Dosage compensation of Z sex chromosome genes in avian fibroblast cells
Genome Biology volume 24, Article number: 213 (2023)
In birds, sex is genetically determined; however, the molecular mechanism is not well-understood. The avian Z sex chromosome (chrZ) lacks whole chromosome inactivation, in contrast to the mammalian chrX. To investigate chrZ dosage compensation and its role in sex specification, we use a highly quantitative method and analyze transcriptional activities of male and female fibroblast cells from seven bird species. Our data indicate that three fourths of chrZ genes are strictly compensated across Aves, similar to mammalian chrX. We also present a complete list of non-compensated chrZ genes and identify Ribosomal Protein S6 (RPS6) as a conserved sex-dimorphic gene in birds.
The molecular mechanisms regulating avian sex determination are not well-understood. Sex in birds is specified genetically, with the female being heterogametic (ZW sex chromosomes) and male homogametic (ZZ sex chromosomes) (Fig. 1A) [1,2,3,4], opposite to eutherian mammals (male XY and female XX). Phylogenomic analyses indicated that Z and X sex chromosomes (chrZ and chrX) evolved separately from different autosomes and likely originated from dimorphic expression of autosomal genes involved in sex differentiation . Suppression of meiotic recombination of chrZ/chrX led to their degeneration into chrW/chrY, respectively . In many non-avian/non-mammalian amniotes (all crocodiles, all sphenodons, some squamates and turtles), sex determination is governed not genetically but rather via environmental factors (e.g., temperature) during critical developmental windows (Fig. 1A) . It has been hypothesized that temperature sex determination was the ancestral mode in amniotes .
A well-studied example of dosage-compensation is XIST (a long non-coding RNA)-mediated chrX condensation and subsequent inactivation in placental mammals [8,9,10]. In birds, however, chrZ does not undergo whole-chromosome condensation or transcriptional inactivation . Estimate of percentage of compensated chrZ genes varies in literature (~20–45%, [12,13,14]). It has been proposed that avian dosage compensation is regulated locally and that lack of compensation of chrZ genes (e.g., DMRT1) is necessary to drive sex-specific germ cell and gonadal differentiation [15,16,17,18,19,20]. In addition to dimorphism in gonadal morphology, birds exhibit secondary sexual dimorphism (e.g., body weight, plumage shape, wattle size and courtship behavior) which is mainly under cell-autonomous regulation via inherited genetic identity and weakly modulated by sex hormones after gonadal differentiation [19, 21,22,23].
Results and discussion
To investigate how sex-specific differentiation in birds is programed cell-autonomously, we compared transcriptional activities in purified male and female embryonic fibroblast cells from seven avian species (the chicken, quail, turkey, blue peafowl, duck, zebra finch and emu), representing all three major clades: the Palaeognathae (emu), Neoaves (zebra finch), and Galloanserae (the rest) (Materials and Methods) (Fig. 1A). Sex was assessed by gonad size , and in each species, at least three embryos (including both male and female) were collected for fibroblast isolation (one embryo per sample). Two different culture conditions (DMEM high glucose and DMEM/F12) (Materials and Methods) were used to control medium-associated expression variation. Cultured cells were passaged five times to reach homogeneity for RNA isolation (Additional file 1: Fig. S1A, showing chicken, peafowl and emu fibroblasts). All cells grew robustly, and no prominent difference was observed between fibroblast cells under different culture conditions or from different origins (individuals, sexes or species), suggesting that our fibroblast cell isolation method and culture protocol are applicable to all birds.
Cap analysis of gene expression (CAGE) maps gene expression by capturing and sequencing 5′ end of mRNA [24, 25]. CAGE-seq can identify transcription start sites (TSSs) with single nucleotide precision and quantify TSS activities free of amplification- or transcript size-introduced biases [26, 27], ideal for analysis of minor expression variations between male and female cells. Applicability of CAGE-seq to chicken samples was reported previously . We decided to use CAGE-seq to investigate sex-biased gene expression in male and female fibroblast cells from these seven species (Fig. 1A; Additional file 1: Fig. S1B; Additional file 2: Table S1; Materials and Methods). CAGE replicates correlated well with each other and with corresponding RNA-seq data used for validation (Additional file 1: Fig. S2, S3).
On average, 8582 TSS peaks representing 7608 genes (Fig. 1B; Additional file 1: Fig. S4; Additional file 3: Table S2) were identified in avian fibroblast cells. Of those, 95.4% (7269 genes) did not show statistically significant difference between the male and female samples (Fig. 1C; Additional file 4: Table S3), and 4.5% (339 genes) showed either male-biased (189) or female-biased (150) differential (sex dimorphic) expression (Fig. 1C; Additional file 4: Table S3). Dimorphism in sex-associated TSS activities had the strongest effect on sample clustering (Fig. 1D; Additional file 1: Fig. S5A), largely due to differential activities of both chrW and chrZ genes in all species studied except for the emu (Fig. 1E; Additional file 1: Fig. S5B, Additional file 5: Table S4) and in agreement with several reported chrW markers (e.g., HINTW, CHDB1, and ATP5A1W) (Fig. 1F) . Although differentially expressed genes were significantly enriched on sex chromosomes (Additional file 6: Table S5), many of them were also located on autosomes (1736 out of 2370, 73.2%; Additional file 4: Table S3), and these genes had an average male/female ratio higher than chrZ genes (Fig. 1G), suggesting that dimorphic expression of chrZ/chrW genes likely control sex specification through autosome genes.
The number of dimorphic genes located on sex chromosomes (Z and W) in each species is shown in Additional file 4: Table S3. In chicken, of the 132 sex chromosome genes with sex specific differential expression, 115 were on the chrZ and 17 on chrW (Fig. 1H; Additional file 4: Table S3). This indicated that of the 387 chrZ genes that were expressed in fibroblast cells, only 30% (115/387) exhibited statistically significant difference between the two sexes, with the rest showing no significant difference. In other birds, the percentages of expressed genes showing statistically significant difference were 6% (23/392) in quail, 35% (123/356) in turkey, 22% (85/380) in duck, 25% (101/411) in zebra finch, and 19% (63/324) in emu (Additional file 4: Table S3). When we excluded peafowl sex chromosome data (due to poor chrZ annotation) , number of genes on chrZ showing no statistical difference between the male and female samples were consistently high in the remaining species (chicken 70%; quail 94%; turkey 65%; duck 78%; zebra finch 75%; and emu 79% [29% when excluding PAR]) (Fig. 1I). These percentages increased further when we applied male/female expression ratio cutoffs (e.g., 1.3-fold and 1.5-fold). Using the most stringent criterion for non-compensation (i.e., genes exhibiting any statistically significant difference between the sexes without applying any ratio cutoff) and not considering PAR in Neognathae species, the average percentage of dosage-compensated chrZ genes was 78% (Fig. 1I). This number is significantly greater than previous estimates for avian chrZ dosage compensation (see introduction) and is comparable with the percentage of X-linked, dosage-compensated genes reported in human fibroblasts [~75% in  and ~70% in ], suggesting that avian dosage compensation works as effectively as in mammals despite the lack of morphologically-distinct chrZ inactivation in birds. Contribution of gametologous genes to differential expression is available in Additional file 7: Table S6, and examples of expression profile are shown in Additional file 1: Fig. S6.
How birds modulate their chrZ gene expression in male versus female cells is unclear. In addition to chromosome-level inactivation in homogametic sex (e.g., female mammals, XX), dosage compensation can be achieved through doubling sex chromosome expression in heterogametic sex (e.g., male Drosophila, XY) , halving sex chromosome expression in homogametic sex (e.g., female nematodes with XX; and male silkworms with ZZ) [34, 35], or a hybrid mechanism (e.g., in monarch butterfly, halving expression in one chrZ segment in homogametic sex and doubling expression in another chrZ segment in heterogametic female) . Our data indicated obvious regionalized distribution of either compensated or non-compensated genes on avian chrZ in the case of emu, in which male-biased chrZ genes appeared to cluster to the stratum 0, whereas PAR (pseudo-autosomal region)-located genes, as expected, did not show sex-specific expression shifts (Additional file 1: Fig. S7A). Stratum 0 and PAR are previously reported features of homomorphic sex chromosome evolution in birds [37, 38]. Autocorrelation analysis revealed weak regionalization of non-compensated chrZ genes in several non-ratite species in comparison to the emu (Additional file 1: Fig. S8).
Sex dimorphic gene expression in birds is known to appear before gonadal differentiation [21, 22] and continue afterwards. However, DMRT1, a chrZ gene and a putative determining factor in testis differentiation, is not expressed in fibroblast cells and does not regulate secondary sex dimorphic characters [19, 20]. Such gonadal factor-independent sex-specific somatic cell specialization is referred to as cell autonomous sex identity (CASI) [23, 39], molecular mechanism of which is unknown. We analyzed our datasets for differentially expressed genes between male and female fibroblast cells in each species (Additional file 8: Table S7 with top 50 highlighted; top 8 shown in Additional file 1: Fig. S9). We noticed four interesting features, both validating previous findings and offering new insight to dosage compensation and CASI. First, one chicken chrZ gene (LOC112530614) exhibited strong inactivation in male fibroblast cells. This gene is located within the MHM1 (male hypermethylation 1) locus (Additional file 1: Fig. S7B) reported to be downregulated in male and active in female [14, 17, 40] and hypothesized to positively regulate female chrZ genes . Although this MHM1 locus is conserved in Galloanserae, it is not universally present in birds [14, 42]. Second, HINTW was listed as a chrW gene expressed in a female-specific manner in the chicken (Fig. 1F), validating previous reports from our and other labs [21, 43,44,45]. Direct- and cross-mapping suggested presence of female-specific HINTW expression in all species analyzed except for the emu (in which the homomorphic nature of its chrZ and chrW precluded us from making an unambiguous prediction). Third, many genes found in this study had not been reported previously and constituted a valuable resource for future functional studies of CASI in each species (Additional file 8: Table S7) (e.g., chicken COL4A1 and AHCY, quail HSPA8, and emu RAD23). Lastly, we uncovered a gene that was highly expressed and universally sex-dimorphic in all seven species (Additional file 1: Fig. S10A). This gene is located on chrZ and encodes RPS6 (ribosomal protein subunit 6) (Fig. 2A). It is extremely conserved evolutionarily (Fig. 2B, Additional file 1: Fig. S10B). Stably higher RPS6 expression in male cells at stages prior to gonadal differentiation (Additional file 1: Fig. S11A) and male-biased expression in all tested avian fibroblast cells indicated by CAGE (Fig. 2C) suggest its universal role in CASI. Differential expression of RPS6 in male and female fibroblast cells was further validated by quantitative PCR using fibroblast RNA from the chicken, quail, duck, and zebra finch (Fig. 2D). Furthermore, RPS6 expression was not affected by tested media conditions (Additional file 9: Table S8), although potential culture media-introduced bias in fibroblast transcriptome cannot be ruled out and awaits further investigation (Additional file 1: Fig. S12).
RPS6 has been implicated in metabolic regulation in mouse embryonic fibroblast cells  and cell cycle regulation in chicken embryonic fibroblast cells , suggesting its involvement in sex-specific metabolic and/or cell-cycle regulation. Supporting a potential role of RPS6 in avian CASI, this gene was reported to play a role in sex-specific vocal learning in zebra finch , and its expression exhibited both cell-autonomous dimorphism and estrogen inducibility in song regions of the finch brain, suggesting that RPS6 promoter is able to integrate both cell-autonomous, sex-specific transcriptional control and sex hormone-mediated systemic regulation. This latter aspect was supported by the presence of estrogen receptor binding sites in avian RPS6 promoters (Fig. 2A). It may also be a conserved feature in mammals, as RPS6 was recently identified as a significant indicator for sex-specific prognosis of diffuse glioma in humans .
In this work, we used embryonic fibroblast cells from seven birds and showed that avian chrZ genes were highly dosage-compensated, similar in its extent to mammalian chrX. Avian and mammalian dosage compensations likely involve different molecular mechanisms. Some chrZ genes were not compensated and may play a role in sex determination. RPS6 was identified as the only chrZ gene exhibiting male-biased expression in all species examined.
Egg collection and fibroblast cell preparation
Avian eggs from seven different species [chicken (Gallus gallus domesticus ), quail (Coturnix coturnix japonica), turkey (Meleagris gallopavo domesticus), blue peafowl (Pavo cristatus), duck (Anas platyrhynchos domesticus), zebra finch (Taeniopygia guttata) and emu (Dromaius novaehollandiae)] were obtained from the following sources in Japan: chicken eggs from Shimojima Farm, Kanagawa; blue peafowl eggs from Kumamoto City Zoo and Botanical Gardens, Kumamoto; quail eggs from Motoki corporation, Saitama; duck eggs from Shiina Hatchery Co., Ltd. , Chiba; turkey eggs from Shimanto Turkey Growers Association, Kochi; emu eggs from Japan Eco System Co., Ltd., Fukuoka; and zebra finch eggs from Hokkaido University, Hokkaido. For fibroblast cell preparation, chicken eggs were incubated at 38.5 °C to reach to stage HH36 . Developmental speed and total incubation time vary among birds . Embryos of all other species were incubated at 37.5–38.5 °C to reach the equivalent stage of chicken HH36. This corresponded approximately to day 10 (D10) in chicken, D9 in quail, D13 in duck and blue peafowl, D14 in turkey, D23 in emu, and D8 in zebra finch. Embryos were then treated by removal of head and neck, limb, skin and fat deposit, and internal organs. The remained trunk portion was trypsinized at 37 °C, followed by neutralization and centrifugation to obtain digested cell pellet. Cell pellet was resuspended with DMEM/10%FBS (FBS Cat# S1820-500 from Biowest, France; DMEM high glucose Cat# 043-30085 from FUJIFILM Wako Chemicals, Japan) and seeded in DMEM/10%FBS or DMEM/F12/10%FBS (DMEM/F12 Cat#042-30795 from FUJIFILM Wako Chemicals, Japan) for expansion.
Total RNA was prepared using commercial reagents (TRIzol, Invitrogen) from each sample of avian embryonic fibroblast cells at the fifth Passage. Five micrograms of RNA from each fibroblast sample was used for nAnTi-CAGE library preparation and subsequent sequencing using the Illumina HiSeq2500 platform (K. K. DNAForm, Yokohama, Japan). Quality of sequenced CAGE reads was checked by using FastQC v0.11.5 and then trimmed with fastx_trimmer -Q33 -l 75 (FASTX Toolkit 0.0.14). Moirai removeN script was applied to remove reads with “N” nucleotides. CAGE reads matching to adapters or rRNA (Moirai defined rRNA for galGal3) were removed with Trimmomatic-0.38 and RNAdust 1.06, respectively. Trimmed CAGE reads were aligned with BWA 0.7.10-r789 and unmapped reads realigned with Hisat2 v2.1.0 to following reference assemblies: chicken - galGal6 (GCF_000002315.6), quail - Coturnix_japonica_2.1 (GCF_001577835.2), turkey - melGal5 (GCF_000146605.3), turkey - MGAL_WU_HG_1.0 (GCA_905368555.1), peacock - GT_SO_6221 (http://gigadb.org/dataset/100559), duck - ZJU1.0 (GCF_015476345.1), zebra finch - bTaeGut2.pat.W.v2 (GCF_008822105.2), emu - droNov1 (GCF_003342905.1), emu - ZJU1.0 (GCA_016128335.1). Alignments were processed into CTSS (CAGE transcription start sites) and CAGE peaks with PromoterPipeline  with default threshold of at least 10 TPM (tags per million) in one of the samples. CTSS and CAGE peaks data were deposited in Zenbu (https://fantom.gsc.riken.jp/zenbu/reports/#Birds_Promoter_Atlas). CAGE libraries of all bird species and Chicken FANTOM data  were realigned to galGal6 and analyzed using the same approach. CAGE peaks were associated with nearby transcripts located on the same strand by using ChIPseeker v1.32.0 package for R. Gene models for MGAL_WU_HG_1.0 were built by using Augustus v3.2.3 with --species=chicken option and hints based on EST and mRNA for melGal5 obtained from UCSC and mapped with blat v.36x2 to MGAL_WU_HG_1.0. Completeness of gene annotation accessed by BUSCO v3.0.2 (76.8% complete). 2kb regions of CAGE peaks located within 100 bp from transcript start were used as a training set for TSSClassifier , and remaining distal intergenic, intronic, exonic, and UTR CAGE peaks were analyzed and classified into “promoters” or “not-promoters” TSS peaks. For the rest of the analysis, we used only promoter region localized (within 3 kb) peaks or other TSS peaks classified as “promoter.” Differential expression analysis was carried by using edgeR v3.38.1 package for R for each species separately on TPM counts using Generalized linear models’ approach. Cross-species comparison was carried out for the purpose of validation (Additional file 1: Fig. S13) through previously described approach  which uses modified variance-stabilizing transformation. Human dermal and mouse embryonic fibroblast samples were obtained from FANTOM6 and 5 repositories, respectively. Orthologs defined by using blastp (v2.2.29) -evalue 1e-5 against chicken protein sequences. Species specific genes were obtained by applying differential expression approach on cross-species normalized counts. To note, peacock genome is assembled to the level of scaffolds and chromosomes information is not available. For Additional file 1: Fig. S5, chicken sex chromosomes were used for Blastn against peacock assembly and the best match scaffold considered as a sex chromosome in peacock. Quail, peacock, and turkey genome assemblies do not include chrW.
Paired end libraries for RNAseq were prepared for each species using 1 female and 1 male samples from each species (a subset of samples used for CAGE-seq) and sequenced on Illumina HiSeq2500 (K. K. DNAForm, Yokohama, Japan). Reads were trimmed against adapter sequences and rRNA with Trimmomatic-0.38, aligned to the genome assemblies with Hisat2 v2.1.0, and counted against gene models with HTSeq v2.0.1. FPKM values were counted with edgeR package for R. Batch effect correction for MDS plot was done by using limma (“removeBatchEffect” function). scRNAseq analysis of published HH4-HH7 and S4-S13 data [54, 55] was carried out by using cellranger-7.0.1 and galGal6 as a reference. Obtained counts were processed through Pagoda2 and Conos packages with default settings to make integrated normalized expression matrix.
Total RNA extracted from fibroblast cells was used to obtain cDNA with SuperScript III Reverse Transcriptase kit (Thermo Fisher). Primers were designed for RPS6 as follows: forward primer #1 matching exon 2 and 3 GGAGTGGAAGGGCTATGTTG, reverse primer #1 matching exon 4 – TTGAACAGCTTGCGGAT, insert size 309 bp; forward primer #2 matching exon 3 GACGTGTCCGCCTTCTGCTC; reverse primer #2 matching exon 4 – TTCCTCACAACATACTGGCG, insert size 268 bp. Two genes with stable expression profiles ARPP19 and YWHAE were selected for control with primers as follows: ARPP19 forward primer on exon 2 – TGAAAGCAAGATACCCTCAT, reverse on exon 3 – TCTTCATCTTTGCTTTAGCC, size 126 bp; YWHAE forward on exon 2 – CAGTGGAAGAAAGAAACCTG, reverse on exon 3 – GAATGAGGTGTTTGTCCAGT, size 216 bp. Efficiency of primers was tested in series of cDNA dilutions 1:10, 1:100, 1:1000, 1:10000, and RPS6 primers #2 with YWHAE as control were selected for qPCR. The primer efficiency calculated as E =( 10^(-1/Slope)-1) × 100 (Additional file 1: Fig. S11B). qPCR was done on LightCycler 96 (Roche) with at least two technical and two biological replicates. Data was analyzed in pcr v1.2.2 package for R. Nucleotide alignment for RPS6 CDS and tree (Neighbor Joining method, Jukes-Cantor distance measure, bootstrap 1k) was created by using CLC Genomic Workbench 20.0.
Availability of data and materials
All transcriptomics data created in this work is deposited in GEO NCBI (GSE213253) as raw and processed expression tables with gene annotations . Genomic views are available through Zenbu-reports  browser https://fantom.gsc.riken.jp/zenbu/reports/#Birds_Promoter_Atlas. Single cell RNAseq data for chicken embryo were accessed through GSE181577  and GSE223189 .
Graves JA. Avian sex, sex chromosomes, and dosage compensation in the age of genomics. Chromosome Res. 2014;22:45–57.
Smith CA, Major AT, Estermann MA. The curious case of avian sex determination. Trends Genet. 2021;37:496–7.
Smith CA. Rowley review. Emu - Austral Ornithology. 2010;110:364–77.
Guioli S, Nandi S, Zhao D, Burgess-Shannon J, Lovell-Badge R, Clinton M. Gonadal asymmetry and sex determination in birds. Sex Dev. 2014;8:227–42.
Kratochvil L, Gamble T, Rovatsos M. Sex chromosome evolution among amniotes: is the origin of sex chromosomes non-random? Philos Trans R Soc Lond B Biol Sci. 2021;376:20200108.
Bachtrog D, Mank JE, Peichel CL, Kirkpatrick M, Otto SP, Ashman TL, Hahn MW, Kitano J, Mayrose I, Ming R, et al. Sex determination: why so many ways of doing it? PLoS Biol. 2014;12: e1001899.
Manuel F-B, Óscar L: Genotypic sex determination systems could be adaptations to extreme temperature environments in reptiles and to endothermy in mammals and birds. bioRxiv 2022:2022.2002.2007.479281.
Cerase A, Pintacuda G, Tattermusch A, Avner P. Xist localization and function: new insights from multiple levels. Genome Biol. 2015;16:166.
Loda A, Heard E. Xist RNA in action: past, present, and future. PLoS Genet. 2019;15: e1008333.
Okamoto I, Nakamura T, Sasaki K, Yabuta Y, Iwatani C, Tsuchiya H, Nakamura SI, Ema M, Yamamoto T, Saitou M. The X chromosome dosage compensation program during the development of cynomolgus monkeys. Science. 2021;374:eabd8887.
Sun D, Maney DL, Layman TS, Chatterjee P, Yi SV. Regional epigenetic differentiation of the Z Chromosome between sexes in a female heterogametic system. Genome Res. 2019;29:1673–84.
McQueen HA, Clinton M. Avian sex chromosomes: dosage compensation matters. Chromosome Res. 2009;17:687–97.
Ellegren H, Hultin-Rosenberg L, Brunstrom B, Dencker L, Kultima K, Scholz B. Faced with inequality: chicken do not have a general dosage compensation of sex-linked genes. BMC Biol. 2007;5:40.
Wright AE, Zimmer F, Harrison PW, Mank JE. Conservation of regional variation in sex-specific sex chromosome regulation. Genetics. 2015;201:587–98.
Mank JE, Ellegren H. All dosage compensation is local: gene-by-gene regulation of sex-biased expression on the chicken Z chromosome. Heredity (Edinb). 2009;102:312–20.
Itoh Y, Melamed E, Yang X, Kampf K, Wang S, Yehya N, Van Nas A, Replogle K, Band MR, Clayton DF, et al. Dosage compensation is less effective in birds than in mammals. J Biol. 2007;6:2.
Arnold AP, Itoh Y, Melamed E. A bird’s-eye view of sex chromosome dosage compensation. Annu Rev Genomics Hum Genet. 2008;9:109–27.
Lee HJ, Seo M, Choi HJ, Rengaraj D, Jung KM, Park JS, Lee KY, Kim YM, Park KJ, Han ST, et al. DMRT1 gene disruption alone induces incomplete gonad feminization in chicken. FASEB J. 2021;35: e21876.
Ioannidis J, Taylor G, Zhao D, Liu L, Idoko-Akoh A, Gong D, Lovell-Badge R, Guioli S, McGrew MJ, Clinton M. Primary sex determination in birds depends on DMRT1 dosage, but gonadal sex does not determine adult secondary sex characteristics. Proc Natl Acad Sci U S A. 2021;118(10):e2020909118.
Smith CA, Roeszler KN, Ohnesorg T, Cummins DM, Farlie PG, Doran TJ, Sinclair AH. The avian Z-linked gene DMRT1 is required for male sex determination in the chicken. Nature. 2009;461:267–71.
Nagai H, Sezaki M, Bertocchini F, Fukuda K, Sheng G. HINTW, a W-chromosome HINT gene in chick, is expressed ubiquitously and is a robust female cell marker applicable in intraspecific chimera studies. Genesis. 2014;52:424–30.
Zhang SO, Mathur S, Hattem G, Tassy O, Pourquie O. Sex-dimorphic gene expression and ineffective dosage compensation of Z-linked genes in gastrulating chicken embryos. BMC Genomics. 2010;11:13.
Zhao D, McBride D, Nandi S, McQueen HA, McGrew MJ, Hocking PM, Lewis PD, Sang HM, Clinton M. Somatic sex identity is cell autonomous in the chicken. Nature. 2010;464:237–42.
Consortium F, the RP, Clst, Forrest AR, Kawaji H, Rehli M, Baillie JK, de Hoon MJ, Haberle V, Lassmann T, et al: A promoter-level mammalian expression atlas. Nature 2014, 507:462-470.
Akalin A, Fredman D, Arner E, Dong X, Bryne JC, Suzuki H, Daub CO, Hayashizaki Y, Lenhard B. Transcriptional features of genomic regulatory blocks. Genome Biol. 2009;10:R38.
Kawaji H, Lizio M, Itoh M, Kanamori-Katayama M, Kaiho A, Nishiyori-Sueki H, Shin JW, Kojima-Ishiyama M, Kawano M, Murata M, et al. Comparison of CAGE and RNA-seq transcriptome profiling using clonally amplified and single-molecule next-generation sequencing. Genome Res. 2014;24:708–17.
Morioka MS, Kawaji H, Nishiyori-Sueki H, Murata M, Kojima-Ishiyama M, Carninci P, Itoh M. Cap analysis of gene expression (CAGE): a quantitative and genome-wide assay of transcription start sites. Methods Mol Biol. 2020;2120:277–301.
Lizio M, Deviatiiarov R, Nagai H, Galan L, Arner E, Itoh M, Lassmann T, Kasukawa T, Hasegawa A, Ros MA, et al. Systematic analysis of transcription start sites in avian development. PLoS Biol. 2017;15: e2002887.
Fridolfsson AK, Cheng H, Copeland NG, Jenkins NA, Liu HC, Raudsepp T, Woodage T, Chowdhary B, Halverson J, Ellegren H. Evolution of the avian sex chromosomes from an ancestral pair of autosomes. Proc Natl Acad Sci U S A. 1998;95:8147–52.
Dhar R, Seethy A, Pethusamy K, Singh S, Rohil V, Purkayastha K, Mukherjee I, Goswami S, Singh R, Raj A, et al. De novo assembly of the Indian blue peacock (Pavo cristatus) genome using Oxford Nanopore technology and Illumina sequencing. Gigascience. 2019;8(5):giz038.
Carrel L, Willard HF. X-inactivation profile reveals extensive variability in X-linked gene expression in females. Nature. 2005;434:400–4.
San Roman AK, Godfrey AK, Skaletsky H, Bellott DW, Groff AF, Harris HL, Blanton LV, Hughes JF, Brown L, Phou S, et al. The human inactive X chromosome modulates expression of the active X chromosome. Cell Genom. 2023;3: 100259.
Lucchesi JC, Kuroda MI. Dosage compensation in Drosophila. Cold Spring Harb Perspect Biol. 2015;7(5):a019398.
Lau AC, Csankovszki G. Balancing up and downregulation of the C. elegans X chromosomes. Curr Opin Genet Dev. 2015;31:50–6.
Rosin LF, Chen D, Chen Y, Lei EP. Dosage compensation in Bombyx mori is achieved by partial repression of both Z chromosomes in males. Proc Natl Acad Sci U S A. 2022;119: e2113374119.
Gu L, Reilly PF, Lewis JJ, Reed RD, Andolfatto P, Walters JR. Dichotomy of dosage compensation along the neo Z chromosome of the monarch butterfly. Curr Biol. 2019;29(4071–4077): e4073.
Vicoso B, Kaiser VB, Bachtrog D. Sex-biased gene expression at homomorphic sex chromosomes in emus and its implication for sex chromosome evolution. Proc Natl Acad Sci U S A. 2013;110:6453–8.
Liu J, Wang Z, Li J, Xu L, Liu J, Feng S, Guo C, Chen S, Ren Z, Rao J, et al. A new emu genome illuminates the evolution of genome configuration and nuclear architecture of avian chromosomes. Genome Res. 2021;31:497–511.
Clinton M, Zhao D, Nandi S, McBride D. Evidence for avian cell autonomous sex identity (CASI) and implications for the sex-determination process? Chromosome Res. 2012;20:177–90.
Teranishi M, Shimada Y, Hori T, Nakabayashi O, Kikuchi T, Macleod T, Pym R, Sheldon B, Solovei I, Macgregor H, Mizuno S. Transcripts of the MHM region on the chicken Z chromosome accumulate as non-coding RNA in the nucleus of female cells adjacent to the DMRT1 locus. Chromosome Res. 2001;9:147–65.
Roeszler KN, Itman C, Sinclair AH, Smith CA. The long non-coding RNA, MHM, plays a role in chicken embryonic development, including gonadogenesis. Dev Biol. 2012;366:317–26.
Itoh Y, Replogle K, Kim YH, Wade J, Clayton DF, Arnold AP. Sex bias and dosage compensation in the zebra finch versus chicken genomes: general and specialized patterns among birds. Genome Res. 2010;20:512–8.
Backstrom N, Ceplitis H, Berlin S, Ellegren H. Gene conversion drives the evolution of HINTW, an ampliconic gene on the female-specific avian W chromosome. Mol Biol Evol. 2005;22:1992–9.
Rogers TF, Pizzari T, Wright AE. Multi-copy gene family evolution on the avian W chromosome. J Hered. 2021;112:250–9.
Smith CA, Roeszler KN, Sinclair AH. Genetic evidence against a role for W-linked histidine triad nucleotide binding protein (HINTW) in avian sex determination. Int J Dev Biol. 2009;53:59–67.
Ruvinsky I, Sharon N, Lerer T, Cohen H, Stolovich-Rain M, Nir T, Dor Y, Zisman P, Meyuhas O. Ribosomal protein S6 phosphorylation is a determinant of cell size and glucose homeostasis. Genes Dev. 2005;19:2199–211.
Belandia B, Brautigan D, Martin-Perez J. Attenuation of ribosomal protein S6 phosphatase activity in chicken embryo fibroblasts transformed by Rous sarcoma virus. Mol Cell Biol. 1994;14:200–6.
Acharya KD, Veney SL. Sexually dimorphic expression and estradiol mediated up-regulation of a sex-linked ribosomal gene, RPS6, in the zebra finch brain. Dev Neurobiol. 2013;73:599–608.
Mohamed E, Kumar A, Zhang Y, Wang AS, Chen K, Lim Y, Shai A, Taylor JW, Clarke J, Hilz S, et al. PI3K/AKT/mTOR signaling pathway activity in IDH-mutant diffuse glioma and clinical implications. Neuro Oncol. 2022;24:1471–81.
Hamburger V, Hamilton HL. A series of normal stages in the development of the chick embryo. J Morphol. 1951;88:49–92.
Nagai H, Mak SS, Weng W, Nakaya Y, Ladher R, Sheng G. Embryonic development of the emu Dromaius novaehollandiae. Dev Dyn. 2011;240:162–75.
Kouno T, Moody J, Kwon AT, Shibayama Y, Kato S, Huang Y, Bottcher M, Motakis E, Mendez M, Severin J, et al. C1 CAGE detects transcription start sites and enhancer activity at single-cell resolution. Nat Commun. 2019;10:360.
Alam T, Agrawal S, Severin J, Young RS, Andersson R, Arner E, Hasegawa A, Lizio M, Ramilowski JA, Abugessaisa I, et al. Comparative transcriptomics of primary cells in vertebrates. Genome Res. 2020;30:951–61.
Williams RM, Lukoseviciute M, Sauka-Spengler T, Bronner ME. Single-cell atlas of early chick development reveals gradual segregation of neural crest lineage from the neural plate border during neurulation. Elife. 2022;11:e74464.
Rito T, Libby ARG, Demuth M, Briscoe J: Notochord and axial progenitor generation by timely BMP and NODAL inhibition during vertebrate trunk formation. bioRxiv 2023:2023.2002.2027.530267.
Deviatiiarov R, Nagai H, Ismagulov G, Stupina A, Wada K, et al.: Dosage compensation of Z sex chromosome genes in avial fibroblast cells. NCBI Gene Expression Omnibus. https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE213253. 2023.
Severin J, Agrawal S, Ramilowski JA, Deviatiiarov R, Shin JW, Carninci P, de Hoon M. ZENBU-Reports: a graphical web-portal builder for interactive visualization and dissemination of genome-scale data. NAR Genom Bioinform. 2023;5:lqad075.
Williams R, Lukoseviciute M, Sauka-Spengler T, Bronner M: Segregation of neural crest specific lineage trajectories from a heterogeneous neural plate border territory only emerges at neurulation. NCBI Gene Expression Omnibus. https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE181577. 2021.
Rito T, Libby A, Demuth M, Briscoe J: Notochord and axial progenitor generation by timely BMP and NODAL inhibition during vertebrate trunk formation. NCBI Gene Expression Omnibus. https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE223189. 2023.
We would like to thank K. Matsushita from Shimanto Turkey Growers Association in Kochi, Japan, for supplying us with turkey eggs.
The review history is available as Additional file 10.
Peer review information
Kevin Pang was the primary editor of this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.
This work was supported by Japan Science and Technology Agency (JST) e-ASIA joint research project grant JPMJSC19E5 and by a Biotechnology for Bird Conservation grant from Revive & Restore during manuscript revision. RD, AS, and OG were supported by Ministry of Science and Higher Education of the Russian Federation grant 075-15-2021-1344.
Ethics approval and consent to participate
This research does not involve human or adult animal subjects. Avian embryonic fibroblast collection was performed following the biosafety and bioethical regulations of the Kumamoto University.
The authors declare no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Fig. S1. Experimental design and the data analysis pipeline. Fig. S2. Correlation between CAGE replicates. Fig. S3. Correlation between CAGE-seq and RNA-seq methods in birds. Fig. S4. Peak distribution and model efficiency estimation. Fig. S5. Sample MDS plots and hierarchical clustering. Fig. S6. Expression profiles for gametologue genes defined by blast between chrW and chrZ genes. Fig. S7. Genomic views. Fig. S8. Autocorrelation of CAGE expression on chromosome. Fig. S9. Top male-female marker genes in avian fibroblast cells. Fig. S10. RPS6 is a universal male-biased marker. Fig. S11. RPS6 expression in chicken embryonic cells and qPCR validation. Fig. S12. Medium composition effect contribution on sex chromosome gene expression in fibroblast cells. Fig. S13. Cross-species comparison of fibroblast gene expression.
Table S1. Embryonic fibroblast samples statistics.
Table S2. Transcribed promoters identification in birds fibroblasts.
Table S3. Numbers of differentially expressed genes between males and females samples
Table S4. Sex chromosome gene expression contribution to sex differences of fibroblast cultures
Table S5. Dimorphic gene enrichment per chromosome (Fisher exact test for DE genes FDR < 0.05 vs expressed genes)
Table S6. Gametologue genes impact on differential expression estimates between male and female fibroblasts
Table S7. CAGE peaks annotation and differential expression analysis (male vs female)
Table S8. Effect of medium on RPS6 expression. Differential expression analysis for DMEM/10%FBS vs DMEM/F12/10%FBS
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Deviatiiarov, R., Nagai, H., Ismagulov, G. et al. Dosage compensation of Z sex chromosome genes in avian fibroblast cells. Genome Biol 24, 213 (2023). https://doi.org/10.1186/s13059-023-03055-z