluorescent in situ hybridization (FISH) to assess replication timing of mammalian chromosomes. This technique allows for the direct comparison of rearranged and un-rearranged chromosomes within the same cell.

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The ...

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present methods for the development of assays to query potentially clinically significant nonsynonymous changes using in vivo complementation in zebrafish. Zebrafish (Danio rerio) are a useful animal system due to their experimental tractability; embryos are transparent to enable facile viewing, undergo rapid development ex vivo, and can be genetically manipulated.1 These aspects have allowed for significant advances in the analysis of embryogenesis, molecular processes, and morphogenetic signaling. Taken together, the advantages of this vertebrate model make zebrafish highly amenable to modeling the developmental defects in pediatric disease, and in some cases, adult-onset disorders. Because the zebrafish genome is highly conserved with that of humans (~70% orthologous), it is possible to recapitulate human disease states in zebrafish. This is accomplished either through the injection of mutant human mRNA to induce dominant negative or gain of function alleles, or utilization of morpholino (MO) antisense oligonucleotides to suppress genes to mimic loss of function variants. Through complementation of MO-induced phenotypes with capped human mRNA, our approach enables the interpretation of the deleterious effect of mutations on human protein sequence based on the ability of mutant mRNA to rescue a measurable, physiologically relevant phenotype. Modeling of the human disease alleles occurs through microinjection of zebrafish embryos with MO and/or human mRNA at the 1-4 cell stage, and phenotyping up to seven days post fertilization (dpf). This general strategy can be extended to a wide range of disease phenotypes, as demonstrated in the following protocol. We present our established models for morphogenetic signaling, craniofacial, cardiac, vascular integrity, renal function, and skeletal muscle disorder phenotypes, as well as others.

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present a systematic approach for developing physiologically relevant, sensitive and specific in vivo assays for interpreting variation in human pathology. Transient genetic manipulation via microinjection of WT and mutant human mRNA and morpholino (MO) antisense oligonucleotides harness the tractability of the developing zebrafish embryo to rapidly assay pathogenic mutations, especially, but not exclusively, in the context of human developmental disorders.

Here,  we present methods for the development of assays to query potentially clinically  significant nonsynonymous changes using in  vivo complementation ...

Rapid and Efficient Zebrafish Genotyping Using PCR with High-resolution Melt Analysis

Zebrafish is a powerful vertebrate model system for studying development, modeling disease, and performing drug screening. Recently a variety of genetic tools have been introduced, including multiple strategies for inducing mutations and generating transgenic lines. However, large-scale screening is limited by traditional genotyping methods, which are time-consuming and labor-intensive. Here we describe a technique to analyze zebrafish genotypes by PCR combined with high-resolution melting analysis (HRMA).&#160;This approach is rapid, sensitive, and inexpensive, with lower risk of contamination artifacts.&#160;Genotyping by PCR with HRMA can be used for embryos or adult fish, including in high-throughput screening protocols.

PCR combined with high-resolution melt analysis (HRMA) is demonstrated as a rapid and efficient method to genotype zebrafish.

Zebrafish is a powerful vertebrate model system for studying development, modeling disease, and performing drug screening. Recently a variety of genetic ...

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

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