Transcriptome-wide association study and eQTL colocalization identify potentially causal genes responsible for human bone mineral density GWAS associations

  1. Basel Maher Al-Barghouthi
  2. Will T Rosenow
  3. Kang-Ping Du
  4. Jinho Heo
  5. Robert Maynard
  6. Larry Mesner
  7. Gina Calabrese
  8. Aaron Nakasone
  9. Bhavya Senwar
  10. Louis Gerstenfeld
  11. James Larner
  12. Virginia Ferguson
  13. Cheryl Ackert-Bicknell
  14. Elise Morgan
  15. David Brautigan
  16. Charles R Farber  Is a corresponding author
  1. Center for Public Health Genomics, School of Medicine, University of Virginia, United States
  2. Department of Biochemistry and Molecular Genetics, School of Medicine, University of Virginia, United States
  3. Department of Radiation Oncology, University of Virginia, United States
  4. Department of Microbiology, Immunology, and Cancer Biology, School of Medicine, University of Virginia, United States
  5. Department of Orthopedics, Anschutz Medical Campus, University of Colorado, United States
  6. Department of Public Health Sciences, School of Medicine, University of Virginia, United States
  7. Department of Mechanical Engineering, Boston University, United States
  8. Department of Mechanical Engineering, University of Colorado Boulder, United States
  9. Department of Orthopaedic Surgery, Boston University Medical Center, United States

Abstract

Genome-wide association studies (GWASs) for bone mineral density (BMD) in humans have identified over 1100 associations to date. However, identifying causal genes implicated by such studies has been challenging. Recent advances in the development of transcriptome reference datasets and computational approaches such as transcriptome-wide association studies (TWASs) and expression quantitative trait loci (eQTL) colocalization have proven to be informative in identifying putatively causal genes underlying GWAS associations. Here, we used TWAS/eQTL colocalization in conjunction with transcriptomic data from the Genotype-Tissue Expression (GTEx) project to identify potentially causal genes for the largest BMD GWAS performed to date. Using this approach, we identified 512 genes as significant using both TWAS and eQTL colocalization. This set of genes was enriched for regulators of BMD and members of bone relevant biological processes. To investigate the significance of our findings, we selected PPP6R3, the gene with the strongest support from our analysis which was not previously implicated in the regulation of BMD, for further investigation. We observed that Ppp6r3 deletion in mice decreased BMD. In this work, we provide an updated resource of putatively causal BMD genes and demonstrate that PPP6R3 is a putatively causal BMD GWAS gene. These data increase our understanding of the genetics of BMD and provide further evidence for the utility of combined TWAS/colocalization approaches in untangling the genetics of complex traits.

Editor's evaluation

Many GWAS studies have been done to understand the genetic contributions to bone density, but very few have managed to pinpoint the gene affected by a polymorphism that caused an observed difference. In this paper, your team shows how scientists can identify causative variants from GWAS studies.

https://doi.org/10.7554/eLife.77285.sa0

Introduction

Osteoporosis, a disease characterized by low bone mineral density (BMD), decreased bone strength, and an increased risk of fracture, affects over 10 million individuals in the United States (Black and Rosen, 2016; Burge et al., 2007). BMD is the single strongest predictor of fracture and a highly heritable quantitative trait (Miller et al., 1999; Ralston and Uitterlinden, 2010; Peacock et al., 2002). Over the last decade, genome-wide association studies (GWASs) have identified over 1100 independent associations for BMD (Morris et al., 2019; Estrada et al., 2012; Kemp et al., 2017). However, despite the success of GWAS, few of the underlying causal genes have been identified (Rocha-Braz and Ferraz-de-Souza, 2016; Sabik and Farber, 2017).

One of the main difficulties in GWAS gene discovery is that the vast majority (>90%) of associations are driven by non-coding variation (Giral et al., 2018; Edwards et al., 2013). Over the last decade, approaches such as transcriptome-wide association studies (TWASs) and expression quantitative trait locus (eQTL) colocalization have been developed which leverage transcriptomic data in order to inform gene discovery by connecting non-coding disease-associated variants to changes in transcript levels (Gusev et al., 2016; Barbeira et al., 2018; Pividori et al., 2020; Giambartolomei et al., 2014; Wen et al., 2017). These approaches have proven successful for a wide array of diseases and disease-associated quantitative traits (Pividori et al., 2020; Bhattacharya et al., 2020; Thom and Voight, 2020). However, the osteoporosis field has lagged behind such efforts, due to the limited number of large-scale bone-related transcriptomic datasets.

In a TWAS, genetic predictors of gene expression (e.g., local eQTL – sets of genetic variants that influence the expression of a gene in close proximity, Nica and Dermitzakis, 2013) identified in a reference population (e.g., the Genotype-Tissue Expression [GTEx] project, Consortium, 2013) are used to impute gene expression in a GWAS cohort. Components of gene expression due to genetic variation are then associated with a disease or disease-associated quantitative trait. Genes identified by TWAS are often located in GWAS associations, suggesting that the genetic regulation of their expression is the mechanism underlying such associations. Several tools, for example, FUSION, PrediXcan, and MultiXcan (Gusev et al., 2016; Gamazon et al., 2015; Barbeira et al., 2019) have been developed to perform TWASs. Most of these tools use GWAS summary statistics, making TWAS widely applicable to large GWAS datasets. In contrast, eQTL colocalization is a statistical approach that determines if there is a shared genetic basis for two associations (e.g., a local eQTL and BMD GWAS locus). Recently, it has been demonstrated that prioritizing genes using both TWAS and eQTL colocalization provides a way to identify genes with the strongest support for causality (Barbeira et al., 2018; Pividori et al., 2020).

The GTEx project has generated RNA-seq data on over 50 tissues across hundreds of individuals (Consortium, 2020). Even though data on the tissues/cell types likely to be most relevant to BMD (bone or bone cells) were not included, the project demonstrated that many eQTL were shared across tissues (Consortium, 2020; Battle et al., 2017). Additionally, it is well known that effects in a wide-array of non-bone cell types and tissues can impact bone and BMD (Fitzpatrick, 2002; Mirza and Canalis, 2015). As a result, we sought to use the GTEx resource in conjunction with TWAS and eQTL colocalization to leverage non-bone gene expression data to identify putatively causal genes underlying BMD GWAS.

Here, we performed TWAS and eQTL colocalization using the GTEx resource and the largest BMD GWAS performed to date to identify potentially causal genes (Morris et al., 2019). Using this approach, we identified 512 genes significantly associated via TWAS with a significant colocalizing eQTL. To investigate the significance of our findings we selected Protein Phosphatase 6 Regulatory Subunit 3 (PPP6R3), the gene with the strongest support not previously implicated in the regulation of BMD, for further investigation. We demonstrate using mutant mice that Ppp6r3 is a regulator of lumbar spine BMD. These results highlight the power of leveraging GTEx data, even in the absence of data from the most relevant tissue/cell types, to increase our understanding of the genetic architecture of BMD.

Results

TWAS and eQTL colocalization identify potentially causal BMD GWAS genes

To identify potentially causal genes responsible for BMD GWAS associations, we combined TWAS and eQTL colocalization using GTEx data (Figure 1A). We began by performing a TWAS using reference gene expression predictions from GTEx (Version 8; 49 tissues) and the largest GWAS performed to date for heel estimated BMD (eBMD) (>1100 independent associations) (Morris et al., 2019; Consortium, 2020). The analysis was performed using S-MultiXcan, which allowed us to leverage information across all 49 GTEx tissues (Barbeira et al., 2019). Our analysis focused on protein-coding genes (excluded non-coding genes). A total of 2156 protein-coding genes were significantly (Bonferroni-adjusted p value ≤0.05) associated with eBMD (Supplementary file 1a).

Figure 1 with 1 supplement see all
Transcriptome-wide association study (TWAS) and expression quantitative trait loci (eQTL) colocalization identify potentially causal bone mineral density (BMD) genome-wide association study (GWAS) genes.

(A) Overview of the analysis. The human image was obtained from BioRender.com. TWAS/colocalization plot for genes in the locus around RUNX2 (B) and RERE (C). The −log10 Bonferroni-adjusted p values from the TWAS analysis (top panel) and the maximum regional colocalization probabilities (RCPs) from the colocalization analyses (bottom panel). Genes alternate in color for visual clarity. Triangles represent RUNX2 (B) and RERE (C). (D) Distribution of prioritized genes within estimated BMD (eBMD) GWAS loci.

Next, we identified colocalizing eQTL from each of the 49 tissues in GTEx using fastENLOC (Pividori et al., 2020; Wen et al., 2017). We identified 1182 colocalizing protein-coding genes with a regional colocalization probability (RCP) of 0.1 or greater (Supplementary file 1b). In total, 512 protein-coding genes were significant in both the TWAS and eQTL colocalization analyses (Table 1 and Supplementary file 1c). Among the identified genes were many with well-known roles in the regulation of BMD, such as RUNX2 (Figure 1B), IGF1, and LRP6, as well as novel genes such as RERE (Figure 1C). Overall, the identified genes had significantly colocalizing eQTL in all 49 GTEx tissues, with eQTL from cultured fibroblasts (132 genes), subcutaneous adipose tissue (117 genes), tibial artery (115 genes), and tibial nerve (114 genes) exhibiting the highest number of significant colocalizations (Supplementary file 1d). TWAS predictors were only generated for genes on autosomes and of the 1103 independent associations identified by Morris et al., 2019, 1097 were autosomal. For each of these, we defined a locus as the region consisting of ±1 Mbp around each association. Of the 1097 loci, almost half (542; 49%) of the loci contained at least one of the 512 prioritized genes. Most loci overlapped one gene (mean = 1.7, median = 1); however, 184 loci overlapped multiple genes, including a locus on Chromosome (Chr.) 20 (lead SNP rs6142137) which contained 9 prioritized genes (Figure 1D and Figure 1—figure supplement 1).

Table 1
Top 10 protein-coding genes significant by colocalization (RCP ≥0.1) and TWAS, sorted by TWAS p value.
GeneTissue with greatest RCPMax. RCPTWAS p value (Bonferroni)
SPTBN1Cells_Cultured_Fibroblasts0.9469<5 × 10−324
CCDC170Spleen0.6582<5 × 10−324
FAM3CArtery_Tibial0.4917<5 × 10−324
SEPT5Skin_Sun_Exposed0.48682.26 × 10−286
FGFRL1Cells_Cultured_Fibroblasts0.16115.31 × 10−272
GREM2Cells_Cultured_Fibroblasts0.99984.31 × 10−257
GPATCH1Whole_Blood0.35643.44 × 10−226
RHPN2Pituitary0.21818.71 × 10−221
BMP4Brain_Cortex0.54685.49 × 10−169
RUNX2Esophagus_Gastroesophageal_Junction0.23721.99 × 10−146

Characterization of genes identified by TWAS/eQTL colocalization

To evaluate the ability of the combined TWAS/colocalization approach to identify genes previously implicated in the regulation of BMD, other bone traits, or the activity of bone cells, we queried the presence of ‘known bone genes’ within the list of the 512 prioritized protein-coding genes. To do so, we created a database-curated set of genes previously implicated in the regulation of bone processes (henceforth referred to as our ‘known bone genes’ list, N = 1399, Supplementary file 1e). Of the 512 genes identified above, 66 (12.9%) were known bone genes, representing a significant enrichment (odds ratio [OR] = 1.72; p = 1.0 × 10−4) over what would be expected by chance (Supplementary file 1f).

We also performed a Gene Ontology (GO) enrichment analysis of the 512 prioritized genes. We observed enrichments in several bone-relevant ontologies, such as ‘ossification’ (p = 3.1 × 10−6), ‘skeletal system development’ (p = 2 × 10−5), and ‘regulation of osteoblast differentiation’ (p = 3.9 × 10−5) (Figure 2A and Supplementary file 1g).

Figure 2 with 1 supplement see all
Transcriptome-wide association study (TWAS) and expression quantitative trait loci (eQTL) colocalization identify Gpatch1 a potentially causal bone mineral density (BMD) genome-wide association study (GWAS) gene.

(A) The top 40 terms from a Gene Ontology analysis of the 512 potentially causal BMD genes identified by our analysis. Terms with clear relevance to bone are highlighted in red. Only terms from the ‘Biological Process’ (BP) subontology are listed, and similar terms were removed for clarity. (B) TWAS/colocalization plot for genes in the locus around GPATCH1 (±1.5 Mbp). The −log10 Bonferroni-adjusted p values from the TWAS analysis (top panel) and the maximum regional colocalization probabilities (RCPs) from the colocalization analyses (bottom panel). Genes alternate in color for visual clarity. Triangles represent GPATCH1. (C) Mirrorplot showing estimated BMD (eBMD) GWAS locus (top panel) and colocalizing GPATCH1 eQTL in whole blood (bottom panel). SNPs are colored by their linkage disequilibrium (LD) with rs11881367 (purple), the most significant GWAS SNP in the locus. (D) Scatterplot of −log10 p values for GPATCH1 eQTL versus eBMD GWAS SNPs. SNPs are colored by their LD with rs11881367 (purple). (E) BMD in Gpatch1 knockdown mice. N = 7 females and N = 4 males for Gpatch1+/− mice, N = 880 females and N = 906 males for Gpatch1+/+ mice. Boxplots indicate the median (middle line), the 25th and 75th percentiles (box) and the whiskers extend to 1.5 * IQR.

To compare our approach with the approach of prioritizing the closest genes to GWAS associations as potentially causal, we quantified the number of genes that were both the closest genes to eBMD GWAS associations and were members of the ‘known bone gene’ list. Of the 863 genes that were the closest genes to eBMD GWAS associations (Supplementary file 1h), 139 were members of the ‘known bone gene list’, representing a more significant enrichment of ‘known bone genes’ than our prioritization approach (OR = 2.56, p = 6.37 × 10−19). Of our 512 prioritized genes, 206 (40%) were also the closest genes to eBMD GWAS associations, with 27 of the remaining 306 prioritized genes (8.8%) being members of the ‘known bone gene’ list.

The International Mouse Phenotype Consortium (IMPC) has recently measured whole body BMD in hundreds of mouse knockouts (Dickinson et al., 2016; Swan et al., 2020). We searched the IMPC database for any of the 512 genes identified by TWAS and eQTL colocalization. Of the 512, 142 (27.7%) had been tested by the IMPC and 64 (12.5% of the 512 prioritized genes, 45% of the 142 IMPC-tested genes) had a nominally significant (p ≤ 0.05) alteration of whole-body BMD in knockout/knockdown mice, compared to controls. Of the 64, 49 (76.5%) were not members of the ‘known bone gene’ list (Figure 2—figure supplement 1A, B).

An example of one of the 64 genes is GPATCH1, located within a GWAS association on human chromosome 19q13.11. GPATCH1 is part of the catalytic step 2 spliceosome, and may be involved in mRNA splicing, and is predicted to enable RNA-binding activity. It is also expressed in bone cells in mouse (Alliance of genome resources, 2022; Lattin et al., 2008). Of all the genes in the region, GPATCH1 had the strongest TWAS association (p = 3.44 × 10−226) (Figure 2B) and the strongest eQTL colocalization (whole blood, RCP = 0.36) (Figure 2B–D). The eQTL and BMD GWAS allele effects for the top SNPs were in the same direction, suggesting that decreasing the expression of GPATCH1 would lead to decreased BMD. BMD data from the IMPC showed that female mice heterozygous for a Gpatch1 null allele had decreased BMD (p = 2.17 × 10−8) (Figure 2E). Together, these data suggest that many of the genes identified by the combined TWAS/colocalization approach are likely causal BMD GWAS genes.

PPP6R3 is a candidate causal gene for a GWAS association on Chr. 11

To identify novel candidate genes for functional validation, we focused on genes with the strongest evidence of being causal. To do so, we increased the colocalization RCP threshold to 0.5, and then sorted genes based on TWAS Bonferroni-adjusted p values. Furthermore, we constrained the list of candidates for functional validation to genes which were not members of the ‘known bone gene’ list or genes with a nominal (p ≤ 0.05) alteration in whole-body BMD as determined by the IMPC. This yielded 137 putatively causal BMD genes (Table 2, Supplementary file 1i, and Figure 2—figure supplement 1C).

Table 2
Top 10 novel protein-coding genes significant by colocalization (RCP ≥0.5) and TWAS, sorted by TWAS p value.
GeneTissue with greatest RCPMax. RCPTWAS p value (Bonferroni)# GTEx tissues with RCP ≥0.5
SPTBN1Cells_Cultured_fibroblasts0.9469<5 × 10−3242
PPP6R3Thyroid0.52915.7 × 10−931
BARX1Colon_Transverse0.77646.36 × 10−631
MEOX2Brain_Nucleus_accumbens_basal_ganglia0.62863.21 × 10−533
REREAdipose_Subcutaneous0.64316.95 × 10−464
SIPA1Nerve_Tibial0.99814.26 × 10−411
CAPZBTestis0.67163.64 × 10−331
B4GALNT3Artery_Aorta0.92412.67 × 10−334
TRPC4APBreast_Mammary_Tissue0.55778.62 × 10−313
AXLMinor_Salivary_Gland0.62059.74 × 10−313

Though it was not on the ‘known bone gene’ list, the first gene ranked by TWAS p value, SPTBN1, has been demonstrated to play a role in the regulation of BMD (Calabrese et al., 2017). The second, PPP6R3, has not been previously implicated in the regulation of BMD. PPP6R3, a regulatory subunit of protein phosphatase 6, which shows ubiquitous expression across tissues in humans (Cristiano, 2020), is located on human Chr. 11 within 1 Mbp of seven independent eBMD GWAS SNPs identified by Morris et al., 2019 (subsequently referred to as ‘eBMD lead SNPs’) (Figure 3A). Of all the protein-coding genes (N = 29) in the ~1.8 Mbp region surrounding PPP6R3, its expression was the most significantly associated with eBMD by TWAS (Bonferroni = 5.7 × 10−93) (Figure 3B). Furthermore, PPP6R3 was the only gene in the region with eQTL (in four GTEx tissues, thyroid, ovary, brain_putamen_basal_ganglia, and stomach with RCPs = 0.53, 0.50, 0.28, and 0.14, respectively) that colocalized with at least one of the eBMD associations (Figure 3B). Based on these data, we chose to further investigate PPP6R3 as a potentially causal BMD gene.

PPP6R3 is a top 10 novel estimated bone mineral density (eBMD) gene.

(A) eBMD genome-wide association study (GWAS) SNPs around the PPP6R3 locus (±1 Mbp). The y-axis represents −log10 eBMD GWAS p values. Highlighted SNPs (black) are the seven lead eBMD GWAS SNPs in the locus. (B) Transcriptome-wide association study (TWAS)/colocalization plot for genes in the locus around PPP6R3 (±1 Mbp). The −log10 Bonferroni-adjusted p values from the TWAS analysis (top panel) and the maximum regional colocalization probabilities (RCPs) from the colocalization analyses (bottom panel). Genes alternate in color for visual clarity. Triangles represent PPP6R3. Mirrorplot of the eBMD locus (C) and PPP6R3 expression quantitative trait loci (eQTL) in thyroid, and fracture locus and PPP6R3 eQTL in thyroid (D). The panels on the right are scatterplots of −log10 p values for PPP6R3 eQTL and eBMD GWAS SNPs (C) and the PPP6R3 eQTL and fracture GWAS SNPs (D). SNPs are colored by their linkage disequilibrium (LD) with rs10047483 (purple), the most significant PPP6R3 eQTL in the locus. Not all genes are shown.

We first determined which of the seven associations colocalized with the PPP6R3 eQTL (Figure 3C). The most significant PPP6R3 eQTL SNP in thyroid tissue (the tissue with the highest RCP) was rs10047483 (Chr. 11, 68.464237 Mbp) (PPP6R3 eQTL p = 6.99 × 10−8, eBMD GWAS p = 1.2 × 10−100) located in intron 1 of PPP6R3. The most significant eBMD lead SNP in the locus was rs11228240 (Chr. 11, 68.450822 Mbp, eBMD GWAS p = 6.6 × 10−101, PPP6R3 eQTL p = 3.7 × 10−6), located upstream of PPP6R3. Consistent with the colocalization analysis, these two variants are in high LD (r2 = 0.941) and rs10047483 does not exhibit strong LD (r2 < 0.104) with any of the other six eBMD lead SNPs in the locus. The eQTL and BMD GWAS allele effects for rs10047483 were opposing, suggesting that a decrease in the expression of PPP6R3 would lead to an increase in BMD.

A recent fracture GWAS identified 14 significant associations, one of which was located in the PPP6R3 region (rs35989399, Chr. 11, 68.622433 Mbp) (Morris et al., 2019). We analyzed the fracture GWAS in the same manner as we did above for eBMD. We found that PPP6R3 expression when analyzed by TWAS was significant for fracture (TWAS Bonferroni-pval = 6.0 × 10−3) and the same PPP6R3 eQTL colocalized with the fracture association (RCP = 0.49 in ovary, RCP = 0.36 in thyroid) (Figure 3D). Together, these data highlight PPP6R3 as a strong candidate for one of the seven eBMD/fracture associations in this region.

PPP6R3 is a regulator of femoral geometry, BMD, and vertebral microarchitecture

To assess the effects of PPP6R3 expression on bone phenotypes, we characterized mice harboring a gene-trap allele (Ppp6r3tm1a(KOMP)Wtsi) (Figure 4A). We intercrossed mice heterozygous for the mutant allele to generate mice of all three genotypes (wild-type (WT, N = 26 females, 21 males), heterozygous (HET, N = 33 females, 36 males), and mutant (MUT, N = 20 females, 21 males)). The absence of PPP6R3 protein in MUT mice was confirmed through western blotting (Figure 4B). Furthermore, we sequenced RNA extracted from the L5 vertebrae of wild-type and mutant mice, and found that PPP6R3 was differentially expressed between the mutant and wild-type mice (padj = 3.67 × 10−104, log2 fold change = −1.63). We did not observe changes in gene expression for any other gene in the locus, including Lrp5 (padj = 0.99, log2 fold change = −0.068), a well-known regulator of bone mass just upstream of Ppp6r3 (Supplementary file 1j).

Figure 4 with 6 supplements see all
Ppp6r3 functional validation shows an effect of genotype on bone mass.

(A) Schematic of the Ppp6r3 gene-trap allele (Ppp6r3tm1a(KOMP)Wtsi). Image obtained from the International Mouse Phenotype Consortium (IMPC). (B) Western blot of Ppp6r3 in experimental mouse spleens. Top left panel shows that PPP6R1 protein (control) levels are not affected by the Ppp6r3 gene-trap allele. Top right panel shows the effect of the gene-trap allele on PPP6R3 protein levels. The two bands are ostensibly due to different PPP6R3 isoforms. Bottom panel shows that PP6C protein (control) levels are not affected by the Ppp6r3 gene-trap allele. For raw blots, refer to Figure 4—source data 1. Least-squares means for spinal (C) and femoral (D) areal BMD (aBMD) dual X-ray absorptiometry (DXA) in Ppp6r3 wild-type (WT), heterozygous (HET), and mutant (MUT) mice. Contrast p values, adjusted for multiple comparisons are presented. *p ≤ 0.05. (E) Representative images of vertebrae for the Ppp6r3 experimental mice. Scale is shown on the bottom right. (F, G) Least-squares means for micro-computed tomography (μCT) measurements in the lumbar spines of Ppp6r3 WT, HET, and MUT mice. Contrast p values, adjusted for multiple comparisons are presented. *p ≤ 0.05. (H, I) Least-squares means for μCT measurements in the femoral midshaft of Ppp6r3 WT, HET, and MUT mice. Contrast p values, adjusted for multiple comparisons are presented. *p ≤ 0.05. Abbreviations: BV/TV – bone volume fraction, vBMD – volumetric bone mineral density, Tot.Ar – total area, Ma.Ar – medullary area. In panels (C, D and F–I), the center points represent the least-square means, and the error bars represent confidence intervals at a confidence level of 0.9. Data presented here are from both male and female mice.

The BMD analyses presented above used heel eBMD GWAS data. We used these data because they represent the largest, most well-powered BMD GWAS to date (Estrada et al., 2012). However, to determine whether perturbation of Ppp6r3 would be expected to impact femoral or lumbar spine BMD in a similar manner, we turned to a smaller GWAS to look at both of these traits. In a GWAS by Estrada et al., 2012, a total of 56 loci were identified for femoral neck (FNBMD) and lumbar spine (LSBMD) BMD. One of the 56 loci corresponded to the same SNPs associated with the PPP6R3 eQTL. The locus was significant for LSBMD; however, it did not reach genome-wide significance for FNBMD (Figure 4—figure supplement 1).

We evaluated BMD at both the femur and the lumbar spine in Ppp6r3tm1a(KOMP)Wtsi mice, with the expectation, based on the above data, that perturbation of Ppp6r3 would have a stronger impact on BMD at the lumbar spine. At approximately 9 weeks of age, we measured areal BMD (aBMD) at the femur and lumbar spine using dual X-ray absorptiometry (DXA). First, we observed no change in body weight at 9 weeks that might impact bone phenotypes (Figure 4—figure supplement 2A). As the above analysis predicted, we observed a significant effect of Ppp6r3 genotype on aBMD at the lumbar spine (WT vs. MUT p = 0.01, Figure 4—figure supplement 2C), but not the femur (WT vs. MUT p = 0.26, Figure 4—figure supplement 2D). It should also be noted, however, that we observed significantly decreased femoral width, but not length, in Ppp6r3 mutant mice (anterior–posterior [AP] femoral width, WT vs. MUT p = 0.02; medial–lateral [ML] femoral width, WT vs. MUT p = 2.2 × 10−6, Figure 4—figure supplement 2B–D).

We further characterized the effects of Ppp6r3 genotype on microarchitectural phenotypes, in spine and femur, via micro-computed tomography (μCT). We observed significant (p ≤ 0.05) decreases in trabecular bone volume fraction (BV/TV, WT vs. MUT p = 0.015, Figure 4—figure supplement 2E, F) and volumetric BMD (vBMD, WT vs. MUT p = 0.015, Figure 4—figure supplement 2G) of the lumbar spine as a function of Ppp6r3 genotype, but found no significant changes in tissue mineral density (TMD, Figure 4—figure supplement 2E), trabecular separation (TbSp), trabecular thickness (TbTh), or trabecular number (TbN) (Figure 4—figure supplement 2F–H). In the femoral midshaft, we observed significant decreases in total area (Tot.Ar, WT vs. MUT p = 0.00724, Figure 4—figure supplement 2H) and medullary area (Ma.Ar, WT vs. MUT p = 0.00903, Figure 4—figure supplement 2I), which are concordant with the aforementioned observed decreases in femoral width in Ppp6r3 mutant mice. We did not observe any changes in trabecular bone parameters in the distal femur.

We also measured Procollagen 1 Intact N-Terminal Propeptide P1NP levels in plasma, and observed a significant increase in P1NP levels in mutant mice (p = 0.00422), suggesting that the observed decrease in bone mass is due to increased bone turnover (Figure 4—figure supplement 2I–K). Finally, to assess the effects of Ppp6r3 genotype on bone matrix composition, we performed periosteal Raman spectroscopy on both the lumbar spines and femurs. We did not observe any significant (p ≤ 0.05) effects of Ppp6r3 genotype on bone matrix composition (Figure 4—figure supplements 36).

Discussion

BMD GWASs have identified over 1100 associations to date. However, identifying causal genes remains a challenge. To aid researchers in further dissecting the genetics of complex traits, reference transcriptomic datasets and computational methods have been developed for the prioritization and identification of causal genes underlying GWAS associations. In this work, our goal was to utilize these data and tools to prioritize putatively causal genes underlying BMD GWAS associations. Specifically, we used the GTEx eQTL reference dataset in 49 tissues to perform TWAS and eQTL colocalization on the largest BMD GWAS. Using this approach, we identified 512 putatively causal protein-coding genes that were significant in both the TWAS and colocalization approaches.

Our approach was inspired by a recent study that used the GTEx resource and a TWAS/eQTL colocalization approach similar to the one we employed. Pividori et al., 2020 recently combined TWAS and eQTL colocalization to GTEx and GWAS data on 4091 traits, including BMD, from the UK Biobank data. A total of 76 protein-coding genes were identified and of the 76, we identified 55 (72.4%) of the same genes in our implementation. There are several reasons for this discrepancy in the number of prioritized genes. First, both studies used a GWAS based on the UK Biobank (Bycroft et al., 2018); however, sample sizes were different. The PhenomeXcan project utilized GWAS data based on the analysis of ~207,000 individuals, whereas we used GWAS data based on the analysis of ~426,000 individuals (Morris et al., 2019; Pividori et al., 2020). Second, the two GWAS studies utilized different association models. Finally, due to the breadth of the PhenomeXcan project, they had a higher multiple-testing burden than we did, which led to different Bonferroni-adjusted p value thresholds (p < 5.49 × 10−10 vs. p ≤ 2.38 × 10−6).

One of many novel genes identified in our study was PPP6R3, which was also identified in the PhenomeXcan project (Pividori et al., 2020). PPP6R3 is a regulatory subunit of protein phosphatase 6 and has been implicated in several cancers (Cristiano, 2020; Stefansson and Brautigan, 2006). In humans, the PPP6R3 protein shows ubiquitous expression across tissues, and may have an important role in maintaining immune self-tolerance (Cristiano, 2020). It is unclear how PPP6R3 may be influencing BMD. However, protein phosphatase 6 has been shown to oppose activation of the nuclear factor kappa-light-chain enhancer of activated B cells (NF-κB) pathway in lymphocytes (Ziembik et al., 2017). Since the NF-κB signaling pathway is highly involved in osteoclastogenesis and bone resorption, it is possible that PPP6R3 may be involved in the regulation of this pathway in osteoclasts (Abu-Amer, 2013). Further studies that characterize the role of PPP6R3, and the effects of its deletion, in bone cells are required to further elucidate its effect on BMD.

The PPP6R3 locus demonstrated a high level of complexity, containing seven independent GWAS associations, at least one of which was also associated with fracture. Interestingly, just upstream of PPP6R3 is LRP5, a WNT signaling co-receptor (Mao et al., 2001). LRP5 is a well-known regulator of BMD and gain and loss of function mutations lead to high bone mass syndrome and osteoporosis pseudoglioma, respectively (Mizuguchi et al., 2004; Boyden et al., 2002; Marques-Pinheiro et al., 2010; Gong et al., 2001). LRP5 expression was not significantly associated with eBMD by TWAS (Bonferroni p = 1), nor did it have a colocalizing eQTL in GTEx tissues (most significant RCP = 1.6 × 10−2 in pancreas). However, another eBMD lead SNP in the region, rs4988321, is a missense mutation in LRP5 (Val667Met) that has been associated with BMD in multiple studies (van Meurs et al., 2008; Brixen et al., 2007; Giroux et al., 2007). While this variant represents an association that is independent of the rs10047483 association (r2 = 0.104), it further highlights the complexity of this locus both in terms of the number of associations and target genes. We believe these data support a model of allelic heterogeneity involving multiple genes (at least LRP5 and PPP6R3) at this locus, similar to other BMD loci identified by GWAS such as the ‘Wnt16’ locus (Chesi et al., 2019).

To determine the effect of Ppp6r3 expression on bone, we characterized bone phenotypes in mice harboring a gene-trap allele (Ppp6r3tm1a(KOMP)Wtsi). Consistent with the observation that the PPP6R3 eQTL SNPs were significantly associated with lumbar spine, but not femoral neck BMD, we observed that Ppp6r3 deletion had a significant effect on lumbar spine BMD, but not femoral BMD. Using μCT, we further characterized the effect of Ppp6r3 deletion on lumbar spine and femoral microarchitecture. We observed significant decreases in trabecular bone volume fraction (BV/TV) and vBMD of the lumbar spine as a function of PPP6R3 genotype. While we did not observe significant effects of Ppp6r3 deletion on trabecular thickness or number, the direction of effects for those phenotypes suggests that the observed decrease in bone volume fraction and BMD may be explained by the cumulative but more subtle effects of Ppp6r3 deletion on trabecular thickness and number. Consistent with our caliper-based observations that femoral width was decreased at the midshaft in Ppp6r3 mutants, we observed a decrease in medullary and total area in the femoral midshaft. We also hypothesized that Ppp6r3 deficiency might lead to defects in matrix composition; however, we did not observe any significant differences. The lack of differences might be due to the fact that we used relatively young mice for these assays.

Our hypothesis regarding the directions of effect of Ppp6r3 expression on BMD based on the eQTL and eBMD/lumbar spine BMD GWAS were opposite to what we observed. There are several reasons that may explain this. First, our hypothesis was based on expression data in non-bone tissues and cell types. Recent studies have shown that the direction of eQTL effects can differ between different cells and tissues within humans (Peters et al., 2016; Mizuno and Okada, 2019). Second, our hypothesis was based on human data, while our functional experiments were performed in mice. Third, we globally deleted Ppp6r3 in mice, as opposed to ablating it in a cell-type-specific manner. Future studies investigating which tissue/cell-type Ppp6r3 is operative in and the generation of conditional Ppp6r3 knockouts will allow us to unravel the precise role of this association and Ppp6r3 in the regulation of bone mass.

As we and others have shown, the use of both TWAS and eQTL colocalization can prioritize putatively causal genes underlying GWAS associations. Here, we have shown the utility of this approach even in the absence of eQTL data from the most phenotype-relevant tissue. However, it is important to highlight the limitations of our analysis. While studies have shown that many eQTL are shared among tissues, the lack of eQTL data in bone and bone cells means that bone-specific eQTL were missed. For example, a study conducted by Mullin et al. performed eQTL colocalization and summary-based Mendelian randomization (SMR) by utilizing GWAS and expression data from osteoclast-like cells, and prioritized several eBMD genes (Mullin et al., 2020). Thirty-eight percent of the colocalizing eQTL and 19% of the SMR genes that they identified overlapped with our 512 prioritized genes, suggesting that we have missed many potential effector genes with eQTL specific to bone cells. In addition, the use of multiple non-bone tissues may have inflated the number of false positives based on coincidence of strong TWAS and eQTL colocalization signals that have no biological impact on bone. Furthermore, the lack of bone transcriptomic data may also explain the observed disparity between our hypothesized and observed direction-of-effect for PPP6R3. It is also important to note that due to the reliance of this approach on eQTL data, genes that affect BMD via non-expression-related mechanisms were not captured. Another limitation of our approach arises from the definition of loci based on linkage disequilibrium (LD). We used a set of previously defined approximately independent LD blocks, derived from a cohort of European individuals, in our fastENLOC analysis (Berisa and Pickrell, 2016). The inexact nature of these data may lead to spurious colocalizations due to mismatches in LD structure between the reference LD blocks and the GWAS/eQTL populations. Additionally, because the GWAS and eQTL data have mismatching LD structures, due to their being derived from cohorts with different ancestries, our analyses, particularly the colocalization analyses, may suffer from reduced power (Hukku et al., 2021). This also raises the related issue of the reduced generalizability of our results in non-European individuals, which brings further attention to the necessity of performing GWASs and providing reference data in diverse and underrepresented populations. Additionally, another issue arises when considering correlations in expression, and predicted expression, between genes in a locus, which may lead to spurious associations in TWAS analyses (Wainberg et al., 2019). Finally, as we show above, our method does not perform as well as prioritizing genes based on their proximity to GWAS associations. However, because our method utilizes systems genetics techniques and data, such as eQTL, we believe that our method prioritizes genes in a more biologically relevant manner. In fact, utilizing the closest gene method alone, PPP6R3 would not have been prioritized as a bone-relevant gene. We suggest that future studies utilize both prioritization techniques, such as taking the closest genes to GWAS associations and cross referencing them with colocalizing and TWAS-associated genes, in order to provide further evidence for functional validation.

In summary, we applied a combined TWAS/colocalization approach using GTEx and identified 512 putatively causal BMD genes. We further investigated PPP6R3 and demonstrated that it is a regulator of lumbar spine BMD. We believe this work provides a valuable resource for the bone genetics community and may serve as a framework for prioritizing genes underlying GWAS associations using publicly available tools and data for a wide range of diseases.

Methods

fastENLOC colocalization

For each of the eBMD and fracture GWASs, we performed colocalization using fastENLOC, by following the tutorial and guidelines available at https://github.com/xqwen/fastenloc (Wen, 2022).

Briefly, for each GWAS, we converted variant coordinates to the hg38 human genome assembly, using the UCSC liftOver tool (minimum ratio of bases that must remap = 1; https://genome.ucsc.edu/cgi-bin/hgLiftOver). We calculated z-scores by dividing GWAS betas by standard errors. We then defined loci based on European LD blocks, as defined based on the results of Berisa and Pickrell, 2016.

Z-scores were then converted to posterior inclusion probabilities using torus (Wen, 2015). Finally, these data were colocalized with fastENLOC for all 49 GTEx V8 tissues, with the ‘-total_variants’ flag set to 14,000,000. Colocalization was performed using pre-computed GTEx multi-tissue annotations, obtained from https://github.com/xqwen/fastenloc; Wen, 2022. Finally, to identify protein-coding genes in the results, we utilized Ensembl’s ‘hsapiens_gene_ensembl’ dataset using biomaRt (version 2.45.8).

S-MultiXcan

We conducted a TWAS by integrating genome-wide SNP-level association summary statistics from an eBMD GWAS (Morris et al., 2019) with GTEx version 8 gene expression QTL data from 49 tissue types. We used the S-MultiXcan approach for this analysis, to correlate gene expression across tissues to increase power and identify candidate susceptibility genes (Barbeira et al., 2019). Default parameters were used, with the exception of the ‘--cutoff_condition_number’ parameter, which was set to 30. Bonferroni-correction of p values was performed on the resultant gene set (22,337 genes), using R’s ‘p.adjust’ function. This was followed by the removal of non-protein-coding genes. The analysis was also performed in the same manner using summary statistics from a fracture GWAS (Morris et al., 2019). Finally, to identify protein-coding genes in the results, we utilized Ensembl’s ‘hsapiens_gene_ensembl’ dataset using biomaRt (Durinck et al., 2009; Durinck et al., 2005).

Creation of the ‘known bone gene’ list

We generated a ‘known bone gene’ set as follows: First, we downloaded Gene Ontology IDs for the following terms: ‘osteo*’, ‘bone’, and ‘ossif*’ from AmiGO2 (version 2.5.13) (Carbon et al., 2009). After removal of non-bone-related terms, we extracted all mouse and human genes related to the GO terms, using biomaRt. From this list, we retained protein-coding genes.

We also used the ‘Human-Mouse: Disease Connection’ database available at the Mouse Genome Informatics website, to download human and mouse genes annotated with the terms ‘osteoporosis’, ‘bone mineral density’, ‘osteoblast’, ‘osteoclast’, and ‘osteocyte’. We used biomaRt to identify the gene biotypes, and retained protein-coding genes. We then used the MGI human-mouse homology table (http://www.informatics.jax.org/downloads/reports/HOM_MouseHumanSequence.rpt) to convert all mouse genes to their human homologs. Finally, we removed genes that were not interrogated in both the colocalization and the TWAS analyses.

GO enrichment analyses

GO analysis was performed for the set of protein-coding genes passing the colocalization threshold RCP ≥0.1 and S-MultiXcan Bonferroni p value ≤0.05, using the ‘topGO’ package (version 2.46.0) in R (Alexa and Rahnenfuhrer, 2021). Enrichment tests were performed for the ‘Molecular Function’, ‘Biological Process’, and ‘Cellular Component’ ontologies, using all protein-coding genes that were subjected to colocalization and MultiXcan analysis as background. Enrichment was performed using the ‘classic’ algorithm with Fisher’s exact test. p values were not adjusted for multiple testing.

LD calculations

LD between variants was calculated using the LDlinkR (version 1.0.2) R package, using the ‘EUR’ population (Myers et al., 2020).

Ppp6r3 knockout mouse generation

The study was carried out in strict accordance with NIH’s Guide for the Care and Use of Laboratory Animals. Additionally, the University of Virginia Institutional Animal Care and Use Committee approved all animal procedures. Ppp6r3 gene-trap mice were generated using targeted embryonic stem cell clones heterozygous for the Ppp6r3tm1a(KOMP)Wtsi gene-trap allele obtained from the International Knockout Mouse Project (KOMP; https://www.komp.org). KOMP ES clones were karyotyped and injected using a XYClone Laser (Hamilton Thorne, Beverly, MA) into B6N-Tyrc-Brd/BrdCrCrl (Charles River, Wilmington, MA) eight-cell stage embryos to create chimeric mice. Resultant chimeras were bred to B6N-Tyrc-Brd/BrdCrCrl mice to obtain germline transmission of the Ppp6r3 gene-trap allele. From a breeding pair of two heterozygous mice, we generated our experimental population through HET × HET matings. Breeder mice were fed a breeder chow diet (Envigo Teklad S-2335 mouse breeder sterilizable diet, irradiated. Product # 7904), and experimental mice were fed a standard chow diet (Envigo Teklad LM-485 irradiated mouse/rat sterilizable diet. Product #7912).

Genotyping of Ppp6r3 mice

DNA for genotyping was extracted from tail clips as follows: tail clips were incubated overnight at 55°C in a solution of 200 µl digestion/lysis buffer (Viagen Direct PCR [tail], Los Angeles, CA) and 1 mg/ml proteinase K (Viagen, Los Angeles, CA). After overnight incubation, tails were heated at 85°C for 45 min, and solutions were subsequently stored at 4°C.

For genotyping, PCR reactions were set up as follows. For each reaction, 1 μl of DNA was mixed with 24 μl of a master mix consisting of 19.5 μl nuclease-free H2O, 2.5 μl 10× PCR reaction buffer (Invitrogen, Waltham, MA), 0.75 μl of MgCl2 (Invitrogen, Waltham, MA), 0.5 μl of 10 mmol Quad dNTPs (Roche Diagnostics GmbH, Mannheim, Germany) 0.25 μl of Platinum Taq DNA polymerase (Invitrogen, Waltham, MA), and 0.25 μl of each primer, diluted to 20 μmol.

Primers: PCR primers were obtained from Integrated DNA Technologies, Coralville, IA.

  • Forward primer: 5′-CAC CTG GGT TGG TTA CAT CC-3′

  • Reverse primer: 5′-GAC CCT GCC TTA AAA CCA AA-3′

The following PCR settings were used:

  • Initialization: 94°C, 120 s

  • Denaturation: 94°C, 30 s (37 cycles)

  • Annealing: 54°C, 30 s (37 cycles)

  • Elongation: 72°C, 35 s (37 cycles)

  • Final elongation: 72°C, 300 s

PCR products were run on a 2% agarose gel for 150 min at 60 V, to distinguish between wild-type, heterozygous and mutant Ppp6r3 mice.

PPP6R3 western blotting

Mouse spleens 20–40 mg in weight were suspended in 1% NP40 buffer (50 mM Tris [pH 8] 100 mM NaCl, 1% NP40, 1 mM EGTA(egtazic acid), 1 mM EDTA(ethylenediaminetetraacetic acid), Protease inhibitor cocktail (04-693-116-001, Roche), 1 mM PMSF(phenylmethylsulfonyl fluoride), 50 mM NaF, 0.2 mM sodium vanadate). The tissue was homogenized by RNase-free disposable pestles (Thermo Fisher #12-141-364) and incubated for 10 min on ice. After brief sonication, the sample was centrifuged for 10 min at 13,000 × rpm at 4°C. The protein concentration in the extract was measured by Bradford assay. 100 µg of sample protein was boiled 5 min in sodium dodecyl sulfate (SDS) sample buffer, loaded in each lane, resolved by gradient SDS–polyacrylamide gel electrophoresis (Bio-Rad #456-1085) and immunoblotted as described in Guergnon et al., 2009. Primary antibodies were diluted 1:1000 (SAPS1 Ab: Thermo Fisher #PA5-44275, SAPS3 Ab: Thermo Fisher #PA5-58405, PP6C Ab: Sigma #HPA050940).

PPP6R3 functional validation

Experimental mice of both sexes were sacrificed at approximately 9 weeks of age (mean age = 61 days). At sacrifice, the right femurs were isolated, and femoral morphology (length and widths in anterior–posterior and medial–lateral orientations) was measured with digital calipers (Mitoyuto American, Aurora, IL).

Femurs were then wrapped in phosphate-buffered saline (PBS)-soaked gauze and stored at −20°C, until analysis. Lumbar vertebrae L3–L5 were also dissected at sacrifice and were wrapped in PBS-soaked gauze and frozen at −20°C. Given our prior experience in measuring bone geometry and microstructure, we used a minimum N = 14 for our analyses, ensuring 80% power to detect a statistically significant effect at an alpha ≤0.05.

Bulk RNA isolation, sequencing, and quantification

We isolated RNA from a randomly chosen subset (n = 16, 4/sex each of WT and MUT) of the available mice.Total RNA was isolated from L5 vertebrae, using the mirVana miRNA Isolation Kit (Life Technologies, Carlsbad, CA). Total RNA-seq libraries were constructed using Illumina TruSeq Stranded mRNA LT sample prep kits. Samples were sequenced to an average of 39 million 2 × 151 bp paired-end reads (total RNA-seq) on an Illumina NextSeq500 sequencer by Psomagen, Inc A custom bioinformatics pipeline was used to quantify RNA-seq data. Briefly, RNA-seq FASTQ files were quality controlled using FASTQC (version 0.11.5) and MultiQC (version 1.11). Reads were trimmed using bbduk (bbmap package version 38.57). Trimmed reads were then aligned to the mm10 genome assembly with HISAT2 (version 2.1), and quantified with Stringtie (version 1.3.3). Read count information was then extracted with a Python script provided by the Stringtie website (prepDE.py).

Bulk RNA differential expression analyses

Using gene count matrices, differential expression was performed using DESeq2 (Love et al., 2014; Version 1.34.0) between wild-type and mutant samples. We used the ‘DESeq’ function from DESeq2, with a design formula of ~sex + genotype. p values were adjusted using the ‘p.value’ function, using the ‘BH’ method.

Dual X-ray absorptiometry

Individual right femurs and the lumbar spine (L5 vertebrae) were isolated from surrounding soft tissues and frozen at −20°C in PBS. DXA was performed on the femurs and lumbar vertebrae using the Lunar Piximus II (GE Healthcare) as described previously by Beamer et al., 2011. In short, 10 isolated bones were placed in the detector field at a time and the samples were analyzed one by one, such that the region of interest (ROI) was set for one specimen at a time for data collection. The ROI for the femurs was on the entire isolated femur. For the spine, was on the entire isolated L5. Care was taken to ensure that the sample orientation was identical for all samples.

μCT and image analysis

All μCT analyses were carried out at the μCT Imaging Core Facility at Boston University using a Scanco Medical μCT 40 instrument (Brütisellen, Switzerland). The power, current, and integration time used for all scans were 70 kVp, 113 μA, and 200 ms, respectively. The L5 vertebrae and right femora were scanned at a resolution of 12 μm/voxel. Two volumes of interest (VOIs) in the L5 were selected for analysis: (1) the entire portion of the L5 vertebra extending from 60 μm caudal to the cranial growth plate in the vertebral body to 60 μm cranial to the caudal growth plate; and (2) only the trabecular centrum contained in the first VOI. Semi-automated-edge detection (Scanco Medical) was used to define the boundary between the trabecular centrum and cortical shell to produce the second VOI. Two VOIs were also analyzed for each femur: (1) a 0.3-mm-long segment of the diaphysis, centered at the mid-point of the bone; and (2) a 1.2-mm-long segment of the distal metaphyseal trabecular compartment. To define the location of the latter, the location of the distal femoral growth plate was determined, and the distal end of the VOI was set at 60 μm proximal to that growth plate. Gaussian filtering (sigma = 0.8, support = 1) was used for partial background noise suppression. A scan of a potassium hydroxyapatite phantom allowed conversion of gray values to mineral density. For segmentation of bone tissue, the threshold was set at a 16-bit gray value of 7143 (521 mgHA/ccm), and this global threshold was applied to all of the samples. For each VOI, the following were calculated: total volume (TV), bone volume (BV), bone volume fraction (BV/TV), BMD, and TMD. BMD was defined as the average density of all voxels in the VOI, whereas TMD was defined as the average density of all voxels in the VOI above the threshold (Bouxsein et al., 2010). For the second VOI, the following additional parameters were calculated: trabecular thickness (Tb.Th), trabecular separation (Tb.Sp), trabecular number (Tb.N), connectivity density (Conn.D), and structure model index (SMI) (Bouxsein et al., 2010).

For the femoral VOI, the following additional parameters were calculated instead: cortical thickness (Ct.Th), total area (Tot.Ar), marrow area (Ma.Ar), maximum and minimum moments of inertia, and polar moment of inertia (Bouxsein et al., 2010).

P1NP collection and quantification

Plasma was collected via submandibular bleeding from isoflurane anesthetized wild-type and mutant PPP6R3 mice (N = 10/sex/genotype). Plasma P1NP levels were measured using commercially available kits from IDS (Gaithersburg, MD), according to the manufacturer’s instructions. The assay sensitivity was 0.7 ng/ml. The intra-assay variation was 6.3%, and the inter-assay variation was 8.5%. All measurements were performed in duplicate.

Raman spectroscopy

Raman spectroscopy was performed using a Renishaw inVia Raman Microscope (Gloucestershire, UK) on each bone sample using a 785-nm-edge red incident laser. A rectangular filled map was created with 3 points in the x-axis and 20 points in the y-axis, for a total of 60 collected points. Each point was exposed 10 times for 6 s per exposure. A custom MATLAB script was used to evaluate the peak position, maximum intensity, peak width, full width at half maximum (FWHM), and the area under each peak. Peak area ratios were calculated for mineral:matrix, carbonate:phosphate, and crystallinity. Furthermore, the standard deviations of peak area ratios were calculated for each mouse, and were further used to evaluate the material heterogeneity in groups.

Statistical analyses

To calculate the enrichment of bone genes in prioritized genes, we performed Fisher’s exact test, using R’s ‘fisher.test’ function, with the alternative hypothesis set as ‘greater’.

For the statistical analysis of the phenotyping results, we calculated least-squares means (lsmeans) using the ‘emmeans’ R package (version 1.5.2.1) (Lenth, 2020). Input for the lsmeans function was a linear model including terms for genotype, weight, and age in days. For sex-combined data, we also added a term for sex. For DXA phenotypes, we included a term for ‘CenterRectX’ and ‘CenterRectY’. For the Raman spectroscopy data, weight and age were not included as terms in the linear model.

We used Tukey’s HSD(honest significance test) test to test for significant differences in lsmeans, for each pair of genotype levels. Tukey’s HSD also controls the family-wise error rate. We performed Welch two-sample t-test’s, using R’s ‘t.test’ function, to quantify differences in P1NP levels between mutant and wild-type mice.

Analyses involving data from the International Mouse Phenotyping Consortium

For the IMPC data, we obtained data using their ‘statistical-result’ SOLR database, using the ‘solrium’ R package (version 1.1.4) (Chamberlain, 2019). We obtained experimental results using the ‘Bone*Mineral*Density’ parameter. We then pruned the resulting data to only include ‘Successful’ analyses, and removed experiments that included the skull. To generate the Gpatch1 boxplot, we obtained raw data using from IMPC’s ‘statistical-raw-data’ SOLR database for Gpatch1, and analyzed the data in the same manner as IMPC, using the ‘OpenStats’ R package (version 1.0.2), using the method = ‘MM’ and MM_BodyWeightIncluded = TRUE arguments (Haselimashhadi et al., 2020). Finally, mouse genes were converted to their human syntenic counterparts using Ensembl’s ‘hsapiens_gene_ensembl’ and ‘mmusculus_gene_ensembl’ datasets through biomaRt.

PhenomeXcan data analysis

We obtained all significant PhenomeXcan gene–trait associations from their paper (https://advances.sciencemag.org/content/6/37/eaba2083), and used data for the ‘3148_raw-Heel_bone_mineral_density_BMD’ phenotype (Pividori et al., 2020). Furthermore, we constrained our search to only include genes that were annotated by the authors as ‘protein_coding’.

LSBMD/FNBMD GWAS analysis

We obtained sex-combined LSBMD and FNBMD GWAS summary statistics from GEFOS (http://www.gefos.org/?q=content/data-release-2012), and then used a custom script that utilized the biomaRt R package to convert variants to their GRCh38 coordinates.

Data availability

eBMD and fracture GWAS summary statistics were obtained from GEFOS, as were the LSBMD and FNBMD GWAS summary statistics. GTEx eQTL data were obtained from the GTEx web portal. Data from the PhenomeXcan project were obtained from Pividori et al., 2020. Statistical data from the IMPC were obtained using an R interface to their SOLR database. Ppp6r3 experimental data are provided on our GitHub (https://github.com/basel-maher/BMD_TWAS_colocalization; Al-Barghouthi, 2022). Mouse-Human homologs were obtained from MGI (http://www.informatics.jax.org/downloads/reports/HOM_MouseHumanSequence.rpt). We also obtained data from the MGI Human-Mouse:Disease Connection database (http://www.informatics.jax.org/diseasePortal). Gene Ontologies were obtained from AmiGO2 (http://amigo.geneontology.org/amigo).

Code availability

Analysis code and the raw data for our Ppp6r3 functional validation analyses are available on GitHub (https://github.com/basel-maher/BMD_TWAS_colocalization, copy archived at swh:1:rev:6aaa8819c2e335013a665e76318dc98aeb9a52ce; Al-Barghouthi, 2022).

Data availability

All data and source code are available on GitHub: https://github.com/basel-maher/BMD_TWAS_colocalization (copy archived at swh:1:rev:6aaa8819c2e335013a665e76318dc98aeb9a52ce).

The following previously published data sets were used
    1. Morris JA
    (2018) GEFOS
    ID release-2018. UK Biobank eBMD and Fracture GWAS Data Release 2018.
    1. Estrada K
    (2012) GEFOS
    ID release-2012. Data Release 2012.
    1. Pividori M
    (2020) hakyimlab
    ID phenomexcan. PhenomeXcan.

References

    1. Cristiano L
    (2020) PPP6R3 (protein phosphatase 6 regulatory subunit 3)
    Atlas of Genetics and Cytogenetics in Oncology and Haematology 01:70657.
    https://doi.org/10.4267/2042/70657
    1. Estrada K
    2. Styrkarsdottir U
    3. Evangelou E
    4. Hsu Y-H
    5. Duncan EL
    6. Ntzani EE
    7. Oei L
    8. Albagha OME
    9. Amin N
    10. Kemp JP
    11. Koller DL
    12. Li G
    13. Liu C-T
    14. Minster RL
    15. Moayyeri A
    16. Vandenput L
    17. Willner D
    18. Xiao S-M
    19. Yerges-Armstrong LM
    20. Zheng H-F
    21. Alonso N
    22. Eriksson J
    23. Kammerer CM
    24. Kaptoge SK
    25. Leo PJ
    26. Thorleifsson G
    27. Wilson SG
    28. Wilson JF
    29. Aalto V
    30. Alen M
    31. Aragaki AK
    32. Aspelund T
    33. Center JR
    34. Dailiana Z
    35. Duggan DJ
    36. Garcia M
    37. Garcia-Giralt N
    38. Giroux S
    39. Hallmans G
    40. Hocking LJ
    41. Husted LB
    42. Jameson KA
    43. Khusainova R
    44. Kim GS
    45. Kooperberg C
    46. Koromila T
    47. Kruk M
    48. Laaksonen M
    49. Lacroix AZ
    50. Lee SH
    51. Leung PC
    52. Lewis JR
    53. Masi L
    54. Mencej-Bedrac S
    55. Nguyen TV
    56. Nogues X
    57. Patel MS
    58. Prezelj J
    59. Rose LM
    60. Scollen S
    61. Siggeirsdottir K
    62. Smith AV
    63. Svensson O
    64. Trompet S
    65. Trummer O
    66. van Schoor NM
    67. Woo J
    68. Zhu K
    69. Balcells S
    70. Brandi ML
    71. Buckley BM
    72. Cheng S
    73. Christiansen C
    74. Cooper C
    75. Dedoussis G
    76. Ford I
    77. Frost M
    78. Goltzman D
    79. González-Macías J
    80. Kähönen M
    81. Karlsson M
    82. Khusnutdinova E
    83. Koh J-M
    84. Kollia P
    85. Langdahl BL
    86. Leslie WD
    87. Lips P
    88. Ljunggren Ö
    89. Lorenc RS
    90. Marc J
    91. Mellström D
    92. Obermayer-Pietsch B
    93. Olmos JM
    94. Pettersson-Kymmer U
    95. Reid DM
    96. Riancho JA
    97. Ridker PM
    98. Rousseau F
    99. Slagboom PE
    100. Tang NLS
    101. Urreizti R
    102. Van Hul W
    103. Viikari J
    104. Zarrabeitia MT
    105. Aulchenko YS
    106. Castano-Betancourt M
    107. Grundberg E
    108. Herrera L
    109. Ingvarsson T
    110. Johannsdottir H
    111. Kwan T
    112. Li R
    113. Luben R
    114. Medina-Gómez C
    115. Palsson ST
    116. Reppe S
    117. Rotter JI
    118. Sigurdsson G
    119. van Meurs JBJ
    120. Verlaan D
    121. Williams FMK
    122. Wood AR
    123. Zhou Y
    124. Gautvik KM
    125. Pastinen T
    126. Raychaudhuri S
    127. Cauley JA
    128. Chasman DI
    129. Clark GR
    130. Cummings SR
    131. Danoy P
    132. Dennison EM
    133. Eastell R
    134. Eisman JA
    135. Gudnason V
    136. Hofman A
    137. Jackson RD
    138. Jones G
    139. Jukema JW
    140. Khaw K-T
    141. Lehtimäki T
    142. Liu Y
    143. Lorentzon M
    144. McCloskey E
    145. Mitchell BD
    146. Nandakumar K
    147. Nicholson GC
    148. Oostra BA
    149. Peacock M
    150. Pols HAP
    151. Prince RL
    152. Raitakari O
    153. Reid IR
    154. Robbins J
    155. Sambrook PN
    156. Sham PC
    157. Shuldiner AR
    158. Tylavsky FA
    159. van Duijn CM
    160. Wareham NJ
    161. Cupples LA
    162. Econs MJ
    163. Evans DM
    164. Harris TB
    165. Kung AWC
    166. Psaty BM
    167. Reeve J
    168. Spector TD
    169. Streeten EA
    170. Zillikens MC
    171. Thorsteinsdottir U
    172. Ohlsson C
    173. Karasik D
    174. Richards JB
    175. Brown MA
    176. Stefansson K
    177. Uitterlinden AG
    178. Ralston SH
    179. Ioannidis JPA
    180. Kiel DP
    181. Rivadeneira F
    (2012) Genome-Wide meta-analysis identifies 56 bone mineral density loci and reveals 14 loci associated with risk of fracture
    Nature Genetics 44:491–501.
    https://doi.org/10.1038/ng.2249
    1. Nica AC
    2. Dermitzakis ET
    (2013) Expression quantitative trait loci: present and future
    Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences 368:20120362.
    https://doi.org/10.1098/rstb.2012.0362

Decision letter

  1. Nele Haelterman
    Reviewing Editor; Baylor College of Medicine, United States
  2. Mone Zaidi
    Senior Editor; Icahn School of Medicine at Mount Sinai, United States
  3. Erika Kague
    Reviewer; University of Bristol, United Kingdom

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Transcriptome-wide Association Study and eQTL colocalization identify potentially causal genes responsible for bone mineral density GWAS associations" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Mone Zaidi as the Senior Editor. The following individual involved in the review of your submission has agreed to reveal their identity: Erika Kague (Reviewer #2).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

The reviewers would like to see more data and discussion on the possible role of LRP5 in the bone phenotype of PPP6R3 mutant mice.

a. Please assess expression levels of Ppp6r3 and Lrp5 in your mutant model.

b. Please include a more in-depth characterization of Ppp6r3 mutant mice:

femoral microCT data, P1NP or TRAP expression.

c. Please discuss whether rs10047483 is in LD with the SNP that is closest to LRP5.

Reviewer #1 (Recommendations for the authors):

The authors have presented the methods, data, analysis, and interpretation clearly. It's refreshing that the authors were transparent about several issues that did not show their approach in the most positive light, i.e. the fact that these approaches were not appreciably better than the "nearest neighbor" approach, the fact that deletion of their candidate gene in mouse resulted in the opposite phenotype from the one predicted. In addition, their explanations for these issues were clear and rational. Overall, the manuscript is interesting and informative and, as such, will be useful for researchers in bone biology as well as for many readers interested in the challenge of candidate gene identification from human GWAS.

Reviewer #2 (Recommendations for the authors):

I have a few comments for the authors:

"Most loci overlapped one gene (mean = 1.7, median = 1); however, 184 loci overlapped multiple genes, including a locus on Chromosome (Chr.) 20 (lead SNP rs6142137) which contained 9 prioritized genes. (Figure 1D)."

1. Figure 1D does not show the respective locus with 9 prioritized genes. It would be helpful to have Figure1E showing the Associated genomic regions and highlighting the 9 prioritized genes as well as what tissues each of them expressed.

"To do so, we created a database-curated set of genes previously implicated in the regulation of bone processes (henceforth referred to as our "known bone genes" list, N=1,399, Supplementary Table 5)."

2. What database was used to annotate the genes as "known bone genes". Could the authors please clarify? Also, why osteoblast transcriptome was not used here?

"we quantified the number of genes that were both the closest genes to eBMD GWAS associations and were members of the "known bone gene" list. "

3. Please provide additional information on what is considered the closest gene (in Mb).

4. Please provide diagrams showing the different steps of gene prioritization (TWAS+eQTL, "known bone genes" within 1Mb) versus (closest gene + "known bone genes") and their overlap ("Of our 512 prioritized genes, 206 (40%) were also the closest genes to eBMD GWAS associations, with 27 of the remaining 306 prioritized genes (8.8%) being members of the "known bone gene" list").

"Together, these data suggest that many of the genes identified by the combined TWAS/colocalization approach are likely causal BMD GWAS ge "

5. Incomplete phrase.

"An example of one of the 64 genes is GPATCH1, located within a GWAS association on human chromosome 19q13.11. "

6. Please provide a brief description of the gene GPATCH1. Any data showing expression of this gene in bone, other than the phenotype from IMPC?

7. In Figure 3A, could the authors please highlight where PPP6R3 is located?

"Together, these data highlight PPP6R3 as a strong candidate for one of the seven eBMD/fracture associations in this region."

8. Please provide a description of the gene PPP6R3 in the Results section as well.

9. rs10047483 is nearby LRP5, is this SNP in LD with the SNPs closest to LRP5? Do the authors suggest that the association to the LRP5 region is polygenic? In the discussion, could the authors provide further examples of complex BMD loci that could help to accept the also role of PPP6R3 in causality? (ex. WNT16 locus by Chesi et al. doi.org/10.1038/s41467-019-09302-x, another example is the FTO locus)

.

10. Figure 4E: Scales are on the bottom left, not right.

11. Would the lack of information to support Ppp6r3 in fracture risk, would be also due to the age at which the analysis of knockout mice was carried out? Could the authors provide a sentence or two to the discussion?

Reviewer #3 (Recommendations for the authors):

While this work makes a solid contribution, the significance is somewhat moderated by the application of established protocols to study eBMD, the limited analysis of 2 genes prioritized using this approach following their systemic ablation, the modest phenotypes of the mutant mice, and the lack of mechanistic studies on candidate gene function. Specifically, one gene was not studied in depth beyond the phenotype available in the IMPC. Deletion of Ppp6r3 causes modest reductions in vertebral BMD measurements by DXA and microCT, but the mechanisms by which the systemic ablation of the Ppp6r3 gene contributes to this mild phenotype are not explored. Additional analysis of Ppp6r3 function in bone cell lines or tissues would be especially important since this 'hit' arose using GTEx, which unfortunately does not include bone tissues and bone cells.

Other points for authors to consider:

The authors acknowledge that the Ppp6r3 variant is near Lrp5, and suggest in the Discussion that the Ppp6r3 variant association is independent of Lrp5. However, given the pivotal and well-known role of Lrp5 in BMD, it is important to document that the effects of the Ppp6r3 gene trap are not affecting Lrp5 expression or function.

Although there is a decrease in femoral width, there is no cortical microCT data for the femur. What might be the cause of this decrease in the width of the femur? Is this a developmental issue? Is there any change in the TRAP or P1NP levels in mutant mice?

It would be great to test whether deletion of PPP6R3 affects the mechanical properties of bone, whether in 3-point bending for the femurs or compression tests for the vertebrae. Does PPP6R3 deletion affect porosity to cause changes in BMD without affecting trabecular parameters?

What is the sex of the mice that were used in this study?

The authors performed pathway analysis for the potentially causal 512 BMD genes and showed GO terms. It would have been informative to include a gene network plot to visualize which genes are shared in these pathways focusing on bone-related pathways or a heatmap plot to show which genes are shared in the bone-related pathways. Additionally, does PPP6R3 belong to any bone-related pathway?

https://doi.org/10.7554/eLife.77285.sa1

Author response

Essential revisions:

The reviewers would like to see more data and discussion on the possible role of LRP5 in the bone phenotype of PPP6R3 mutant mice.

a. Please assess expression levels of Ppp6r3 and Lrp5 in your mutant model.

This is a great critique. We performed RNA-seq on RNA isolated from L5 vertebrae from wild-type and Ppp6r3 mutants. We choose vertebrae for this analysis given the stronger bone phenotype (Figure 4); however, we would expect similar results independent of the bone site analyzed. Ppp6r3 expression was the most significantly differentially expressed (P=3.67 x 10-104, log fold change = -1.63). As would be expected, it was downregulated in Ppp6r3 mutants (as described in Methods, the mice used in our analysis harbor a gene-trap allele, so while no protein was detected in homozygous mutant mice, transcript would be expected to be present at a low level as observed). We did not observe a difference in Lrp5 expression or any other gene in proximity (or on the same chromosome) of the gene-trap allele. We have added these data to the manuscript (pg. 8, line 294) and supplemental data file 1j.

b. Please include a more in-depth characterization of Ppp6r3 mutant mice:

femoral microCT data, P1NP or TRAP expression.

We agree these additional phenotypes should be addressed. We performed femoral microCT and found significant differences in Total Area and Medullary Area consistent with our caliper-based measures of femoral size. No differences were observed for other cortical or any of the trabecular (distal femur) bone parameters. Additionally, we measured P1NP levels in plasma and observed a significant increase in Ppp6r3 mutant mice, potentially suggesting high bone turnover in mutant mice. We did not have enough plasma to measure a marker of bone resorption such as TRAP or CTx. We have added these new data to the manuscript (pg. 9, line 336-342), Figure 4 and Figure 4 —figure supplement 2.

c. Please discuss whether rs10047483 is in LD with the SNP that is closest to LRP5.

Of the seven lead eBMD GWAS variants, rs10047483 (lead PPP6R3 eQTL variant) is only in LD with rs11228240 (r2=0.94) and accounts for the significant colocalization between the PPP6R3 eQTL and eBMD association. Rs10047483 is NOT in LD (r2<0.11) with any of the additional six eBMD lead GWAS variants, including the ones in close proximity to LRP5. These data are stated in the Results section of the manuscript (pg. 7, line 268-277).

Reviewer #2 (Recommendations for the authors):

I have a few comments for the authors:

"Most loci overlapped one gene (mean = 1.7, median = 1); however, 184 loci overlapped multiple genes, including a locus on Chromosome (Chr.) 20 (lead SNP rs6142137) which contained 9 prioritized genes. (Figure 1D)."

1. Figure 1D does not show the respective locus with 9 prioritized genes. It would be helpful to have Figure1E showing the Associated genomic regions and highlighting the 9 prioritized genes as well as what tissues each of them expressed.

We have added Figure 1 —figure supplement 1 which highlights the locus on Chr. 20. Supplementary file 1c contains the tissues, RCPs and TWAS pvals of the genes.

"To do so, we created a database-curated set of genes previously implicated in the regulation of bone processes (henceforth referred to as our "known bone genes" list, N=1,399, Supplementary Table 5)."

2. What database was used to annotate the genes as "known bone genes". Could the authors please clarify? Also, why osteoblast transcriptome was not used here?

Good point. We list the databases used in the Methods section entitled, “Creation of the “known bone gene” list”.

We wanted to use genes that had a functional connection to bone, which includes those active in multiple bone cell types and parameters. The bone transcriptome will just show expression in Obs but that doesnt really mean they are “Ob specific”.

"we quantified the number of genes that were both the closest genes to eBMD GWAS associations and were members of the "known bone gene" list. "

3. Please provide additional information on what is considered the closest gene (in Mb).

We used the closest gene list published in the Morris et al. BMD GWAS. For each of the 1103 associations identified, the closest gene was defined as the gene whose transcription start site was the shortest distance to the lead variant.

4. Please provide diagrams showing the different steps of gene prioritization (TWAS+eQTL, "known bone genes" within 1Mb) versus (closest gene + "known bone genes") and their overlap ("Of our 512 prioritized genes, 206 (40%) were also the closest genes to eBMD GWAS associations, with 27 of the remaining 306 prioritized genes (8.8%) being members of the "known bone gene" list").

Good idea. We added Figure 2 —figure supplement 1 which includes requested diagrams.

"Together, these data suggest that many of the genes identified by the combined TWAS/colocalization approach are likely causal BMD GWAS ge "

5. Incomplete phrase.

Thanks! This has been corrected.

"An example of one of the 64 genes is GPATCH1, located within a GWAS association on human chromosome 19q13.11. "

6. Please provide a brief description of the gene GPATCH1. Any data showing expression of this gene in bone, other than the phenotype from IMPC?

We now provide a brief description of GPATCH1 and cite studies demonstrating it is expressed in multiple bone cell-types.

7. In Figure 3A, could the authors please highlight where PPP6R3 is located?

This has been added to Figure 3A.

"Together, these data highlight PPP6R3 as a strong candidate for one of the seven eBMD/fracture associations in this region."

8. Please provide a description of the gene PPP6R3 in the Results section as well.

A brief description of PPP6R3 function has been added.

9. rs10047483 is nearby LRP5, is this SNP in LD with the SNPs closest to LRP5? Do the authors suggest that the association to the LRP5 region is polygenic? In the discussion, could the authors provide further examples of complex BMD loci that could help to accept the also role of PPP6R3 in causality? (ex. WNT16 locus by Chesi et al. doi.org/10.1038/s41467-019-09302-x, another example is the FTO locus)

Great points. With regards to the LD, as described above in response to essential revision #3, no, rs10047483 is not in LD with the BMD lead SNPs overlapping LRP5. Yes, we believe the data support a model of alleleic heterogeneity involving multiple genes (at least LRP5 and PPP6R3) within this locus. We have noted the similarity, in terms of complexity, between this locus and the Wnt16 locus in the discussion.

10. Figure 4E: Scales are on the bottom left, not right.

This has been corrected.

11. Would the lack of information to support Ppp6r3 in fracture risk, would be also due to the age at which the analysis of knockout mice was carried out? Could the authors provide a sentence or two to the discussion?

We did not directly measure biomechanical strength of bones from Ppp6r3 knockout mice; however this could be the focus of future research. We did measure bone matrix composition and, yes, we agree age could be a factor. We have added a brief note in the discussion related to this.

Reviewer #3 (Recommendations for the authors):

While this work makes a solid contribution, the significance is somewhat moderated by the application of established protocols to study eBMD, the limited analysis of 2 genes prioritized using this approach following their systemic ablation, the modest phenotypes of the mutant mice, and the lack of mechanistic studies on candidate gene function. Specifically, one gene was not studied in depth beyond the phenotype available in the IMPC. Deletion of Ppp6r3 causes modest reductions in vertebral BMD measurements by DXA and microCT, but the mechanisms by which the systemic ablation of the Ppp6r3 gene contributes to this mild phenotype are not explored. Additional analysis of Ppp6r3 function in bone cell lines or tissues would be especially important since this 'hit' arose using GTEx, which unfortunately does not include bone tissues and bone cells.

Other points for authors to consider:

The authors acknowledge that the Ppp6r3 variant is near Lrp5, and suggest in the Discussion that the Ppp6r3 variant association is independent of Lrp5. However, given the pivotal and well-known role of Lrp5 in BMD, it is important to document that the effects of the Ppp6r3 gene trap are not affecting Lrp5 expression or function.

Great point. As shown above in the response to essential revision #1, we did not observe a difference in expression of Lrp5 in vertbraes from Ppp6r3 mice.

Although there is a decrease in femoral width, there is no cortical microCT data for the femur. What might be the cause of this decrease in the width of the femur? Is this a developmental issue? Is there any change in the TRAP or P1NP levels in mutant mice?

We have added cortical microCT data of the femur and we did confirm the decrease in bone size. It is unclear if the phenotype is developmental or the mechanism. We also measured P1NP and, surprisingly, we observed a significant increase in mutant mice. In the context of bone mass, this may suggest decreased bone mass due to high turnover. We do want to stress that our goal for this project was using TWAS/colcoalzation to identify candidate genes and then provide additional support for select genes in the form of experimental validation. Now that we have identified Ppp6r3 and confirmed its involvement in bone, we have additional experiments planned to determine how Ppp6r3 impacts bone and all the points you bring up will be addressed.

It would be great to test whether deletion of PPP6R3 affects the mechanical properties of bone, whether in 3-point bending for the femurs or compression tests for the vertebrae. Does PPP6R3 deletion affect porosity to cause changes in BMD without affecting trabecular parameters?

This is a great point. Unfortunately, we don’t have bones that we could use to measure biomechanical strength and we didn’t measure porosity. Again, you bring up great points that will be critical to address to understand the mechanisms underlying the role of Ppp6r3 in bone that will be addressed in future studies.

What is the sex of the mice that were used in this study?

Thanks for catching this important omission! All studies included both male and female mice. We now specify this in the results, Figure 4 legend, and methods (under PPP6R3 functional validation).

The authors performed pathway analysis for the potentially causal 512 BMD genes and showed GO terms. It would have been informative to include a gene network plot to visualize which genes are shared in these pathways focusing on bone-related pathways or a heatmap plot to show which genes are shared in the bone-related pathways. Additionally, does PPP6R3 belong to any bone-related pathway?

We provide all 1140 nominally significant GO enrichments for the 512 genes in Supplementary file 1g, including which of the 512 genes belong to each enriched GO term. Unfortunately, we are not exactly sure what you mean by “network plot”?

No, PPP6R3 does not belong to any explicitly bone related gene ontologies.

https://doi.org/10.7554/eLife.77285.sa2

Article and author information

Author details

  1. Basel Maher Al-Barghouthi

    1. Center for Public Health Genomics, School of Medicine, University of Virginia, Charlottesville, United States
    2. Department of Biochemistry and Molecular Genetics, School of Medicine, University of Virginia, Charlottesville, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9816-8044
  2. Will T Rosenow

    Center for Public Health Genomics, School of Medicine, University of Virginia, Charlottesville, United States
    Contribution
    Formal analysis
    Competing interests
    No competing interests declared
  3. Kang-Ping Du

    Department of Radiation Oncology, University of Virginia, Charlottesville, United States
    Contribution
    Resources, Investigation
    Competing interests
    No competing interests declared
  4. Jinho Heo

    Department of Microbiology, Immunology, and Cancer Biology, School of Medicine, University of Virginia, Charlottesville, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  5. Robert Maynard

    Department of Orthopedics, Anschutz Medical Campus, University of Colorado, Aurora, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  6. Larry Mesner

    1. Center for Public Health Genomics, School of Medicine, University of Virginia, Charlottesville, United States
    2. Department of Public Health Sciences, School of Medicine, University of Virginia, Charlottesville, United States
    Contribution
    Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  7. Gina Calabrese

    Center for Public Health Genomics, School of Medicine, University of Virginia, Charlottesville, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  8. Aaron Nakasone

    Department of Mechanical Engineering, Boston University, Boston, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  9. Bhavya Senwar

    Department of Mechanical Engineering, University of Colorado Boulder, Boulder, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  10. Louis Gerstenfeld

    Department of Orthopaedic Surgery, Boston University Medical Center, Boston, United States
    Contribution
    Supervision, Funding acquisition, Writing – review and editing
    Competing interests
    No competing interests declared
  11. James Larner

    Department of Radiation Oncology, University of Virginia, Charlottesville, United States
    Contribution
    Resources, Supervision
    Competing interests
    No competing interests declared
  12. Virginia Ferguson

    Department of Mechanical Engineering, University of Colorado Boulder, Boulder, United States
    Contribution
    Supervision, Funding acquisition, Writing – review and editing
    Competing interests
    No competing interests declared
  13. Cheryl Ackert-Bicknell

    Department of Orthopedics, Anschutz Medical Campus, University of Colorado, Aurora, United States
    Contribution
    Supervision
    Competing interests
    Reviewing editor, eLife
  14. Elise Morgan

    Department of Mechanical Engineering, Boston University, Boston, United States
    Contribution
    Supervision, Funding acquisition, Writing – review and editing
    Competing interests
    No competing interests declared
  15. David Brautigan

    Department of Microbiology, Immunology, and Cancer Biology, School of Medicine, University of Virginia, Charlottesville, United States
    Contribution
    Resources, Supervision
    Competing interests
    No competing interests declared
  16. Charles R Farber

    1. Center for Public Health Genomics, School of Medicine, University of Virginia, Charlottesville, United States
    2. Department of Biochemistry and Molecular Genetics, School of Medicine, University of Virginia, Charlottesville, United States
    3. Department of Public Health Sciences, School of Medicine, University of Virginia, Charlottesville, United States
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Methodology, Writing – original draft, Project administration, Writing – review and editing
    For correspondence
    crf2s@virginia.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6748-4711

Funding

National Institute of Arthritis and Musculoskeletal and Skin Diseases (AR071657)

  • Louis Gerstenfeld

National Center for Research Resources (S10RR021072)

  • Jinho Heo
  • Elise Morgan

The funders had no role in study design, data collection, and interpretation, or the decision to submit the work for publication.

Acknowledgements

Research reported in this publication was supported in part by the National Institute of Arthritis and Musculoskeletal and Skin Diseases of the National Institutes of Health under Award Number AR071657 to CRF, LCG, and EFM, and by the National Center for Research Resources of the National Institutes of Health under Award Number S10RR021072 to EFM. BMA-B. was supported in part by a National Institutes of Health, Biomedical Data Sciences Training Grant (5T32LM012416). The authors acknowledge Wenhao Xu (University of Virginia) and the Genetically Engineered Mouse Models (GEMM) core for their technical assistance in generating the Ppp6r3 gene-trap mice. We thank Clifford J Rosen and Phuong T Le (MaineHealth Institute for Research) for performing the P1NP assay. We thank the IMPC for accessibility to BMD data in knockout mice (https://www.mousephenotype.org/). The data used for the analyses described in this manuscript were obtained from the IMPC SOLR database on 3/8/21. The Genotype-Tissue Expression (GTEx) Project was supported by the Common Fund of the Office of the Director of the National Institutes of Health, and by NCI, NHGRI, NHLBI, NIDA, NIMH, and NINDS. The data used for the analyses described in this manuscript were obtained from the GTEx Portal on 6/30/20.

Ethics

Our animal research protocol was reviewed and approved by the University of Virginia (UVA) Institutional Animal Care and Use Committee (IACUC), approved protocol #3741-12-20, titled 'Generation and characterization of mouse models of osteoporosis'. All experimental work was carried out under our approved UVA Institutional Biosafety Committee protocol #640-08.

Senior Editor

  1. Mone Zaidi, Icahn School of Medicine at Mount Sinai, United States

Reviewing Editor

  1. Nele Haelterman, Baylor College of Medicine, United States

Reviewer

  1. Erika Kague, University of Bristol, United Kingdom

Publication history

  1. Preprint posted: October 13, 2021 (view preprint)
  2. Received: January 24, 2022
  3. Accepted: November 16, 2022
  4. Version of Record published: November 23, 2022 (version 1)

Copyright

© 2022, Al-Barghouthi et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

Metrics

  • 63
    Page views
  • 9
    Downloads
  • 0
    Citations

Article citation count generated by polling the highest count across the following sources: Crossref, PubMed Central, Scopus.

Download links

A two-part list of links to download the article, or parts of the article, in various formats.

Downloads (link to download the article as PDF)

Open citations (links to open the citations from this article in various online reference manager services)

Cite this article (links to download the citations from this article in formats compatible with various reference manager tools)

  1. Basel Maher Al-Barghouthi
  2. Will T Rosenow
  3. Kang-Ping Du
  4. Jinho Heo
  5. Robert Maynard
  6. Larry Mesner
  7. Gina Calabrese
  8. Aaron Nakasone
  9. Bhavya Senwar
  10. Louis Gerstenfeld
  11. James Larner
  12. Virginia Ferguson
  13. Cheryl Ackert-Bicknell
  14. Elise Morgan
  15. David Brautigan
  16. Charles R Farber
(2022)
Transcriptome-wide association study and eQTL colocalization identify potentially causal genes responsible for human bone mineral density GWAS associations
eLife 11:e77285.
https://doi.org/10.7554/eLife.77285

Further reading

    1. Computational and Systems Biology
    2. Neuroscience
    Sergio Oscar Verduzco-Flores, Erik De Schutter
    Research Article Updated

    How dynamic interactions between nervous system regions in mammals performs online motor control remains an unsolved problem. In this paper, we show that feedback control is a simple, yet powerful way to understand the neural dynamics of sensorimotor control. We make our case using a minimal model comprising spinal cord, sensory and motor cortex, coupled by long connections that are plastic. It succeeds in learning how to perform reaching movements of a planar arm with 6 muscles in several directions from scratch. The model satisfies biological plausibility constraints, like neural implementation, transmission delays, local synaptic learning and continuous online learning. Using differential Hebbian plasticity the model can go from motor babbling to reaching arbitrary targets in less than 10 min of in silico time. Moreover, independently of the learning mechanism, properly configured feedback control has many emergent properties: neural populations in motor cortex show directional tuning and oscillatory dynamics, the spinal cord creates convergent force fields that add linearly, and movements are ataxic (as in a motor system without a cerebellum).

    1. Computational and Systems Biology
    2. Immunology and Inflammation
    Mingyao Pan, Bo Li
    Short Report Updated

    T cells are potent at eliminating pathogens and playing a crucial role in the adaptive immune response. T cell receptor (TCR) convergence describes T cells that share identical TCRs with the same amino acid sequences but have different DNA sequences due to codon degeneracy. We conducted a systematic investigation of TCR convergence using single-cell immune profiling and bulk TCRβ-sequence (TCR-seq) data obtained from both mouse and human samples and uncovered a strong link between antigen-specificity and convergence. This association was stronger than T cell expansion, a putative indicator of antigen-specific T cells. By using flow-sorted tetramer+ single T cell data, we discovered that convergent T cells were enriched for a neoantigen-specific CD8+ effector phenotype in the tumor microenvironment. Moreover, TCR convergence demonstrated better prediction accuracy for immunotherapy response than the existing TCR repertoire indexes. In conclusion, convergent T cells are likely to be antigen-specific and might be a novel prognostic biomarker for anti-cancer immunotherapy.