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  • Research article
  • Open Access

Sexually dimorphic gene expression emerges with embryonic genome activation and is dynamic throughout development

Contributed equally
BMC Genomics201516:295

https://doi.org/10.1186/s12864-015-1506-4

  • Received: 28 July 2014
  • Accepted: 30 March 2015
  • Published:

Abstract

Background

As sex determines mammalian development, understanding the nature and developmental dynamics of the sexually dimorphic transcriptome is important. To explore this, we generated 76 genome-wide RNA-seq profiles from mouse eight-cell embryos, late gestation and adult livers, together with 4 ground-state pluripotent embryonic (ES) cell lines from which we generated both RNA-seq and multiple ChIP-seq profiles. We complemented this with previously published data to yield 5 snap-shots of pre-implantation development, late-gestation placenta and somatic tissue and multiple adult tissues for integrative analysis.

Results

We define a high-confidence sex-dimorphic signature of 69 genes in eight-cell embryos. Sex-chromosome-linked components of this signature are largely conserved throughout pre-implantation development and in ES cells, whilst the autosomal component is more dynamic. Sex-biased gene expression is reflected by enrichment for activating and repressive histone modifications. The eight-cell signature is largely non-overlapping with that defined from fetal liver, neither was it correlated with adult liver or other tissues analysed. The number of sex-dimorphic genes increases throughout development. We identified many more dimorphic genes in adult compared to fetal liver. However, approximately two thirds of the dimorphic genes identified in fetal liver were also dimorphic in adult liver. Sex-biased expression differences unique to adult liver were enriched for growth hormone-responsiveness. Sexually dimorphic gene expression in pre-implantation development is driven by sex-chromosome based transcription, whilst later development is characterised by sex dimorphic autosomal transcription.

Conclusion

This systematic study identifies three distinct phases of sex dimorphism throughout mouse development, and has significant implications for understanding the developmental origins of sex-specific phenotypes and disease in mammals.

Keywords

  • Sexual dimorphism
  • Pre-implantation embryo
  • Chromatin
  • Gene expression
  • Epigenetics
  • Development
  • Mouse
  • Mammalian
  • Sex-specific

Background

Sex determines anatomical, physiological and behavioural development in mammals. This developmental divergence arises as a consequence of sex-chromosome complement and is largely, although, not exclusively mediated through the organisational and activational effects of sex-specific hormones [1]. In adulthood, sex-specific gene expression is widespread in somatic tissues [2]. Consequentially, sex influences a plethora of complex traits that do not directly relate to reproductive roles. Exemplifying this, many diseases exhibit sex bias in prevalence or severity [3], and association of genetic variants with disease states is sex-dependent [4,5].

The ‘Four Core Genotypes’ mouse model has been used to dissect the contribution of sex-chromosome complement and the organisational and activational effects of sex-specific hormones [6], revealing that both hormones and chromosomal complement exert independent and divergent effects over metabolic and behavioural phenotypes [7,8]. Furthermore, animal models of the Developmental Origins of Health and Disease hypothesis reveal differential outcomes of early environmental insults dependent on sex [9], suggesting that sex should be considered as third parameter in any gene-environment interactions. Despite this, the origin and nature of these sex based differences are largely unexplored.

There is evidence that sexually dimorphic gene expression arises prior to gonadal differentiation, even in pre-implantation blastocysts, with both sex-linked and autosomal genes affected [10,11]. This dimorphism could determine the effects of specific environmental factors on long term developmental outcomes. Indeed, embryo culture medium induces long-term effects on glucose homeostasis in a sexually dimorphic manner [12]. Despite the relevance to human health and reproductive technologies, the origins and developmental dynamics of mammalian sexual dimorphism have not been characterized in detail. Here we report a high confidence signature of sexually dimorphic genes in the mouse coincident with embryonic genome activation using unmanipulated embryos, and from fetal and adult liver and performed comparative analyses using data from other sources. Our findings demonstrate that dimorphic expression emerges in the early cleavage embryo and is highly dynamic throughout development. Furthermore, we show that this is reflected in the chromatin structure. Through defining the nature and developmental origins of sexual dimorphism we provide a background for interpreting gene-environment interactions in directing developmental outcomes.

Results

Sexually dimorphic gene expression emerges very early in development

Eight-cell embryos provide the first snap-shot in mice after the initiation of embryonic genome activation at the 2 cell stage, when maternal transcripts have been depleted [13,14]. We isolated 12 eight-cell embryos for a discovery set and a further 12 for validation. Importantly, the embryos were produced by natural mating of inbred C57BL/6 J mice, without super-ovulation or in vitro culture, thereby providing a unique opportunity to examine sexually dimorphic expression in vivo. We generated transcriptome profiles for individual embryos using an adapted single-cell RNA-seq protocol (sequencing statistics are given in Additional file 1: Table S1). To our knowledge, this is the largest number of genome-wide transcriptome profiles from individual mouse embryos to date. We detected 13 469 transcripts expressed across all eight cell embryos, which is greater than previous microarrays of blastocysts, but less than other RNA-seq data from eight cell embryos that was sequenced at greater depth [11,14]. The sex of the embryos was determined by plotting expression of the Y chromosome gene, Eif2s3y and the X chromosome gene, Xist, as both are expressed in a sex-specific manner in murine blastocysts [10]. A clear separation of 2 distinct groups consisting of 6 female and 6 male embryos for the discovery set was found (Additional file 2: Figure S1). We then sought to identify genes that were differentially expressed between male and female embryos, revealing 69 genes with a genome-wide corrected p-value < 0.1, (Benjamini-Hochberg correction) and −0.5 < log2 (male/female) > 0.5 (Table 1). The majority of sex dimorphic differences (78%) originated from the X and Y chromosomes (51/69 and 3/69, respectively; Figure 1A), whilst ~22% (15/69) were autosomal in origin (Figure 1B). All X-linked genes were more expressed in female eight-cell embryos. Sex-dimorphic autosomal genes were equally distributed amongst the sexes.
Table 1

Genes defined as sexually dimorphic from eight-cell embryos

Gene name

CHR

Start

End

Strand

baseMean

log2FoldChange

Xist

chrX

103460373

103483233

-

275.66

−9.01

Ube1y1

chrY

818713

844224

+

232.95

9.02

Ddx3y

chrY

1260715

1286613

-

153.51

8.49

Bex1

chrX

136213972

136215513

-

168.51

−6.46

B230206F22Rik

chrX

103560910

103623754

-

34.38

−6.73

Sms

chrX

157443954

157492046

-

6792.54

−1.32

Eif2s3y

chrY

1010612

1028598

+

224.03

5.94

Rbm3

chrX

8138975

8145802

-

3756.52

−1.06

Fthl17

chrX

9033486

9034333

+

18.80

−5.67

Huwe1

chrX

151803282

151935417

+

2245.04

−0.81

Hprt

chrX

52988078

53021660

+

16923.45

−1.10

Eif2s3x

chrX

94188709

94212651

-

3306.76

−0.81

Syap1

chrX

162856843

162888462

-

2009.79

−1.13

Yy2

chrX

157566119

157568985

-

237.80

−2.07

Eif1ax

chrX

159372195

159385699

+

1826.30

−1.01

Ccng1

chr11

40748552

40755286

-

403.37

1.88

D7Ertd715e

chr7

59969577

59974431

-

365.59

−1.63

Atp6ap2

chrX

12587759

12617051

+

1103.98

−1.02

Uba1

chrX

20658421

20683179

+

4718.65

−0.82

Ddx3x

chrX

13281022

13293983

+

4503.61

−0.96

Timm8a1

chrX

134537258

134541629

-

3804.40

−1.20

Tsr2

chrX

151087094

151096543

-

2495.42

−0.76

Med14

chrX

12675371

12761973

-

725.57

−1.36

Usp38

chr8

80980733

81014906

-

1115.47

−0.99

Rhox5

chrX

37754608

37808878

+

27.14

−3.92

Gnl3l

chrX

150983133

151017322

-

910.96

−0.74

Kdm6a

chrX

18162667

18279358

+

481.77

−0.98

Als2cr11

chr1

59050506

59094900

-

9.24

4.02

Aen

chr7

78895927

78908833

+

513.62

0.99

Bcap31

chrX

73686183

73716175

-

1554.31

−0.63

Rbbp7

chrX

162760372

162779090

+

4036.08

−0.81

Acsl4

chrX

142317993

142390535

-

1262.01

−1.08

Idh3g

chrX

73778963

73786897

-

2177.00

−0.71

Lamp2

chrX

38405064

38456455

-

1278.91

−1.03

Rbmx

chrX

57383348

57393036

-

321.01

−0.91

Porcn

chrX

8193850

8206525

-

303.00

−1.11

Ssr4

chrX

73787028

73790828

+

798.15

−0.82

Otud5

chrX

7841831

7874858

+

728.02

−0.71

1810030O07Rik

chrX

12654884

12673546

-

134.89

−1.85

Atrx

chrX

105797615

105929372

-

707.08

−0.85

Prps1

chrX

140456603

140476140

+

8178.23

−0.77

Aifm1

chrX

48474944

48513426

-

387.98

−1.18

Gm14511

chrX

8975712

8976559

-

31.79

−3.30

Mapkapk2

chr1

131053704

131097543

-

46.62

2.72

Wdr45

chrX

7722249

7728201

+

2311.35

−0.85

Cox7b

chrX

106015700

106022450

+

116.54

−1.37

Mad2l1bp

chr17

46147385

46153551

-

1191.38

−0.92

Tmem56

chr3

121202010

121263316

-

542.41

−1.12

Rpl10

chrX

74270816

74273135

+

14774.94

−0.80

Cybasc3

chr19

10577723

10589830

+

465.93

0.85

Mtf1

chr4

124802549

124849800

+

512.08

−0.75

Cks2

chr13

51645232

51650662

+

2503.02

−0.66

Gm14458

chrX

8985900

8986755

-

5.13

−3.53

Cdca7

chr2

72476219

72486890

+

867.49

0.67

Pgrmc1

chrX

36598225

36606079

+

414.68

−1.00

Thoc2

chrX

41794994

41911901

-

381.21

−1.06

Cited1

chrX

102247440

102251769

-

3344.44

−0.53

Ypel5

chr17

72836704

72851195

+

2403.16

−0.70

Cdk16

chrX

20688493

20699877

+

802.78

−0.70

Dnajc28

chr16

91614257

91618999

-

10.24

3.47

Fmr1

chrX

68678555

68717961

+

587.22

−1.20

Gla

chrX

134588169

134601005

-

476.84

−1.06

Hnrnph2

chrX

134601286

134607054

+

879.15

−0.74

Mpp1

chrX

75109733

75130949

-

430.95

−1.07

Msl3

chrX

168654118

168673902

-

355.99

−1.12

Tbc1d25

chrX

8154472

8176181

-

258.99

−1.15

Trappc1

chr11

69323986

69325793

+

118.07

1.26

Utp14a

chrX

48256934

48282449

+

619.81

−0.73

Xiap

chrX

42067836

42109664

+

449.57

−0.86

Dimorphic genes were defined from RNA-seq data generated from 6 male and 6 female eight-cell embryos (adjusted p-value < 0.1, (Benjamini-Hochberg correction) and −0.5 < log2(male/female) > 0.5). TSS = transcriptional start site.

Figure 1
Figure 1

Identification and biological replication of sexually dimorphic genes identified in eight-cell embryos. Transcripts were defined as sex dimorphic from RNA-seq data using a genome-wide corrected p-value < 0.1 and −0.5 < log fold change (male/female) > 0.5 (A) Sexually dimorphic X-linked (51) and Y-linked (3) genes identified in the discovery set of six male and six female eight-cell embryos. (B) Autosomal genes (15) identified in the discovery set of six male and six female eight-cell embryos. Transcripts more highly expressed in a particular sex are indicated in red, those less expressed are indicated in blue. Rows representing individual transcripts were scaled to mean = 0, standard deviation = 1 (C) Average log fold change (male/female) for the discovery set (x-axis) plotted against log fold change (male/female) for an independent set of eight female and four male embryos (r = 0.95, p-value < 2.2 × 10−16). X-linked genes are indicated in red, Y-linked genes are indicated in blue and autosomal genes are indicated in green.

Substantial variation between individual embryos is apparent (Figures 1A,B, Additional file 2: Figure S2), and may reflect small differences in the timing of embryonic genome activation. However, we validated sex-biased expression of these genes in another 12 eight-cell embryos isolated in an independent experiment (8 female, 4 male) (Figure 1C). Biological replicates showed a strong directional correlation (ρ = 0.95, p-value < 2.2x10−16), with all but one of the 69 genes showing the same sex-based expression bias, even though transcripts show average log(male/female) values small in magnitude. Three non-coding transcripts were included in the eight-cell signature, consistent with a role for non-coding RNAs in early embryo development [15]. Two of these were involved in X inactivation, Xist and B230206F22Rik (also known as Ftx) [16,17]. The third, D7Ertd715e is located immediately 3′ to the Snrpn/Snurf imprinted cluster on chromosome 7, but its function is unknown. We performed KEGG analysis of X-linked genes, using detected X chromosome genes as background and separately for the autosomal genes, using all detected autosomal genes as background. Neither analysis revealed enrichment after Bonferroni correction (p < 0.05).

The dynamics of sexual dimorphism at different developmental stages

Having defined a high confidence, replicated gene expression signature from eight-cell embryos, we explored the temporal dynamics of these genes with respect to sex in pre- and post-implantation development. We obtained publically available transcriptome data derived from multiple stages (late two-, four- and sixteen-cell) of mouse pre-implantation development [14]. Not all embryonic stages profiled by Deng et al., were included because sex could not be determined (earlier than late two-cell), only one sex was sampled, or the sex ratio was highly skewed (e.g. blastocysts). Several factors make using this data for defining stage-specific signatures problematic. The embryos profiled by Deng et al. are F1 hybrids of two genetically diverse inbred mouse strains (CAST/EiJ females × C57BL/6 J males). As such, embryo sex is directly confounded by genotype. They were also produced using super-ovulation, which may influence embryo development [18]. Furthermore, only small numbers of embryos were profiled at each stage (<5 total, Table 2), reducing the confidence of signatures defined de novo. Nonetheless, the data is useful for comparative analyses using the high confidence signature we defined from eight-cell embryos. To ensure that the coverage and dynamic range between these datasets is comparable, as we used different RNA-seq protocols, we first compared our eight-cell male embryos (n = 6) to eight-cell male embryos profiled by Deng et al., (n = 4), establishing a high degree of correlation between these profiles (Additional file 2: Figure S3, ρ =0.84, p-value < 2.2 × 10−16). To get a more expansive view of the developmental dynamics of dimorphic expression, we additionally generated embryonic stem (ES) cells derived from eight-cell embryos (2 male, 2 female) using the 2i method [19]. The 2i methodology results in ES cell lines that are transcriptionally and epigenetically similar to pre-implantation epiblast [20-22]. This was confirmed by a strong correlation of the 2i ESC transcriptome to recently published inner cell mass single-cell profiles (Additional file 2: Figure S4, ρ = 0.70, p-value < 2.2 × 10−16) [23]. To profile post-implantation development we generated RNA-seq profiles from 17.5 dpc (days post-coital) fetal and adult liver (12 male, 12 female at each stage). We selected liver, because sex-based differences are well characterised and prolific in adults [24]. The ES cell, fetal and adult transcriptomes were all from a C57BL/6 J background.
Table 2

Sample characteristics and sources for comparative analyses

Sample type

Number of males

Number of females

Data type

Source

2-cell embryos

2

3

Single cell RNA-seq

[14]

4-cell embryos

1

3

Single cell RNA-seq

[14]

8-cell embryos

5

0

Single cell RNA-seq

[14]

8-cell embryos (discovery)

6

6

Whole embryo RNA-seq

This study

8-cell embryos (replication)

4

8

Whole embryo RNA-seq

This study

16-cell embryos

2

2

Single cell RNA-seq

[14]

2i ES cells

2

2

RNA-seq

This study

2i ES cells

2

2

ChIP-seq

This study

Fetal liver

12

12

RNA-seq

This study

Placenta

2

4

Microarray

[28]

Adult liver

12

12

RNA-seq

This study

Adult liver

165

169

Microarray

[24]

Adult adipose

165

169

Microarray

[24]

Adult brain

165

169

Microarray

[24]

Adult muscle

165

169

Microarray

[24]

Taking the two-, four- and sixteen-cell embryo data generated by Deng et al., together with our ES cell, fetal and adult liver data, we next examined whether the dimorphic expression of the 69 genes defined in eight-cell embryos is conserved throughout development. Unfortunately, Deng et al., only sampled males at the eight-cell embryo stage, precluding a direct comparison at this developmental time-point. Of the 69 signature genes, only one gene located on the Y chromosome (Eif2s3y) was expressed in a sexually dimorphic manner (logFC > 0.5) across all profiles, from 2 cell embryos onwards. From the four-cell stage onwards, a substantial proportion of transcripts are of embryonic origin [13,14] and the sexually dimorphic expression of the other Y chromosome genes (Ddx3y, Ube1y1) is established. The sex-specific expression of the majority of X chromosome genes is established from the two- to four-cell stage and maintained throughout pre-implantation development and in the ES cells (Figure 2A). However, the majority of the sex-chromosome genes within this signature do not show dimorphic expression in fetal or adult liver.
Figure 2
Figure 2

Sex biased expression of sexually dimorphic genes defined in eight-cell embryos across pre-implantation development, fetal and adult tissues. Sex-biased expression is represented as log fold change (male/female). Box plots represent mean with interquartile range. Whiskers are defined by extremes within 1.5X the interquartile range and additional points outside this range are shown as circles. (A) X-linked genes (red). Data for the placenta is restricted to 45/51 transcripts for which data was available (B) Autosomal genes (green) more expressed in male embryos. Data for the placenta is restricted to 7/8 transcripts for which data was available (C) Autosomal genes (green) more expressed in female embryos. Data for the placenta is restricted to 6/7 transcripts for which data was available.

Interestingly, the 15 autosomal genes that are sexually dimorphic in eight-cell embryos did not show consistent directional changes at other time-points in pre-implantation development. As these genes were expressed at comparable levels to the sex-chromosome encoded transcripts, it seems unlikely that an increased signal to noise ratio is responsible for this. Despite not showing dimorphism at other pre-implantation stages, 14/15 of these genes show biological validation at the eight-cell stage (Figure 1C), suggesting that this stage-specific autosomal sexual dimorphism is bona fide. The expression level of these genes seems to be very dynamic throughout pre-implantation development (Additional file 2: Figure S5). Similar pre-implantation stage-specific expression has also been shown for genes identified as dimorphic in bovine blastocysts [11]. Indeed, global autosomal gene expression reaches a nadir at the eight-cell stage (Additional file 2: Figure S6). Collectively, we show that sex-chromosome genes show largely consistent sexual dimorphism throughout pre-implantation development and in ground state pluripotent stem cells, regardless of genetic background, but that dimorphism of autosomal genes is more temporally dynamic.

X-linked dimorphic genes escape paternal X inactivation in pre-implantation development

The emergence of sexually dimorphic expression of Xist at the four-cell stage coincides with the initiation of transcriptional inactivation of the paternal X chromosome in mice [25]. Higher X-linked expression in females must therefore reflect either an up-regulation of these genes from the maternal X chromosome in females, or escape from paternal X chromosome inactivation in the pre-implantation stages. Consistent with the escape from silencing of the paternal X we find that the majority of X-linked sex dimorphic genes are located distally from the X inactivation centre (Additional file 2: Figure S7), confirming previous observations [26]. Indeed, allelic data generated by Deng et al., confirms that 35/38 (~92%) genes that had adequate allele-specific information were expressed from both maternal and paternal X chromosomes at one or more developmental time-points (four-, sixteen- or early blastocyst) [14]. Although there is some suggestion that inter-specific crosses may have altered X inactivation, reassuringly, two of the three transcripts solely of paternal origin are known to be involved in silencing the chromosome from which they are transcribed, Xist and B230206F22Rik (also known as Ftx) [16,17]. The other transcript (Gla) showed paternal-specific expression at the four-cell stage, with allelic data for later stages not available. Notably, silencing of paternal expression of this transcript has previously been shown to be initiated after the eight-cell and completed only by the blastocyst stage [27].

Sex-biased expression of most X chromosome genes that are dimorphic in eight-cell embryos, with the exception of Xist, and Eif2s3x is erased (−0.5 < logFC > 0.5) in fetal and adult liver (Figure 2A), but female-biased expression was retained for 5/51 X-linked genes in publically available gene expression array data from late gestation placenta [28], at log(male/female) < −0.5. As extra-embryonic tissues in the mouse demonstrate imprinted inactivation of the paternal X chromosome, we asked if dimorphic X-linked expression is more similar in general between these two tissues. Using a cut-off of log(male/female) < −0.5, we identified 79 X-linked genes showing female biased expression in placenta and found that there was a trend towards female-biased expression of these genes in the eight-cell embryos (t test, mean = −0.14, p-value = 0.046). Consistent with re-activation of the inactive paternal X chromosome in the epiblast-like 2i ES cells, the female bias in expression of the dimorphic X-linked genes identified from eight-cell embryos is maintained and expanded upon (Additional file 2: Figure S8). In fetal and adult somatic tissues, a different set of X chromosome genes are dimorphic.

Sex-biased expression is correlated with post-translational histone modifications

Specific post-translational histone modifications are associated with particular transcriptional states and genomic features. We sought to explore the relationship of sexually dimorphic expression in pre-implantation development with chromatin features. To address this, we generated genome-wide profiles for three post-translational histone modifications in our 2i ES cells using chromatin immunoprecipitation combined with deep sequencing (ChIP-seq). 12–20 million mapped reads were generated for each mark (Additional file 1: Table S1). Determining average profiles for the 2000 most high- or low-expressed genes confirmed the expected enrichment of H3K4me3 at active gene promoters, whilst H3K27me3 was depleted (Additional file 2: Figure S9). Both H3K27me3 and H3K9me3 were enriched at repetitive elements and H3K9me3 was associated with imprinted genes, confirming the distribution of these modifications is typical of ground state pluripotency [20]. Genes located on the X chromosome conform to genome-wide distribution patterns.

Given the association of H3K4me3 with transcriptional activity, we next asked whether the X-linked genes identified as sex-dimorphic in the eight-cell embryos are more enriched for H3K4me3 in female 2i ES cells. Dimorphic expression of 46/51 of these genes is conserved in the ES cells. Before assessing relative enrichment for X-linked dimorphic genes, we first needed to account for the difference in chromosome dosage between the sexes. When ChIP-seq reads were counted in 3 kb windows surrounding the TSS, we were able to show that all three histone marks show relative enrichment in females, as expected. We developed a model allowing us to correct for chromosome dosage in subsequent analyses (see Methods). Interestingly, after applying the model for dosage correction, we found that sex-dimorphic genes show female-biased enrichment of H3K4me3 surrounding the TSS (FEfemale = 1.47; p-value = 0.002), whilst H3K9me3 (FEfemale = 0.72; p-value = 0.028) and H3K27me3 (FEfemale = 0.63; p-value = 0.005) show greater enrichment in the males (Figure 3). This is consistent with enrichment of H3K4me3 in female compared to male ES cells at the X-linked Rhox6/9 genes which is lost upon differentiation and loss of expression [29].
Figure 3
Figure 3

X-linked genes more expressed in female eight-cell embryos show female-biased enrichment for H3K4me3 and depletion of H3K9me3 and H3K27me3 in 2i embryonic stem (ES) cells. After correcting for sex differences in X chromosome dosage, X-linked genes that do not show sex-biased enrichment of post-translational histone modifications should fit the model (black line). All X-chromosome genes were used as background (grey points). Enrichment was calculated for the 3 kb surrounding the transcriptional start site from ChIP-seq data generated from 2i ES cells. Dimorphic X-linked genes identified from eight-cell embryos are more expressed in females (orange points). (A) H3K4me3 is relatively enriched in females for the dimorphic genes (FEfemale = 1.47; p-value = 0.002) (B) H3K9me3 is relatively depleted in females for the dimorphic genes (FEfemale = 0.72; p-value = 0.028) (C) H3K27me3 is relatively depleted in dimorphic genes (FEfemale = 0.63; p-value = 0.005). Significance for fold enrichment was calculated using permutation tests.

Although power for de novo calling is limited by the number of biological replicates for the ES cells, we were able to identify sex-biased expression (genome-wide corrected p-value < 0.1, −0.5 < logFC > 0.5) for use in correlative analyses. Whilst the X-linked genes identified as sex-dimorphic in eight-cell embryos show evidence for escape from paternal X inactivation, they represent only a subset of the X-linked genes identified as sex-dimorphic in ES cells, where the paternally silenced X has been reactivated (Additional file 2: Figure S8). Consistent with what was found for the transcripts identified from eight-cell embryos, we observe that transcriptional activity of X-linked genes defined from ES cells is associated with enrichment of H3K4me3 in the 3 kb surrounding the TSS (Figure 4A; t-test p-value < 2.2x10−16 (mean = −0.136)). However, the ES cell defined X-linked dimorphic genes differ from the eight-cell embryo defined subset in that whilst male cells show H3K9me3 enrichment (Figure 4B; t-test p-value < 2.2x10−16 (mean = 0.619)), male specific H3K27me3 enrichment is very weak (Figure 4C; t-test p-value = 0.04 (mean = 0.037)), suggesting differential regulation of these gene subsets at the chromatin level, despite both maternal and paternally inherited alleles in both subsets being transcriptionally active in the ES cells.
Figure 4
Figure 4

Histone post-translational modifications associated with the transcriptional start site (TSS) of sexually dimorphic genes in 2i embryonic stem cells. Transcripts were defined as sex dimorphic from RNA-seq data using a genome-wide corrected p-value < 0.1 and −0.5 < log fold change (male/female) > 0.5. The average log fold change (male/female) is shown on the x axis. Enrichment for specific post-translational histone modifications was calculated for the 3 kb surrounding the TSS from ChIP-seq data generated from 2i ES cells (y axes). Correlations for sex dimorphic X-linked genes and TSS (A) H3K4me3 enrichment (t-test p-value < 2.2 × 10−16 (mean = −0.136)) (B) H3K9me3 enrichment (t-test p-value < 2.2 × 10−16 (mean = 0.619)) (C) H3K27me3 enrichment (t-test p-value = 0.04 (mean = 0.037)). Correlations for dimorphically expressed autosomal genes and TSS (D) H3K4me3 enrichment (ρ = 0.57, p-value < 2.2 × 10−16) (E) H3K9me3 enrichment (ρ = 0.0052, p-value = 0.89) (F) H3K27me3 enrichment (ρ = −0.077, p-value = 0.036).

A positive correlation between expression and H3K4me3 enrichment in the 3 kb surrounding the TSS of sex-dimorphic autosomal genes expressed in ES cells was found (Figure 4D; ρ = 0.57, p-value < 2.2x10−16). TSS H3K9me3 enrichment does not correlate with gene expression, although is enriched in males (Figure 4E; ρ = 0.0052, p-value = 0.89), whilst H3K27me3 is very weakly anti-correlated (Figure 4F; ρ = −0.077, p-value = 0.036). The autosomal genes identified in the eight-cell embryos were not called as significantly dimorphic in the ES cells, although they were still expressed. Furthermore, as there were so few sexually dimorphic autosomal genes in the eight-cell signature, we could not make correlations with the ESC histone marks. Our findings suggest that sex-biased gene expression associates with enrichment of H3K4me3 at the TSS in the sex that has higher expression. This relationship is true for both X-linked and autosomal genes.

Sexual dimorphism is more pronounced in adult compared to fetal liver

Sexually dimorphic expression of the eight-cell signature genes in liver at adult and fetal stages did not recapitulate that found in pre-implantation development (Figure 2). To investigate the inter-relationship of sex-biased gene expression at later developmental stages, we identified a signature of 1488 dimorphic genes within adult liver (genome-wide corrected P < 0.1, −0.5 < logFC > 0.5; Additional file 3: Table S2). Autosomal genes account for ~96% of the dimorphic genes, in contrast to what is observed at pre-implantation stages (Additional file 2: Figure S8). KEGG analyses identified many enriched pathways, with drug metabolism as most significant, consistent with previous findings (Additional file 4: Table S3) [30]. Reassuringly, our data strongly correlated with previously published adult liver dimorphic differences, despite the different profiling platforms and genetic backgrounds between the two datasets (ρ = 0.82; p-value < 2.2x10−16) [24]. On the premise of this, we established a cut off (−0.5 < logFC > 0.5) by which we could define a signature for the other tissues; adipose, muscle and brain, profiled by Yang et al. Consistent with previous analyses we found that there was not a significant correlation for sex-dimorphic expression across tissues (data not shown), whilst X-linked genes showed modest female-biased expression differences [31]. Furthermore, the signature defined from eight-cell embryos did not show directional consistency in any of the adult tissues (data not shown).

We also defined a sexually dimorphic signature of 394 genes from late gestation fetal liver (genome-wide corrected p-value < 0.1, −0.5 < logFC > 0.5; Additional file 3 Table S2). As with adult liver, the majority (~94%) of sex-biased differences were autosomal in origin (Additional file 2: Figure S8). KEGG analysis identified 8 signaling and disease-associated pathways that were enriched in fetal liver after Bonferroni correction p-value < 0.05; (Additional file 4: Table S3). Using the same criteria, we identified 3.8X as many sex-dimorphic transcripts in adult compared to fetal liver. There was substantial overlap between the two developmental stages, with 72% (264/369) of autosomal, 67% (14/21) of X-linked and 100% (4/4) of Y-linked genes identified as dimorphic in fetal liver also being identified as dimorphic in adult liver (Additional file 3: Table S2), collectively representing a 11.95 fold enrichment (p-value < 0.001), when all expressed genes are considered as background. Yet, this enriched subset of dimorphic transcripts that is common across both fetal and adult liver do not show directional consistency across datasets (Figure 5) (X-squared = 2.07, p-value = 0.15). Our findings suggest that sexually dimorphic gene expression is present in late gestation liver, but is further expanded upon in the equivalent adult tissues, with some sexual-dimorphism being developmental stage specific. Consistent with previous findings, we did not find any up-regulation of X chromosome expression to equate with the level of autosomal gene expression, thereby refuting Ohno’s hypothesis [32]. The expression level of the X chromosome relative to autosomal genes was remarkably similar in males and females (1.84X and 1.82X, respectively for adult liver), but showed some variation according to developmental stage (1.43X and 1.43X, respectively for fetal liver).
Figure 5
Figure 5

Sexually dimorphic expression across fetal and adult liver does not show directional consistency. Transcripts were defined as sex dimorphic from RNA-seq data using a genome-wide corrected p-value < 0.1 and −0.5 < log fold change (male/female) > 0.5. X-linked genes are shown in red, Y-linked genes are shown in blue and autosomal genes are shown in green (A) The average log fold change (male/female) for sex dimorphic genes defined from fetal liver (x axis) correlated with expression in the adult liver (y axis); X-linked (ρ = 0.094, p-value = 0.70), autosomal (ρ = 0.068, p-value = 0.19). (B) The log fold change (male/female) for sex dimorphic genes defined from adult liver (x axis) correlated with expression in the fetal liver (y axis) X-linked (ρ = 0.013, p-value = 0.93), autosomal (ρ = −0.042, p-value = 0.13).

Sex-based expression differences in adult liver are known to be largely dependent on differences in growth hormone regulation in response to sex-specific gonadal hormones [33,34]. To test if growth hormone responsiveness underlies the differences in dimorphic expression between fetal and adult liver, we asked if the dimorphic genes present in adult, but not fetal liver were enriched for previously identified growth hormone responsive transcripts [35]. Indeed, adult-specific sex-dimorphic genes had a 5.4X enrichment (p < 0.001). Collectively, our results suggest that sex-biased expression of some genes in liver is intrinsic, but that sex differences in growth hormone induces dimorphism of additional genes. Our data suggests that this latter mechanism is not yet operational in late gestation.

Discussion

We present a whole-genome view of in vivo sexually dimorphic gene expression throughout mouse development, revealing that sex-specific expression biases in the embryo, fetus and adult are largely distinct.

Our study of dimorphism in pre-implantation development provides advances on previous work, by sampling prior to any cellular differentiation and eliminating culture induced artifacts [11,36,37]. By defining a high-confidence signature from eight-cell embryos we could then use this signature to probe other pre-implantation developmental stages despite the data being underpowered for defining sex dimorphic signatures de novo. We find that pre-implantation development is characterised by conserved dimorphism of sex-chromosome-linked genes, predominantly from the X chromosome [38]. Female-biased expression of some X-linked genes arises in two cell embryos, around the time embryonic genome activation is initiated in the mouse. Whether this is indicative of differences in the rate at which maternally inherited transcripts are degraded amongst sexes, or otherwise results from nascent transcription is unknown [14]. However, at least in the case of Fthl17, an X-linked maternally imprinted gene, transcription from the paternally inherited chromosome in the early embryo is responsible for female only expression [36]. Some of the X-linked genes in our eight-cell signature have been shown to escape paternal X inactivation in extra-embryonic tissues [39,40]. Allelic information suggests that for many of these genes, escape from paternal X silencing might also underlie their dimorphism in pre-implantation development. Consistent with previous findings, absolute paternal expression is either associated with the establishment of X inactivation (with the exception of Fthl17), whilst partial expression is more common amongst genes distal from the X inactivation centre [26] These genes also show a selective depletion of H3K27me3 in female ES cells, in contrast to sex-dimorphic X-linked genes that are not dimorphic in the eight cell embryos, even though the transcriptional behaviour between these subsets is similar in the ES cells. The difference between gene subsets escaping imprinted X inactivation in the pre-implantation embryo and placenta might be accounted for by progressive silencing of the paternal X chromosome throughout the pre-implantation period [27]. Similar X-linked dimorphism may not be conserved in humans, however, as regulation of X-inactivation is substantially different between species [41].

Autosomal genes also show dimorphism in pre-implantation stages, but sex-specific expression of these genes seems to be temporally restricted. Although we did not see conservation of these differences at other pre-implantation stages, we demonstrate validation in two independent cohorts of eight-cell embryos. The identification of dimorphic autosomal genes in eight-cell embryos implies regulation by sex-specific trans acting factors. In epiblast-like ES cells, autosomal genes highly expressed in one sex had corresponding enrichment for H3K4me3 around the TSS, consistent with what has been observed for strongly sex-biased genes in mouse liver [42]. Our eight-cell dimorphic signature included both X-linked transcription factors, e.g. YY2, and chromatin modifiers e.g. Kdm6a [37]. Interestingly, Kdm6a, has been shown to regulate specific targets, such as the Rhox6/9 cluster in a sex-specific manner in ES cells [29], whilst its Y-encoded homolog (Uty), does not completely recapitulate its function [43,44]. Sex-chromosome complement might otherwise influence autosomal gene transcription through the inactive X chromosome influencing heterochromatic gene silencing in trans [45].

In line with previous findings, we found a large number of dimorphically expressed genes in adult liver that were not conserved across other adult tissues. Expanding on this, we also show that directionality was not conserved with the pre-implantation signature. We did, however, identify a subset of genes that show dimorphic expression in both fetal and adult liver. KEGG pathway analysis revealed dimorphism in common pathways relating to metabolism and stage-specific pathways relating to signal transduction in the liver at these two stages. Many additional genes were dimorphic in adults, possibly due to the activational effects of gonadal hormones. Consistent with this hypothesis we show that adult-specific liver dimorphic genes are enriched for growth hormone responsiveness [35]. Although gonadal hormones are produced in late gestation, the regulation of pituitary growth hormone secretion by gonadal hormones is minimal prior to puberty in mice [46].

Conclusions

Our findings reveal that there are core transcriptional differences between the sexes that are consistent throughout pre-implantation development. Most of the genes identified in eight-cell embryos do not retain significantly dimorphic expression in fetal and adult tissues. Although sex-dimorphic expression is highly tissue-specific, a subset of genes is conserved across fetal and adult liver suggesting that sex-biased expression of this subset may be due to sex-chromosome complement, whilst differences are more likely driven by sex-specific physiology, which differs according to the stage of development. Although at a gene level we see a dramatic difference across pre-implantation, fetal and adult development, pathway level analysis reveals some conservation. By providing a genome-wide view of sex-dimorphic expression from post-fertilisation to adult, we hope to improve the understanding of the underlying molecular biology of sexually dimorphic phenotypes.

There is substantial evidence to suggest that sex-specific responses to environmental stimuli can occur prior to exposure to sex-specific hormones, and in some instances, even when the exposure is restricted to the previous generation [47]. Sex-chromosome complement can drive dramatic effects, as exemplified by the epigenome-wide differences in male and female murine ES cells when cultured in the presence of serum or defined medium [22,48]. Such effects may have lasting impact on developmental trajectories and disease risk [12]. Through characterising sexually dimorphic gene expression through a developmental trajectory, we reveal novel aspects of sex-specific biology and the inter-relationship of key phases across development. Our findings provide a platform for future work exploring the role of sex in moderating gene-environment interactions and highlight the importance of incorporating sex in studies of common disease and interventions [9,49].

Methods

Animal maintenance and tissue isolation

All animal procedures were conducted in accordance with the Home Office Animals (Scientific Procedures) Act 1986 (Project License number - 70/6693). C57BL/6 J mice were obtained from Charles River Laboratories, UK. All animals were maintained on standard laboratory chow and a 12 hr light/dark cycle. Male mice were housed with virgin females overnight. The detection of a vaginal plug the following morning was considered 0.5 dpc. Females were killed at 2.5 dpc and eight-cell embryos isolated from the fallopian tubes by flushing. Alternatively, females were killed at 17.5 dpc and embryonic liver was collected and snap frozen with liquid nitrogen. Adult animals were killed at 15–20 weeks of age and tissues collected and snap frozen. All animals were killed between 10 am and 12 pm.

Single cell isolation from 8 cell embryos

Single embryos were transferred to acidic Tyrode’s solution to remove the zona pellucida, then washed in PBS-BSA (1 mg/mL) and dissociated into single cells as described previously [50]. All cells from a single embryo were used to generate a single RNA-seq library. For the discovery set, embryos of both sexes were derived from 4 independent litters, embryos used for biological replication were derived from an Additional 5 independent litters. The sires of the replicate litters were exposed to in utero protein restriction.

Derivation of embryonic stem (ES) cell lines

ES cells were derived from eight-cell embryos as previously described [51]. Briefly, isolated eight-cell embryos were cultured in KSOM medium supplemented with (2i): mitogen-activated protein kinase inhibitor (PD0325901, 1 μM) and glycogen synthase kinase-3 inhibitor (CHIR99021, 3 μM). After two days, embryos were transferred to NDiff227 medium with 2i and LIF and allowed to develop into blastocysts. The trophectoderm was lysed by immunosurgery, and the ICM (inner cell mass) of each embryo plated in NDiff227 supplemented with 2i and LIF and for expansion to generate 2i ES cell lines.

Generation and sequencing of RNA-seq libraries

The eight-cell embryo RNA-seq libraries were generated using a single-cell protocol adapted from [50]. Further details are provided in Supplementary Information.

Generation and sequencing of ChIP-seq libraries

The chromatin immunoprecipitation (ChIP) assay was performed according to previously published protocols with minor modifications [52]. Chromatin was sonicated to get fragments of 100 to 700 bp and immunoprecipitated with the following antibodies anti-H3K27me3 (07–449, Millipore), anti-H3K4me3 (39159, Active Motif) and anti-H3K9me3 (ab8898, Abcam). ChIP-seq libraries were prepared using the Illumina ChIP-seq librarary prep kit, according to the manufacturers’ instructions.

Analysis

All analysis was performed on UCSC reference genome mm10 and the gene annotation file downloaded from Tophat website (downloaded on September 25th 2013).

RNA-Seq analysis

All RNA-Seq data was mapped using Tophat v2.0.4 with Bowtie 2 v2.1.0 and samtools v0.1.18 using default settings. The mean insert sizes and standard deviations were calculated in silico using Picard Tools v1.98 from 1,000,000 reads. Duplicates were filtered using Picard Tools and reads were assigned to a gene using HTSeq. Differential analysis was performed using DESeq2 v1.4.5 which uses a generalised linear model in which counts are modelled using a negative binomial distribution. Genome wide corrected p-values were calculated using Benjamini-Hochberg multiple testing adjustment procedure. Normalised variance stabilizing transformed counts were used for all plots and further analysis. Existing public data was extracted as raw counts and analysed with DESeq2 in the same manner as our data.

ChiP-Seq analysis

All ChiP-Seq data was mapped using Bowtie 2 v2.1.0 with default settings. Coverage across the genome was calculated using genomeCoverageBed from Bedtools v2.17.0 and converted to BigWig format using bedGraphToBigWig downloaded from the UCSC website. For correlation with RNA-Seq data the average coverage over +/−1.5 kbp of the TSS of each gene was calculated using bigWigAverageOverBed.

Chromosome dosage model

To account for the difference in chromosome dosage between the sexes we defined a model based on the distribution of reads for the 2 male samples. We first removed the Y chromosome and then counted the (average) total number of reads (across the 2 samples) within a 3 kb window around the TSS of each of the genes across all remaining chromosomes including chrX (N total ). We then calculated the average total number of reads on the X chromosome (N x ). A scaled read density for chromosome X genes was then calculated:
$$ {S}_i=\frac{2{N}_i}{N_{total}+{N}_x} $$
Where N i represents the number of reads for gene i on chromosome X. We can then compare this to the actual (average) read density for the X chromosome of the female samples calculated as:
$$ {A}_i=\frac{N_i}{N_{ftotal}} $$
where N ftotal is the total number of reads (excluding the y-chromosome) for the female samples and N i represents the number of reads for gene i on chromosome X.

Data access

ChIP-seq data and RNA-seq data are available in the NCBI Gene Expression Omnibus (GSE59222), and will be made public upon acceptance of the manuscript for publication.

Notes

Declarations

Acknowledgments

VKR is supported by grants from the Biotechnology and Biological Sciences Research Council, UK (BB/M012494/1), VKR and CG by (BB/G00711/X/1). MLH is supported by a Research Council UK Academic Fellowship. RL is supported by EU-FP7 BLUEPRINT. We thank King’s College London FWB Genomics Centre for performing high throughput sequencing.

Authors’ Affiliations

(1)
Centre for Genomics and Child Health, Blizard Institute, Barts and The London School of Medicine and Dentistry, 4 Newark Street, London, E1 2AT, UK

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