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Chicken Genome what are the LGE 'chromosomes'?

Chicken Genome what are the LGE 'chromosomes'?



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The chicken genome identifies two "LGE" sequences in the chicken genome. Are these distinct chromosomes or some highly variable sequence from the genome that is put in a separate sequence? I'm thinking they are not really separate chromosomes… It would be great to know a little bit about their biology. the papers didn't seem to help me.


LG stands for "linkage group". It seems the Chicken Genome Sequence group (Hillier et al., 2004) allocated several linkage groups (alleles or genes which tend to be inherited together) to the microchromosomes (tiny chromosomes typical of birds and reptiles), in this case called "linkage group E64" and "linkage group E22… ". There are a load more microchromosomes which are the ones numbered 28-31 and 33-38 which haven't yet had their sequences resolved (Burt, 2007).

References:

  • Burt, D.W. (2007) Emergence of the Chicken as a Model Organism: Implications for Agriculture and Biology. Poultry Science. 86 (7), 1460 -1471. Available from: [Accessed: 8 February 2012].
  • Hillier, L.D.W., Miller, W., Birney, E., Warren, W., Hardison, R.C., Ponting, C.P., Bork, P., Burt, D.W., Groenen, M.A.M. & Delany, M.E. (2004) Sequence and comparative analysis of the chicken genome provide unique perspectives on vertebrate evolution. Nature. 432 (7018), 695-716.

Chicken Genome what are the LGE 'chromosomes'? - Biology

The data should help us understand better our own biology and may give us fresh insight on avian-borne diseases such as salmonella and bird flu.

It could also lead to a step-change in the food industry with the development of more productive and healthier birds.

The International Chicken Sequencing Consortium reports its work in Nature.

"The chicken is the first bird as well as the first agricultural animal to have its genome sequenced and analysed," said Richard Wilson, of the Washington University School of Medicine in St Louis, US, and a lead researcher on the project.

The primary subject for the study was the red jungle fowl ( Gallus gallus ), the wild species from which domestic poultry was bred several thousand years ago.

The consortium's investigation shows the chicken to have approximately one billion base-pairs, or bonded "letters", of DNA. This compares with roughly three billion found in mammals, such as the human.

The analysis reveals that just 2.5% of the human code can be matched to chicken DNA.

It is an important finding. This small portion contains genes that have been largely preserved over the 310 million years since humans and birds shared a common ancestor.

"We believe that the bits of the genome that are most resilient to change are those that have been most crucial to our survival throughout evolutionary history," said Chris Ponting, from Oxford University, UK, who has been comparing the chicken and human data.

"This 2.5% corresponds to 70 million letters of DNA and among these is where we can look first for mutations linked to human disease. In effect, the chicken genome has helped us condense the human genome to something more manageable."

The chicken has long had important roles in science. Developmental biologists have used it to study embryonic growth.

Biomedical researchers have also made important advances in immunology and cancer research by studying chickens. The first tumour virus and cancer gene were identified in chicken research.

All of these areas will be advanced by knowledge of the bird's genome.

The new data may give science an insight into the genetics of resistance, something that would perhaps help researchers develop better vaccines or identify the poultry strains least likely to be susceptible to pathogens.

"What this research does give us is an incredible set of tools to study the genetic variation of these birds," said Ewan Birney, from the European Bioinformatics Institute in Cambridge, UK.

"We know there's a lot of difference between different strains of chicken and different types of birds in the way they transmit these diseases, but we don't know which genes are really involved in helping prevent transmission of, say, the flu virus," he told BBC News.

"With the genome and the genetic tools that that gives us, we'll have a much better platform to do this sort of research in the future."

Other researchers expect there to be big pay-offs for agriculture, too, with the possibility of identifying the underlying biochemical drivers of traits such as bigger eggs and tastier, leaner meat.

On a pure research level, though, there are some real gems in the chicken genome.

These include the realisation that the birds have a keen sense of smell. Scientists can also see genes related specifically to feathers, claws and scales - code sequences that are absent in humans.


Background

The existence of males and females in sexually reproducing organisms and the associated difference in phenotypic optima between sexes imposes an intergenomic conflict in both the evolution of gene sequences and of gene expression. With the exception of the minority of the genome being confined to one sex, as for Y-chromosome sequences, there is thus a trade-off in the evolutionary genetic interests of the two sexes. One way organisms might respond to such sexual antagonism is to evolve sex-biased gene expression, in which the fixation of a sexually antagonistic allele (beneficial in one sex whilst being costly to the other) is followed by the evolution of modifiers to down-regulate gene expression in one sex [1]. It is increasingly recognized, using transcriptome profiling, that a significant proportion of the protein-coding genome has differential expression levels in males and females [2–4]. Many of these genes would be sex-biased in one or a few tissues only [4], so the total number of genes found to be sex-biased typically increases with number of tissues analysed data from Drosophila melanogaster [3] and mice [4] indicate that as much as 50% of all protein-coding genes might be subject to sex-specific regulation of mRNA expression. Moreover, experimental work in Drosophila melanogaster confirms the frequent genomic occurrence of sexually antagonistic alleles and their response to selection [5, 6].

In line with theoretical predictions for the probability of fixation of sexually antagonistic mutations [7], it has been observed that genes with sex-biased expression are non-randomly distributed in the genome. For example, male-biased genes expressed in somatic tissue of nematodes, flies and mammals are underrepresented on the X chromosome, and the same applies to genes expressed post meiosis in germ line [8–11]. In birds, male-biased genes are over-represented on the Z chromosome [12–14]. It has been shown experimentally in Drosophila melanogaster that the X is unusual when it comes to genes conferring sexual antagonism [15].

An obvious alternative explanation for the observation of sex-differential expression of sex-linked genes derives from the fact that gene dose differs between sexes. However, it is well known that organisms have evolved various mechanisms for equilibrating the expression of X-linked genes in males and females (dosage compensation), including X chromosome inactivation in mammals, up-regulation of gene expression on the single X chromosome of Drosophila males and down-regulation of gene expression of both X chromosomes of Caenorhabditis elegans hermaphrodites [16, 17]. With the exception of individual genes that escape dosage compensation [18], sex-linked gene dose should therefore not be expected to lead to overall differences in expression levels between males and females. Intriguingly, in birds the status of dosage compensation is unclear [19, 20]. Early work of sex-linked plumage traits in chicken and other bird species provided no evidence for a compensating mechanism [21], and this was followed by the landmark observation of a double dose of the Z-linked liver enzyme aconitase expressed in males compared to females [19]. Moreover, the absence of sex chromatin and the synchronous replication of the two Z chromosomes in males indicate that there is no Z chromosome inactivation [22, 24, 25]. More recent studies using real-time PCR experiments have added a further dimension to the question because the pattern that emerges is a heterogeneous one, with several examples of genes expressed at similar levels in the two sexes [23, 25, 26]. Importantly, a recent microarray-based study of global gene expression in somatic tissue has indicated that dosage compensation of sex-linked genes in chicken is less effective than is the case in mammals [27]. To study this in some further detail we have taken a genome-wide microarray approach to analyse sex-biased gene expression in both somatic tissue and gonads of chicken. Our data suggest that, overall, dosage compensation does not occur in chicken, meaning that the majority of sex-linked genes is expressed at lower levels in females than in males and that, in females but not in males, the expression levels of sex-linked genes are generally lower than of autosomal genes.


Results

Expression profiling of blastoderms and embryonic day 4.5 gonads reveals at least partial cell autonomous molecular sexual differentiation in chicken

Deep transcriptome sequencing was used to profile gene expression at two developmental time points in males and females 12-h blastoderms (Hamburger and Hamilton stage 1) and day 4.5 embryonic gonads (stage 26) [25]. The rationale for using these times points was our focus on sex determination. Blastoderms represent the earliest accessible post-laying developmental stage, prior to primitive streak formation and gastrulation. This stage was chosen to specifically address the question of cell autonomous molecular sex differentiation pre-dating morphological differentiation. The second tissue, embryonic gonads at day 4.5, represents the time when the gonads are still morphologically identical in each sex ('bipotential').

Sequenced read-pairs were mapped to the chicken genome, (galGal3), using the TopHat 1.3.1 software [26]. The overlap of read-pairs with Ensembl genes was then counted. Differential expression analysis was undertaken by testing the female counts against male counts at both time-points using edgeR [27], with a false discovery rate (FDR) <0.05. Genes known to be expressed sexually dimorphically in E4.5 gonads served as positive controls. For example, DMRT1 and AMH are known to be male upregulated by approximately two-fold in E4.5 gonads, and FOXL2 is expressed only in female gonads at E5.0. Meanwhile, both Aromatase and SOX9 are expressed after E4.5, and were expected to be non-dimorphic in our datasets [28]. These patterns were confirmed in the RNA-seq (see Additional file 1, Figure S1), validating the sequencing results.

Our annotation-based differential expression analysis [29] revealed hundreds of genes differentially expressed between males and females in both tissues (362 in blastoderms and 357 in the gonads) (Figure 1A, and Additional file 2 and 3). This indicated robust sexually dimorphic gene expression pre-dating gonadal development, and supports the notion of cell autonomous sexual differentiation at the molecular level in chicken. In the blastoderm, most of the genes upregulated in males were Z-linked (85%), with a smaller but significant proportion annotated to autosomes (12%) (Figure 1B). This indicated that the Z chromosome is not fully dosage compensated, with the mean expression of Z-linked genes 1.6-fold higher in males compared with females (see Additional file 1 Figure S2), as reported previously [30–32]. Meanwhile genes upregulated in females were annotated to the W chromosome (38%), autosomes (39%), or to the Un_random chromosome (21%) (Figure 1B). The latter represents a virtual chromosome of un-assembled and un-localised chicken sequence. A similar trend was observed in the E4.5 gonads (Figure 1B). Notably, a very small number Z linked sequences were female-biased, in both tissues (Figure 1B). These derived from the MHM locus (Male Hypermethylated), a curious sequence that has previously been reported to be female specific and hypothesised to play a role in localised dosage compensation (upregulation of some neighbouring Z genes in females) [33, 34].

RNA-seq analysis of embryonic chicken blastoderms and embryonic day 4.5 (stage 26) gonads. (A) Differentially expressed annotated genes (Ensembl-based). Number of genes showing either male-biased differential expression (FDR <0.05 blue) or female-biased expression (FDR <0.05 red) in blastoderms and E4.5 gonads. (B) Chromosomal allocation of differentially expressed genes, based on annotated gene data. In blastoderms, female biased genes were located on the W or W_random chromosome (red), the autosomes (grey), or the un-assembled Un_random chromosomes (green). One Z-linked gene showed female-biased expression (aqua). The vast majority of male-biased genes were Z-linked (aqua), and autosomal (grey). Two were on Un_random chromosomes and zero on the W chromosome. A similar pattern was observed in the gonads, with the majority being Z-linked in males, and W-linked or on the Un_random in females. (C) Bar graph illustrating the number of genes with sexually dimorphic expression in at least one tissue on the W, Z, autosomal and Un_random chromosomes. Genes were tested for different patterns of sexually dimorphic expression between tissues, and are grouped as to whether they show a significantly larger female: male ratio difference in the blastoderms (purple), the gonads (yellow) or whether no significant difference is observed (aqua) (FDR <0.05) (See also Additional file 4, Table S1c). (D) Bar graph of genes of genes that are significantly sexually dimorphic in at least one tissue on the W, Z, autosomal and Un_random chromosomes, similar to Fig. 1C). Here, genes are grouped based on a change in average expression between tissues. Shown are the number of genes which are significantly more highly expressed in the blastoderms (purple), higher in the gonads (yellow), and no significant change (blue) (FDR <0.05) (See also Additional file 4, Table S1c).

Some genes involved in sexual differentiation of the gonads begin to be dimorphically expressed between the sexes only in the gonads (for example, FOXL2). These genes therefore showed a different pattern of sexual dimorphism from blastoderm to gonad that is, the female: male ratio was significantly different between tissues. To identify additional genes showing this difference in sexual dimorphism, we used a robust statistical test for the difference in female: male fold change between tissues (see Additional file 1 and 4). Of all of the 43 W-linked Ensembl sequences we detected, none of these genes showed a different pattern of sexual dimorphism (Figure 1C). For Z-genes, of which 262 were differentially expressed in at least one tissue, only three genes (1%) showed a significantly different pattern of sexual dimorphism between tissues (Figure 1C). This included two MHM locus genes that show a large increase in female expression in the gonads, and the Endothelial Tyrosine Kinase (TEK) gene that was more dimorphic in blastoderm (ENSGALG00000018479, ENSGALG00000023324, ENSGALG00000001840). However, 40 autosomal genes (25%) and 22 Un_random genes (45%) showed a different pattern of sexual dimorphism between tissues (Figure 1C). These data indicate that sex-specific molecular pathways that manifest in the blastoderms differ to those in gonads, primarily due to difference in autosomal gene expression. Given that sex chromosome genes did not generally deviate in their female: male ratio between the two tissues, it is interestingly to speculate how these genes can then activate different downstream genes in blastoderms and gonads. In the case of DMRT1, a known testis determinant that is differentially expressed with a similar ratio in both tissues, we found that the average expression level ((male+female)/2) of this gene dramatically increased from blastoderm to gonad. We therefore tested all genes for significant differential expression between tissues, regardless of sex (see Additional files 1 and 4). Indeed, a number of sexually dimorphically expressed W and Z genes also showed differential expression between the tissues, that is, W-genes - 23 (53%), Z-genes - 194 (74%) (Figure 1D and Additional file 4). Those genes upregulated in the gonads (Figure 1D and Additional file 4) are therefore interesting candidate gonadal sexual differentiation genes. Altogether, the data indicate that the relative expression levels of dimorphic sex-linked genes could explain their ability to regulate different downstream genes in different tissues.

To shed light on the molecular pathways that might underlie cell-autonomous sexual differentiation, we first screened the datasets for genes implicated in gonadal sex differentiation. A list was compiled of 117 genes previously linked tovertebrategonadogenesis (see Additional file 5). The set of genes differentially expressed between male and female blastoderms was significantly enriched with these gonadogenesis genes (P = 0.0098, Fisher's exact test), mostly due to non-compensated Z-linked genes (for example, 17βHSDB4 (ENSGAL00000002187), DMRT1 (ENSGAL00000010160) and CFC1B (ENSGAL00000012623)). Only one differentially expressed autosomal 'implicated gonadal gene' was found differentially expressed in this tissue (VNN1, ENSGALG00000013992) (Additional file 2). No genes previously proven to have a role in gonadal sex differentiation were found among the W-linked sequences. Taken together, these data indicate that sex chromosome genes do not activate known sexual differentiation pathways in blastoderms of either sex.

In contrast to the blastoderms, the gonads showed a different set of differentially expressed autosomal genes, many of which have a known link to gonadal sex differentiation, such as FOXL2, Anti-Müllerian Hormone (AMH), INHA (Inhibin-A) and HSP70 (ENSGALG00000011715). This indicates that the sex chromosomes have initiated developmental programs specifically associated with gonadal sex differentiation at embryonic day 4.5 (stage 25), 1 to 2 days prior to the onset of gonadal sexual differentiation (stages 29-30).

These findings show that different sexually dimorphic molecular pathways are engaged by the sex chromosomes in the two different tissues examined here. However, most of the differentially expressed genes detected in blastoderms and gonads had no previous connection to sexual differentiation. To characterise potential pathways activated in female versus male tissues, we assessed gene ontology of all genes that were exclusively differentially expressed in only blastoderms or gonads, using the DAVID programs [35]. The top three clusters of GO terms for each group are shown in (Additional file 1, Figure S10). Given the low number of genes, very few GO terms showed significant enrichment when we corrected for multiple testing (Benjamini), however numerous genes involved in cell stress and DNA damage repair showed sex differential expression in the blastoderm. Various members of the hepatic fibrosis pathway were also differentially expressed (A2M and Col3A1 upregulated in females, and IL1R2 and EGF in males). Sex-specific gene expression in the liver has been previously described, involving >1,000 genes that affect a wide range of biological processes [36]. For genes specifically differentially expressed in the gonads, the top GO terms included neuroactive ligand-receptor interaction (P value = 0.0081). These genes include the GABA receptor alpha 4 and two glutamate receptors. Among the list of autosomal genes expressed in the day 4.5 gonads in a sexually dimorphic fashion were several transcription factors not previously linked to sex per se (four upregulated in females and six in males) (Additional file 2). These data reveal the engagement of different molecular pathways at the two tissues assayed.

As mentioned previously, Z- and W-linked genes generally showed similar dimorphic expression in both blastoderms and gonads (Figure 1C), while many autosomal genes were only differentially expressed in one tissue (Figure 1C). In addition, a significant proportion of genes annotated to the Un_Random chromosome showed a stable dimorphic expression profile (Figure 1C). These unassigned sequences may in fact map to sex chromosomes, especially the poorly annotated W chromosome. Evidence for this conclusion was supported by their sex expression ratios, which were similar to those of the sex chromosome genes (Additional files 2 and 3). As the chicken W sex chromosome and its potential role in avian sex determination is currently poorly understood, we exploited the RNA-seq data to definitively annotate these sequences and to address W-linked gene expression.

Annotation of the W chromosome

Given the robust W-linked gene expression in both blastoderms to E4.5 gonads, we considered that this chromosome might play an important role in sex determination and cell autonomous molecular sex differentiation. However, the chicken W sex chromosome is currently incompletely annotated and its sequence is only partially assembled. In light of the challenges associated with sequencing the W chromosome, we investigated the W transcriptome and its potential role in avian sex determination in more detail. Two approaches were used to construct full-length open reading frames for the W transcriptome, genome-guided and genome independent (de novo) assembly.

Potential mis-annotated W genes were initially identified by their female specific expression (Ensembl) (see Methods and Materials). However, a significant fraction of reads also mapped outside annotated genes. Most notably, 62% of reads that mapped to the Un_random chromosome were not in annotated genes (Additional file 1, Figure S3). We hypothesised that some of these sequences were unannotated W-linked contigs. In order to identify unannotated genes, we extended our analysis of the RNA-seq data using a genome guided transcript assembly procedure, Cufflinks [37]. The Cufflinks analysis was performed by mapping to the chicken genome (Galgal4), and a set of chicken transcripts was created by running Cufflinks 1.3.0 on the mapped reads (see Additional file 1). A significant proportion of expressed W genes identified through the Cufflinks analysis encoded retroviral elements with at least one open reading frame that showed homology to a retroviral protein (with an e-value <0.001, using BLAST). Together with pseudogenes, these sequences were filtered out of subsequent analysis (see Methods and Materials and Supplementary Methods in Additional file 1).

This analysis allowed the compilation of 26 W-linked genes (Table 1). Most of these genes have previously been suggested or confirmed to be W-linked [22], but we identified two novel W-linked sequences, TXN-like and SUB1-W. In addition, two W-linked genes, RPL17-W and HNRPK-W, lie on the Un_random chromosome as described in [22] and confirmed by us. Located within the intronic region of these genes are two annotated small nucleolar RNAs (SNORD58-W-1 and SNORD58-W 2) and one microRNA (mIR-7b-W) that have not been documented to be W-linked. Give the location of the host gene and the presence of a gametologue on the Z chromosome, these genes should be reassigned to the W chromosome.

During both the Ensembl and Cufflinks analyses, it was noted that in at least 12 cases, multiple identified transcripts encoded the same putative protein (Suppl. Table 3). While in some cases, such as HINT-W and FAF, this was due to multiple copies of the gene in the genome, in many cases there was a single copy of the gene, but it had been split across non-contiguous or gapped regions of the genome. Genome-guided assemblies such as Cufflinks are limited by the quality of the genome and transcripts cannot be assembled across segments of the genome which are not correctly scaffolded or which contain gaps in the assembled sequence. This is particularly true for the Un_random and W_random chromosomes, which contain unassembled fragments of the genome. To address this issue we performed a genome independent (de novo) transcript assembly using the ABySS software [38, 39] and used the de-novo assembled transcripts to reassemble different Cufflinks genes together. This enabled genes previously assigned to different regions of a chromosome or even across different chromosomes to be joined together into a single gene (see Additional file 1 for details). Figure 2 shows the results of such analysis for five representative genes, RASA-W, ST8SIA-W, GOLPH-3-W, ZSWIM6-W and NEDD4-W (the remaining assembled W transcripts are given in Figure S4 see Additional file 1). Deduced complete transcripts are shown together with the sequences annotated by Ensembl and those derived from Cufflinks and ABySS assemblies. For example, the RASA-W transcript was assembled by joining seven sequences previously assigned partly to the W and partly to various fragments of the Un_random and W_random chromosome.

Delineation of complete W-linked transcript sequence. Complete transcript sequences for all W-expressed genes were determined using a combined approach of assembling transcripts from Cufflinks, the Abyss de-novo assembly and the latest chicken annotation data (Ensembl). An example of the open reading frame for five W-linked genes is shown. The hatched rectangles represent the different genomic regions to which sequences could be aligned: the W/W_random chromosome (red), Un_random chromosome (green), autosomes (grey) and gaps represent absent genomic sequence. The coloured bars below show the corresponding transcripts defined by Ensembl (aqua), Cufflinks (yellow) and ABySS (blue) analyses. The plots along the top represent the read coverage for the female gonadal sample (black) and the blastoderm sample (grey).

Using this approach, the open reading frames of W-linked mRNAs could be completely assembled, with the exception of one gene, NEDD4-like-W. This resulted in the characterisation of ten genes where previously only part of their ORF was known, due to the poor assembly of the W chromosome. Of these genes, we estimate that, on average, 60% of open reading frame sequence was previously missing from that available in either Ensembl or on Genbank (Additional file 6). Subsequent expression analysis of W genes was carried out by mapping reads to the newly assembled complete cDNA sequences. Final FPKM values (Fragments Per Kilobase of exon per Million fragments mapped) are presented in Table 1.

The complete W-linked transcripts identified in this RNA-seq screen showed gene ontologies with varying functions, none of which are patently associated with sexual differentiation (Table 1 and Table S3 - Additional file 6). However, the lists included genes specifying proteins associated with transcriptional regulation (ZSWIM6-W, ZNF532-W, MIER3-W, BTF3-W, SUB1-W), signalling (SMAD2-W, SMAD7-W, RASA1-W), the ubiquitination pathway (UBAP2-W, UBE2R2-W), the antioxidant thioredoxin (TXN-like-W), the ATP synthase, ATP5A1-W, and the aberrant nucleotide-binding protein, HINTW. As noted above, most W-linked genes did not show significant expression changes between the two tissues examined (Table 1). The most highly expressed genes across both tissues were HINT-W, RPL7-W, ATPA5A1-W, BTF3-W, VCP-like-W and hnRNKP.

Confirmation of female-restricted expression and W-linkage

The RNA-seq data demonstrated that the chicken W sex chromosome harbours more genes than previously thought and that these genes show robust transcriptional activity. All W genes were expressed in both blastoderms and E4.5 gonads. To validate the RNA-seq, quantitative RT-PCR was carried out on four representative genes, using W-gene specific primers. PCR amplification was detected in female but not male blastoderm and gonadal RNA samples (Figure 3A and 3B), thus confirming female specific expression. For some of these genes, whole mount in-situ hybridisation also confirmed female gonad-specific expression (Additional file 1, Figure S5). FISH mapping was used to validate W linkage, showing a single signal in female but not male cells (Figure 3B-E and Additional file 1, Figure S6). In addition, for predicted or novel genes not yet assigned to the W, PCR analysis confirmed that they are present specifically in female but not male chicken genomic DNA (see Materials and Methods and Figure S6 in Additional file 1).

Validation of RNA-seq by quantitative RT-PCR and Confirmation of W-linkage. (A) Blastoderm expression analysis of four representative W genes, KCMF-W, RASA-W, MIER3-W and ZNF532. Expression was detectable in females (red) but not in males (blue). Normalised W gene expression is shown mean +/- SEM n = 3 ** P <0.05. (B-E) FISH mapping of genes identified by RNA-seq to the W sex chromosome in female chicken metaphase spreads. BAC clones were used as probes. (B) BAC clone Ch261-113E6 (ZNR-W, BTF3-W) (red) and BAC Ch261-178N8 (RASA1-W, BTF3-W) (green). (C) BAC clone Ch261-107E4 (HNRPK2-W, GOLPH3-W (red). (D) BAC clone Ch261-60P24 (ZNF532-W, SnoR58-W) (red). (E) BAC clone Ch261-114G22 (UBE2R2-W, RASA1-W, SnoR121A-W) (red). Metaphase chromosomes are stained with DAPI (blue). A single signal was detected in each case and only in female cells, confirming W linkage.

Conservation and relative expression of W-linked genes and their Z gametologues

The complete W-linked transcripts assembled in this study were used to screen for homologues on the Z chromosome (gametologues). All W-linked genes with the exception of FAF (Female Associated Factor) were found to have gametologues on the Z. There was high sequence homology between almost all W- and Z-linked copies at both the DNA level (average of 88.4% identity, Table 1) and the protein level (average of 90.3% identity, Table 1). An exception was the gene HINTW, which showed 41% sequence and 48.5% amino acid homology with its Z gametologue (Table 1). Evolution of novel function among W-linked genes was examined by calculating the rates of synonymous and non-synonymous substitution for each Z-W gene pair [40] and is represented in Table 1, and as a sliding window across each gene pair in Figure S7 (Additional file 1). The dN/dS value for HINTW was the highest of all W genes (0.6) indicating that it has undergone the least amount of purifying selective pressure.

The combined expression of the W and Z gametologues was assessed in both blastoderms and gonads and is shown in Figure 4. For virtually all expressed W-linked genes, the Z gametologue was also expressed, and in both tissues. (The exception was FAF, which lacks a Z gametologue). Total expression from the W and Z gametologues in females was in most cases comparable to the expression of the two Z-linked copies in males, where typically the Z and W contributed equally to the total expression in females (Figure 4A, B). This suggests that most W/Z gametologues in the chicken embryo effectively operate in an autosomal-like fashion (having two functional copies in both male and female). However in some cases, the combined W/Z gene expression in females was significantly higher than the total Z-linked expression in males and was primarily due to W transcription. In the blastoderm, this was the case for HINT, SMAD2 and MIER-3(Figure 4A). In E4.5 gonads, female expression was higher for HINT, MIER3, the putative transcription factor ZSWIM6, VCP-like (Valosin-containing protein) and ST8SIA3 (a sialyltransferase-like gene) (Figure 4B).

Expression levels of W/Z gametologue pairs. Expression of W-linked genes (red) compared to their Z-linked gametologues (blue), for blastoderms (A) and gonads (B). The total combined expression of gametologue pairs is shown for females (red - W/blue - Z, left bar in pair) and males (ZZ -blue only, right bar in pair). The shaded data are shown on an adjusted FPKM scale (inset). Genes with significantly different expression between the sexes are identified (* P <0.01).


Construction of a radiation hybrid map of chicken chromosome 2 and alignment to the chicken draft sequence

Background: The ChickRH6 whole chicken genome radiation hybrid (RH) panel recently produced has already been used to build radiation hybrid maps for several chromosomes, generating comparative maps with the human and mouse genomes and suggesting improvements to the chicken draft sequence assembly. Here we present the construction of a RH map of chicken chromosome 2. Markers from the genetic map were used for alignment to the existing GGA2 (Gallus gallus chromosome 2) linkage group and EST were used to provide valuable comparative mapping information. Finally, all markers from the RH map were localised on the chicken draft sequence assembly to check for eventual discordances.

Results: Eighty eight microsatellite markers, 10 genes and 219 EST were selected from the genetic map or on the basis of available comparative mapping information. Out of these 317 markers, 270 gave reliable amplifications on the radiation hybrid panel and 198 were effectively assigned to GGA2. The final RH map is 2794 cR6000 long and is composed of 86 framework markers distributed in 5 groups. Conservation of synteny was found between GGA2 and eight human chromosomes, with segments of conserved gene order of varying lengths.

Conclusion: We obtained a radiation hybrid map of chicken chromosome 2. Comparison to the human genome indicated that most of the 8 groups of conserved synteny studied underwent internal rearrangements. The alignment of our RH map to the first draft of the chicken genome sequence assembly revealed a good agreement between both sets of data, indicative of a low error rate.


Background

Genomes sizes (as measured by the DNA mass per diploid nucleus) are smaller on average in birds than in other tetrapod classes, and genome sizes within the class Aves show less variation than those of other tetrapod classes [1,2]. It has been proposed that reduced genome size in birds represents an adaptation to the high rate of oxidative metabolism in birds, which results primarily from the demands of flight [1-4]. Cell size and nuclear genome mass are correlated in vertebrates, and cell sizes of birds are smaller than those of mammals [1]. Smaller cells are advantageous in an animal with a high rate of oxidative metabolism because a smaller cell has a greater surface area per volume of cytoplasm, thus facilitating gas exchange.

An alternative to the hypothesis that the reduced genome size is adaptive is the hypothesis that it resulted from an event of genomic DNA loss that was fixed in the ancestor of all birds due to genetic drift. The fixation of even a deleterious mutation is possible if the population undergoes an extreme bottleneck [5]. Some authors have argued that such a bottleneck may have occurred in the ancestor of birds at the end of the Cretaceous period [6], although this conclusion is not consistent with recent molecular evidence placing the radiation of the avian orders well prior to that time [7].

In order to decide whether genome reduction in birds was adaptive or due to a random event, Hughes and Hughes [8] compared the lengths of corresponding introns of orthologous chicken (Gallus gallus) and human (Homo sapiens) genes. They found that corresponding introns were significantly shorter in chickens, indicating that numerous independent deletions have occurred in the introns of birds. These results support the hypothesis that genome size reduction in birds is adaptive, since it is unlikely that such a large number of independent deletion events were due to chance alone. Additional evidence in support of the adaptive hypothesis is provided by the observation that a secondary increase in genome size has occurred in avian lineages which have become flightless or have reduced flying ability [9].

It has been suggested that an important factor in genome size reduction in birds has been that birds have lower levels of repetitive DNA than other vertebrates [10]. Genomes of mammals and reptiles are estimated to consist of about 30�% repeats, while those of birds have been estimated to consist of only 15�% repeats [10-12]. In birds chromosomes are of two types: a minority of macrochomosomes (3𠄶 μm in length) and a larger number of microchromosomes (0.5𠄲.5 μm in length). In the chicken, there are six pairs of macrochromosomes, and thirty-three pairs of microchromosomes [13]. There is a high rate of chiasma formation on avian microchromosomes, and this may be an adaptation that ensures correct pairing of these chromosomes during meiosis and mitosis [14]. Burt [10] proposed that the avoidance of repeats in the avian genome may in turn be an adaptation that enhances the probability of chiasma formation between homologous microchromosomes. This hypothesis and the hypothesis that genome size reduction represents an adaptation to flight are not mutually exclusive, since both factors may be at work simultaneously. Consistent with Burt's hypothesis, Wicker et al. [15] reported that in the chicken genome the ratio of repeats to protein-coding genes is higher on macrochromosomes than on minochromosomes.

The sequencing of a substantial portion of the chicken genome has made it possible to examine quantitatively the distribution of repeating sequences on different chromosomes in the genome. Here we compare the distribution of repeats on 28 sequenced autosomes of chicken with that on the 22 human autosomes in order to test the hypothesis that reduction in repeat density in the avian genome has occurred as a result of adaptive evolution.


Status of the current preliminary genome assemblies

Preliminary assemblies for alligator and crocodile are available. The assembly for alligator additionally uses information from a 120× physical coverage, Illumina 1.5 kbp mate-pair library. The current crocodile assembly was generated with 80× coverage from a 380 bp paired-end Illumina library. The statistics for the length and contiguity of these two assemblies are shown in Table 1. These assembly statistics are on par with other early stage de novo assemblies using short read data [7, 70].

To obtain early estimates of potential TE content, we analyzed the current assemblies using RepeatMasker and a custom repeat library. The library consisted of all vertebrate TEs identified in RepBase [71] and a set of potential TEs identified by applying RepeatScout [72] to both raw 454 data and to the current assemblies (D. Ray, unpublished data). Consistent with earlier studies [59, 73, 74], much of the repetitive content of the genome comprises non-long terminal repeat (non-LTR) retrotransposons from the CR1 family (Figure 3). There is also high content of Chompy-like miniature inverted-repeat transposable elements (MITEs) [75], Penelope retrotransposons, ancient short interspersed repetitive elements (SINEs), and satellite/low complexity regions. Overall, 23.44% of the alligator and 27.22% of the crocodile genome assemblies are annotated as repetitive compared with 50.63% seen in humans. Thus, this preliminary analysis provides further evidence that these reptilian genomes might be easier to assemble than typical mammalian genomes due to their lower repeat content.

The size of different repeat families classified in our current alligator and crocodile assemblies. Despite more long-distance insert libraries for alligator, more repeats were found in the crocodile assembly. This strongly suggests that crocodiles have more repeats than do alligators, and perhaps the difference will become even more striking as the crocodile assembly improves.

We also examined GC content across the assemblies (Figure 4). Alligators and crocodiles appear to have a higher mean GC content than many other vertebrates. Additionally their large standard deviation in GC content across contigs is similar to that of birds and mammals, suggesting that their base composition is heterogeneous and likely contains GC-rich isochores. This is unlike the situation in the lizard (Anolis) and frog (Xenopus), which lack strong isochores based upon analyses of genomic data [76], or the turtle Trachemys scripta, which appears to lack strong isochores based upon analyses of expressed genes [77]. However, these results are consistent with previous analyses of ESTs that suggested the existence of GC-rich isochores in the alligator genome [62, 77]. Thus, these crocodilian genome data extend the results of the previous analyses and confirm the genome-wide nature of GC-content heterogeneity in crocodilian. We expect improved crocodilian genome assemblies to further illuminate the details of isochore structure in reptiles.

The distribution of GC proportion across several species. Note that alligators and crocodiles have a higher overall proportion of GC than many other vertebrates, as predicted by early BAC-end scans [42]. Abbreviation: SD standard deviation.


Reviewers' comments

Reviewer's report 1

Igor Zhulin, University of Tennessee and Oak Ridge National Laboratory

This is an interesting discovery of novel viral families in the chicken genome, which accounts for more than 2% of the genome sequence. I do not have any major concerns regarding this paper and support its publication however, I would like to offer some comments for authors' consideration, mainly regarding the clarity and presentation.

Authors' response

We are grateful to the reviewer Dr. Igor Zhulin for providing a number of very specific and constructive comments regarding the clarity and presentation of the manuscript. We revised the paper according to his suggestions.

Reviewer's report 2

Itai Yanai, Harvard University

I support publication of this manuscript.


Open Research

This Whole Genome Shotgun project has been deposited at DDBJ/ENA/GenBank under the accession WUCP00000000. The version described in this paper is version WUCP01000000. Raw read sequences generated in the de novo sequencing have been deposited in the Sequence Read Archive (SRA) at NCBI under the project access ion PRJNA380312. Published genome data used in the analyses can be found under the following accession codes: G. gallus (GRCg6a [ftp://ftp.ensembl.org/pub/release-96/fasta/gallus_gallus/dna/]) M. gallopavo (UMD2 [ftp://ftp.ensembl.org/pub/release-96/fasta/Meleagris_gallopavo/dna/]). More dataset, as well as the pipelines and scripts, can be found in figshare https://figshare.com/projects/Phasianus_colchicus_genome_sequencing_and_assembly/88112.

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Turkey Genetics 101

I love watching the turkeys on Martha&rsquos Vineyard. They travel in small family groups of two parents with chicks and adolescents, coalescing into larger tribes.

When it rains, wild turkeys go about their business, pecking at food &ndash I&rsquove yet to see one raise it&rsquos mouth and drown. And they have feelings. My daughter and I once watched as 4 turkeys stood around a comrade who&rsquod just been run over, clearly distraught. None left, even as cars went by.

I saw a family of 7 perched on a branch in size order, crapping in unison. And one early morning I turned a corner and faced a lawn of turkeys. I froze and counted them &ndash 42 &ndash a little afraid, because the hens are quite enormous. They turned and stared as I quietly backed away.

Like humans, a few individuals can become deranged. On Father&rsquos Day in 2008, for example, a &rdquohostile, feral turkey&rdquo chased two cops onto their patrol car. A man who had raised the unfortunate Tom tried to help his pet when the officers shot and killed the terrorist turkey. Later, neighbors and a UPS delivery person corroborated the police view that Tom was indeed violent, and there was much concern that the &ldquosociopath with feathers&rdquo had transmitted his errant behavioral genes to the next generation.

For the most part, the turkeys of Martha&rsquos Vineyard are oblivious to gaping humans and the occasional yapping canine. Maybe that&rsquos because the island&rsquos somewhat limited biodiversity offers no natural predators, just a human population that surges in the summer with many folk who aren&rsquot used to sharing space with large, wild birds.

I haven&rsquot checked the DNA of the turkeys of Martha&rsquos Vineyard, but I&rsquod bet their immune systems are a lot tougher than those of the barnyard variety. Instead of the ginormous &ldquobreasts&rdquo (why do we speak of breasts in birds, who do not lactate?) that cause the broad-breasted &ldquoindustrial&rdquo white turkey to topple over, these wild turkeys have more dark meat (more myoglobin), which enables them to suddenly soar up into the treetops at a clap of thunder, and to have flown to the island in the first place, reaching speeds, according to PETA, of 55 mph.

Having followed the turkeys of Martha&rsquos Vineyard for years, and not particularly liking to eat their brethren, I thought I&rsquod compile a listicle of a dozen facts about the genetics of Meleagris gallopavo.

1. The domesticated turkey originated in Mexico in about 800 BC. Their races, like races in humans, are defined by a superficial characteristic &ndash plumage color &ndash rather than full gene-based ancestry. Only a half dozen or so of the turkey&rsquos 16,000 genes contribute to plumage color, just like a handful of our 20,000 genes impart skin color. All commercial lines descend from one ancestral group, although the American Poultry Association recognizes 7 breeds.

2. Like human chromosomes, turkey chromosomes were initially described rather vaguely. A 1931 paper describes 4 big J-shaped chromosomes, 2 big rods, 3 short rods, and 29 globes. Today we know the genome is splayed out over 39 pairs of autosomes. Like all birds, males have two Z chromosomes and females one Z and one W, somewhat the opposite of humans, in whom females are the homogametic sex (XX).

3. The turkey genome project got underway at Virginia Tech in 2008, and the sequence of a hen named Nici was published in 2010. Nici means &ldquoNicholas inbred,&rdquo after famed turkey farmer George Nicholas who, at Nicholas Turkey Breeding Farms in Sonoma, California, turned the wild bird into today&rsquos barely recognizable top-heavy product of extreme artificial selection. Conventional agriculture fosters far more profound change than the one-gene-at-a-time tweaking of GMOs.

4. Nici was the spawn of 9 generations of full-sib matings. The brother-sister matings left just enough heterozygosity to provide interesting gene variants, but not enough diversity to gum up the sequencers and assemblers. Having the turkey genome sequence will enable breeders to select traits based on genotype rather than phenotype, which can theoretically help to preserve some of the valuable traits hidden in the recessive state.

5. The turkey genome is 1.1 billion bases. Unlike the rush to sequence the human genome, with the turkey interest is more on annotation &ndash figuring out what genes do and how that can make farmers money &ndash than in accumulating sequences from different individuals.

6. The turkey was the fifth farm animal to have its genome done, following the pig, cow, sheep, and chicken. The duck was done in 2013.

7. Not surprisingly, the turkey genome is similar to that of the chicken, differing most obviously by some two dozen chromosomal inversions. The turkey genome sequence is being used to fill in some gaps in the chicken genome sequence, although at 1.8 SNPs per kilobase, turkeys have a less diverse genomes than do chickens, which have 5.5. The reason: the ancestral chicken population was much larger than the ancestral turkey population. The turkey genome has five regions of exceptional genetic uniformity, and the mitochondrial genome is also much less diverse.

8. An autosomal recessive condition of turkeys is red blood cells that have two nuclei.

9. Turkeys have superb vision. With five types of the visual pigment rhodopsin, 7 types of photoreceptors, and 4 types of cones, they can see into the ultraviolet. We can&rsquot. They also have a type of T cell receptor apparently unique to them.

10. Industrial turkeys are more likely to suffer from non-inherited diseases than inherited ones. The list is long. Being smashed into close quarters can trigger feather picking (auto and allo), stampeding, and a hardness of the underfoot called bumblefoot. Poor nutrition can soften bones and enlarge hocks. Poisons include milkweed and aflatoxin.

Turkeys can give us Newcastle disease (viral respiratory), Chlamydia psittaci (bacterial respiratory), tuberculosis, and typhoid. They also get a host of fungal conditions (thrush, brain, skin, lungs), protozoan woes (blackhead), more viral (flu, bluecomb, tumors) and bacterial infections (erysipelas, cholera, pox, and a malodorous infection of the navel), gross worms (redworm, flukes, round worms, tapeworms) and of course lice, ticks, and mites. Highly selective breeding teamed with overuse of antibiotics has pummeled their immune systems. Industrialized turkeys are particularly susceptible to aflatoxin poisoning from fungus growing on feed corn, which causes liver cancer in humans. A glutathione s-transferase gene variant that detoxes aflatoxin, found in wild turkeys, has been bred out of their domesticated relatives.

11. Intense selection for a huge white breast and ultra-accelerated growth, reaching &ldquomarket weight&rdquo in 131 days compared to 185 among my avian friends on Martha&rsquos Vineyard, kills heart and skeletal muscle cells, collapses legs, deforms skeletons, and decimates immunity.

12. Five natural &ldquoheritage&rdquo varieties of turkeys had more red blood cells, proteins, T cell responses, and disease resistances compared to industrial birds. Heritage turkeys also make more vitamin C, which birds synthesize and therefore don&rsquot need to consume, and the vitamin enhances immunity. The heritage birds enjoy &ldquolower mortality rate, the ability to mate naturally, excellent hatchability, active foraging, intelligence, and overall attractiveness. The only parameters on which the industrial strains excel are feed conversion and rate of gain,&rdquo according to one review.

Meleagris gallopavo is a beautiful animal. No wonder Ben Franklin wanted the turkey to be the national bird.


Watch the video: Χρωμοσώματα και χρωματίνη (August 2022).