BackgroundFace morphology is strongly determined by genetic factors. However, only a small number of genes related to face morphology have been identified to date. Here, we performed a two-stage genome-wide association study (GWAS) of 85 face morphological traits in 7569 Koreans (5643 in the discovery set and 1926 in the replication set).ResultsIn this study, we analyzed 85 facial traits, including facial angles. After discovery GWAS, 128 single nucleotide polymorphisms (SNPs) showing an association of P < 5 × 10− 6 were selected to determine the replication of the associations, and meta-analysis of discovery GWAS and the replication analysis resulted in five genome-wide significant loci. The OSR1-WDR35 [rs7567283, G allele, beta (se) = −0.536 (0.096), P = 2.75 × 10− 8] locus was associated with the facial frontal contour; the HOXD1-MTX2 [rs970797, A allele, beta (se) = 0.015 (0.003), P = 3.97 × 10− 9] and WDR27 [rs3736712, C allele, beta (se) = 0.293 (0.048), P = 8.44 × 10− 10] loci were associated with eye shape; and the SOX9 [rs2193054, C allele, beta (se) (ln-transformed) = −0.007 (0.001), P = 6.17 × 10− 17] and DHX35 [rs2206437, A allele, beta (se) = −0.283 (0.047), P = 1.61 × 10− 9] loci were associated with nose shape. WDR35 and SOX9 were related to known craniofacial malformations, i.e., cranioectodermal dysplasia 2 and campomelic dysplasia, respectively. In addition, we found three independent association signals in the SOX9 locus, and six known loci for nose size and shape were replicated in this study population. Interestingly, four SNPs within these five face morphology-related loci showed discrepancies in allele frequencies among ethnic groups.ConclusionsWe identified five novel face morphology loci that were associated with facial frontal contour, nose shape, and eye shape. Our findings provide useful genetic information for the determination of face morphology.Electronic supplementary materialThe online version of this article (10.1186/s12864-018-4865-9) contains supplementary material, which is available to authorized users.

Present work investigated glucomannan (GM) and xylan distribution in poplar xylem cells of normal- (NW), opposite- (OW) and tension wood (TW) with immunolocalization methods. GM labeling was mostly detected in the middle- and inner S(2) (+S(3)) layer of NW and OW fibers, while xylan labeling was observed in the whole secondary cell wall. GM labeling in vessels of NW and OW was much weaker than in fibers and mostly detected in the S(2) layer, whereas slightly stronger xylan labeling than fibers was detected in the whole secondary cell wall of vessels. Ray cells in NW and OW showed no GM labeling, but strong xylan labeling. These results indicate that GMs and xylans are spatially distributed in poplar xylem cells with different concentrations present in different cell types. Surprisingly, TW showed significant decrease of GM labeling in the normal secondary cell wall of gelatinous (G) fibers compared to NW and OW, while xylan labeling was almost identical indicating that the GM and xylan synthetic pathways in fibers have different reaction mechanisms against tension stress. Unlike fibers, no notable changes in GM labeling were detected in vessels of TW, suggesting that GM synthesis in vessels may not be affected by tension stress. GM and xylan was also detected in the G-layer with slightly stronger and much weaker labeling than the normal secondary cell wall of G-fibers. Differences in GM and xylan distribution are also discussed for the same functional cells found in hardwoods and softwoods.

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Localization of homogalacturonan (HG) and xyloglucan epitopes in developing and mature pit membranes from different pit types in xylem of Populus tremula L. × P. tremuloides Michx. (hybrid aspen) and Populus tremula L. (European aspen) was investigated using immunogold labeling. Pit types not mediated by ray parenchyma (intervessel- and fiber pits) showed significant developmental changes in HG epitope localization. Both low- and high methyl-esterified HG epitopes (recognized by LM19 and LM20, respectively) were detected in developing pit membranes of intervessel- and fiber pits until late stages of xylem formation, whereas no HG- and high methyl-esterified HG epitopes were detected in mature intervessel (except for annulus regions of pit membranes)- and mature fiber pit membranes, respectively. In contrast, no notable developmental changes in HG epitope localization were detected in pit types mediated by ray parenchyma (vessel-ray-, ray- and fiber-ray pits) during pit maturation. Vesselray- and fiber-ray pits showed abundant low- and high methyl-esterified HG epitopes in pit membranes, while ray pits showed presence of primarily low methyl-esterified HG epitope during all stages of pit development including at maturity. With xyloglucan (recognized by LM15), specific developmental changes in epitope localization were detected in vessel-ray pits. Xyloglucan epitope was detected in developing vessel-ray pit membranes, but was almost absent in mature pit membranes. Instead, xyloglucan was detected in the protective layers of vessel-ray pits showing completely different localization pattern than homogalacturonan, which was only detected in pit membranes. Together, our results suggest that the chemistry of pit membranes varies depending on both the developmental stage and pit type.

We investigated the spatial and temporal distribution of xylans in the cell walls of differentiating earlywood tracheids of Cryptomeria japonica using two different types of monoclonal antibodies (LM10 and LM11) combined with immunomicroscopy. Xylans were first deposited in the corner of the S(1) layer in the early stages of S(1) formation in tracheids. Cell corner middle lamella also showed strong xylan labeling from the early stage of cell wall formation. During secondary cell wall formation, the innermost layer and the boundary between the S(1) and S(2) layers (S(1)/S(2) region) showed weaker labeling than other parts of the cell wall. However, mature tracheids had an almost uniform distribution of xylans throughout the entire cell wall. Xylan localization labeled with LM10 antibody was stronger in the outer S(2) layer than in the inner layer, whereas xylans labeled with LM11 antibody were almost uniformly distributed in the S(2) layer. In addition, the LM10 antibody showed almost no xylan labeling in the S(1)/S(2) region, whereas the LM11 antibody revealed strong xylan labeling in the S(1)/S(2) region. These findings suggest that structurally different types of xylans may be deposited in the tracheid cell wall depending on the developmental stage of, or location in, the cell wall. Our study also indicates that deposition of xylans in the early stages of tracheid cell wall formation may be spatially consistent with the early stage of lignin deposition in the tracheid cell wall.