Global and gene specific DNA methylation changes during zebrafish development
DNA methylation is dynamic through the life of an organism. Previous studies have primarily focused on DNA methylation changes during very early embryogenesis. In this study, global and gene specific DNA methylation in zebrafish (Danio rerio) embryos, larvae and adult liver were compared. The percent methylation of cytosines was low in 2 to 4.3 hours post fertilization (hpf) zebrafish embryos and was consistently higher in zebrafish older than 6 hpf. Furthermore, quantitative real-time PCR (qPCR) results showed relatively high DNA methyltransferase 1 (dnmt1) and low glycine N-methyltransferase (gnmt) mRNA expression in early embryogenesis. By studying methylation patterns and gene expression of five developmentally important genes, namely vasa, Ras-association domain family member 1 (rassf1), telomerase reverse transcriptase (tert), c-jun and c-myca, we found that the timing of changes in DNA methylation patterns was gene specific, and changes in gene expression were not necessarily correlated with the DNA methylation patterns.
1. Introduction<!>2.1. Zebrafish care<!>2.2 Genomic DNA isolation<!>2.3 Global DNA methylation measurement<!>2.4 Exon, intron identification and gene homology<!>2.5 CpG island identification and classification<!>2.6 Sodium bisulfite sequencing<!>2.7 Quantitative reverse transcription real time PCR (qPCR)<!>2.8 Statistical analysis<!>3.1 Constitutive global DNA methylation changes during development<!>3.2 Constitutive zebrafish dnmt1 and glycine N-methyltransferase (gnmt) mRNA expression during development<!>3.3 De novo DNA methylation in the vasa promoter during zebrafish development<!>3.4 Constitutive vasa mRNA expression during zebrafish development<!>3.5 Constitutive DNA methylation and gene expression of rassf1, tert, c-jun, and c-myca during development<!>3.6 Efficiency of sodium bisulfite treatment<!>3.7 CG loss in genomic DNA<!>4. Discussion<!>5. Conclusion
<p>DNA methylation is the best studied epigenetic mechanism, and the only one that can covalently modify DNA. DNA methylation is mostly found at cytosine residues that are followed by guanines (CpG sites) (Gruenbaum et al., 1981), and approximately 2–5% of all cytosines are methylated in mammalian genomes (Fuke et al., 2004). CpG islands (CGI) are regions with densely clustered 'CG' dinucleotides that are typically unmethylated in active genes. DNA methylation patterns are established during early development by de novo DNA methyltransferases (dnmts). The already established methylation pattern can be copied from the parental strand into the daughter strand after each cell division by a semiconservative maintenance dnmt (Razin and Riggs, 1980). Therefore, DNA methylation patterns are stably memorized and have long-term impact on an organism's phenotype without any changes in the genotype.</p><p>However, DNA methylation patterns are also dynamic during development. In mammals, DNA methylation patterns experience dramatic changes in the inner cell mass (lineage-specific) during early development and in the early primordial germ cells (PGCs, germ-line specific) after sex determination (Anway and Skinner, 2006). After fertilization, both paternal and maternal genomes undergo a rapid global loss of methylation and hit the lowest point by the blastocyst stage (Mayer et al., 2000;Sanz et al., 2010). At the morula and blastocyst stages, extensive acquisition of de novo DNA methylation occurs and eventually about 70% or more of CpGs are rapidly remethylated (Shi and Wu, 2009). A second round of genome-wide methylation reprogramming occurs in PGCs after sex determination. The methylation level remains low in the PGCs until de novo methylation occurs during germ cell maturation and gametogenesis (Rousseaux et al., 2005;Lees-Murdock and Walsh, 2008). Currently, the significance of genome-wide methylation reprogramming is not well understood but disruptions in this highly coordinated process may contribute to developmental defects or embryo death (Haaf, 2006). DNA methylation patterns are believed to be established in early embryogenesis, but modest gains and losses of CpG methylation were observed in the transition of embryonic stem cells to differentiated cells (Meissner et al., 2008; Mohn et al., 2008). Although most epigenetic marks are maintained after birth, DNA methylation patterns are still changeable by external environment interventions, and the epigenome is the most plastic during early postnatal development (Fraga et al., 2005;Bjornsson et al., 2008).</p><p>At the other end of the life spectrum, DNA methylation changes are part of normal aging and tend to associate with age-onset pathologic phenotypes (Gravina and Vijg, 2010). The expression and activity of dnmt1 decline and errors of methylation maintenance increase in aging individuals (Lopatina et al., 2002). As a result, global hypomethylation is detected in most vertebrate tissues with aging (Wilson et al., 1987; Barbot et al., 2002; Bollati et al., 2009). The relaxation of epigenetic control of genes contributes to abnormal gene expression (Bennett-Baker et al., 2003), functional decline in organs, and progression of age-related diseases (Wang et al., 2008).</p><p>Zebrafish are an important model organism for developmental biology and related scientific fields because its genome has approximately 70% homology to human genes (Howe et al., 2013). Mechanisms of DNA methylation in fish are generally conserved with that of mammals (Walter et al., 2002; Varriale and Bernardi, 2006; Aniagu et al., 2008; Zemach et al., 2010). For example, most of the effector proteins in the methylation machinery have been identified in zebrafish (Goll and Halpern, 2011). In the past decade, a few studies have investigated the DNA methylation changes during early embryogenesis in zebrafish. After initial debates, it is now clear that global DNA demethylation and reestablishment are conserved in fish (McGowan and Martin, 2002; Mhanni and McGowan, 2004; MacKay et al., 2007). However, there are no studies so far to measure the DNA methylation changes in later developmental stages and aging adult zebrafish. Therefore, in this study, we measured the global and gene specific DNA methylation in embryos, larvae and adult livers (1.5-year old).</p><p>For specific gene studies, five important developmental genes, i.e. vasa, Ras association domain family member 1 (rassf1), telomerase reverse transcriptase (tert), c-jun and c-myca, were selected for sodium bisulfite sequencing and gene expression analysis. Expression of vasa is essential for germline development and is used as a marker gene to distinguish the germ cell (including PGCs) population (Fujiwara et al. 1994; Castrillon et al., 2000). The tumor suppressor gene rassf1 was selected because its promoter methylation has strong correlation with gene expression in mammals, and it is one of the most frequently silenced genes by promoter hypermethylation in many cancer types (Richter et al., 2009). Tert is a catalytic subunit of telomerase, and its promoter activity is partially regulated by DNA methylation and histone modulation (Kyo et al., 2008). Both c-jun and c-myca are proto-oncogenes essential for embryogenesis because genetic deletion of either of them can lead to embryonic lethality (Eferl et al., 1999; Soucek and Evan, 2010). The target regions of the five genes represent sequences with different CpG richness, and they are located on five different chromosomes. Importantly, the CGI distribution, splicing and protein homology of the five genes are well conserved between zebrafish and mammals (Supplemental Table 1 and Supplemental Figure 1).</p><p>In this study, a statistically significant increase in global methylation was found by 6 hours post fertilization (hpf) in zebrafish. Yet, a unique timing of methylation pattern establishment in the five specific genes, vasa, rassf1, tert, c-jun and c-myca, was observed. Finally, there was an inverse relationship between DNA methylation and gene expression in vasa. Our results indicate that the process of DNA remethylation is dynamic, complicated, and time-specific for individual genes. Future studies are needed to understand the biological significance of global DNA methylation during early development.</p><!><p>AB line wild-type zebrafish (Danio rerio) were purchased from Zebrafish International Resource Center (ZFIN, Eugene, OR, USA) and raised under the approved IACUC protocol. Fish were kept in Aquatic Habitats ZF0601 Zebrafish Stand-Alone System (Aquatic Habitats, Apopka, FL, USA) with zebrafish water (pH 7.0–7.5, 60 parts per million (ppm), Instant Ocean, Cincinnati, OH) at 28 °C (range 24–30 °C), 14:10 light-dark cycle. Adult fish were fed twice daily with tropical flake fish food and live brine shrimp. Larvae were fed with ArteMac-0 powdered food (20–80 micron size, Bio-Marine, Hawthorne, CA, USA) and/or live brine shrimp depending on their age.</p><!><p>To determine global DNA methylation status through sequential time points, genomic DNA from zebrafish embryos (2, 3.3, 4.3, 6, 11.7, 24, and 36 hpf; n=3 pools, 30 or 50 embryos/pool), hatching early-larvae (48, 60, and 72 hpf; n=3 pools, 10 early-larvae/pool), larvae (96 hpf; n=3 pools, 10 larvae/pool), and adult female and male livers (n=3, no pooling) was extracted with DNAzol (Molecular Research Center, Cincinnati, OH, USA) according to manufacturer's protocol except the following modifications: homogenized embryos were digested overnight with proteinase K (1.8 mg/mL) to eliminate the high content of yolk protein; PolyAcryl carrier (Molecular Research Center) was used to isolate small quantities of DNA such as in embryos; and DNA was further purified with the DNA Clean & Concentrator kit (ZYMO Research, Irvine, CA, USA). For bisulfite sequencing, genomic DNA from zebrafish embryos (n=3 pools, 30–50 embryos/pool), larvae (n=3 pools, 10 larvae/pool) and livers (n=3, no pooling) was isolated with DNeasy Blood & Tissue Easy Kit (Qiagen, Valencia, CA, USA) according to manufacturer's protocol. For both methods, DNA was treated with RNAase to remove RNA contaminants. DNA concentrations were quantitated with Nanodrop 2000 (Thermo Scientific, Wilmington, DE, USA).</p><!><p>Global DNA methylation was measured with the MethylFlash™ Methylated DNA Kit (Epigentek Group, Farmingdale, NY, USA, Cat. No. P-1034) by following the manufacturer's protocol. The OD intensity of the colorimetric reaction was measured by a Biotek ELx800 plate reader (Winooski, VT, USA). The OD intensity of each sample was normalized to 100 ng of DNA. The cytosine methylation percentage was calculated by using the formula: cytosine methylation % = [(OD sample – OD negative control/100ng)/(OD positive control – negative control) x 2/5ng] X 100; 100 ng is the amount of DNA per sample, 2 is a factor to normalize 5-methylcytosine in the positive control to 100%, and 5 ng is the amount of positive control, which contains 50% unmethylated and 50% methylated cytosine. Three biological replicates were measured in duplicate.</p><!><p>The information of exon, intron, and splice variants of the specific genes were found on Ensembl (http://useast.ensembl.org/index.html). Gene homologs or protein pairwise alignments were done on NCBI HomoloGene (http://www.ncbi.nlm.nih.gov/homologene) or SDSC Biology WorkBench.</p><!><p>CpG islands were found by using the Methyl Primer Express v1.0 (Applied Biosystems, Foster City, CA, USA). The criteria to determine CpG islands were set as follows: DNA region length longer than 300 bp but shorter than 2000 bp; greater than 50% C+G content; and observed CpG/expected CpG of 0.65 (Gardiner-Garden and Frommer, 1987). Promoters or regions were further classified into three categories to distinguish strong CpG islands, weak CpG islands and sequences with low abundance of CpG sites (Takai and Jones, 2002). High CpG promoters/regions (HCP) contain a 500 bp sequence with GC content above 55%, CpG ratio above 0.75, and a CpG observed to expected ratio greater than 0.6. Intermediate CpG promoters/regions (ICP) contain a region below 500 bp and have moderate CpG richness (GC content less than 55%) with CpG ratio between 0.48 and 0.75 and have a CpG observed to expected ratio between 0.4 and 0.6. Low CpG promoters/regions (LCP) do not have a region of 500 bp with a CpG ratio above 0.48 or CpG observed to expected ratio above 0.4 (Weber et al., 2007).</p><!><p>Bisulfite sequencing was performed as described in (Fang et al., 2013). Genomic DNA was treated with sodium metabisulfite and cleaned with EZ Bisulfite DNA Clean-up Kit (ZYMO Research). Previously validated bisulfite specific primers were used for PCR amplification (Fang et al., 2013). PCR products were ligated into pGEM® T Easy Vector System (Promega, Madison, WI, USA) and cloned into DH5 E. coli competent cells (Invitrogen, Grand Island, NY, USA). Each treatment or time-point had three biological samples and at least eight white colonies from each sample were selected for sequencing. Plasmid DNA was isolated in 96-well format using a modified alkaline-lysis extraction procedure with Qiagen reagents and sequenced in the ABI 3730xl sequencer using BigDye v3.1 Terminator/Buffer Ready Rxn Cycle Sequencing kit (Applied Biosystems). Sequence data were analyzed with DNAstar (SeqMan, Madison, WI, USA). CpG methylation percentage was calculated as (total number of methylated CpG)/(number of CpG sites in each gene X number of colonies sequenced).</p><!><p>RNA from embryos (20–50 embryos/pool), larvae (10 larvae/pool) or livers (no pooling) was isolated with RNAzol (Molecular Research Center) and purified with RNeasy Mini Kit (Qiagen) by following the manufacturer's protocols. Total RNA was reverse transcribed to cDNA by using TaqMan® Reverse Transcription Reagents (Applied Biosystems). Relative abundance of target genes to 18S rRNA transcripts in the cDNA libraries was determined by qPCR with SYBR®Green and validated qPCR primers in a GeneAmp 7500 Sequence Detection System (Applied Biosystems) (Fang et al., 2013).</p><!><p>Results were analyzed by GraphPad Prism 5.0 (GraphPad Software, La Jolla, CA, USA) and presented as mean ± standard error of the mean (SEM). Statistical differences between treatment groups were determined using one-way ANOVA followed by Neuman–Keulls post hoc test (p < 0.05). Data of the qPCR were calculated with the 2−ΔΔCT method. Statistical differences between treatments or time-points were determined on the linearized 2−ΔCT values. The correlation coefficient between vasa promoter methylation and gene expression was calculated using parametric correlation analysis (Pearson).</p><!><p>The average percentage of methylated cytosines was 0.61% in 2–4.3 hpf zebrafish embryos (Figure 1). The methylation increased significantly to an average of 2.06% in 6 to 96 hpf whole zebrafish, and adult female and male livers. Additionally, the percent of methylated cytosines between 4.3 hpf embryos and adult male liver was not significantly different.</p><!><p>Transcripts of dnmt1 mRNA were maternally transferred into the eggs with high abundance in unfertilized zebrafish eggs (Figure 2A). Transcription of dnmt1 was initially high and then dropped significantly at 4 hpf and remained at a low level until a transient increase at 60 hpf (pec-fin stage).</p><p>The expression of gnmt was low throughout early embryo development (Figure 2B). Between 12 and 24 hpf, gnmt mRNA expression rose significantly by 5-fold. Peak gnmt expression was at 60 hpf, but like dnmt1, expression also decreased again at 96 hpf.</p><p>Comparing the relative expression between dnmt1 and gnmt, dnmt1 was maternally deposited about 7-fold more in unfertilized eggs compared to gnmt (Figure 2C). Starting at 24 hpf, gnmt exceeded dnmt1 expression by 1.6-fold. At 96 hpf, gnmt expression was about 5.6-fold higher than dnmt1.</p><!><p>The potential methylation target region of the vasa promoter is illustrated in Figure 3 and further explained in Table 1. Only ~6% of the 5 CpG sites in the vasa promoter were methylated in zebrafish embryos aged from 3.3 to 8 hpf (Figure 4A and Supplemental Figure 2). By 12 hpf, the CpG methylation percentage increased significantly to 37.5%. Methylation continued to increase with development to 68.3% at 48 hpf and 79.8% at 96 hpf. The promoter was also highly methylated in the 1.5-year old female livers (76.8%).</p><!><p>Transcripts of vasa mRNA were maternally deposited into the oocyte with high abundance in unfertilized eggs (Figure 4B). After fertilization, vasa expression significantly increased at 3.3 hpf, indicating embryonic transcription of vasa mRNA. The mRNA began to drop slightly at 4 hpf, and the highest relative decrease of vasa mRNA expression occurred at 12 hpf. The decrease in expression continued, and the lowest expression was detected at 96 hpf which corresponded to only 0.2% of the vasa transcription in the 3.3 hpf embryos.</p><p>Consistent with theory, the trends of vasa DNA methylation and mRNA expression during embryogenesis were opposite from each other (Figure 4C). As the DNA methylation increased, vasa expression decreased significantly. The Pearson correlation coefficient of vasa promoter methylation with gene expression was −0.885 (P<0.00001).</p><!><p>The target regions of these four genes are illustrated and described in Figure 3 and Table 1. The 26 CpG sites located in the first promoter (CGI-1) of rassf1 were hypomethylated at 3.3 hpf in mid-blastula embryos, 96 hpf larvae, and 1.5-year old female zebrafish livers (Figure 5A and Supplemental Figure 3A) and not significantly different. In contrast, the CpG sites in the second promoter (CGI-2) of rassf1 were highly methylated (Figure 5B and Supplemental Figure 3B). The percentage of CpG methylation reached 98.0% at 3.3 hpf and remained highly methylated at 5 hpf (96.5%), 6 hpf (98.5%), and 96 hpf (92.5%). In adult livers, the methylation percentage dropped significantly to 73.3%. The rassf1-001 mRNA increased significantly at 5 hpf, peaked at 96 hpf, and then decreased significantly in adult livers (Figure 6A). Similarly, total rassf1 mRNA expression was low at 3.3 hpf, then increased dramatically at 5 hpf, but the total expression dropped significantly later (Figure 6B). The percentage of rassf1-001 represented in the total rassf1 expression at different time-points is shown in Figure 6C.</p><p>For tert promoter, all the CpG sites except CpG #1 were highly methylated in all the time-points tested (Supplemental Figure 4). Neither the methylation pattern nor the percentage was different between the three time-points (Figure 7A). However, tert gene expression was significantly different (Figure 7B). Expression of tert at 96 hpf was 3.5-fold higher than that at 3.3 hpf. Tert expression in adult livers was extremely low, and was only 5.3% and 1.5% of the expression in 3.3 hpf embryos and 96 hpf larvae, respectively.</p><p>The 36 CpG sites located in the CGI in the c-jun first exon were hypomethylated (Figure 7C and Supplemental Figure 5). Neither the methylation pattern nor the percentage was different between the three time-points. However, like tert, gene expression was significantly different (Figure 7D). Expression of c-jun at 96 hpf was 379-fold higher compared to the expression at 3.3 hpf. In adult livers, expression was lower than 96 hpf larvae, but still 39-fold higher than the expression in 3.3 hpf embryos.</p><p>The 26 CpG sites located in the CGI in the c-myca second exon were also hypomethylated (Figure 7E and Supplemental Figure 6). The overall methylation percentage was statistically higher in the 3.3 hpf embryos than the percentage in 96 hpf larvae and adult livers. As a possible result of this hypomethylation, c-myca expression at 96 hpf was higher than that at 3.3 hpf (Figure 7F).</p><!><p>The rate of conversion of non-CpG cytosines was examined to evaluate the efficiency of bisulfite conversion. Three PCR products (eight colonies chosen from each product) derived from vasa (207 bp), rassf1 CGI-1 (314 bp), rassf1 CGI-2 (438 bp), tert (233 bp), c-jun (425 bp) and c-myca (444 bp) were sequenced. The amplified regions have 43, 94, 51, 101 and 91 non-CpG cytosines, respectively. The efficiencies of bisulfite conversion were all above 99.8% (Supplemental Table 2). This demonstrates that the sodium bisulfite conversion of cytosines was very efficient and did not generate artifacts due to incomplete conversion.</p><!><p>When alignments were done between converted Genbank sequences (non-CpG C was converted to T) and our sequence data in DNAstar, dinucleotide polymorphisms were observed in rassf1, tert and c-myca due to CG losses in our fish. In rassf1 CGI-2, 36.2% of the CpG #16 was mutated to TT (Table 2). In tert, CpG #7 was completely changed to TA. In c-myca, 81.0% of CpG #26 was changed to TT.</p><!><p>In this study, global and gene specific DNA methylation was measured at different developmental time-points in zebrafish. Our data showed that overall about 0.53%, 0.54%, 0.75% of the cytosines were methylated in the genome of 2, 3.3, and 4.3 hpf embryos, respectively (Figure 1), which increased approximately four times at 6 hpf and later life stages. These results are consistent with Mhanni and McGowan (2004) and Wu and coworkers (2011) who showed that cytosine methylation increased from 2 and 3.3 hpf to 6 hpf. Liver tissue from 1.5 year old zebrafish was used to represent differentiated adult conditions. It should be noted that both percent methylation and gene expression would be expected to be variable between adult tissues, however, adult liver global cytosine methylation was not statistically different than embryo and larval results after 6 hpf.</p><p>We measured the expression of two DNA methylation related enzymes during embryogenesis aiming to associate their expression profiles with DNA methylation changes. Due to the genome duplication events, there are eight known dnmt orthologs in zebrafish (Shimoda et al., 2005; reviewed in Goll and Halpern, 2011). We have focused on dnmt1 in this study. Of the dnmt orthologs, there is only a single copy of dnmt1 that is highly similar to human dnmt1. This is also the most studied dnmt ortholog in zebrafish, whereas less is known about the rest of dnmts though their differential activity may be involved in tissue-specific differentiation (Wu et al., 2011). On the other hand, zebrafish has only one copy of gnmt. Gnmt is the enzyme responsible for the transfer of a methyl group from S-adenosylmethionine (SAM) to glycine forming S-adenosylhomocysteine (SAH) and sarcosine, thus it preserves the ratio of SAM/SAH, a sensitive indicator of cellular methylation capacity. Constitutively, we found that both dnmt1 and gnmt were maternally transferred mRNAs and expressed throughout development (Figure 2). However, their expression profiles were distinct. The average expression of dnmt1 was high from 8-cell (1.25 hpf) to mid-blastula (3.3 hpf) stage and was approximately 5-fold higher than the average expression from sphere (4 hpf) to early larvae (96 hpf) stage. In contrast, gnmt expression was low in the beginning and increased significantly from 24 hpf. Expression of dnmt1 was higher than gnmt until 24 hpf when gnmt expression began to exceed dnmt1. Although dnmt1 is dominantly a maintenance methyltransferase, recent studies reported that dnmt1 also plays an active role in de novo methylation (Fatemi et al., 2002; Athanasiadou et al., 2010). Low expression of gnmt during early embryogenesis could be expected to help preserve a high SAM/SAH ratio, whereas dnmt1 will consume SAM as a methyl donor to remethylate DNA. Therefore, the relatively low expression of gnmt together with high expression of dnmt1 may contribute to DNA remethylation in early zebrafish embryos.</p><p>In addition to measuring the global DNA methylation changes, we have also studied the DNA methylation patterns of individual genes during zebrafish development. Five important developmental genes, i.e. vasa, rassf1, tert, c-jun and c-myca, were selected for sodium bisulfite sequencing analysis, which is able to generate single nucleotide resolution of individual CpG methylation status.</p><p>The vasa promoter is a low CpG promoter (LCP) (Table 3) and would be predicted to be highly methylated, which was supported by our results. The de novo DNA methylation in the vasa promoter did not occur in the blastula stage (3.3 hpf) when the majority of the methylation patterns are typically established. In fact, the vasa promoter did not begin to be methylated until 11 hpf and reached its plateau methylation level by 48 hpf. The DNA methylation pattern was thereafter not different than the livers of adult zebrafish. Relative hypomethylation of vasa at 3.5 hpf (mid-late blastula) compared to differentiated ZF-4 zebrafish fibroblast cells was also found by Lindeman and coworkers (2010). Consistent with the increasing promoter methylation during development, vasa mRNA expression decreased continuously over the same period. The most dramatic decline of vasa mRNA occurred at 12 hpf, the same time-point when the promoter methylation percentage first became significantly higher than the previous stages. The opposite trends of DNA methylation and gene expression suggest that vasa promoter methylation status was associated with zygotic vasa expression. This is also consistent to our previous finding that benzo[a]pyrene –treated zebrafish larvae (96 hpf) had decreased promoter methylation in vasa and increased vasa gene expression (Fang et al., 2013). A consideration worth noting is that in these studies the genomic DNA was extracted from whole embryos whereas the vasa mRNA is assumed to be predominately synthesized by the PGCs that represent only a small fraction compared to somatic tissues. Therefore, in future studies it will be informative to extract genomic DNA from isolated PGCs to further substantiate the relationship between vasa methylation and gene expression.</p><p>The single copy of rassf1 gene can generate multiple transcript variants by using two different CGI promoters, and this is well conserved among human, rat, mouse and zebrafish (Supplemental Figure 1). Zebrafish rassf1 has five transcripts and possibly three CpG rich promoters crossing each TSS (Supplemental Figure 7). Due to the high identity between the five transcripts, it was a challenge to design specific qPCR primers to target each one of the rassf1 variants. It was only possible to measure the transcription of rassf1-001 by designing primers located within exon 1 (Fang et al. 2013). We also measured the total expression of all the rassf1 variants simultaneously using the primer pairs located at exon 5 and 6. Our results showed that rassf1 CGI-1 was almost 100% unmethylated, which is similar to the normal methylation pattern in the first promoter of human rassf1 (Peters et al., 2007). In contrast, rassf1 CGI-2 was heavily methylated (>92%). In contrast to the pattern noted for vasa, the methylation pattern for rassf1 CGI-2 was established during early embryogenesis, and the methylation percentage decreased in aging fish livers. However, there was no apparent correlation between changes of promoter methylation patterns and rassf1 mRNA expression.</p><p>The proximal promoter of tert was highly methylated (except in CpG #1), and its consistent methylation pattern was also established by 3.3 hpf. Although the methylation patterns were unchanged, tert mRNA expression was significantly different at the three time-points tested. The expression of tert was the highest in 96 hpf larvae when extensive cellular proliferation required tert to add telomere ends to the newly-replicated DNA. On the contrary, tert expression was the lowest in adult livers presumably because the aging animals were losing telomerase activity (Anchelin et al., 2011). The results from rassf1 and tert contradicted the general notion that promoter DNA methylation and gene expression are inversely correlated (Kass et al., 1997). Others too have found that some promoter methylation patterns do not correlate well with gene expression (Weber et al., 2007). Therefore, for rassf1 and tert, DNA methylation may not be the dominate factor regulating their gene expression during development. Accordingly, histone modifications and miRNAs play important roles in regulating transcription of rassf1 and tert (Li et al., 2012; Daniel et al., 2012).</p><p>The majority of previous studies have focused on promoter DNA methylation, however, recent studies found the importance of gene body methylation. In plants, gene body DNA methylation was positively correlated with gene transcription (Zhang et al., 1999; Zilberman et al., 2007). However, conflicting results were found in vertebrates. Some studies reported that actively transcribed genes have extensive DNA methylation within their gene bodies, supporting the positive correlation between methylation and transcription (Lister et al., 2009; Rauch et al., 2009). Yet, results from in vitro studies showed that methylation of the gene body was sufficient to repress gene expression in the absence of promoter methylation (Keshet et al., 1985; Graessmann et al., 1994). Another study found that methylation of alternative promoters discovered within gene bodies was associated with gene silencing (Maunakea et al., 2010). Therefore, gene body methylation is important for transcriptional regulation. Here we measured the gene body methylation in two genes, namely, c-jun and c-myca.</p><p>We found that the CpG sites were nearly 100% unmethylated in the coding region of c-jun, and the methylation patterns were maintained from 3.3 hpf to 1.5 year-old zebrafish liver. However, the c-jun mRNA expression was significantly different in distinct life stages. This indicates that the low methylation in the c-jun gene body is permissive to gene expression, but the actual magnitude is likely determined by other factors such as miRNAs and histone modification (Yamaguchi et al., 2005; Kappelmann et al., 2012).</p><p>Due to genome duplication, zebrafish have two copies of c-myc. We studied c-myca because it is phylogenetically closer to mammalian c-myc (Supplemental Table 1). The target region was in the second exon, which is within the coding region. The results showed that the methylation percentage of c-myca in 3.3 hpf embryos was significantly higher than in 96 hpf larvae and adult livers. When demethylation occurred, gene expression was significantly higher at 96 hpf compared to 3.3 hpf, supporting the theory that gene body methylation also has a reverse correlation with gene expression for c-myca.</p><p>From our results, the relationship between CpG density and methylation outcome agrees with the earlier study done in human promoters (Weber et al., 2007). Weber and coworkers found that LCP were almost exclusively methylated, ICP were less likely to be methylated (21%), and the majority of HCPs (97%) had very low methylation. Our results showed a similar trend. For example, the LCP in vasa was highly methylated in larvae and liver; some ICPs, like rassf1 CGI-1 and c-myca gene body, were hypomethylated and some ICPs, like rassf1 CGI-2 and tert, were hypermethylated; the HCP in c-jun gene body was hypomethylated. Work in zebrafish found that at (3.3 hpf) and after (5.3 hpf) the midblastula transition there was an increase in methylated HCPs relative to LCPs (Andersen et al., 2012). By comparing the constitutive DNA methylation and mRNA expression in the five genes, we noticed that promoter hypermethylation in vasa and tert were associated with low mRNA expression (Figure 8). In contrast, hypomethylation in rassf1 CGI-1, intragenic c-jun and c-myca were linked to high mRNA expression. The high content of unmethylated cytosines within a gene body may increase the copy number of mRNA being transcribed and improve the mRNA stability (Duan and Antezana, 2003; Bauer et al., 2010).</p><p>In this study, we found that evolutionary CpG loss was apparent in the zebrafish genome. Similar to the finding in mammals, CpG mutations preferentially target the methylated cytosines in zebrafish. For example, CpG losses were identified in both the heavily methylated rassf1 CGI-2 and tert promoters. Even the CpG loss in c-myca was located at the end of the target region which had a higher methylation percentage. More instances of CpG loss will likely be discovered as genome-wide methylation studies become more widely used.</p><!><p>While global DNA methylation begins to increase after 3.3 hpf in zebrafish embryos and reached a plateau between 6 and 96 hpf, the timing of establishment of promoter methylation patterns was gene specific. Of the five genes studied, we found that de novo methylation of rassf1, tert, c-jun, and c-myca occurred by 3.3 hpf during early embryogenesis. However, in the vasa promoter, the methylation pattern was established at later stages of development. In addition, changes in gene expression during embryo-larval development did not correlate with DNA methylation patterns in each gene except for vasa and c-myca. Therefore, as the role of DNA methylation and gene expression in development is more extensively studied, it will be important to investigate the changes at gene and CpG island specific resolution as well as consider other mechanisms of epigenetic modulation.</p>