Recently, studies from a number of independent laboratories have demonstrated that ethanol has the capacity to bring epigenetic changes to contribute to the development of FASD (Kim and Shukla 2005 ; Kaminen‐Ahola et al . 2010a , b ; Downing et al . 2011 ; Zhou et al . 2011a ; Subbanna et al . 2013b , 2014a , b ; Subbanna and Basavarajappa 2014 ). Epigenetic modifications of genomic DNA and histone proteins are critical in orchestrating the transcriptome of different cell types and their developmental potentials (Reik 2007 ; Suzuki and Bird 2008 ; Ma et al . 2010 ). Abnormal changes in histone modifications and/or DNA methylation play a major role in modulating gene expression and cellular functions that result in long‐lasting altered phenotypes (Vaissiere et al . 2008 ) and several human developmental disorders (Campuzano et al . 1996 ; Petronis 2003 ; Makedonski et al . 2005 ; Ryu et al . 2006 ; Warren 2007 ; Gavin and Sharma 2010 ). Studies from several laboratories have demonstrated that exposure to ethanol at various developmental stages is associated with genome‐wide/gene‐specific alterations in histone modifications (Kim and Shukla 2005 ; Park et al . 2005 ; Pal‐Bhadra et al . 2007 ; Moonat et al . 2013 ; Subbanna et al . 2013b ), changes in DNA methylation (Garro et al . 1991 ; Haycock and Ramsay 2009 ; Liu et al . 2009 ; Ouko et al . 2009 ; Downing et al . 2011 ; Zhou et al . 2011b ), and long‐lasting altered phenotypes reminiscent of fetal alcohol syndrome (Kaminen‐Ahola et al . 2010b ). Collectively, these observations suggest that ethanol has the ability to act as a potent epigenetic modulator and induce deficits in neuronal differentiation (Veazey et al . 2013 ) and possibly maturation leading to learning and memory deficits (Izumi et al . 2005 ; Noel et al . 2011 ; Wilson et al . 2011 ; Sadrian et al . 2012 ; Subbanna et al . 2013a , 2014a ; Subbanna and Basavarajappa 2014 ) as observed in human FASD (Mattson et al . 2011 ; Lebel et al . 2012 ; Norman et al . 2013 ). Based on these interesting facts, this study was undertaken to evaluate the mechanisms related to DNA methylation using a mouse model of FASD which induces widespread activation of caspase 3 immediately after ethanol exposure in P7 mice. We document one of the possible novel mechanisms through which DNA methylation was reduced in the mouse model of FASD. In addition, P7 cannabinoid receptor type 1 (CB1R) null mice that exhibit no ethanol‐induced activation of caspase 3 are resistant to ethanol‐induced impairment of DNA methyltransferases DNMT1 or DNMT3A or DNA methylation.

Alcohol drinking during pregnancy is an intractable health problem worldwide and a leading cause of intellectual disability in Western nations (Mattson et al . 2011 ). The range of dysfunctions associated with alcohol exposure during development is collectively termed fetal alcohol spectrum disorder (FASD) and is characterized by widespread neuropsychological defects (Mattson and Riley 1998 ; Mattson et al . 1998 ) that involve hippocampal and neocortex dysfunctions (Mattson et al . 1996 ; Clark et al . 2000 ; Bookstein et al . 2001 ), including deficits in learning and memory (Goodman et al . 1999 ; Mattson et al . 1999 ). FASD is a major public health crisis with an estimated incidence rate as high as 2–5% in the United States and several Western European countries (May et al . 2009 ). Rodents are the most commonly used animal models for FASD research; however, their gestational period is much shorter than that of human beings (18–23 days for mice/rats), and in a significant amount of third trimester equivalents (Bayer et al . 1993 ) brain development takes place following birth in these species (Tran et al . 2000 ; Cronise et al . 2001 ). In rodent models, the brain is particularly sensitive to ethanol between post‐natal days 6 and 10 (P6–10) due to the fact that the beginning of the second week is a critical period of synaptic development (Marchal and Mulle 2004 ; Lanore et al . 2010 ). A single episode of binge‐like ethanol exposure on P7 was shown to induce robust activation of caspase 3 (a marker for neurodegeneration) in several brain regions (Ikonomidou et al . 2000 ; Saito et al . 2010 ; Wilson et al . 2011 ; Sadrian et al . 2012 ; Subbanna et al . 2013b ), perturb local and interregional brain circuit integrity in the olfacto‐hippocampal pathway (Wilson et al . 2011 ; Sadrian et al . 2012 ) resulting in impaired learning and memory task performance in adulthood (Subbanna et al . 2013a , 2014a ; Subbanna and Basavarajappa 2014 ) as observed in human FASD (Mattson et al . 2011 ; Lebel et al . 2012 ; Norman et al . 2013 ). So far, there are no effective treatments for FASD because our understanding of the molecular cause of FASD is limited.

Unless indicated otherwise, the experiments involved an equal number of pups per treatment and were performed in triplicate. All of the data are presented as the mean ± SEM. A statistical comparison of the data was performed by either a Student's t test or one‐way analysis of variance (anova) or a two‐way anova with Bonferroni's post hoc test. In all of the comparisons, p < 0.05 was considered to indicate statistical significance. The statistical analyses were performed using the Prism software (GraphPad, San Diego, CA, USA).

For DNA methylation assay, 4–24 h after the first saline or ethanol injection, pups were killed by decapitation, neocortex and hippocampus were dissected, flash frozen and stored at −80°C. Genomic DNA was isolated from hippocampus and neocortex by Qiagen kit (Qiagen Sciences, MD, USA). The levels of global DNA methylation were determined using Epigentek (Farmingdale, NY, USA) methylflash methylated DNA quantification kit (Colorimetric) (Epigentek, Farmingdale, NY, USA) according to the manufacturer's instructions. 5‐methylcytosine (5‐mC) fraction of DNA was detected using capture and detection antibodies and then quantified by reading the absorbance at 450 nm. Synthetic unmethylated (50% of cytosine) and methylated (50% of 5‐mC) DNA oligomers (Epigentek, Farmingdale, NY, USA) were used as a negative and positive control, respectively. The percentage of 5‐mC was calculated using the formula provided in the kit procedure and were normalized to percentage of control (The graphs represent the global DNA methylation levels multiplied by an arbitrary factor to set the control to 100).

For the qPCR studies, 4–24 h after the first saline or ethanol injection, pups were killed by decapitation, neocortex and hippocampus were dissected, flash frozen, and stored at −80°C. The samples were subjected to qPCR as described before (Subbanna et al . 2013a , b ) using Mm00599763_m1 (dnmt1), Mm00432881_m1 (dnmt3a), and 4352932 (Gapdh)] from Applied Biosystems. GAPDH was used as an endogenous mRNA control. Three independent runs were carried out for each set of samples. For each run, triplicate reactions were carried out for each sample. Data obtained were analyzed with the use of SDS2.4 software (Life Technology, Grand Island, NY, USA). The amount of target (dnmt1 and dnmt3a) normalized to endogenous reference (Gapdh) and relative to a calibrator was given by‐2ΔΔCt.

For western blot analysis, 4–24 h after the first saline or ethanol injection, pups were killed by decapitation, hippocampus and neocortex were dissected, flash frozen, and stored at −80°C. Homogenates from the hippocampus and neocortex of the pups were processed and subjected to immunoblot as described previously (Basavarajappa and Subbanna 2014 ; Basavarajappa et al . 2014 ). Developmental brain samples were prepared by scarifying the P0–P90 mice according to their date of birth, respectively. Blots were Ponceau S stained to confirm equal loading in each lane and were incubated with primary antibody anti‐rabbit‐DNMT3A (monoclonal, #3598, 1 : 1000), anti‐rabbit‐DNMT1 (monoclonal, #5032, 1 : 1000), anti‐rabbit CC3 (Asp175) (polyclonal, #9661, 1 : 1000), and anti‐mouse‐β‐actin (monoclonal, #3700, 1 : 1000) (Cell Signaling, Danvers, MA, USA) for 3 h at 25°C or overnight at 4°C and processed as previously described by our laboratory (Basavarajappa et al . 2008 ). Incubation of blots with a secondary antibody (goat anti‐mouse peroxidase conjugate, #AP 124P, 1 : 5000, Millipore Corporation, Bedford, MA, USA; goat anti‐rabbit, #AP132P, 1 : 5000, Millipore) alone did not produce any bands.

Eight hours after the first dose ethanol/saline injection, the pups were anesthetized with isoflurane (1–3%) and perfused with a solution containing 4% paraformaldehyde and 4% sucrose in 0.05 M cacodylate buffer (pH 7.2). The free‐floating sections containing hippocampus and cortex were immunostained using anti‐rabbit cleaved caspase 3 (Asp175) (CC3) (polyclonal, #9661, 1 : 500, Cell Signaling, Danvers, MA, USA) by the ABC reagents (Vectastain ABC Elite Kit, Vector Labs, Burlingame, CA, USA) with a peroxidase substrate (DAB) kit (Vector Labs) as described previously (Subbanna et al . 2013a , b ). The primary antibodies were omitted from the reactions as a control for secondary antibody specificity. All photomicrographs were taken through a 2.5× or 40× objective with a Nikon Eclipse TE2000 inverted microscope attached to a digital camera (DXM1200F, Morrell Instrument Company, Melville, NY, USA).

In our previous studies, pre‐administration of G9a/G9a‐related protein (GLP) inhibitor, Bix‐01294 (Bix, 2‐(Hexahydro‐4‐methyl‐1H‐1,4‐diazepin‐1‐yl)‐6,7‐dimethoxy‐N‐[1‐(phenylmethyl)‐4‐piperidinyl]‐4‐quinazolinamine trihydrochloride) (Cayman, MI, USA)], CB1R antagonist (SR, SR141716A, gift from RBI) and broad‐spectrum caspase inhibitor, and quinoline‐Val‐Asp(Ome)‐CH2‐O‐phenoxy (Q‐VD‐OPh) (SM Biochemicals, Anaheim, CA, USA) 30 min before first dose ethanol treatment prevented the activation of caspase 3 in P7 mice (Subbanna et al . 2013a , b , 2014a , b ; Subbanna and Basavarajappa 2014 ). Therefore, we used Bix or SR or Q‐VD‐OPh to inhibit caspase 3 activation in this study. Bix and SR were dissolved separately in 10 μL of ethanol followed by a two or three drops of Tween 80 (10 μL) and further volume was made up with sterile saline solution. The optimum dose of Bix or SR (1 mg/kg) solution was administered by s. c. injection at a volume of 5 μL/g body weight 30 min before first ethanol administration. The Bix or SR vehicle solution was injected as a control. The optimum dose of Q‐VD‐OPh (1 mg/kg) was administered by s. c. injection at a volume of 5 μL/g body weight 30 min before first dose ethanol treatment. Q‐VD‐OPh was dissolved in sterile saline solution. Sterile saline was used as vehicle in these experiments. Mice were kept with the dams until the pups were killed and their brains removed 4–24 h after the first saline/ethanol injection. Bix or SR or Q‐VD‐OPh treatment did not alter P7 ethanol‐induced intoxication (sleeping time) at the time of brain harvest (Subbanna et al . 2013a , b ). Bix or SR or Q‐VD‐OPh alone treated P7 mice looked like saline‐treated mice and did not cause any inflammation or bleeding in any of the organs (Subbanna et al . 2013a , b ). The brains were processed for several analyses, as described below.

An ethanol treatment paradigm, which has been previously shown to induce robust apoptotic neurodegeneration with no lethality in P7 mice (Olney et al . 2002 ), was used in this study. Half of the pups (male and female) in each litter were treated subcutaneously (s. c.) with saline and the other half with ethanol at P7 (based on the day of birth) (2.5 g/kg s. c. at 0 h and again at 2 h) as described previously by our laboratory (Wilson et al . 2011 ; Sadrian et al . 2012 ; Subbanna et al . 2013a , b ). Blood ethanol levels (BEL) in pup serum were then determined using a standard alcohol dehydrogenase‐based method (Lundquist 1959 ). In time kinetic studies, saline was injected for 0‐h ethanol treatment.

Blocking CB1 receptor with SR141617A (SR) rescues ethanol‐induced degradation of DNA methyltransferases (DNMT1) and DNMT3A proteins and impaired DNA methylation in the neonatal mouse brain. (a and b) Western blot analysis of DNMT1 and DNMT3A proteins and β‐actin (loading control) in hippocampal and neocortical nuclear extracts from the four (S + V, E + V, S + SR, and E + SR) groups ( n = 15 pups/group). P7 mice were treated with ethanol for 8 h, and SR was pre‐treated for 30 min before the ethanol treatment. (c) Global DNA methylation quantification was performed in hippocampal and neocortical DNA from saline (S), ethanol (E), S + SR, or E + SR groups ( n = 10 pups/group) (* p < 0.05 vs. S + V; # p < 0.05 vs. E + V. Error bars, SEM. (two‐way anova with Bonferroni's post hoc test). HP, hippocampus; NC, neocortex.

In our previous studies, both SR and CB1RKO provided protection against ethanol‐induced activation of caspase 3 without altering ethanol metabolism (Subbanna et al . 2013a , 2014a ). We used a specific CB1R antagonist (SR) or CB1RKO mice in our next experiment to further examine the inhibition of DNA methylation and the levels of the associated proteins DNMT1 and DNMT3A by ethanol are due to widespread activation of caspase 3. Our previous findings suggested that maximum inhibition of caspase 3 activation was found at 1 mg/kg (Subbanna et al . 2013a ). Thus, we used SR at 1 mg/kg in our current studies. Our findings suggest that SR pre‐administration before ethanol treatment completely rescued the loss of DNMT1 and DNMT3A protein levels in both the brain regions ( p < 0.05) (Fig. 6 a and b). Statistical analysis using two‐way anova with Bonferroni's post hoc tests indicates significant influence of ethanol (vs. saline) (hippocampus: DNMT1, F 1,20 = 20, p < 0.05; DNMT3A, F 1,20 = 18, p < 0.05; neocortex: DNMT1, F 1,20 = 22, p < 0.05; DNMT3A, F 1,20 = 15, p < 0.05) and a significant interaction between ethanol and SR pre‐treatment (hippocampus: DNMT1, F 1,20 = 14, p < 0.05; DNMT3A, F 1,20 = 12, p < 0.05; neocortex: DNMT1, F 1,20 = 23, p < 0.05; DNMT3A, F 1,20 = 16, p < 0.05). Pre‐treatment of SR or vehicle alone fails to alter DNMT1 or DNMT3A protein levels ( p > 0.05). Similarly, decrease in DNA methylation due to ethanol treatment in P7 mice was also prevented by SR pre‐treatment. Statistical analysis using two‐way anova with Bonferroni's post hoc tests confirmed a significant influence of ethanol (vs. saline) (hippocampus: F 1,20 = 27, p < 0.05; neocortex: F 1,20 = 25, p < 0.05). A significant interaction between ethanol and SR treatment was also evident (hippocampus: F 1,20 = 10, p < 0.05; neocortex: F 1,20 = 18, p < 0.05). SR or vehicle pre‐treatment failed to reduce DNA methylation ( p > 0.05) (Fig. 6 c). Statistical analysis of one‐way anova with Bonferroni's post hoc tests suggested that, consistent with SR treatment, the CB1R KO provided protection against the P7 ethanol‐induced decrease in DNMT1 and DNMT3A protein levels in both the brain regions (Fig. 7 a and b) (hippocampus: DNMT1, F 1,20 = 18, p < 0.05; DNMT3A, F 1,20 = 13, p < 0.05; neocortex: DNMT1, F 1,20 = 23, p < 0.05; DNMT3A, F 1,20 = 12, p < 0.05). Similarly, CB1RKO mice provided protection against the P7 ethanol‐induced decrease in DNA methylation in the hippocampus ( F 1,20 = 28, p < 0.05) and neocortex ( F 1,20 = 26, p < 0.05) (one‐way anova with Bonferroni's post hoc tests) (Fig. 7 c).

In our previous studies, we have shown that pharmacological inhibition of G9a/GLP by Bix before ethanol treatment prevents the activation of caspase 3 in P7 mice without affecting ethanol metabolism (Subbanna et al . 2013b , 2014b ; Subbanna and Basavarajappa 2014 ). In this study, we examined whether the reduced levels of DNMT1 and DNMT3A protein observed after ethanol treatment could be rescued by Bix (1 mg/kg, optimum dose) pre‐treatment. Bix administration before ethanol treatment prevented DNMT1 and DNMT3A protein loss in both the brain regions (Fig. 5 a and b). Statistical analysis by two‐way anova with Bonferroni's post hoc analysis showed a significant effect of ethanol (vs. saline) (hippocampus: DNMT1, F 1,28 = 14, p < 0.05; DNMT3A, F 1,28 = 9, p < 0.05; neocortex: DNMT1, F 1,28 = 16, p < 0.05; F 1,28 = 16, p < 0.05). A significant interaction between ethanol and Bix treatment was also evident (hippocampus: DNMT1, F 1,28 = 11, p < 0.05; DNMT3A, F 1,28 = 17, p < 0.05; neocortex: DNMT1, F 1,28 = 14, p < 0.05; DNMT3A, F 1,28 = 12, p < 0.05). The saline and saline‐Bix groups were failed to affect significantly ( p > 0.05). In our next experiment, we determined whether DNA methylation that was reduced after ethanol treatment could be rescued by Bix‐pre‐treatment. Comparison of two‐way anova with Bonferroni's post hoc test indicated a significant influence of ethanol treatment (vs. saline) (hippocampus: F 1,28 = 22, p < 0.05; neocortex: F 1,28 = 24, p < 0.05). A significant interaction between ethanol and Bix treatments was also evident (hippocampus; F 1,28 = 18, p < 0.05; neocortex; F 1,28 = 20, p < 0.05) (Fig. 5 c). These findings suggest that Bix prevents the ethanol‐induced decrease in DNA methylation by restoring caspase 3‐mediated reduction of DNMT1 and DNMT3A protein levels in P7 mice.

Pharmacological inhibition of caspase 3 rescues P7 ethanol‐induced degradation of DNA methyltransferases (DNMT1) and DNMT3A proteins as well as DNA methylation in the neonatal mouse brain. (a and b) P7 Mice pre‐treated for 30 min with Q‐VD‐OPh (1 mg/kg) or vehicle were exposed to ethanol and DNMT1 and DNMT3A proteins levels were determined in nuclear extracts of hippocampus (HP) and neocortex (NC) by a western blot analysis. β‐actin was used as a loading control. (c) Global DNA methylation quantification was performed in HP and NC DNA from saline (S), ethanol (E), S + Q‐VD‐OPh, or E + Q‐VD‐OPh groups ( n = 10 pups/group) (* p < 0.05 vs. S; # p < 0.05 vs. E. Error bars, SEM. (two‐way anova with Bonferroni's post hoc test).

We hypothesized that the time‐dependent reduction of DNMT1 and DNMT3A protein may be due to the action of ethanol‐activated caspase 3 in P7 mice as activated caspase enzymes degrades many proteins within the cell (Fischer et al . 2003 ; Kamada et al . 2005 ; Subbanna et al . 2013b ; Subbanna and Basavarajappa 2014 ). In these studies, we used a third‐generation dipeptidyl broad‐spectrum caspase inhibitor (Q‐VD‐OPh). Our previous in vivo study have shown that Q‐VD‐OPh (Renolleau et al . 2007 ) effectively prevents the generation of CC3 fragment in P7 mice without any detectable toxicity [for references, see (Renolleau et al . 2007 ; Subbanna et al . 2013b )]. It also prevented dimethylated histone H3 lysine 9 and total histone H3 protein degradation in ethanol‐treated P7 mice (Subbanna et al . 2013b ). Administration of optimum dose of Q‐VD‐OPh (1 mg/kg) before ethanol treatment did not alter the BELs (0.41 ± 0.3 g/dL at 3 h, gradually reduced to 0.23 ± 0.08 g/dL at 9 h after the first ethanol injection), indicating that Q‐VD‐OPh does not modulate ethanol metabolism as observed in our previous studies (Subbanna et al . 2013b ). Pre‐administration of Q‐VD‐OPh before ethanol treatment inhibited the DNMT1 and DNMT3A protein (Fig. 4 a and b) degradation in both the brain regions (hippocampus: DNMT1, F 1,20 = 23, p < 0.05; DNMT3A, F 1,20 = 17, p < 0.05; neocortex: DNMT1, F 1,20 = 18, p < 0.05; DNMT3A, F 1,20 = 17, p < 0.05). The saline and saline‐Q‐VD‐OPh pre‐treatment fail to alter DNMT1 and DNMT3A protein levels ( p > 0.05). Pre‐administration of Q‐VD‐OPh before ethanol treatment rescued impaired DNA methylation (Fig. 4 c) in both the brain regions (hippocampus, F 1,20 = 25, p < 0.05; neocortex, F 1,20 = 28, p < 0.05) (two‐way anova with Bonferroni's post hoc test).

To examine the mechanism by which ethanol may reduce DNA methylation, we examined the influence of ethanol exposure on DNMT1 and DNMT3A protein expression in the hippocampus and neocortex obtained 4–24 h after the first saline or ethanol injection. Ethanol decreased DNMT1 and DNMT3A protein levels in the hippocampus (Fig. 3 a) (DNMT1; F 3,28 = 18, p < 0.05; DNMT3A; F 3,28 = 28, p < 0.05) and neocortex (Fig. 3 b) (DNMT1; F 3,28 = 16, p < 0.05; DNMT3A; F 3,28 = 22, p < 0.05) at 8–24 h (after first ethanol injection) time points compared to the saline control (0 h) (one‐way anova with Bonferroni's post hoc test). We also measured mRNA levels to understand whether a reduced DNMT1 and DNMT3A protein level is due to the suppression of their transcription. The results reveal that decreased DNMT1 mRNA levels by ethanol were found only at the 24‐h time point in the hippocampus (Fig. 3 c) ( F 3,28 = 20, p < 0.05). However, in neocortex, decrease in DNMT1 mRNA levels was found at 8 h, but it was enhanced at 24 h ( F 3,28 = 25, p < 0.05) (Fig. 3 d). We found reduced DNMT3A mRNA levels in the hippocampus ( F 3,28 = 15, p < 0.05) at the 8‐ and 24‐h time points. In neocortex, decrease in DNMT3A mRNA levels was found only at 24 h ( F 3,28 = 12, p < 0.05) (one‐way anova with Bonferroni's post hoc test). Control Gapdh mRNA levels remained unchanged in any of the brain regions examined (data not shown). Over all, ethanol treatment in P7 mice reduced the protein more severely compared to mRNA levels except in neocortex.

We measured the levels of DNMT1 and DNMT3A, enzymes involved in DNA methylation (Biniszkiewicz et al . 2002 ; Feng et al . 2010 ), during various stages of brain development. DNMT1 antibody recognized a major band running at 200 kDa (predicted to run at ~183 kDa) as observed in several previous studies (Robertson et al . 2000 ; Liu et al . 2003 ). DNMT3A antibody recognized a major band running below 140 kDa marker and above the 100 kDa marker (estimated at ~130 kDa) as observed in our previous study (Subbanna et al . 2014b ). There is a discrepancy between the observed and predicted molecular weight of DNMT3A (predicted to run at ~101 kDa). The post‐translational modification can impact the mobility of a protein in sodium dodecyl sulfate–polyacrylamide gel electrophoresis gels. The post‐translational modifications and/or features of DNMT3A that result in the reduced mobility (and thus higher observed molecular weight) have not been well characterized in the literature. Several studies included data demonstrating recombinant and endogenous cellular DNMT3A runs well above the predicted 101 kDa (Takeshima et al . 2006 ; Chang et al . 2011 ; Suetake et al . 2011 ; Subbanna et al . 2014b ). We then examined the developmental pattern of DNMT1 and DNMT3A protein. Neocortex nuclear and cytosolic protein extracts at several developmental stages were used in western blot analysis. Nuclear DNMT1 ( F 8,45 = 150, p < 0.05) and DNMT3A ( F 8,45 = 160, p < 0.05) protein levels were substantially higher during synaptic development (synaptogenesis) compared to the levels in the adult brain (P90) (Fig. 2 a). Similar expression of DNMT1 ( F 8,45 = 110, p < 0.05) and DNMT3A ( F 8,45 = 90, p < 0.05) were also found in cytosolic protein extracts (Fig. 2 b) (one‐way anova with Bonferroni's post hoc test).

We then examined the DNA methylation in the nuclear changes experienced by apoptotic neurons in ethanol‐treated P7 mice. First, we measured global DNA methylation levels using methylflash methylated DNA quantification kit. Assay specificity was determined using synthetic unmethylated DNA (contains 50% of cytosine) (negative control) (synthetic unmethylated DNA) and methylated DNA (contains 50% of 5‐methylcytosine) (positive control) (synthetic methylated DNA). The amount of methylated DNA was found to be directly proportional to the optical density (OD) intensity (Fig. 1 c). The optical density was used with the kit's included formulas to calculate the global DNA methylation levels. We measured global DNA methylation in hippocampal and neocortical DNA obtained 4–24 h after the first saline or ethanol injection. Global DNA methylation was reduced after ethanol exposure (8 and 24 h) compared to saline treatment (0 h) in the hippocampus ( F 1,20 = 58, p < 0.05) and neocortex ( F 1,20 = 62, p < 0.05) (one‐way anova with Bonferroni's post hoc test) (Fig. 1 d).

Ethanol exposure induces apoptotic neurodegeneration and reduces DNA methylation in the P7 mouse brain. (a) Coronal brain sections (hippocampus and retrosplenial cortex) from saline‐ and ethanol‐treated animals were immunostained with an anti‐rabbit CC3 antibody. The white arrows indicate CC3‐positive neurons in the hippocampus and retrosplenial cortex, respectively. Scale bars = 200 μm. The respective images were enlarged to show CC3‐positive cells (*). The scale bars represent 50 μm. CC3‐positive cells were counted in the hippocampus and the retrosplenial cortex ( n = 10 pups/group). (b) Western blot analysis of CC3 using cytosolic extracts (20 μg) of hippocampal and neocortical samples obtained 4–24 h after the first saline or ethanol injection ( n = 15 pups/group). Ponceau S staining confirmed equal loading and β‐actin were used as loading controls. * p < 0.05 versus 0 h or saline control (one‐way anova with Bonferroni's post hoc test). For 0‐h ethanol group, saline was injected. (c) Global DNA methylation assay specificity was determined using synthetic unmethylated DNA (contains 50% of cytosine) (negative control) [synthetic unmethylated DNA (SUD)] and methylated DNA (contains 50% of 5‐methylcytosine) (positive control) (synthetic methylated DNA). The optical density (OD) was used with the kit's included formulas to calculate the global DNA methylation levels. S, saline; E, ethanol. (d) Global DNA methylation quantification in hippocampal (HP) and neocortical (NC) DNA obtained 4–24 h after the first saline or ethanol injection. For 0‐h ethanol group, saline was injected. Error bars, SEM. Error bars, SEM. HP, hippocampus; NC, neocortex. (* p < 0.05, n = 10 pups/group) (one‐way anova with Bonferroni's post hoc test).

We injected P7 mice with ethanol (2.5 g/kg, sc at 0 h and again at 2 h) and measured BELs at two time points. This experimental paradigm resulted in BELs of 0.45 ± 0.4 g/dL at 3 h that was gradually reduced to 0.24 ± 0.07 g/dL at 9 h after the first ethanol injection. We determined neurodegeneration using cleaved caspase 3 antibody (generation of CC3 as a marker for neurodegeneration) in the brains of P7 mice 8 h after first dose of ethanol or saline treatment. As shown in Fig. 1 a, CC3 immunoreactivity was found throughout the forebrain [hippocampus, ( F 1,11 = 68, p < 0.05) and cortex ( F 1,11 = 180, p < 0.05) regions] in the ethanol‐exposed P7 brains (one‐way anova with Bonferroni's post hoc test). Using western blot analysis, we measured CC3 protein levels in hippocampal and neocortical protein cytosolic extracts obtained 4–24 h after the first saline or ethanol injection. Statistical analysis using one‐way anova with Bonferroni's post hoc tests suggests that 8 and 24 h after first dose of postnatal ethanol treatment in P7 mice significantly enhanced CC3 protein levels in both the hippocampus ( F 3,28 = 73, p < 0.05) and neocortex ( F 3,28 = 75, p < 0.05) (Fig. 1 b) compared to 0 h (saline control). In all our time kinetic studies, saline was injected for 0‐h ethanol treatment.

Discussion

In this study, using a mouse model of FASD, we demonstrate for the first time that the mechanism by which post‐natal ethanol impairs DNMT1 and DNMT3A followed by DNA methylation in hippocampus and neocortex, two brain regions that are important for learning and memory (Vann and Albasser 2011). This was accomplished through differential transcriptional regulation of both genes and caspase 3‐mediated degradation of DNMT1 and DNMT3A, which are produced at significantly higher rate during early brain development compared to that in the mature mouse brain. Overall, our current results are consistent with previous findings demonstrating that DNMT1 and DNMT3A are the most highly expressed early in young age and their levels drop significantly by adulthood in most brain regions (Robertson et al. 1999; Feng et al. 2005; Brown et al. 2008; Simmons et al. 2013). Our results are also supported by one more study in which DNMT1 and DNMT3A protein expression were found in both the nuclear and cytosolic fractions of the mouse brain (Chestnut et al. 2011).

The control of gene transcription, via dynamic modification of DNA methylation, plays an essential function in the regulation of gene expression. The active modification of DNA methylation occurs in a tissue‐ and time‐specific way and appears to be vital for embryonic development (Matsuda and Yasutomi 1992; Kakutani et al. 1996; Martin et al. 1999). Exposure to ethanol during embryonic development has been shown to induce changes in DNA methylation (Garro et al. 1991; Haycock and Ramsay 2009; Liu et al. 2009; Ouko et al. 2009; Downing et al. 2011; Zhou et al. 2011b) and is implicated in the etiology of numerous developmental anomalies (Ouko et al. 2009; Kaminen‐Ahola et al. 2010b). Using a mouse model of FASD, our study specifically suggests that ethanol exposure during post‐natal development induces widespread activation of caspase 3 that leads to the loss of DNMT1 and DNMT3A proteins, thereby causing DNA hypomethylation. Furthermore, ethanol‐activated caspase 3 also cleaves some nuclear proteins such as dimethylated (lysine 9) histone proteins (Subbanna et al. 2013b) in neonatal mice. Our findings are in line with another study, in which maternal ethanol consumption (9–11th day of pregnancy) decreases the activity of DNMT in 12th day of fetal mice, resulting in DNA hypomethylation (Garro et al. 1991). However, these findings are in contrast with another study in which ethanol treatment was shown to decrease expression of cell cycle genes through enhanced DNMT activity followed by DNA hypermethylation in neural stem cells (NSCs) (Hicks et al. 2010). In yet another study, repeated post‐natal ethanol exposure of P2‐10 mice also enhanced DNA methylation in P20 rats (Otero et al. 2012). Despite this observation, the DNMT1 transcription is reduced by ethanol treatment in both NSCs (Hicks et al. 2010) and rat sperm (Bielawski et al. 2002) a finding that supports our data.

In this study, ethanol took a long time (24 h) to reduce the expression of the methyltransferase genes encoding for DNMT3A and DNMT1 proteins that are important for de novo and the ‘maintenance’ of DNA methylation, respectively, in the hippocampus. But, in the cortex, ethanol even increased the expression of the DNMT1 gene at the time point where ethanol induces the highest activation of caspase 3. This increase may be a compensatory response to the continuous presence of ethanol and suppression of DNMT3A, and this needs to be investigated further in future studies. However, a similar contrasting effect of ethanol exposure in NSCs and fetal brain on the expression of DNMT1 mRNA and protein has been reported (Hicks et al. 2010). In an embryonic ethanol model, ethanol enhanced the expression of DNMT3A and 3B, but not DNMT1 (Mukhopadhyay et al. 2013) genes. A recent study reported that ethanol exposure significantly impaired DNA methylation (5‐mC staining) in both the dorsal and the ventral neural tube and that led to developmental delay (Zhou et al. 2011c). Together, these data suggest that ethanol, which induces the activation of caspase 3 in neonatal mice, can also impact the cellular DNA methylation machinery by modulating DNMT proteins and, thus playing a significant role in the maturation of synaptic circuits during development. Several studies have revealed that DNA methylation is essential to various types of synaptic plasticity and learning and memory behaviors in adult animals (Weaver et al. 2004; Miller and Sweatt 2007; Roth et al. 2009; Feng et al. 2010; Miller et al. 2010). Together, these observations suggest that DNA methylation is important during post‐natal development; therefore, its reduction, even for a short period, may cause long‐lasting synaptic dysfunction in adult mice, as observed in similar mice models of FASD (Izumi et al. 2005; Wilson et al. 2011; Sadrian et al. 2012; Subbanna et al. 2013a; Subbanna and Basavarajappa 2014).

The cellular proteins that drive biological processes are regulated at different levels, including transcription, translation, post‐translational modification, and degradation. In steady‐state conditions, cellular proteins are dynamic and most of them are continuously synthesized and degraded. Protein degradation mechanisms adjust protein levels in response to both internal and environmental stimuli. In the nucleus, presence of many caspase 3 substrates was described (Fischer et al. 2003) and were implicated in the nuclear morphological changes that occur in most of the apoptotic cells (Kamada et al. 2005). Consistent with this notion, low concentrations of ethanol induce significantly lower amounts of caspase 3 activation in P7 mice and enhance DNMT3A levels without any proteolytic degradation (Subbanna et al. 2014b). Thus, our study suggests that DNMT1 and DNMT3A proteins may act as substrates for widely activated caspase 3 due to high concentration of ethanol in the developing brain. Accordingly, caspase 3 inhibition rescues loss of DNMT1 and DNMT3A as well as DNA methylation in ethanol‐exposed P7 mice. However, our study does not exclude other possible mechanisms of DNMT1 and DNMT3A proteins degradation in ethanol‐treated P7 mice. Similar degradation of DNA methyltransferases by ethanol exposure during the embryonic period was observed (Mukhopadhyay et al. 2013). However, this degradation is mediated through the ubiquitin–proteasome pathways (Mason et al. 2012). DNMT1 protein stability was also shown to be regulated by ubiquitin‐mediated proteosomal degradation, although the enzymes responsible for the ubiquitination state of DNMT1 have not been reported (Agoston et al. 2005). In addition, DNMT1 stability may also be controlled through other post‐translational modification mechanisms such as phosphorylation and methylation (Sun et al. 2007; Esteve et al. 2009; Wang et al. 2009). It is also possible that reduced DNMT1 and DNMT3A protein levels found in this study may be a result of post‐translational modification by ubiquitination followed by degradation through the action of the ubiquitin‐26S proteasome system. Such possibilities warrant future investigation. In our previous study, pre‐administration of G9a/GLP inhibitor (Bix) before ethanol treatment prevented the activation of caspase 3 as well as caspase 3‐mediated degradation of dimethylated H3K9 (Subbanna et al. 2013b). Furthermore, pre‐administration of Bix before ethanol treatment also prevented long term potentiation (LTP) as well as learning and memory deficits in adult mice (Subbanna and Basavarajappa 2014). In this study, pre‐administration of Bix also prevented the degradation of DNMT1 and DNMT3A proteins and restored DNA methylation. Similarly, pre‐administration of CB1R antagonists (SR), which was shown to prevent ethanol‐induced activation of caspase 3 in neonatal mice (Subbanna et al. 2013a), also prevented the degradation of DNMT1 and DNMT3A proteins and rescued DNA methylation. Interestingly, the ablation of the CB1R, which provides protection against ethanol‐induced activation of caspase 3 in neonatal mice (Subbanna et al. 2013a), also restored the levels of DNMT1, DNMT3A proteins and DNA methylation that had been reduced by ethanol. These observations may suggest that observed reduction of DNA methylation for a short period during a critical period of synaptic development (Marchal and Mulle 2004; Lanore et al. 2010) can have a long‐lasting impact on genome regulation and synaptic circuit formation leading to neurobehavioral abnormalities as observed in mice and human FASD (Mattson et al. 2011; Lebel et al. 2012; Norman et al. 2013). This is partly because both the pharmacological blockade of G9a (through use of Bix inhibitor) and CB1R (through use of SR) and the use of CB1R null mice led to the protection against P7 ethanol‐induced activation of caspase 3 but also prevented long‐lasting neurobehavioral abnormalities in adult mice (Subbanna et al. 2013a,b, 2014a; Subbanna and Basavarajappa 2014).

Hypomethylation of CpG islands, especially those islands colocalized with gene promoter regions, is generally associated with gene expression. Impaired DNMT activity and the resulting reduction in DNA methylation levels as well as DNA binding proteins were shown to regulate several genes associated with neurodegenerative disorders in adults [for references see (Lagali and Picketts 2011)]. Therefore, it is possible that during early brain development, ethanol‐induced DNA hypomethylation may differentially regulate the transcription of genes encoding for survival factor (Kokubo et al. 2009) such as activity‐regulated cytoskeleton‐associated protein (Arc) (Subbanna et al. 2014a) or CB1R or G9a expression (Subbanna et al. 2013a,b), inducing a delay in neuronal development (Chen et al. 2013) and deficits in learning and memory in adult animals (Noel et al. 2011; Wilson et al. 2011; Sadrian et al. 2012; Subbanna et al. 2013a, 2014a; Subbanna and Basavarajappa 2014). Future studies using chromatin immunoprecipitation coupled with genome‐wide analysis will further unveil the impact of impaired DNMT1 and DNMT3A proteins and DNA methylation on specific gene expression and function in ethanol teratogenesis.

In summary, pharmacological inhibition of caspase 3 by Q‐VD‐OPh or caspase 3 inactivation by means of Bix or SR treatment prior to ethanol treatment prevents impairment of DNMT1 and DNMT3A proteins and DNA methylation. P7 CB1R null mice, which exhibit no ethanol‐induced activation of caspase 3, exhibit no impairment of DNMT1, DNMT3A, and DNA methylation. This experimental evidence elucidates a novel potential molecular mechanism underlying ethanol teratogenesis, through which ethanol may impair DNA methylation. These impairments, which occur within cells of the developing brain, may have a distinct role in causing long‐lasting deficits in learning and memory, as observed in human FASD (Mattson et al. 2011; Lebel et al. 2012; Norman et al. 2013) as well as animal models of FASD (Noel et al. 2011; Wilson et al. 2011; Sadrian et al. 2012; Subbanna et al. 2013a, 2014a; Subbanna and Basavarajappa 2014), and should be explored in future studies.