General

No statistical methods were used to predetermine the sample size, and the experiments were not randomized. The investigators were not blinded to the allocation during experiments and the outcome assessment.

Ethics statement

Animal care and all experimental procedures were approved by the Animal Experiment Committee, Graduate School of Bioagricultural Sciences, Nagoya University (approval number: 2017030238). Experiments were conducted according to the Regulations on Animal Experiments in Nagoya University.

Animals

To obtain the tissue, genomic DNA, and/or mRNA samples or construct a reference family for genetic linkage analysis, commercial female quails were purchased from a local hatchery (Cyubu-kagaku-shizai, Nagoya, Japan), and quail strains (WE, NIES-L, and NIES-Fr) [16–18] were supplied by the National BioResource Project Chicken/Quail, Nagoya University, Japan. The commercial quail and the NIES-L and NIES-Fr strains exhibited wild-type brown-speckled eggshells, whereas the WE strain exhibited white eggshells. All quails were unvaccinated and maintained individually with free access to water and a commercially available diet. Most quails were maintained under a 14:10 h light/dark photoperiod; however, some were maintained under a 16:8 h light/dark photoperiod for use with quantitative polymerase chain reaction (qPCR) analysis. Room temperature (RT) was maintained at approximately 25°C. Egg-laying quails that were 16–48 weeks old were used for the animal experiments. Notably, a superficial layer of brown pigment is formed on the eggshell at approximately 21–22 h following the last oviposition in Japanese quail [19–21]. Oviducts were collected after decapitation of quail following isoflurane anesthesia. After all experiments, quail were decapitated after isoflurane anesthesia.

mRNA sequencing and differential gene expression analysis

To identify the differentially expressed genes (DEGs) between the wild-type and mutant uteri and detect the genes that are specifically upregulated in the uterus (uterus-specific genes) compared to the other six organs (heart, liver, intestine, kidneys, lungs, and brain) and two tissues (muscle and isthmus), we performed mRNA sequencing of the uteri and isthmuses from the wild-type and mutant quail. For preparing tissue samples, females of both the commercial quail (exhibiting wild-type eggshells) and the WE mutant strain were used (n = 6 for both). To obtain mucosal tissues from the isthmuses and uteri just before the secretion of eggshell pigments, female quails were sacrificed 19–20 h after the last oviposition. Then, the isthmuses and uteri were isolated from the oviducts and opened with scissors with the lumen side up. The tissues were rinsed with phosphate-buffered saline (PBS). Subsequently, ice-cold PBS containing 40 mM dithiothreitol was used as a buffer. Mucosal surfaces were scraped from the uteri and isthmuses by rubbing softly with an interdental brush, and the separated tissues were collected by centrifugation at 400 × g and 4°C. Preliminary histological observation of the uteri and isthmuses after treatment confirmed that the epithelia were mainly separated from the mucosal folds (S3 Fig).

For the extraction of total RNA, 50 mg of the tissue pellet was dissolved in 1 mL of TRI Reagent (Molecular Research Center, Cincinnati, OH, USA) and total RNA was purified using the RNeasy Mini Kit (Qiagen, Hilden, Germany). RNA quality was assessed using Bioanalyzer Pico Chips (Agilent Technologies, Santa Clara, CA, USA). RNA samples with integrity numbers greater than 7.6 were used for constructing libraries. We isolated poly(A)+ RNA from 1 ng of total RNA using the NEBNext Poly(A) mRNA Magnetic Isolation Module (NEB, Ipswich, MA, USA), following the manufacturer’s instructions. We converted oligo(dT)-selected RNA into a cDNA library for mRNA sequencing using the NEBNext Ultra Directional RNA Library Prep Kit for Illumina (NEB), according to the manufacturer’s instructions. The library was sequenced on an Illumina HiSeq 1500 platform by paired-end sequencing (100 bp).

We trimmed the adapter sequences from the reads using Trimmomatic software v0.33 [22], and then mapped the reads to the reference genome (Accession code: GCF_001577835.1) using TopHat2 v2.1.0 [23]. Read counts per gene and fragments per kilobase of transcript per million mapped reads (FPKM) were calculated using Cuffdiff v2.2.1 [24] and Cufflinks v2.2.1 [25]. FPKM values were then converted into transcripts per kilobase million. We obtained the mRNA sequencing data of other organs and tissues of 15-month-old male quails from the Sequence Read Archive (Bio Project number: PRJNA296888), which were obtained by the analysis of six RNA samples for each organ or tissue on an Illumina HiSeq 2500 platform using paired-end sequencing (100 bp). The count data of the other organs and tissues were generated using the same procedure used in this study to obtain the mRNA sequencing data of the uteri and isthmuses of quail. The average number of mapped read pairs is also shown in S1 Data.

We then compared the gene expression profiles between the uterus and other organs and tissues of the wild-type quail. Before detecting DEGs, we excluded genes from the dataset whose counts fell below the threshold (1) in any sample. Then, we performed likelihood ratio tests using the glmLRT function in EdgeR v3.20.9 [26]. In DEG analysis, the full model for count data was compared to the reduced model, in which the object coefficient was set to 0. P-values were adjusted for comparison of the expression levels between the uterus and each organ or tissue of quail using the Benjamini–Hochberg method. Genes whose counts were significantly larger (adjusted p < 0.05, fold change > 2) in the uterus were classified as genes that were specifically upregulated in the uterus. Hierarchical clustering was performed using the heatmap.2 function in the gplots package of R v.3.4.3 using Ward’s method with the Euclidean distance metric.

Construction of a reference family, double-digest restriction site-associated DNA (ddRAD) sequencing, and association test

For genetic linkage mapping of the causative genomic region, we constructed a reference family and obtained genome-wide single nucleotide polymorphism (SNP) markers by ddRAD sequencing, and then performed an association test using these SNP markers and the eggshell color phenotype data. To construct the reference family for genetic linkage analysis, one male of the WE strain and one female of the NIES-L strain that exhibited wild-type eggshells were used as parents. The F₂ hybrids were generated by crossing an F₁ hybrid male with three F₁ hybrid females. Genomic DNA was extracted from the red blood cells of the parents and 99 F₂ hybrid females using the DNeasy Blood & Tissue Kit (Qiagen). The library was constructed according to a previously described method [27]. After digesting 100 ng of DNA from each sample using EcoRI and MseI (NEB), adapters containing barcode sequences were ligated to the ends of the DNA fragments. DNA fragments ranging from 300–500 bp were collected using Pippin Prep (Sage Science, Beverly, MA, USA) and purified using AMPure XP beads (Beckman, Brea, CA, USA). The purified fragments were amplified (six cycles) with PCR primer sets containing index sequences using Phusion High-Fidelity DNA polymerase (NEB). Library quality was validated using 2100 Bioanalyzer with an Agilent High-Sensitivity DNA Kit (Agilent Technologies). The library was sequenced on an Illumina HiSeq 1500 platform by paired-end sequencing (100 bp) at the Functional Genomics Facility, National Institute for Basic Biology, Japan.

Sequence data were demultiplexed using their barcodes, cleaned by removing the low quality reads, and trimmed to 95 or 100 bp using Stacks v1.44 [28]. The processed reads were aligned to the reference sequence using Bowtie 2 v2.2.8 [29]. SNPs were assigned by aligning paired-end reads to the reference sequence using Stacks. SNPs with a depth of coverage of less than 8× were counted as missing data. After eliminating SNPs that deviated from the Hardy–Weinberg equilibrium (p < 0.000001) or had call rates of less than 80%, an association test was performed using Fisher’s exact test implemented in PLINK v1.90 [30]. Sequences of the adapters and primers are shown in S1 Data.

Whole-genome resequencing

To obtain SNP markers and candidate mutations around the locus responsible for the white eggshell phenotype, we performed whole-genome resequencing of the parent quails used to generate F₁ hybrids in genetic linkage analysis, as described above. Genomic DNA was extracted from the red blood cells of the parent quails. Library construction and paired-end sequencing were performed using an Illumina HiSeq X Ten platform (150 bp) at BGI Shenzhen. After trimming the reads using Trimmomatic, the reads were mapped to the reference sequence (Accession code: GCF_001577835.1) using BWA-MEM v0.7.15-r1140 [31]. Subsequently, variants on chromosome 6 were called using Picard v1.119 (http://broadinstitute.github.io/picard/) and the Genome Analysis Toolkit [32], according to a variant call pipeline (https://github.com/gencorefacility/variant-calling-pipeline). We filtered out bases with low Phred-scaled quality scores (less than 30) and reads with low mapping quality (less than 30) using the Genome Analysis Toolkit. Genotypes with individual coverage of less than 10× or more than 70× were filtered out using SnpSift v4.3t [33], and sequence variants that were found in the candidate causative region (candidate region) were annotated using SnpEff v4.3t [33]. To narrow down the candidate region, we searched for candidate genomic intervals for informative SNP markers that could reveal the parental origins of chromosomal regions of the F₂ progeny. To identify causative sequence variants around the candidate region, we extracted sequence variants that were homozygous in the WE strain and then filtered out those found in the NIES-L strain. We searched for structural variants around the candidate region using Integrative Genomics Viewer v2.6.3 [34] and Pindel v0.2.5b9 [35]. Nucleotide sequences were determined using an ABI PRISM 3130 DNA Analyzer (Applied Biosystems, Foster City, CA, USA) after cycle-sequencing reactions using the BigDye Terminator v1.1 Cycle Sequencing Kit (Applied Biosystems). Primer sequences are listed in S1 Table.

qPCR analysis

To compare the gene expression levels of target genes between the wild-type and mutant uteri, qPCR analysis was performed using RNA samples, which were used for mRNA sequencing (n = 5 for both the commercial quail and WE strain). qPCR analysis was also performed using RNA samples extracted from the uterine tissues of female quail that were not used for mRNA sequencing (n = 5 for both the commercial quail and WE strain). Tissue collection and RNA extraction for these additional 10 RNA samples were conducted using the same method as that used for mRNA sequencing; however, quails were maintained under a 16:8 h light/dark photoperiod, and uterine tissues were collected 17 h after oviposition [36]. Total RNA (500 ng) was reverse transcribed into cDNA in a 10 μL reaction mixture using oligo(dT) and the ReverTra Ace qPCR RT Master Mix with a gDNA Remover (Toyobo, Osaka, Japan). PCR amplification (0.1 μL cDNA in a 10 μL reaction mixture) was performed using the Thunderbird SYBR qPCR Mix (Toyobo) and the StepOnePlus Real-Time PCR System (Applied Biosystems, Carlsbad, CA, USA). Melt curve analysis was performed immediately after the amplification. β-actin was used as an endogenous control. Each experiment was performed in triplicate. For all primer sets, the amplification efficiency was 92–112%, and the correlation coefficient was greater than 0.97. The primer sequences and PCR conditions are listed in S1 Table.

Histology

To investigate the accumulation of PPIX in the uterine mucosa of wild-type and mutant quails, uteri were collected from female quails (n = 6 for both the commercial quail and WE strain) 19–20 h after oviposition and fixed overnight in Bouin’s solution. The fixed tissues were dehydrated in a series of graded ethanol, immersed in a 1:1 ethanol/xylene solution, and subsequently embedded in paraffin. The tissues were sectioned to a thickness of 4 μm and mounted onto ovalbumin-coated glass slides. After deparaffinization, the sections were stained with Mayer’s hematoxylin and eosin or left unstained.

Transmission electron microscopy (TEM)

Ultrastructural analyses of the mucosal epithelia of the wild-type and mutant uteri were performed using TEM to investigate the distribution of transport vesicles in apical cells of the uterine mucosa. Uterine samples were collected from female quails (n = 1 for both the commercial quail and WE strain) 19–20 h after oviposition, fixed overnight in 2% glutaraldehyde (in 0.1 M phosphate buffer) at 4°C, rinsed with 0.1 M phosphate buffer overnight at 4°C, and post-fixed in 2% osmium tetroxide (in deionized water [DW]) for 2 h at 4°C. The fixed specimens were dehydrated in a series of graded ethanol (50, 70, 90, 100, 100, and 100%) for 15 min at RT, immersed in propylene oxide for 30 min, followed by immersion in a propylene oxide and epoxy resin mixture for 2 h at RT, and finally embedded in gelatin capsules with epoxy resin for two days at 60°C. Ultrathin sections of 80 nm thickness were cut using an ultramicrotome with diamond knives and mounted on copper grids with a mesh size of 200 μm. The sections were stained with 2% uranyl acetate (in DW) for 15 min and lead staining solution for 5 min at RT. TEM was performed using an H-7600 TEM (Hitachi, Tokyo, Japan) operated at an accelerating voltage of 100 kV.

In situ hybridization

To investigate the distribution of the transcripts of target genes in the uterine mucosa of wild-type and mutant quails, in situ hybridization analyses were performed. For the synthesis of RNA probes, partial sequences of cDNAs of the target genes were obtained by PCR amplification (see S1 Table for primer sequences and S2 Data for partial sequences of cDNAs) and then inserted into the pGEM-T Easy Vector System (Promega, Madison, WI, USA). RNA probes were synthesized in vitro using SP6 RNA polymerase and plasmid DNA digested with SphI (5’-aminolevulinate synthase 1 (ALAS1)) and ApaI (hephaestin-like 1 (HEPHL1) and paired-like homeodomain 3 (PITX3)). Uterine tissues were collected from female quails (n = 2 for both the commercial quail and the WE strain) at 19‒20 h after ovulation, fixed with 4% paraformaldehyde (PFA) overnight, and dehydrated sequentially with 25, 50, and 75% MeOH/PBT (PBS, 0.1% Tween 20), and 100% MeOH for 5 min each. After dehydration, the tissues were immersed in 100% EtOH and 100% xylene for 1 h each. Subsequently, the tissues were embedded in Tissue-Tek Paraffin WaxII60 (Sakura, Tokyo, Japan) and 10 μm sections were prepared using a Leica RM2125 microtome (Leica, Wetzlar, Germany). Sections were dried on glass slides overnight and then hydrated sequentially with 100% xylene, 100% EtOH, 90% EtOH, 70% EtOH, and PBT for 5 min each. After the hydration process, the slides were immersed in 1 μg/mL proteinase K in PBT for 7 min at 37°C. Slides were washed thrice with PBT for 5 min and then fixed with 4% PFA for 20 min. The sections were hybridized with 1 μg/mL of DIG-labeled RNA in the hybridization buffer (50% formamide, 5× saline-sodium citrate [SSC], 1 mg/mL total RNA, 100 μg/mL heparin, 0.1% Tween 20, 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate [CHAPS], and 10 mM ethylenediaminetetraacetic acid [EDTA]) overnight at 68°C. The slides were washed twice with 50% formamide in 2× SSC for 30 min. After cooling, the slides were incubated with the anti-DIG antibody (1:1,000; Roche, Penzberg, Germany) overnight. Then, the slides were washed thrice with Tris-buffered saline with 0.1% Tween 20 for 5 min and the color was developed in AP buffer (100 mM Tris-HCl, 100 mM NaCl, 10 mM MgCl₂, 4.5 μL/mL NBT [Roche], and 3.5 μL/mL BCIP [Roche], pH 9.5).

Gene Ontology (GO) term enrichment analyses

To infer biological processes that are involved in eggshell pigment production and those affected in the mutant uterus, GO term enrichment analyses were performed using the overrepresentation test (released 2021-02-24) of the Protein Analysis Through Evolutionary Relationships (PANTHER) Classification System (annotation version 16.0 released 2020-12-01) [37]. The Gallus gallus database was used as the reference. PANTHER GO-Slim Biological Process was used as the annotation dataset. P-values of Fisher’s exact test were adjusted using the Benjamini–Hochberg method. GO terms were considered significantly enriched if they had a false discovery rate of < 0.05.

Biochemical assay to evaluate the PPIX-forming ability of the uterus

To test whether the insufficiency of 5’-aminolevulinic acid (ALA) or its precursor causes low PPIX production, we compared the PPIX-forming abilities of the uterine homogenates of wild-type and mutant quails in the presence of high concentrations of ALA. Uterine tissues were collected from female quails (n = 1 for both the commercial quail and WE strain) 15–20 h after oviposition and homogenized in ice-cold PBS containing 0.05% Triton-X100. After centrifugation at 20,000 × g and 4°C for 5 min, the supernatants of each homogenate were incubated in the presence of 0, 1, or 2 mM ALA (Wako, Osaka, Japan) in a 96-well plastic plate at 39°C for 20 h (three reactions were performed for each concentration of ALA in 110 μL homogenates). The absorbance was measured at 405 nm using an ARVOx4 2030 Multilabel Reader (PerkinElmer, Waltham, MA, USA) because PPIX exhibits maximum light absorbance at 409 nm. Supernatants incubated without ALA supplementation were used as controls. The total protein concentrations of the supernatants were determined using the Bradford method, and the mean absorbance per 1 mg/mL total protein concentration was calculated for each sample at each ALA concentration.

Statistical analysis

R v.3.4.3 (R Core Team) and MS Excel (Microsoft Corp., Redmond, WA, USA) were used for all statistical analyses. For the comparison of expression levels by qPCR analysis, Welch’s two-tailed t-test was performed using delta Ct values. Welch’s two-tailed t-test was also used to test the differences in the absorbance (at 405 nm) of uterine homogenates between the wild-type and mutant quails. The significance (alpha) level of the statistical tests was set to 0.05.

Histological analysis of the uterine mucosae of the wild-type and mutant quails

We performed gross and histological examination of the uterine mucosae of wild-type and mutant quails, which lay eggs with normal and white eggshells, respectively (Fig 1A; S4 Fig). The uterine mucosa of the wild-type quail exhibited a dark brown color, and that of the mutant quail was pale brown (Fig 1B). Histological examination revealed that brown pigment granules were accumulated in the apical cells of the mucosal epithelium of the uterus in the wild-type quail before anticipated secretion (19–20 h after oviposition), whereas few brown pigment granules were observed in the mutant quail (Fig 1C and 1D). Instead, pink-colored, eosinophilic granules accumulated in the cytoplasm of apical cells in the mutant, suggesting that transport vesicles containing few or no PPIX were formed even in the mutant cells. Electron microscopic observation of the uterine mucosa sections also showed that transport vesicles were accumulated in the apical cells of both the wild type and mutant (S5 Fig).

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Fig 1. White eggshell phenotype and low accumulation of brown pigments in the uterine mucosa of the mutant quail.

(A) Wild-type and mutant white eggs. (B) Wild-type and mutant uteri were opened with the lumen side up. (C) Schematic diagram of the uterine mucosa. Epithelium of the uterine mucosa consists of alternatingly arranged apical and basal cells. Pigment granules containing protoporphyrin IX (PPIX) molecules accumulate in the luminal side of the cytoplasm of apical cells before secretion. Lamina propria is located on the inner side of the uterine mucosa. (D) Sections of uterine mucosae obtained from wild-type and mutant quails just before pigment secretion (left and middle panels, HE staining; right panels, no staining). Ep and LP indicate epithelium and lamina propria, respectively. Brown pigments accumulated in the epithelium of the uterine mucosa of the wild-type quail, but not in that of the mutant quail (left panels). Higher-magnification images of mucosal folds (middle panels) indicate that brown pigment granules accumulated in the wild-type apical cells, but not in the mutant apical cells (black arrowheads); however, pink-colored, eosinophilic granules accumulated in the luminal side of the mutant apical cells. Eosinophilic granules also accumulated more centrally in both the wild-type and mutant apical cells (white arrowheads). Sections without staining clearly demonstrate that brown pigments were accumulated in the luminal side of the apical cells (arrows) of the wild-type quail, but not in the apical cells of the mutant quail. Scale bars: 1 cm in (A, B), 100 μm in (D).

https://doi.org/10.1371/journal.pone.0265008.g001

Characterization of the mutant uterus by gene expression analysis and identification of uterus-specific genes

To characterize the mutant uterus at the gene expression level, we performed mRNA sequencing analysis using the uterine tissues of wild-type and mutant quail and compared gene expression profiles between the wild-type and mutant uteri. The results showed that 261 out of approximately 17,000 expressed genes were differentially expressed (adjusted p < 0.05, fold change < 0.5 or > 2) (Fig 2A; S3 Data). These DEGs consisted of 148 and 113 genes that showed upregulated and downregulated expression, respectively, in the mutant (Fig 2A; S3 Data). We then focused on changes in the expression levels of genes known to be involved in heme and iron metabolism [11] and found that ALAS1 [38], HEPHL1 [39], and transferrin receptor protein 1 (TFRC) [40] were downregulated in the mutant uterus (Fig 2A; S2 Table). The expression levels of these genes in the mutant decreased to 20% (ALAS1), 0.1% (HEPHL1), and 41% (TFRC) of those in the wild type (S2 Table).

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Fig 2. Identification of genes that are specifically upregulated in the wild-type uterus and downregulated in the mutant uterus.

(A) Visualization of the result of differential gene expression analysis using wild-type and mutant uterine tissues. Dots indicate the expression levels of genes. Differentially expressed genes (DEGs) [adjusted p < 0.05, fold change (log₂ scale) > 1 or < −1] are highlighted. Genes that were downregulated in the mutant and specifically upregulated in the wild-type uteri and transferrin receptor protein 1 (TFRC) are labeled. (B) Gene expression levels of the 382 genes whose expression levels were higher in the uterus than in the other six organs and two tissues, visualized by log-transformed transcripts per kilobase million (TPM) values. (C) The number of downregulated genes in the mutant uterus, uterus-specific genes in the wild type, and genes showing both expression patterns. (D, E) Comparison of the expression levels of 5’-aminolevulinate synthase 1 (ALAS1) (D) and hephaestin-like 1 (HEPHL1) (E) in the wild-type uterus with those in other wild-type organs and tissues and the mutant uterus, using mRNA sequencing data. Each bar indicates the relative value of the expression level of the gene in the wild-type uterus when the expression level in each organ or tissue and the mutant uterus is defined as 1. Values in (D, E) are not indicated by the log scale.

https://doi.org/10.1371/journal.pone.0265008.g002

To identify uterus-specific genes, we compared gene expression profiles between the uterus and the other six organs and two tissues. The results showed that 382 genes were specifically upregulated in the uterus (Fig 2B; S4 Data). The expression levels of genes related to heme and iron metabolism in the wild-type uterus were compared to those in other organs and tissues (S3 Table). Eleven of the DEGs, including ALAS1 and HEPHL1, were also included in the uterus-specific genes (Fig 2C; S3 and S4 Tables; S3 and S4 Data). The expression levels of ALAS1 and HEPHL1 were 4–22 times and 171–1,205 times higher, respectively, in the uterus than in other organs and tissues (Fig 2D and 2E; S4 Table). Significant downregulation of ALAS1 and HEPHL1 in the mutant uterus was confirmed by qPCR analysis (Welch’s two-sided t-test, p < 0.01) (S6 Fig; S5–S11 Data). The results of GO term enrichment analyses of the uterus-specific genes and DEGs are shown in S5 and S6 Tables, respectively. GO “biological process” terms, such as extracellular matrix organization and cation homeostasis, were overrepresented in both groups.

Genetic linkage mapping of the responsible locus

The white eggshell phenotype of the mutant quail is inherited in an autosomal recessive manner [14]. However, the causative mutation, which we refer to as white eggshell (we), remains unknown. Therefore, to identify this mutation, we performed chromosomal mapping of the locus by genetic linkage analysis using 99 F₂ progeny obtained by mating three pairs of F₁ progeny derived from crossing one mutant male (the WE strain) with one wild-type female (the NIES-L strain). SNP markers were obtained by ddRAD sequencing of the genomic DNA of the 99 F₂ progeny and their parents. The average number of aligned read pairs and phenotype data are shown in S1 Data. An association test using a total of 15,200 SNP markers revealed a significant association of 235 SNP markers on chromosome 6 (Fisher’s exact test, adjusted p < 0.01) with the white eggshell phenotype (Fig 3A; S12 Data). The marker SNP201288 displayed the highest association (Fig 3A) and logarithm of the odds (LOD) score and the lowest recombination frequency (Fig 3B and 3C; S12 Data); its genotypes in the F₂ progeny were fully concordant with the eggshell color phenotype (Fig 3D). Two SNP markers, SNP201255 (located adjacent to SNP201288) and SNP202246, were separated by a large genomic interval (2,230 kb) (Fig 3E). SNP202246 was located far (2,179 kb) from SNP201288 and displayed discordance between genotypes and phenotypes in eight F₂ progeny (Fig 3D).

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Fig 3. Genetic linkage mapping of the we locus.

(A) Association test using 15,200 single nucleotide polymorphism (SNP) markers. x-axis, genomic coordinates of SNPs; y-axis, negative logarithm of p-values from Fisher’s exact test. Blue line indicates the Bonferroni-corrected p-value of 0.01. SNP201288 displayed the strongest association with the phenotype. (B) Logarithm of the odds (LOD) scores between the we locus and all SNP markers on chromosome 6. SNP201288 and flanking SNPs (SNP201255 and SNP202246) display high LOD scores. (C) LOD scores and recombination frequencies of three SNPs near the we locus. (D) Genotypes of SNPs near the we locus in 99 F₂ individuals. SNP201288 shows a complete concordance between the genotypes and eggshell colors in the F₂ progeny. Discordance between the genotypes and eggshell colors was observed for SNP201255 in F₂ individual #3398 and for SNP202246 in eight F₂ progeny shown by brackets, including the F₂ individual #4040. (E) Genetic and physical maps of SNP markers around the we locus. Genetic distances between the we locus and flanking markers are indicated by parentheses. Positions of SNP202482 and SNP202389 are inverted between the genetic and physical maps, possibly due to an assembly error in the draft genome. (F) Genotypes of Chr6-we1 and Chr-we2 are concordant and discordant, respectively, with the eggshell color, in the F₂ individual #4040. Recombination in the interval between SNP markers is indicated by “×”.

https://doi.org/10.1371/journal.pone.0265008.g003

To narrow down the candidate region, we carried out whole-genome resequencing of the parental quail, which was used for genetic linkage analysis, and obtained sequence variant information on chromosome 6. Sequence data of 33 and 45 GB were obtained for the father (WE strain) and mother quail (NIES-L strain), respectively (S1 Data). The average depth of coverage of chromosome 6 was approximately 16 and 23 for the father and mother quails, respectively (S1 Data). Structural variants, including copy number variation, large deletions or insertions, and inversions, could not be found around the candidate region. Almost all informative SNP markers were detected in the vicinity of SNP201288 and SNP202246 (S13 Data). Among them, the genotypes of Chr6-we1 and Chr6-we2 were concordant and discordant, respectively, with the eggshell color phenotype of the F₂ progeny (#4040) (Fig 3F; S7 Fig; S7 Table). Furthermore, Chr6-we-a, which was detected in a 2,030 kb genomic region between Chr6-we1 and Chr6-we2, displayed a concordance between the genotypes and phenotypes of the eight F₂ progeny (S7 Fig; S7 Table). However, the genotype of Chr6-we-a in the mutant was also observed in the quail strain NIES-Fr, which exhibited wild-type eggshells (S7 Table). Therefore, we eliminated the 2,058 kb region between Chr6-we1 and SNP202246 from the 2,230 kb candidate region, narrowing down the candidate region to the remaining 172 kb region located between SNP201255 and Chr6-we1.

Identification of the causative mutation and causative gene around the we locus

To identify the candidate mutation, we searched for sequence variants (SNPs and insertion–deletion mutations [indels]) that were homozygous in the father quail (mutant) but not found in the mother quail (wild type). The results showed that 957 candidate sequence variants were detected in the 172 kb candidate region (Fig 4A and 4B; S8 Table; S13 Data). The 172 kb candidate region harbored seven protein-coding genes (PITX3, GBF1, TMEM150A, NFKB2, PSD, FBXL15, and CUEDC2) and two non-coding genes (LOC107315847 and LOC107315848) (Fig 4C). Six missense variants were included in the 957 sequence variants (S8 Table; sheets 5 and 6 in S13 Data); however, none were predicted to have adverse effects on the functions of the protein-coding genes, including GBF1, NFKB2, PSD, and CUEDC2 (S9 Table). Furthermore, although it is difficult to precisely estimate the effect of sequence variants, the remaining sequence variants were not considered to be related to functions (sheet 6 in the S13 Data). These results raised the possibility that the mutation affects the expression level of the gene, rather than its function.

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Fig 4. Distribution of sequence variants around the we locus and expression analysis of genes around the 172 kb candidate region.

(A) Number of sequence variants that were homozygous in the father quail (WE strain), but not in the mother quail (NIES-L strain), around the we locus. White arrowheads indicate the locations of the SNP markers. (B) Frequencies of SNPs and insertion–deletion mutations (indels) within the 172 kb candidate region. (C) Eight genes and a part of paired-like homeodomain 3 (PITX3) that are harbored in the 172 kb candidate region (green arrows). (D) Relative expression levels of 15 genes (located around the 172 kb candidate region) in the mutant uteri against those in the wild-type uteri. NA denotes genes whose expression was not detected in the wild-type uteri. Expression levels of six genes (enclosed in squares) substantially decreased in the mutant uteri (less than 60–70% of those in the wild-type uteri). (E) Graphs showing the results of qPCR analysis of PITX3 using RNA samples (n = 5 for both wild-type and mutant quails) that were not used for mRNA sequencing. The expression levels in the wild-type uterus are shown relative to those in the mutant uterus defined as 1. The expression level in each sample is shown by a circle. **p < 0.01.

https://doi.org/10.1371/journal.pone.0265008.g004

Therefore, we predicted that the recessive mutation suppresses the expression of the responsible gene, because mutations that enhance gene expression levels are most likely to be dominant and not recessive. Therefore, we searched for genes on chromosome 6 that were downregulated in the mutant uteri using our mRNA sequencing data. Two genes, PRKG1 and C6H10orf71, were significantly downregulated in the mutant (adjusted p < 0.05, fold change < ½) (S10 Table; sheet 5 in S3 Data). However, they do not seem to be causative genes because both are located far (>~1 Mb) from the 172 kb candidate region. Thus, we focused on the genes located within the 172 kb candidate region and in the region up to 100 kb on either side (nucleotide positions 14.23–14.61 Mb), of which six genes (PIK3AP1, LOC107315928 [2-hydroxyacylsphingosine 1-beta-galactosyltransferase-like], PITX3, TMEM150A, PSD, and CUEDC2), were substantially downregulated in the mutant (their expression was less than approximately 70% of that in the wild type) (Fig 4D). PITX3 encodes a transcription factor involved in the normal development, differentiation, and maintenance of cells in other tissues [41–44]. The other five genes encode proteins that may function in molecular and cellular processes other than transcriptional regulation, such as galactosylceramide biosynthesis and signal transduction [45–49]. Because PITX3 is likely involved in a tissue-specific transcriptional regulation process, we expected that this gene could be a strong candidate. qPCR analysis using mRNA samples that were used for mRNA sequencing also showed non-significant but substantial downregulation of PITX3 in the mutant uterus (S8 Fig; S5 and S14 Data). We observed significant downregulation of PITX3 in the mutant uterus by qPCR analysis using additional RNA samples (Welch’s two-sided t-test, p < 0.01) (Fig 4E; S5 and S15 Data). The difference between the qPCR analyses may be due to the unstable expression of PITX3. However, these results consistently indicate that PITX3 is downregulated in the mutant.

Distribution analysis of PITX3, ALAS1, and HEPHL1 transcripts in the uterine mucosa

Subsequently, to identify tissues that produce PPIX, we focused on two uterus-specific genes, ALAS1 and HEPHL1, and PITX3, and examined the expression patterns of these genes in sections of the uterine mucosa of wild-type and mutant quail. We investigated the distribution of PITX3, ALAS1, and HEPHL1 transcripts in the uterine mucosa of wild-type and mutant quail. These three genes were expressed in apical cells of the mucosal epithelium and cells of the lamina propria (LP), the latter of which are located adjacent to the basal layer of the mucosal epithelium, in the wild type (upper panels of Fig 5). In contrast, hybridization signals were very weak for all three genes in both apical cells and LP cells in the mutant (lower panels of Fig 5). Notably, the hybridization signals of ALAS1 were more intense in the LP cells than in the apical cells of the wild-type quail (Fig 5A). These results raise the possibility that the mucosal epithelium and lamina propria are involved in PPIX production.

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Fig 5. Distribution of PITX3, ALAS1, and HEPHL1 transcripts in uterine mucosal tissues.

(A–C) Light micrographs of the sections of mucosal folds of the wild-type (upper panels) and mutant (lower panels) uteri. PITX, ALAS1, and HEPHL1 were expressed in the apical cells on the luminal side of the epithelia (Ep) of the wild-type uteri, and their expression was observed as bluish hybridization signals (upper panels, black arrowheads). Hybridization signals were also observed in the cells of the lamina propria (LP) (upper panels, white arrowheads). ALAS1 was highly expressed in the cells of LP. The hybridization signals of three genes in the luminal side of the Ep (lower panels, black arrowheads) and LP cells (lower panels, white arrowheads) of the mutant were weak compared to those of the wild type.

https://doi.org/10.1371/journal.pone.0265008.g005

Comparison of the PPIX-forming abilities between the wild-type and mutant uterine homogenates

Finally, to test the causal relationship between low ALAS1 expression and low PPIX production in the mutant uterus, we examined whether the PPIX-forming ability in uterine homogenates of the mutant quail could be recovered by supplementation with a large amount of ALA. The results showed that homogenates from mutant uteri could produce the same amount of PPIX as those from wild-type uteri in both 1 mM and 2 mM concentrations of ALA (Welch’s two-sided t-test, P ≥ 0.05) (Fig 6; S16 Data), indicating that the restricted availability of ALA or its precursors due to downregulated ALAS1 expression is the main cause of the low PPIX production.

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Fig 6. Comparison of the PPIX-forming abilities of the uterine homogenates between the wild-type and mutant quails.

Absorbance of uterine homogenates incubated in the presence of different concentrations of 5’-aminolevulinic acid (ALA). Bars indicate the mean absorbance at 405 nm per 1 mg/mL total protein. Dots indicate the absorbance of each sample. Error bars indicate the standard error of the mean.

https://doi.org/10.1371/journal.pone.0265008.g006