Developmental Biology 376 (2013) 51–61

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Developmental Biology

0012-16

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n Corr

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journal homepage: www.elsevier.com/locate/developmentalbiology

Acute dietary zinc deficiency before conception compromises oocyteepigenetic programming and disrupts embryonic development

X. Tian, F.J. Diaz n

Center for Reproductive Biology and Health and Department of Animal Science, The Pennsylvania State University, University Park, PA 16802, USA

a r t i c l e i n f o

Article history:

Received 3 September 2012

Received in revised form

12 January 2013

Accepted 15 January 2013Available online 21 January 2013

Keywords:

Zinc

Oocyte

Embryo

Methylation

06/$ - see front matter & 2013 Elsevier Inc. A

x.doi.org/10.1016/j.ydbio.2013.01.015

esponding author.

ail address: fjd10@psu.edu (F.J. Diaz).

a b s t r a c t

Recent findings show that zinc is an important factor necessary for regulating the meiotic cell cycle and

ovulation. However, the role of zinc in promoting oocyte quality and developmental potential is not

known. Using an in vivo model of acute dietary zinc deficiency, we show that feeding a zinc deficient

diet (ZDD) for 3–5 days before ovulation (preconception) dramatically disrupts oocyte chromatin

methylation and preimplantation development. There was a dramatic decrease in histone H3K4

trimethylation and global DNA methylation in zinc deficient oocytes. Moreover, there was a 3–20 fold

increase in transcript abundance of repetitive elements (Iap, Line1, Sineb1, Sineb2), but a decrease in

Gdf9, Zp3 and Figla mRNA. Only 53% and 8% of mature eggs reached the 2-cell stage after IVF in animals

receiving a 3 and 5 days ZDD, respectively, while a 5 day ZDD in vivo reduced the proportion of 2-cells

to 49%. In vivo fertilized 2-cell embryos cultured in vitro formed fewer (38%) blastocysts compared to

control embryos (74%). Likewise, fewer blastocyst and expanded blastocyst were collected from the

reproductive tract of zinc deficient animals on day 3.5 of pregnancy. This could be due to a decrease in

Igf2 and H19 mRNA in ZDD blastocyst. Supplementation with a methyl donor (SAM) during IVM

restored histone H3K4me3 and doubled the IVF success rate from 17% to 43% in oocytes from zinc

deficient animals. Thus, the terminal period of oocyte development is extremely sensitive to

perturbation in dietary zinc availability.

& 2013 Elsevier Inc. All rights reserved.

Introduction

Development of healthy embryos and progeny is criticallydependent on the quality of the gametes and an optimal uterineenvironment. The quality of the maternal epigenome, in particular,contributes significantly to promoting optimal embryonic devel-opment and postnatal health (Corry et al., 2009; Hales et al., 2011).The epigenome of the oocyte is dramatically remodeled duringoogenesis. As the oocyte nears ovulation, major changes in chro-matin structure and biochemistry take place to prepare forfertilization and embryonic development. These changes includereorganization of the chromatin from a non-surrounded nucleolus(NSN) to a surrounded nucleolus (SN) configuration where thechromatin becomes tightly packed around the nucleolus (Debeyet al., 1993; Mattson and Albertini, 1990; Zuccotti et al., 1995).Chromatin modifications are an important component of epige-netic programming in the oocyte. These include methylation ofhistone proteins and methylcytosine residues on the DNA. GlobalDNA methylation is low in early oogenesis and peaks as oocytesreach full size (Kageyama et al., 2007; Kim et al., 2003). DNA

ll rights reserved.

methylation is necessary to establish imprinted (monoallelic) geneexpression (Bartolomei and Tilghman, 1997). Maternal imprintsare established during oogenesis at selected imprint controlregions (ICR) (Lucifero et al., 2004, 2002). Most imprinted genesare hypermethylated on the maternal allele, with fewer, such asthe Igf2/H19 loci, hypermethylated on the paternal allele (Daviset al., 1999; Tremblay et al., 1997). Relative hypomethylation in thegrowing oocyte helps to maintain a high rate of transcription(Bouniol-Baly et al., 1999; De La Fuente and Eppig, 2001; De LaFuente et al., 2004). High transcription in oocytes is necessary toproduce and store the large quantity of maternal RNAs andproteins (Schultz and Wassarman, 1977; Sternlicht and Schultz,1981; Wassarman et al., 1979) that are needed as maternal factorsuntil embryonic genome activation. However, because of theextensive hypomethylation, many of the transcripts produced inthe oocyte are of repetitive sequences, such as the mouse transcript(MT) family of retrotransposons, which accounts for up to 14% oftranscripts in fully-grown oocyte (Evsikov et al., 2004; Mehlmannet al., 2004; Peaston et al., 2004, 2007). Later in oogenesis,transcription of repetitive elements including, intracisternalA particle (Iap), long interspersed nuclear element 1 (Line1) andshort interspersed nuclear element (SINEb) are silenced by DNAmethylation (Lees-Murdock et al., 2003; Rollins et al., 2006; Walshet al., 1998). Failure to re-methylate repetitive elements can lead to

X. Tian, F.J. Diaz / Developmental Biology 376 (2013) 51–6152

aberrant expression and/or genome instability including theincrease in DNA double-strand breaks (Hedges and Deininger,2007). DNA methylation also causes global transcriptional silen-cing of cellular genes in fully-grown oocyte (Bouniol-Baly et al.,1999; De La Fuente and Eppig, 2001; Miyara et al., 2003). Histonemethylation and acetylation are another type of epigenetic mod-ification regulated during oocyte growth and maturation. Acetyla-tion of histone 3 (K9 and K18), histone 4 (K5 and K12) increase asthe oocyte nears ovulation and the chromatin rearranges into a SNconfiguration (Kageyama et al., 2007). Likewise di- and tri-methylation of histone H3K4 and H3K9 increases in fully-grownoocytes (Kageyama et al., 2007). In the one-celled zygote, histonemethylation progressively increases in the paternal pronucleus asthe paternal genome is reprogrammed after fertilization (Arneyet al., 2002; Lepikhov and Walter, 2004; Liu et al., 2004). Deletionof genes encoding histone or DNA methyltransferase enzymesseverely compromises oocyte development and fertility (Andreu-Vieyra et al., 2010; Howell et al., 2001; Kaneda et al., 2004). Thus,chromatin methylation is an important component of epigeneticprogramming during oogenesis.

Zinc has recently been recognized as an important factorrequired for completion of meiosis and egg activation in vitro(Bernhardt et al., 2011; Kim et al., 2010; Kong et al., 2012; Suzukiet al., 2010; Tian and Diaz, 2012) and for follicle rupture andcompletion of meiosis in vivo (Tian and Diaz, 2012). Numerousstudies have shown that zinc deficiency during pregnancy causesabnormal embryo and fetal development and poor progeny health(Apgar, 1985; Keen et al., 2003; Uriu-Adams and Keen, 2010).In other tissues, zinc deficiency decreases histone and DNAmethylation (Breksa Iii and Garrow, 2002; Wallwork andDuerre, 1985). However, the consequence of zinc depletion duringthe final period of oogenesis on chromatin methylation, fertiliza-tion and preimplantation development has not been examined.Thus, given that, (1) zinc deficiency is known to decreasedchromatin methylation in other tissues and (2) chromatin methy-lation is increasing in the oocyte during antral follicular develop-ment, we hypothesize that preconception zinc deficiency willdecrease chromatin (DNA and histone) methylation and impairfertilization and preimplantation development. The findings pro-vide evidence in support of this hypothesis and establish theperiod of final oocyte growth as a critical transition in oocyteepigenetic programming that is sensitive to perturbation in wholebody zinc homeostasis.

Materials and methods

Animal model of zinc deficiency

Female CD1 mice (Mus musculus) were obtained from the researchcolony of the investigators. To study effects of acute in vivo zincdeficiency before ovulation on oocyte epigenetic programming andembryonic development, newly weaned 18 day old mice werehoused on wire-bottom racks in polycarbonate cages and given acontrol diet (29 mg zinc/kg) based on AIN76 (MP Biomedicals, Solon,OH) or a zinc deficient diet (ZDD), which is the same diet as thecontrol with zinc omitted from the mineral mix (o1 mg zinc/kg).Diets were given for 3 or 5 days before ovulation. A 5 day treatmentwas chosen because in previous (Tian and Diaz, 2012) and prelimin-ary work (data not shown, longer treatments with a zinc deficientdiet cause a decrease in ovulation. A 3 day treatment was used insome experiments to show a dosage effect of a shorter dietarytreatment. Animals were primed with 5 IU eCG (National Hormoneand Peptide Program, NIDDK) 2 days before the end of the dietarytreatment period. Fully-grown GV stage oocytes were collected bygentle puncture of follicles with 25 ga needles. To collect mature eggs,

animals were primed with eCG for 48 h and induced to ovulate with5 IU hCG. Ovulated oocytes were collected from the oviduct 13 hlater. To examine embryonic development, animals were induced toovulate with eCG/hCG and bred to a fertile male. Only animals with amating plug on the morning after mating were used for subsequentanalyses. Preimplantation embryos were flushed from the reproduc-tive tract on days 1.5 (2 cell) and 3.5 (blastocysts). Importantly, at thetime of hCG, animals receiving a zinc-deficient diet were switchedback to a control diet for the duration of the experiment. To examinefertilization ability in vitro, female mice were superovulated at theend of a 3 or 5 days dietary treatment as described above. Ovulatedoocytes were recovered from the oviduct and fertilized in vitro asdescribed previously (O’Brien et al., 2003). Briefly, expandedcumulus–oocyte complexes were released from the oviduct anddirectly transferred to fertilization dish containing sperm(�10�106/ml) collected from the tail of the epididymis of a fertilemale. The medium used was MEM with 3 mg/ml BSA as describedpreviously (O’Brien et al., 2003). Complexes were incubated for 5 h,washed of excess sperm and cultured for an additional 20 h (24 htotal). At the end of culture, the proportion of 2-cell embryos wasdetermined as a measure of fertilization. Pronuclear stage embryoswere collected 8 h after fertilization in vitro. Animals were main-tained according to the Guide for the Care and Use of LaboratoryAnimals (Institute for Learning and Animal Research). All animal usewas reviewed and approved by the IACUC committee at ThePennsylvania State University.

Immunofluorescence

Immunofluorescence staining of control and zinc-deficientoocytes and fertilized eggs was conducted essentially as describedpreviously (Tian and Diaz, 2012; Tian et al., 2010). Briefly, COCwere collected from eCG-primed mice (48 h) and manuallydenuded by gentle pipetting to remove the cumulus cells. Oocytesand embryos from in vitro fertilization experiments were fixedimmediately in 4% PFA for 30 min. After fixing, oocytes andembryos were permeabilized with PBS (0.05% Tween and 0.5%Triton-X) for 15 min at room temperature followed by washing(3� ) in PBST (1% BSA)and then blocked in goat serum blockingbuffer (PBS, 2% goat serum, 1% BSA, 0.1% Triton-X and 0.05%Tween 20) for 1 h followed immediately by incubation with anti-trimethylated histone H3K4 (1:400, Cell Signaling) or anti-dimethylated histone H3K4 (1:400, Cell Signaling) for 1 h. For5-methylcytosine (5-Mec) detection, oocytes and embryos werefixed for 15 min with 4% PFA and treated with 2 N HCl for 30 minand then neutralized with Tris–HCl (PH¼8) for 15 min. Afterwashing in PBST, cells were immunostained with antibodyagainst 5-methylcytosine (1:200, Calbiochem) for 1 h. After wash-ing in PBST, oocytes and embryos were incubated with goat anti-rabbit IgG Alexa Fluor 594 or goat anti-mouse IgG Alexa Fluor 546for 1 h followed by washing with PBST and was mounted in DAPIanti-fade gold (Invitrogen) on glass slides with etched rings toprevent them from being ruptured by the cover slip. Slides wereimaged with a fluorescent microscope (AxioScope 2 Plus, Leica,Bannockburn, IL). Fluorescence intensity was determined withImage J software. All values within a replicate were normalized tothe average integrated pixel density of the control oocytes.

Isolation of total RNA and qPCR

Total RNA was isolated from 40 to 50 oocytes and 10 blastocystsusing the RNAeasy micro or mini kits (Qiagen, Valencia, CA),respectively. Total RNA was reverse transcribed into cDNA asdescribed previously (Diaz et al., 2006) using the Quantitek cDNAsynthesis kit (Qiagen, Valencia, CA). Quantification of Gdf9, Bmp15,Figla, Zp3, Nobox, Igf2, H19, Ddx4, Nr4a1 and Pou5f1 mRNAs was

Table 1Primer sequences used for qPCR.

Genesymbol

Forward Reverse

Igf2 AGGGGAGCTTGTTGACACG GGGTATCTGGGGAAGTCGTC

H19 CATGTCTGGGCCTTTGAA TTGGCTCCAGGATGATGT

Ddx4 CCCATTGTATTAGCAGGACGA GCGACTGGCAGTTATTCCAT

Nr4a1 CACAGCTTGGGTGTTGATGT GCTCCTTCAGACAGCTAGCAA

Gdf9 CTACAATACCGTCCGGCTCT CAAGTGTTCCATGGCAGTCA

Bmp15 ACACAGTAAGGCCTCCCAGA GATGAAGTTGATGGCGGTAAA

Figla ACAGAGCAGGAAGCCCGTA GTCAGAGGGTCTGCCACTGT

Nobox AGGGACGTTCCTGGCAGT GCTGCTTGCTTGGTAGTCCT

Iap ACAAGAAAAGAAGCCCGTGA GCCAGAACATGTGTCAATGG

Line1 GAGACATAACAACAGATCCTGA AACTTTGGTACCTGGTATCTG

Sineb1 GTGGCGCACGCCTTTAATC GACAGGGTTTCTCTGTGTAG

Sineb2 GAGATGGCTCAGTGGTTAAG CTGTCTTCAGACACTCCAG

Mt TGTTAAGAGCTCTGTCGGATGTTG ACTGATTCTTCAGTCCCAGCTAAC

Rpl19 TTCAAAAACAAGCGCATCCT CTTTCGTGCTTCCTTGGTCT

Fig. 1. Effect of zinc deficiency on chromatin methylation in GV-stage oocytes. Represe

(H3K4me2) (A), histone H3K4 trimethylation (H3K4me3) (B) and DNA methylation (C) i

5 days. Red¼methylated histone or 5-methylcytosine, Blue¼DNA (DAPI). *Po0.05, N¼

DNA methylation.

X. Tian, F.J. Diaz / Developmental Biology 376 (2013) 51–61 53

conducted using gene specific primers (Table 1) and Rpl19 mRNA asthe normalizer as described previously (Livak and Schmittgen, 2001;Tian and Diaz, 2012). Only one product of the appropriate size wasidentified for each set of primers and all amplification products weresequenced to confirm specificity. Primers for repetitive elements(Iap, Line1, Sneb1, Sineb2 and Mt) were validated previously (Su et al.,2012). Experiments were repeated 3–5 times and values shown arethe mean 7SEM.

Statistical analysis

Results from ovulation rate, the number of 2-cell stageembryos, blastocysts and qPCR were analyzed by either Student’st-test or two-way ANOVA followed by Tukey’s HSD post hoc test ifa positive F test was detected. Proportional data was transformed(arcsine) before analysis. The JMP 7.1 statistical analysis software(SAS, Cary, NC) and Microsoft Excel were used for analyses.

ntative images and nuclear fluorescence intensity of histone H3K4 dimethylation

n GV stage oocytes from animals receiving a control or zinc deficient diet (ZDD) for

3–4 (10–20 oocytes/replicate). Scale bar¼50 mM. (A) H3K4me2, (B) H3Kme3 and (C)

X. Tian, F.J. Diaz / Developmental Biology 376 (2013) 51–6154

Results

Zinc deficiency causes epigenetic defects in oocytes

To establish that oocyte epigenetic programming is altered inzinc deficient oocytes, we used immunofluorescence staining andmeasurements of nuclear fluorescence intensity in GV-stageoocytes from control and zinc deficient animals to measuredifferences in chromatin methylation. Dimethylation of H3K4was not altered by a 5 day treatment with a zinc deficient diet(Fig. 1A). However, little or no staining was observed fortrimethylated histone H3K4 in zinc deficient oocytes (Fig. 1B).To determine if DNA methylation might also be affected by zincdeficiency, an antibody against 5-methylcytosine (5-MeC) wasused to detect global DNA methylation. Surprisingly, DNA methy-lation was also dramatically reduced in zinc deficient oocytescompared to control oocytes (Fig. 1C).

Preconception zinc deficiency causes aberrant gene expression

in oocytes

Epigenetic defects, particularly reduced DNA methylationcould interfere with global transcriptional silencing and/or silen-cing of repetitive elements in oocytes. The expression of severalhighly abundant and important oocyte transcripts was measuredby qPCR as an indirect measure of transcriptional silencing.Concentration of Gdf9 and Figla mRNA were decreased by �50%and while Zp3 mRNA was only moderately decreased ((P¼0.08),Fig. 2A) in zinc deficient oocytes. The concentration of Bmp15 andNobox mRNA were not altered by zinc deficiency in oocytes.In contrast, there were significant increases in concentrations of

Fig. 2. Effect of zinc deficiency on transcript expression in GV-oocytes. (A)

Relative (fold change) concentration of oocyte-specific transcripts (Gdf9, Bmp15,

Zp3, Figla, and Nobox) in control and ZDD oocytes. (B) Relative (fold change)

concentration of repetitive elements (Iap, Line1, Sineb1, Sineb2 and MT) in control

and ZDD oocytes. *Po0.05, #P¼0.08, N¼4 (10 oocytes/replicate).

transcripts for various repetitive elements. Iap transcriptsincreased more than 20 fold while Line1, Sineb1 and Sineb2

transcripts increased 2–3 fold in zinc deficient oocytes. Mt

transcripts did not differ in zinc deficient oocytes compared tocontrols (Fig. 2B).

Fertilization potential in vitro is decreased by preconception zinc

deficiency

To test for detrimental effects of zinc deficiency (3 or 5 days)on fertilization that are independent of effects on the oviduct anduterus, the fertilization potential of control and zinc-deficientoocytes was determined in vitro. Ovulation rate did not differ ineither control of zinc deficient groups treated for 3 or 5 days(Fig. 3A). However, fertilization rate (proportion of mature eggsthat progress to the 2-cell stage) was dramatically decreased inthe ZDD group from 70% to 52% in the 3 day treatment group andfrom 83% to 8% in the 5 day treatment group (Fig. 3B–D). Thedramatic collapse in fertilization rate after a 5 day zinc deficienttreatment in fertilized eggs was caused by a failure to reachmetaphase II (no polar body, 79%) or by a failure of pronucleusfusion (pronuclei present, 21%) (Fig. 3E and F).

To determine if chromatin methylation is altered by zincdeficiency in fertilized eggs, oocytes from control or zinc-deficient animals (5 days) were fixed 8 h after fertilizationin vitro. Embryos were immunostained with an antibody recog-nizing H3K4me3. Embryos from control animals showed thetypical dimorphic pattern of intense immunostaining forH3K4me3 in the maternal pronucleus with little staining evidentin the paternal pronucleus (Fig. 4). The fertilized zinc deficienteggs showed disorganized chromatin characteristic of meioticfailure. However, the sperm nucleus did manage to enter theoocyte indicating that some early steps in fertilization still occurin zinc deficient oocytes.

In vivo fertilization and blastocyst development is impaired by

preconception zinc deficiency

In contrast to the dramatic decrease in fertilization ratein vitro, fertilization in vivo was not as dramatically affected byzinc deficiency. In vivo fertilization rate was 95% in controlanimals, but was reduced to 49% in ZDD animals (Fig. 5A andB). However, the 2-cell embryos from zinc deficient animals wereless competent than control embryos. Embryos were cultured inKSOM medium to determine the rate of blastocyst formation.Only 38% of 2-cell embryos from zinc deficient animals reachedthe blastocyst stage after in vitro culture compared to 74% ofcontrol embryos (Fig. 5C and D). To test if blastocyst developmentin vivo is also affected, animals were fed a control diet or zincdeficient diet for 3 and 5 days before ovulation and mated to afertile male as above. On day 3.5 after mating, embryos wereflushed from the reproductive tract. There was no difference inthe total number of embryos recovered from the reproductivetract in either experiment (3 day, control 3875, ZDD 4678;5 day, control 1973, ZDD 1972; N¼4–5). However, the propor-tion of blastocyst and expanded blastocysts were reduced, whilethe proportion or non-blastocyst (one cell to morula) and earlyblastocysts were increased in zinc deficient animals (Fig. 6). Thedecrease in proportion of blastocysts and expanded blastocystscould be related to a much lower level of Igf2 mRNA in zincdeficient blastocyst embryos (Fig. 7). Expression of the linkedtranscript, H19, was also decreased in zinc deficient embryos, butthe non-imprinted transcripts, Ddx4 (vasa) and Nr4a1 were notaffected by zinc deficiency in blastocyst embryos (Fig. 7). Impor-tantly, transcripts analyzed in Fig. 7 were measured in fully-formed blastocyst embryos from control and ZDD animals. Thus,

Fig. 3. Effect of zinc deficiency on in vitro fertilization. The number of ovulated oocytes (A), proportion of 2-cell embryos (B) and representative images (C) and (D) 24 h

after in vitro fertilization from animals receiving a control (C) or zinc deficient diet (ZDD) (D) for 3 or 5 days. Representative images of oocytes with meiotic failure (79%)

(E) and pronuclear arrest (21%) (F) in ZDD fertilized eggs. Blue¼DNA, Red¼Actin. *Significant difference by Student’s t-test, Po0.05, (N¼4; 50–60 eggs/replicate).

Proportion of 2-cell embryos were transformed (arcsine) before analysis. Scale bar¼50 mM (panels 3C–D) and 100 mM (panels 3E–F). (A) Ovulation Rate, (B) Fertilization

Rate, (C) Control embryos, (D) ZDD (5 days) embryos, (E) ZDD: Meiotic failure (79%) and (F) ZDD: pronuclear arrest (21%).

X. Tian, F.J. Diaz / Developmental Biology 376 (2013) 51–61 55

the decrease in Igf2/H19 mRNA is not due to an obvious differencein developmental stage.

Supplementation with SAM during IVM restores histone H3K4me3

and improves fertilization potential of zinc deficient oocytes

The dramatic decrease in chromatin methylation in oocytes fromanimals receiving a zinc deficient diet is alleviated by supplementa-tion with the methyl donor S-adenosylmethionine (SAM). Mice weregiven a zinc deficient diet for five before ovulation. The COC werecollected from animals primed with eCG and were cultured in MEM-alpha medium alone (control), medium only (ZDD) or medium

supplemented with 100 mM SAM (ZDDþSAM) for 24 h. At the endof culture, oocytes were fixed and immunostained for histoneH3K4me3. As expected, control oocytes had robust H3K4me3, butzinc deficient oocytes did not (Fig. 8). However, zinc deficient oocytescultured with SAM had levels of H3K4me3 similar to control oocytes(Fig. 8). To test if SAM can increase the proportion of zinc-deficientoocytes progressing to the 2-cell stage, COC were collected from eCG-primed animals fed a control or zinc deficient diet for 5 days andmatured in vitro. Control embryos have a high fertilization rate of 95%which was reduced to 19% in zinc deficient oocytes (Fig. 9). However,zinc deficient oocytes matured in the presence of SAM had a 45%fertilization rate (Fig. 9).

Fig. 4. Effect of zinc deficiency on histone H3K4 trimethylation in fertilized eggs.

Representative images of fluorescent immunostaining for histone H3K4 trimethy-

lation 8 h after in vitro fertilization of ovulated eggs from animals receiving a

control (A) or zinc deficient diet (ZDD) (B) for 5 days. (N¼3, 10–20 oocytes/

replicate). Red¼trimethylated histone, Blue¼DNA (DAPI), Green¼Actin. Scale

bar¼50 mM. (A) control and (B) ZDD.

X. Tian, F.J. Diaz / Developmental Biology 376 (2013) 51–6156

Discussion

Nutritional deficiencies during pregnancy can impair developmentand postnatal health by altering specific epigenetic marks (Burdgeet al., 2007; Cooney et al., 2002; Kim et al., 2009; Waterland and Jirtle,2003; Wu et al., 2004). Previously, we reported that dietary zincdeficiency caused multiple problems in ovarian function during theperiovulatory period, including failure to complete meiosis and a lackof cumulus expansion and ovulation (Tian and Diaz, 2012). Theexperimental design and findings of the current study are summar-ized in Fig. 10 and shows that maternal zinc deficiency during a verynarrow window just before ovulation disrupts oocyte epigeneticprogramming including a decrease in DNA and histone methylationand associated increase in expression of repetitive elements. Theseepigenetic defects along with previously shown meiotic defects (Tianand Diaz, 2012) severely compromise fertilization and preimplanta-tion embryonic development. However, whether zinc deficiency actsdirectly on the oocyte or indirectly through other mechanisms todecrease chromatin methylation in the oocyte is not known. In vitrosupplementation with S-adenosylmethionine, a methyl donor,restored histone methylation and improved fertilization rate of zinc

deficient eggs. Our challenge now is to fully uncover the precisepathways and proteins affected by zinc deficiency that lead todecreased oocyte developmental potential. Understanding thesezinc-mediated mechanisms will provide new insights into the reg-ulation of oocyte quality and fertility.

Experiments using dietary zinc deficiency during early preg-nancy have produced mixed results on fertility. In rats fed a zincdeficient diet on days 1–4 of gestation, fewer blastocyst embryosformed (Hurley and Shrader, 1975) suggesting delayed growth.However, an earlier study from the same group, using a similartreatment period (days 1–5 of gestation), found that a zincdeficient diet caused only minor effects on fertility and embryonicdevelopment at birth (Hurley et al., 1971). In mice, a zinc deficientdiet given one day before mating to one day after mating (3 days)did not disrupt blastocyst development (Peters et al., 1991). Theseobservations in rodents suggest that any defect induced by short-term (3–4 days) dietary zinc deficiency in early pregnancy can berescued by subsequent restoration of dietary zinc. In contrast tothese earlier studies, treatment with a zinc deficient diet duringthe 3–5 days before conception caused a dramatic decrease inmaturation, fertilization and preimplantation development. Pre-vious work clearly shows that zinc is essential for completion ofmeiosis I (Bernhardt et al., 2011; Kim et al., 2010; Tian and Diaz,2012). In the present study, we show a decrease in the fertiliza-tion rate of 25% and 90% after a 3 and 5 days treatment with a zincdeficient diet. Much of this decrease is due to a failure tocomplete meiosis, but the observed incidence of pronuclearformation (21%) and the appearance of partially decondensedsperm heads in the cytoplasm of fertilized zinc deficient eggssuggests that early steps in fertilization can still occur. Similarly,oocytes matured in the presence of a zinc chelator, TPEN, andfertilized in vitro progressed to the pronuclear stage and exhib-ited calcium oscillations, but did not divide to form 2-cellembryos (Kim et al., 2010). Clearly zinc deficient eggs, whetherin vivo or in vitro, can undergo early stages of fertilization butbecome arrested before the 2-cell stage. The pronuclear stagearrest in zinc deficient oocytes could also be the result ofpremature egg activation. Normally, zinc increases duringmaturation and is required for establishment of metaphase IIarrest. At fertilization, zinc is actively exported from the eggwhich allows resumption of the cell cycle and egg activation(Bernhardt et al., 2012; Kim et al., 2011). Interestingly, our resultsshow that zinc deficient eggs are more susceptible to fertilizationfailure in vitro than in vivo. What causes this difference infertilization potential is unknown, but does suggest that womenwith marginal or acute zinc deficiency may have lower successrates during ART procedure and may benefit from additional zincsupplementation. In addition to meiotic and fertilization defects,zinc deficient oocytes have reduced developmental competenceto the blastocyst stage after fertilization. On day 1.5 after mating,only 43% of 2-cell embryos from zinc deficient eggs reached theblastocyst stage when cultured in vitro compared to 95% ofcontrols embryos. On day 3.5 after mating there were fewerblastocyst and expanded blastocysts in zinc deficient animalscompared to controls suggesting a slower rate of development.The decreased expression of the growth promoting gene, Igf2

could account for some of the defects in preimplantation devel-opment, but how dietary zinc deficiency results in decreased Igf2/

H19 expression in the embryo remains to be determined. Collec-tively, these results firmly establish that preconception zincstatus is an important determinant of maturation, fertilizationand preimplantation development.

The findings support the hypothesis that preconception diet-ary zinc deficiency decreases oocyte developmental potential, butthe mechanism responsible for this effect is unknown. Thedecrease in oocyte H3K4me3 could account for many of the

Fig. 5. Effect of zinc deficiency on in vivo fertilization and blastocyst formation. Proportion of 2-cell embryos (A), representative images (B) of embryos recovered on day

1.5 of pregnancy from animals receiving a control or zinc deficient diet for 5 days. Proportion of blastocyst embryos (C) and representative images (D) of 2-cell embryos in

panel A, cultured for 3 days in KSOM medium. *Significant by Student’s t-test, Po0.05, N¼4–5 animals. Scale bar¼250 mM (panel B), 500 mM (panel D).

X. Tian, F.J. Diaz / Developmental Biology 376 (2013) 51–61 57

defects observed in zinc deficient oocytes. Recently, the lysinemethyltransferase protein, mixed lineage leukemia 2 (MLL2), wasshown to catalyze the trimethylation of H3K4 in oocytes (Andreu-Vieyra et al., 2010). Deletion of MLL2 in oocytes abolished bothtri- and dimethylation, but not monomethylation of H3K4. Theseepigenetic changes were associated with ovulation failure, lack oftranscriptional silencing, increased expression of repetitive ele-ments, and infertility. This phenotype is remarkably similar to ourdietary model of zinc deficiency where there is ovulation failure(Tian and Diaz, 2012), a decrease in H3K4 trimethylation and anincrease in expression of repetitive elements (present study).MLL2 is a zinc binding protein (Bach et al., 2008) and could bedirectly affected by a lack of zinc in the oocyte. However, thishypothesis remains to be tested directly.

Methylation of DNA on cytosine residues is another epigeneticmodification that is important for oocyte programming and wasdecreased by dietary zinc deficiency. DNA methylation is low ingrowing oocytes, but increases steadily as the oocyte nears ovula-tion (Kageyama et al., 2007; Kim et al., 2003). High levels of DNAmethylation are established at imprinting control regions ofimprinted loci (Bartolomei and Tilghman, 1997). DNA methylationis also essential for suppression of repetitive elements in the oocyte(Lees-Murdock et al., 2003; Rollins et al., 2006). Failure to silencerepetitive sequences can cause aberrant expression of nearby genes(Dolinoy et al., 2007) and/or genome instability (Hedges andDeininger, 2007). For example, methylation of the intracisternal

A-particle (Iap) prevents inappropriate expression of the nearbyAgouti gene. However, IAP becomes active when hypomethylatedand drives ectopic expression of the Agouti gene and associatedincrease in obesity and other metabolic defects in mice (Blewittet al., 2006; Morgan et al., 1999). In the oocyte, repetitive elementsoften serve as alternative promoters for other genes (Evsikov et al.,2004; Mehlmann et al., 2004; Peaston et al., 2004, 2007). GlobalDNA methylation was decreased significantly in zinc deficientoocytes and this likely contributed to the failure to represstranscription of repetitive elements. On the other hand, the lackof dramatic effects on cellular transcripts (Fig. 4A) suggests that aglobal transcriptional de-repression did not occur in zinc deficientoocytes. Further studies using transcriptional run-on assay, a directmeasure of transcriptional activity, are needed to determine theeffects of zinc deficiency on global transcription in the oocyte.Nevertheless, aberrant gene expression caused by increased activ-ity of repetitive elements together with decreased DNA and histonemethylation could be a major cause of developmental failure ofzinc deficient oocytes.

An open question is how does dietary zinc deficiency causemethylation defects in the oocyte. Methylation of DNA andhistone proteins requires the biosynthesis of the universal methyldonor, S-adenosylmethionine or (SAM) (Loenen, 2006). Duringmethylation reactions, SAM is converted to S-adenosyl homocys-teine (SAH), which is reversibly converted to homocysteine andthen to methionine. Methionine then serves as the substrate to

Fig. 6. Effect of zinc deficiency on preimplantation development in vivo.

(A) Proportion of embryos at different developmental stages including, non-

blastocysts (single cell to morula), early blastocyst, blastocyst and expanded

blastocyst collected from control and 3 day ZDD groups on 3.5 days after mating.

(B) Representative images of embryos from the 3 day treatment group. (C)

Proportion of embryos at different developmental stages collected from animals

in the 5 day treatment group. (C) Representative images of embryos from the

5 day treatment group (Control and ZDD). *Significant by Student’s t-test, Po0.05,

N¼4–5 animals, group. Scale bar¼100 mM. (A) Blastocyst 3 day ZDD (%), (B)

Brightfield, (C) Blastocyst 5 dat ZDD (%) and (D) Brightfield.

Fig. 7. Effect of zinc deficiency on blastocyst gene expression. Relative (fold

change) concentration of Igf2, H19, Ddx4 and Nr4a1 mRNA in blastocyst embryos

recovered from animals receiving a control and ZDD for 5 days. Relative levels of

mRNA measured by qPCR using the 2-ddct method. *Significant difference by

Student’s t-test, Po0.05, N¼4 (10 blastocyst/replicate).

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regenerate SAM. The recycling of homocysteine to SAM is undercontrol of two zinc-dependent enzymes, betaine–homocysteinemethyltransferase (BHMT) and 5-methyl tetrahydrofolate

homocysteine methyltransferase (MTR) (Koutmos et al., 2008;Loenen, 2006). BHMT and MTR catalyzed the transfer of methylgroups to homocysteine to re-generate methionine. This is thekey step in the re-cycling of SAM from methionine. Both SAH andhomocysteine are potent inhibitors of methylation reactions (DeCabo et al., 1995; de la Haba and Cantoni, 1959; Hoffman et al.,1980). Therefore, the efficient conversion of homocysteine tomethionine is essential to prevent build-up of these inhibitorymetabolites. Inhibition of BHMT decreases SAM and increaseshomocysteine concentration in the liver (Strakova et al., 2011),thus not only decreasing methyltransferase substrate (SAM), butalso increasing a potent inhibitor of methyltransferase activity(SAH). Similarly, zinc deficiency decreases DNA and histonemethylation in the liver (Breksa Iii and Garrow, 2002; Wallworkand Duerre, 1985), possibly by limiting SAM availability formethylation reactions and/or increasing inhibition by SAH/homo-cysteine. The full complement of enzymes involved in methionineand SAM biosynthesis are expressed in oocytes (Benkhalifa et al.,2008; Ikeda et al., 2010). We propose a working hypothesis thatzinc deficiency inhibits production of SAM in the oocyte resultingin reduced methylation capacity and hence reduced DNA andhistone methylation. Another possibility is that SAM produced inthe liver, which is a major site for SAM biosynthesis, could supplySAM to the oocyte through the circulation. Thus, it is equallypossible that zinc deficiency could inhibit hepatic SAM biosynth-esis and thus indirectly affect SAM concentration in the oocyte.Further work is needed to distinguish between these two possi-bilities. Regardless of whether SAM in the oocyte is producedlocally or originates from the liver, supplementation of oocytesfrom zinc deficient animals with SAM during maturation restoredhistone methylation and greatly improved fertilization success.Thus, SAM depletion appears to be one way that dietary zincdeficiency degrades oocyte quality. There is evidence in theliterature that regulation of SAM/SAH metabolism has a signifi-cant effect on fertility. High homocysteine concentrations infollicular fluid are associated with decreased oocyte developmen-tal potential in women undergoing assisted reproduction (Ebischet al., 2006; Steegers-Theunissen et al., 1993) and in women withPCOS (Berker et al., 2009). Whether high follicular fluid SAHconcentrations are associated with epigenetic alterations is notknown, but certainly warrants further research.

Zinc deficiency is common in many parts of the world(Wuehler et al., 2005) and in minority and poor populations inthe US (Schneider et al., 2007). Impaired liver function or liverdamage caused by alcoholism also results in systemic zincdeficiency (Flynn et al., 1981; Sullivan and Lankford, 1965). Ourfindings show that acute dietary zinc deficiency before ovulationdramatically decreases oocyte quality and developmental poten-tial and is associated with defects in epigenetic programmingpossibly caused by local or systemic SAM depletion. These

Fig. 8. (A) Effect of SAM supplementation H3K4me3 nuclear fluorescence intensity. Representative fluorescence images of histone H3K4me3 in GV oocytes from animals

receiving a control (B) or zinc deficient diet (ZDD) (C) and cultured in medium alone or medium containing 100 mM SAM (D) for 24 h.*Significant difference by ANOVA

followed by Tukey’s HSD test, Po0.05, N¼3 replicates with 10–15 oocytes/group. Scale bar¼50 mM. (A) Histone H3K4me3, (B) Control, (C) ZDD and (D) ZDD þ SAM.

Fig. 9. SAM rescue during IVF. Proportion of 2-cell embryos (A) and representative images (B) of in vitro matured and fertilized oocytes from control, ZDD and ZDD animals

supplemented with SAM (100 mM) during IVM (16 h). *Significant difference by ANOVA followed by Tukey’s HSD test, Po0.05, N¼3 replicates with 20–30 COC/group.

Scale bar¼50 mM. (A) Fertilization (%) and (B) Brightfield.

Fig. 10. Schematic showing timing of treatment in relation to endpoints examined. Dietary treatments were applied to growing oocytes just entering antral stage of

development around 18 days of age. Treatments were stopped 48 h after the end of eCG, which encompasses the period of antral follicular growth. Effects on maturation,

fertilization and preimplantation development were determined. Previous research showed that TPEN treatment disrupts completion of meiosis I, but effects of in vivo zinc

deficiency on fertilization and preimplantation development have not been reported.

X. Tian, F.J. Diaz / Developmental Biology 376 (2013) 51–61 59

X. Tian, F.J. Diaz / Developmental Biology 376 (2013) 51–6160

observations and the previously shown effects of zinc deficiencyon the meiotic cell cycle (Bernhardt et al., 2012; Kim et al., 2011,2010; Kong et al., 2012; Tian and Diaz, 2012) could lead toimproved methods of assisted reproductive procedures by mod-ulating zinc and/or SAM availability during oocyte maturationand fertilization. The present findings add further evidence thateven transient dietary deficiencies can affect fertility. Futureresearch will examine how zinc deficiency leads to reduces SAMbiosynthesis and whether other metabolic pathways are affected.Finally, it will be very interesting to determine the impact ofpreconception zinc deficiency on postimplantation developmentand postnatal health.

Funding

Research was supported by start-up funds from the College ofAgricultural Sciences at The Pennsylvania State University andNIH HD057283 to FJD.

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