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

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.

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. 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.

Figure 1—figure supplement 1. Chromosome 20 contains a locus with nine prioritzed genes.

Transcriptome-wide association study (TWAS)/colocalization plot for genes in the chromosome 20 locus (lead (single nucleotide polymorphism)SNP rs6142137). 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) are shown. Genes alternate in color for visual clarity. Traingles represent the nine prioritized genes in the locus.

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.

Gene	Tissue with greatest RCP	Max. RCP	TWAS p value (Bonferroni)
SPTBN1	Cells_Cultured_Fibroblasts	0.9469	<5 × 10−324
CCDC170	Spleen	0.6582	<5 × 10−324
FAM3C	Artery_Tibial	0.4917	<5 × 10−324
SEPT5	Skin_Sun_Exposed	0.4868	2.26 × 10−286
FGFRL1	Cells_Cultured_Fibroblasts	0.1611	5.31 × 10−272
GREM2	Cells_Cultured_Fibroblasts	0.9998	4.31 × 10−257
GPATCH1	Whole_Blood	0.3564	3.44 × 10−226
RHPN2	Pituitary	0.2181	8.71 × 10−221
BMP4	Brain_Cortex	0.5468	5.49 × 10−169
RUNX2	Esophagus_Gastroesophageal_Junction	0.2372	1.99 × 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. 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.

Figure 2—figure supplement 1. Schematic of prioritization pipeline.

(A) Venn diagram showing the overlaps in data in the general gene prioritization pipeline. Overlapping transcriptome-wide association study (TWAS)/colocalization genes are termed ‘prioritized genes’. (B) Venn diagram showing overlap between the significant (p ≤ 0.05) S-MultiXcan TWAS genes and the prioritization datasets. (C) Venn diagram showing overlap between the fastENLOC colocalizing genes and the prioritization datasets.For all data, the black values indicate values obtained when a fastENLOC colocalization threshold of 0.1 is used, while the red values are those that are obtained when a fasENLOC colocalization threshold of 0.5 is used.

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.

Gene	Tissue with greatest RCP	Max. RCP	TWAS p value (Bonferroni)	# GTEx tissues with RCP ≥0.5
SPTBN1	Cells_Cultured_fibroblasts	0.9469	<5 × 10−324	2
PPP6R3	Thyroid	0.5291	5.7 × 10−93	1
BARX1	Colon_Transverse	0.7764	6.36 × 10−63	1
MEOX2	Brain_Nucleus_accumbens_basal_ganglia	0.6286	3.21 × 10−53	3
RERE	Adipose_Subcutaneous	0.6431	6.95 × 10−46	4
SIPA1	Nerve_Tibial	0.9981	4.26 × 10−41	1
CAPZB	Testis	0.6716	3.64 × 10−33	1
B4GALNT3	Artery_Aorta	0.9241	2.67 × 10−33	4
TRPC4AP	Breast_Mammary_Tissue	0.5577	8.62 × 10−31	3
AXL	Minor_Salivary_Gland	0.6205	9.74 × 10−31	3

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.

Figure 3. 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 (Ppp6r3^{tm1a(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 (p_adj = 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 (p_adj = 0.99, log₂ fold change = −0.068), a well-known regulator of bone mass just upstream of Ppp6r3 (Supplementary file 1j).

Figure 4. 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.

Figure 4—figure supplement 1. LSBMD and FNBMD genome-wide association study (GWAS) SNPs in the PPP6R3 locus.

(A) GWAS SNPs for LSBMD in the PPP6R3 locus. SNPs in red are significant PPP6R3 expression quantitative trait loci (eQTL) in thyroid tissue. The dashed line represents the genome-wide significance level (p value = 5 × 10⁻⁸). (B) GWAS SNPs for FNBMD in the PPP6R3 locus. SNPs in red are significant PPP6R3 eQTLin thyroid tissue. The dashed line represents the genome-wide significance level (p value = 5 × 10⁻⁸). (C) Mirroplot of LSBMD SNPs and PPP6R3 eQTL SNPs are colored by their linkage disequilibrium (LD) with rs10047483 (purple), the most significant PPP6R3 eQTL in thyroid. In the LSBMD panel, the most significant SNP is highlighted in purple, as rs10047483 was not assayed. (D) Mirroplot of FNBMD SNPs and PPP6R3 eQTL SNPs are colored by their LD with rs10047483 (purple), the most significant PPP6R3 eQTL in thyroid. In the FNBMD panel, the most significant SNP is highlighted in purple, as rs10047483 was not assayed.

Figure 4—figure supplement 2. Ppp6r3 functional validation.

(A) Weight. (B) Anterior–posterior (AP) femoral width. (C) Medial–lateral (ML) femoral width. (D) Femoral length (FL). (E) Lumbar spine tissue mineral density (TMD). (F) Lumbar spine trabecular separation (TbSp). (G) Lumbar spine trabecular thickness (TbTh). (H) Lumbar spine trabecular number (TbN). (I) P1NP serum concentration in sex-combined samples. (J) P1NP serum concentration in male samples. (K) P1NP serum concentration in female samples. Panels (E–H) are measured by micro-computed tomography (μCT). In panels A–H, least-square means are plotted. p values are contrast p values, adjusted for multiple comparisons. In panels I–K, p values are calculated by t-test. Asterisks represent significance (p ≤ 0.05).

Figure 4—figure supplement 3. Raman spectroscopy in femur, means.

(A–C) Least-square means for mean(mineral:matrix) in sex-combined, female, and male samples, respectively. (D–F) Least-square means for mean(carbonate:phosphate) in sex-combined, female, and male samples, respectively. (G–I) Least-square means for mean(crystallinity) in sex-combined, female, and male samples, respectively. Contrast p values, adjusted for multiple comparisons, are presented.

Figure 4—figure supplement 4. Raman spectroscopy in spines, means.

(A–C) Least-square means for mean(mineral:matrix) in sex-combined, female, and male samples, respectively. (D–F) Least-square means for mean(carbonate:phosphate) in sex-combined, female, and male samples, respectively. (G–I) Least-square means for mean(crystallinity) in sex-combined, female, and male samples, respectively. Contrast p values, adjusted for multiple comparisons, are presented.

Figure 4—figure supplement 5. Raman spectroscopy in femurs, standard deviation.

(A–C) Least-square means for sd(mineral:matrix) in sex-combined, female, and male samples, respectively. (D–F) Least-square means for sd(carbonate:phosphate) in sex-combined, female, and male samples, respectively. (G–I) Least-square means for sd(crystallinity) in sex-combined, female, and male samples, respectively. Contrast p values, adjusted for multiple comparisons, are presented.

Figure 4—figure supplement 6. Raman spectroscopy in spines, standard deviation.

(A–C) Least-square means for sd(mineral:matrix) in sex-combined, female, and male samples, respectively. (D–F) Least-square means for sd(carbonate:phosphate) in sex-combined, female, and male samples, respectively. (G–I) Least-square means for sd(crystallinity) in sex-combined, female, and male samples, respectively. Contrast p values, adjusted for multiple comparisons, are presented.

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 Ppp6r3^{tm1a(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 3–6).

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 (Ppp6r3^{tm1a(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 Ppp6r3^{tm1a(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-Tyr^c-Brd/BrdCrCrl (Charles River, Wilmington, MA) eight-cell stage embryos to create chimeric mice. Resultant chimeras were bred to B6N-Tyr^c-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).

Funding Statement

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

Funding Information

This paper was supported by the following grants:

• National Institute of Arthritis and Musculoskeletal and Skin Diseases AR071657 to Louis Gerstenfeld.

• National Center for Research Resources S10RR021072 to Jinho Heo, Elise Morgan.

Additional files

Supplementary file 1. Supplementary data files associated with this manuscript.

(a) 2156 protein-coding genes that are significant by S-MultiXcan transcriptome-wide association study (TWAS) analysis (Bonferroni p value ≤0.05). Columns are Ensembl ID, gene name, the nominal S-MultiXcan p value, the number of tissues available to S-MultiXcan, the number of independent components of variation among the tissues, and the Bonferroni-adjusted p value. (b) 1182 protein-coding genes that are significant in the fastEnloc colocalization analysis (regional colocalization probability [RCP] ≥0.1). Columns are Ensembl ID, signal cluster name from the expression quantitative trait loci (eQTL) analysis, the gene name, the RCP, and the colocalizing Genotype-Tissue Expression (GTEx) tissue. (c) The 512 protein-coding genes that are significant by both TWAS and colocalization (Bonferroni p value ≤0.05 and RCP ≥0.1). Columns are Ensembl ID, signal cluster name from the eQTL analysis, gene name, the RCP, the colocalizing GTEx tissue, the nominal S-MultiXcan p value, and the Bonferroni-adjusted p value. (d) Number of the 512 significantly colocalizing genes per GTEx tissue. Columns are the GTEx tissue and the number of unique colocalizing eQTL in the relevant tissue. (e) The ‘known bone gene’ list. Columns are gene name and Ensembl ID. (f) The 66 genes that are significant by both TWAS and colocalization (Bonferroni p value ≤0.05 and RCP ≥0.1), and are also members of the ‘known bone gene’ list. Columns are gene name and Ensembl ID. (g) Gene Ontology (GO) enrichments for the 512 protein-coding genes that are significant by both TWAS and colocalization (Bonferroni p value ≤0.05 and RCP ≥0.1). Only GO enrichments with a p value ≤0.05 are included. Columns are the GO IDs, GO terms, p values, the GO subontology (BP – Biological Process, CC – Cellular Component, MF – Molecular Function), and the Ensembl IDs for the genes that are members of the GO ontology. p values were calculated using a one-sided Fisher’s exact test, and were not adjusted for multiple comparisons. (h) The 863 genes closest to estimated bone mineral density (eBMD) genome-wide association study (GWAS) associations. These data were obtained from Morris et al., 2019. (i) 137 novel putatively causal BMD genes, after increasing RCP ≥0.5, and removing genes that were members of the ‘known bone gene’ list and genes with a nominal (p ≤ 0.05) alteration in BMD as determined by the International Mouse Phenotype Consortium (IMPC). Columns are Gene name, Ensembl ID, colocalization RCP, the tissue with the highest RCP, the S-Multixcan Bonferroni-adjusted p value, and the number of tissues that were significantly colocalizing. Note that TLN2 appears twice due to having the same RCP in two tissues. (j) Results of the differential expression analysis performed on RNA isolated from vertebrae. The analysis results show the difference in expression between mutant and wild-type mice. The columns are the Ensembl IDs, gene names, baseMean (the average of normalized count values, taken over all samples), log2 fold change, standard error of the log2 fold change estimate, the Wald statistic, the test p value, and the BH-adjusted p value.

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 datasets were used:

Morris JA. 2018. UK Biobank eBMD and Fracture GWAS Data Release 2018. GEFOS. release-2018

Estrada K. 2012. Data Release 2012. GEFOS. release-2012

Pividori M. 2020. PhenomeXcan. hakyimlab. phenomexcan

GTEx Consortium 2020. GTEx V8. gtexportal. V8

IMPC Consortium 2021. IMPC. mousephenotype. mousephenotype