αKlotho Regulates Age-Associated Vascular Calcification and Lifespan in Zebrafish

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aKlotho Regulates Age-Associated Vascular Calcification and Lifespan in Zebrafish Graphical Abstract

Authors Ajeet Pratap Singh, Maria X. Sosa, Jian Fang, ..., Samuel M. Cadena, Mark C. Fishman, David J. Glass

Correspondence [email protected]

In Brief aKlotho regulates mineral homeostasis and affects lifespans in mammals. Singh et al. show that a loss of aklotho in zebrafish results in reduced lifespans and vascular calcification in the outflow tract of the heart. Vascular calcification is associated with an upregulation of bone remodeling pathways and osteoclast differentiation.

Highlights d

Zebrafish aklotho mutants display reduced lifespans

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The aklotho phenotype occurs later in zebrafish than in mice

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Zebrafish aklotho mutants display adult-onset vascular calcification Calcification coincides with an increase in osteoclast differentiation pathways

Singh et al., 2019, Cell Reports 28, 2767–2776 September 10, 2019 ª 2019 Novartis Institutes for Biomedical Research. https://doi.org/10.1016/j.celrep.2019.08.013

Cell Reports

Report aKlotho Regulates Age-Associated Vascular Calcification and Lifespan in Zebrafish Ajeet Pratap Singh,1 Maria X. Sosa,1 Jian Fang,1 Shiva Kumar Shanmukhappa,2 Alexis Hubaud,1 Caroline H. Fawcett,1 Gregory J. Molind,1 Tingwei Tsai,1 Paola Capodieci,3 Kristie Wetzel,3 Ellen Sanchez,1 Guangliang Wang,1 Matthew Coble,1 Wenlong Tang,1 Samuel M. Cadena,5 Mark C. Fishman,4 and David J. Glass1,5,6,* 1Zebrafish Group, Chemical Biology and Therapeutics, Novartis Institutes for Biomedical Research, 181 Massachusetts Avenue, Cambridge, MA 02139, USA 2Preclinical Safety, Novartis Institutes for Biomedical Research, 250 Massachusetts Avenue, Cambridge, MA 02139, USA 3DAx/Discovery and Translational Pharmacology, Novartis Institutes for Biomedical Research, 181 Massachusetts Avenue, Cambridge, MA 02139, USA 4Harvard Department of Stem Cell and Regenerative Biology, Harvard University, 7 Divinity Ave, Cambridge, MA 02138, USA 5Age-Related Disorders Group, Chemical Biology and Therapeutics, Novartis Institutes for Biomedical Research, 181 Massachusetts Avenue, Cambridge, MA 02139, USA 6Lead Contact *Correspondence: [email protected] https://doi.org/10.1016/j.celrep.2019.08.013

SUMMARY

The hormone aKlotho regulates lifespan in mice, as knockouts die early of what appears to be accelerated aging due to hyperphosphatemia and soft tissue calcification. In contrast, the overexpression of aKlotho increases lifespan. Given the severe mouse phenotype, we generated zebrafish mutants for aklotho as well as its binding partner fibroblast growth factor-23 (fgf23). Both mutations cause shortened lifespan in zebrafish, with abrupt onset of behavioral and degenerative physical changes at around 5 months of age. There is a calcification of vessels throughout the body, most dramatically in the outflow tract of the heart, the bulbus arteriosus (BA). This calcification is associated with an ectopic activation of osteoclast differentiation pathways. These findings suggest that the gradual loss of aKlotho found in normal aging might give rise to ectopic calcification. INTRODUCTION Systemic factors that regulate aging are of interest due to their potential as novel drug targets in preventing or slowing down age-related decline in animal health. aKlotho, a molecular scaffold protein, is considered an anti-aging hormone that regulates mineral homeostasis in mammals (Chen et al., 2018; Kuro-o, 2013; Kuro-o et al., 1997; Kurosu et al., 2006, 2005; Lindberg et al., 2014; Shimada et al., 2004). It is one of the few systemic secreted factors whose loss is sufficient to induce premature morbidity and mortality that resembles accelerated aging (Kuro-o et al., 1997), and its overexpression extends lifespans (Kurosu et al., 2005). It is therefore of interest to understand

the cellular and molecular mechanisms by which aKlotho regulates the aging process. The mouse knockout models of aklotho are difficult to study; animals die by
812 weeks of age and are difficult to maintain (Ferna´ndez et al., 2018; Kuro-o et al., 1997; unpublished data). aklotho loss-of-function mice develop normally until about 3 or 4 weeks of age and then begin to display age-related conditions, including ectopic calcification, arteriosclerosis, osteoporosis, and reduced lifespans (Kuro-o et al., 1997). It was suggested that the extended lifespan in mice overexpressing aklotho is due to a suppression of insulin and the insulin-like growth factor-1 signaling (Kurosu et al., 2005), although it is likely that other pathways are involved. Recently, it was shown that an increase in autophagy levels could delay or prevent early mortality in aklotho mutant mice (Ferna´ndez et al., 2018). aKlotho acts as a co-receptor for fibroblast growth factor-23 (FGF23) (Urakawa et al., 2006). Knockouts of fgf23 have a similar accelerated aging phenotype to that of aklotho and show a perturbation in vitamin D metabolism (Shimada et al., 2004). aKlotho can also be released from the cell surface. In such settings, it can still heterodimerize with FGF23, functioning as a co-ligand in forming a high-affinity activator of FGF receptor signaling (Erben, 2018). Fish diverged from tetrapods approximately 400 million years ago (Daeschler et al., 2006; Romer, 1967). There are fundamental differences in physiology between fish and terrestrial vertebrates owing to unique demands of aquatic versus terrestrial environments. In terms of renal function and mineral and fluid homeostasis, they have evolved different physiologies. For example, renal function in freshwater fish serves partly to prevent overhydration, whereas in mammals, it is designed to prevent dehydration. We therefore examined if the roles of aKlotho, a renal hormone, would be conserved in aging and mineral homeostasis. The zebrafish also provide a tractable system for measuring certain behaviors, including physical activity. The zebrafish genome encodes one aklotho and one fgf23 (Mangos et al., 2012; Sugano and Lardelli, 2011). Consistent with

Cell Reports 28, 2767–2776, September 10, 2019 ª 2019 Novartis Institutes for Biomedical Research. 2767 This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

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mammalian studies, zebrafish aklotho expression is detected in multiple organs, including adult kidneys (Mangos et al., 2012). Zebrafish fgf23 is expressed in corpuscles of Stannius, a teleost-specific, kidney-associated endocrine gland involved in mineral homeostasis (Elizondo et al., 2010; Mangos et al., 2012). We generated knockouts of aklotho and fgf23 in zebrafish in order to understand the mechanistic link between aging and the aKlotho/FGF23 pathway. RESULTS Zebrafish Mutants in Both aklotho and fgf23 Have EarlyOnset Mortality We targeted the aklotho gene using the CRISPR/Cas9 method (Irion et al., 2014), generating mutations in two background ze€ and AB, and identified aklotho alleles carrying brafish strains, Tu frameshift mutations and early stop codons (Table S1). We also targeted fgf23 because the function of aKlotho in mammals depends in large part upon its binding to FGF receptors and recruiting FGF23 to activate FGF signaling (Chen et al., 2018; Kurosu et al., 2006). We find that aklotho/ and fgf23/ mutant zebrafish display essentially indistinguishable phenotypes (Figure 1). As adults of about 5 months of age, they develop emaciated bodies, tattered fins, and an opaque overgrowth on the eyes (aklotho/ mutant; Figures 1A, 1B, 1D, and 1E; fgf23/ mutant; Figures 1C, 1F, and S1A–S1F). In addition, female aklotho/ and fgf23/ mutants displayed protruding eyes (Figures S1G–S1L). The onset of mortality in mutant colonies began around 45 months post-fertilization (mpf; survival curves in Figures 1G–1J; p < 0.001), compared to wild-type strains of zebrafish, which live for 35 years (Carneiro et al., 2016; Gerhard et al., 2002). Both aklotho/ and fgf23/ mutant fish appear morphologically comparable to wild-type siblings at 23 mpf (Figures S1M–S1P) and are fertile as young adults, allowing us to breed homozygotes. Among adult progeny (
3 mpf) obtained by inbreeding aklotho heterozygotes, we recovered homozygous mutants in ratios consistent with Mendelian inheritance. Among 281 siblings raised together until adulthood, we obtained 70 wild-type siblings (25%), 130 heterozygous siblings (46%), and 79 homozygous aklotho mutants (28%). Among 241 adult zebrafish obtained from breeding parents fgf23 heterozygotes, we obtained 66 wild-type siblings (27%), 128 heterozygous siblings (52%), and 50 homozygous mutants (20%). This indicates that aklotho/ and fgf23/ mutants have no survival disadvantage until adulthood, even when raised with wild-type siblings. In order to probe the timing of more subtle aspects of physical decline, we analyzed the behavior of the aklotho/ and fgf23/

zebrafish in two settings: a circular arena (Figure 2K) and an arena resembling their home tank (Figure 2L). Although spontaneous behavior in zebrafish is intrinsically variable, aklotho/ and fgf23/ mutants demonstrated reduced activity in both behavioral paradigms at 5 and 6 mpf, corroborating the physical evidence of decline at this time (Figures 1M–1P). We conclude from these data that there is an adult-onset, agerelated decline in the body condition in both aklotho/ and fgf23/ mutants. Although it is difficult to align developmental frameworks between species, the adult-onset decline in both aklotho/ and fgf23/ mutant zebrafish appears to be proportionally later than described for the mouse aklotho mutants (Kuro-o et al., 1997; unpublished data). Vascular Calcification and Inflammation across Organs in aklotho/ Mutant Zebrafish In order to understand the phenotype at the cell and tissue level, we performed comprehensive histopathological analysis by H&E staining on sections of 5-month-old wild-type and aklotho/ mutant fish (Figure 2; N = 3 males each). In aklotho/ fish, there was widespread calcification and inflammation. Within the integument of aklotho/ fish, there was a reduction in the number of mucosal cells and necrosis in areas of the epidermis and dermis, with a mineralization of the dermal vasculature (Figure 2A). Furthermore, in aklotho/ mutants, there was calcification of medium- to small-size blood vessels in the skeletal muscles (arrow in Figure 2B), accompanied by a degeneration and fibrosis of adjacent skeletal muscles with immune cell infiltration (Figure 2B). Mineralization was often in a concentric pattern in the affected areas. The gill arch demonstrated bone overgrowth (hyperostosis) with chondrodysplasia of the gill arch (Figure 2C) and a loss of normal architecture of filaments and lamellae due to blunting, fusion, and necrosis of the lamellae epithelium, along with immune cell infiltration. In the aklotho/ zebrafish, calcification was particularly striking within the walls of the bulbus arteriosus (BA) (the outflow tract of the heart) (Figures 2D and 2E). The BA is composed of smooth muscles and is lined by the endothelium, and its elasticity is believed to buffer pulsatile blood flow to the thin-walled capillaries of the gills (Farrell, 1979; Grimes and Kirby, 2009). To validate calcification in the BA, we used alizarin red, a stain for calcification (Walker and Kimmel, 2007). The BA in aklotho/ mutants is prominently stained with alizarin red in contrast to wild-type animals (Figure 2D; insets), confirming calcification. Calcification was also observed in the bile duct of the livers of aklotho/ fish (arrow in Figure 2F). The kidneys of aklotho/ mutants appeared comparable to the wild-type controls (Figure 2G). Within the skeletal system, there were multifocal areas of hyperostosis and

Figure 1. Zebrafish aklotho and fgf23 Mutants

€ wild-type strain, (B) aklotho (klD5), and (C) fgf23 (fgf23ins1) mutant in Tu € background; (D) AB (A–F) Body condition of aklotho and fgf23 mutant males at 5 mpf: (A) Tu wild-type strain, (E) aklotho (klD5), and (F) fgf23 (fgf23D11) mutant in AB background. (G–J) Survival curves for (G) aklotho (n = 36 background controls, 32 mutants; p < 0.0001) and (H) fgf23 mutants (n = 14 wild-type siblings, 14 mutants; p < 0.0001) € background and for (I) aklotho (n = 24 wild-type siblings, 21 mutants; p = 0.0001) and (J) fgf23 mutants (n = 60 background controls, 68 mutants; p < 0.0001) in Tu in AB background. Log-rank (Mantel-Cox) test for statistical analysis on survival curves in GraphPad Prism. (K and L) Analysis of speed (cm) in the (K) circular arena and (L) home-tank arena. Age (m; mpf) on x axis; n = number of fish. (M and N) aklotho mutants and wild-type controls in (M) circular and (N) home-tank arena. (O and P) fgf23 mutants and wild-type controls in (O) circular and (P) home-tank arena. Statistical analysis using unpaired t test in GraphPad Prism. See also Figure S1 and Table S1.

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Figure 2. Vascular Calcification and Inflammation in aklotho Mutants H&E staining on paraffin sections from 5-month-old €) and aklotho (klD5) males. wild-type control (Tu Shown are (A) skin (arrows indicate mucous cells in wild type); (B) muscle (arrows indicate vascular calcification); (C) gills; (D) heart (arrow indicates calcification in the BA); (D0 and D00 ) alizarin redstained whole-mount hearts (arrows indicate the BA); (E) BA (arrow indicates calcification); (F) liver (arrows indicate bile-duct); and (G) kidney (arrows indicate glomeruli). (H) Bright-field images of 5-mpf wild-type (left) and klD5 (right) hearts. TRAP staining on (I) whole mount and (J) cryosection of 5-mpf hearts. See also Figure S2.

chondrodysplasia. These changes were prominent in the caudal region (Figure S2A). Frequently, regions of bone overgrowth were accompanied by areas of dystrophic calcification, connective tissue proliferation, and immune cell infiltration (Figure S2A). Vascular calcification was the most prominent phenotype. In fact, unprocessed and unstained BAs appear opaque white in aklotho/ (Figure 2H), indicating severe calcification. fgf23/ mutants phenocopy aklotho/ mutants—the BAs in fgf23/ mutants are prominently stained with alizarin red, in contrast to wild-type animals (Figure S2B). It has been shown that calcification in zebrafish is accompanied by an increase in osteoclast activity (Apschner et al., 2014). Consistent with this, we observe strong Tartrate-resistant acid phosphatase (TRAP) staining in the outflow tract of the aklotho/ mutant hearts (Figures 2I and 2J), indicating the presence of osteoclasts in the BA. Regulation of Osteogenesis in BAs of aklotho/ Mutants In order to understand the molecular mechanisms underlying ectopic calcification in aklotho/ mutants, we performed an 2770 Cell Reports 28, 2767–2776, September 10, 2019

RNA sequencing (RNA-seq) analysis of the kidney, heart (including BA), and gills at 3 mpf, when mutants appeared comparable phenotypically to wild-type siblings, and at 5 mpf, when aklotho/ mutants became phenotypically distinct from wild-type siblings (n = 8 animals per genotype and condition; 4 males and 4 females). In the wild-type siblings, aklotho expression is highest in the kidneys (Figure 3A[3]). As expected, a significant downregulation of aklotho expression is observed in aklotho/ mutants at both 3 and 5 mpf (Figure 3A[1]). In wildtype fish, fgf23 expression is detected in the kidneys and, surprisingly, in the gills (Figure 3B[2]). In aklotho/ mutant gills, fgf23 is the most significantly upregulated gene at 3 mpf, indicating a dysregulation of the aKlotho/FGF23 axis (Figure 3B[2]). At 3 mpf, there were only modest changes from the wild-type siblings in the aklotho/ patterns of gene expression in the kidney, heart, and gills (Figures 3C–3G; Table S2) and no statistically significant change at the pathway level (Figures 4A and S3A; Tables S3 and S4). At 5 mpf, in the kidneys, which were histologically indistinguishable from the wild type (Figure 2G), the main noticeable change was an upregulation of genes involved in the metabolism of the heme (Figure 4A). In the gills, at 5 mpf, a time of widespread disorganization based on histopathology, there was an increase in the expression of genes of the extracellular matrix (ECM) organization pathway (Figure 4A). There is minimal overlap between genes differentially regulated at 3 and 5 mpf in an organ (Figures S3B and S3C). Next, we analyzed tissue samples for pathway level changes in the transcriptome by a hypergeometric test (Figure 4A; Table S3) and gene set enrichment analysis by a weighted Kolmogorov-Smirnov test (Figure S3A; Table S4). Both tests revealed that at 5 mpf, multiple pathways were dysregulated in aklotho/ mutant tissues. In heart samples collected for this analysis, two out of eight hearts showed visual signs of calcification in the BA at 3 mpf. All eight hearts displayed calcification in

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the BA at 5 mpf, indicating an adult-onset progressive vascular calcification. At 3 mpf, we did not observe statistically significant changes in the pathway level analysis (Figure 4A). However, at the individual gene level, aklotho mutant hearts displayed an upregulation in genes involved in bone formation and remodeling, such as matrix metallopeptidase-9 (mmp9), mmp13, and osteopontin (secreted phosphoprotein 1/spp1) (Figures 4B, 4C, 4E, and S4A) (Page-McCaw et al., 2007; Standal et al., 2004). In mammals, mmp9 and mmp13 are required for the transition from cartilage into bone (Page-McCaw et al., 2007; Stickens et al., 2004). SPP1 is known to be an inhibitor of calcification (Standal et al., 2004). At 5 mpf, multiple pathways involved in bone formation, bone remodeling, osteoclast activity, ECM remodeling, and inflammation are upregulated in aklotho/ mutant hearts (Figure 4). spp1 is the most significantly upregulated gene in aklotho/ mutant hearts at 5 mpf (Figure 4E). We validated spp1 expression using qPCR on dissected BAs and observed an
600-fold enrichment in spp1 transcript in aklotho/ mutants at 5 mpf (Figure S4C). entpd5a (ectonucleoside triphosphate diphosphohydrolase 5a), an osteoblast marker in zebrafish (Huitema et al., 2012), is also upregulated in aklotho/ mutant hearts at this stage (Figure 4D). However, we do not observe an upregulation of the conventional markers of the osteoblast lineage in aklotho/ mutant hearts, including runx2, sp7, col10a1, and col1a2 (Huitema et al., 2012; Vijayakumar et al., 2013; Yang et al., 2011). The qPCR analysis for runx2a and runx2b on dissected BAs showed a modest increase in runx2b levels and no change in runx2a levels at 5 mpf (Figure S4C). Recent studies have identified a role for osteolectin/clec11a and integrin-a11/itga11 signaling in osteoblast differentiation and the maintenance of adult skeletal bone mass (Shen et al., 2019; Yue et al., 2016); both clec11a and itga11a are upregulated in aklotho/ mutant hearts at 5 mpf (Figure S4B). Thus, our analysis reveals an upregulation of genes involved in ECM remodeling and bone formation that could explain the observed ectopic vascular calcification. Calcification is remodeled and counter-regulated by the activity of hematopoietic stem cell-derived osteoclasts. The RANK/ RANKL/OPG pathway is required for osteoclast differentiation from hematopoietic lineage (Edwards and Mundy, 2011; Novack and Teitelbaum, 2008; Teitelbaum and Ross, 2003). In aklotho/  mutant hearts, key members of this pathway (Figure S4D) and osteoclast-enriched enzymes such as ctsk (encoding Cathepsin K) and acp5 (encoding TRAP) are upregulated at 5 mpf (Figures 4F and 4G), suggesting an increase in osteoclast activity in the ectopically mineralized region. In order to localize the ongoing transcriptional activity in the heart, we performed RNAscope analysis using spp1 probe; this analysis revealed a highly localized spp1 expression in the BAs of aklotho/ mutants (Figure 4H).

DISCUSSION The aKlotho/FGF23 pathway appears to play a role in the aging of zebrafish. Both aklotho/ and fgf23/ zebrafish display early-onset morbidity, beginning at about 4 or 5 months of age, accompanied by spinal deformities, loss of fin integrity, and widespread ectopic calcification, especially of the outflow tract of the heart. Soft-tissue calcification increases with advancing age in humans, and vascular calcification is associated with an increase in atherosclerosis and cardiovascular mortality (Leopold, 2013; McClelland et al., 2006; Shaw et al., 2015; Thompson et al., 2013). The mechanisms that lead to soft-tissue calcification remain poorly understood. It has been suggested (Hortells et al., 2017, 2018; Persy and D’Haese, 2009; Pillai et al., 2017), but debated (O’Neill and Adams, 2014), that cardiovascular calcification actually reflects the osteogenic cell fate change of vascular smooth muscle. Here, we find that in the absence of aKlotho/FGF23, vascular tissue in the BA changes its pattern of gene expression to resemble that of bone: a pro-osteogenic reorganization of the ECM may promote the observed calcification phenotype in smooth muscle cells of the BA. This leads to a surge in antiosteogenic mechanisms, including a local differentiation of osteoclasts. The effect of aKlotho/FGF23 signaling on vascular calcification is likely non-cell autonomous. aklotho is primarily expressed in kidneys, whereas fgf23 is expressed in kidneys and gills. Interestingly, a homozygous missense mutation of aKlotho was reported in a human; this mutation resulted in severe tumoral calcinosis including ectopic calcifications, indicating the zebrafish model is predictive of the human condition (Ichikawa et al., 2017). The BA is an elastic, valveless cardiac outflow tract in teleosts that is believed to act as a windkessel to protect the delicate gill vasculature from large variations of pressure generated by the ventricle (Farrell, 1979; Grimes and Kirby, 2009; Maldanis et al., 2016). The calcification of the BA may compromise its elasticity, leading to large fluctuations in blood pressure in the gill vasculature and a consequent loss of the gill architecture. Thus, vascular calcification may be the primary cause of the onset of morbidity in aklotho/ and fgf23/ zebrafish. This has striking parallels with chronic kidney disease in humans: vascular calcification is considered to contribute to mortality in patients with chronic kidney disease (Go et al., 2004; Mizobuchi et al., 2009), and aKlotho treatment has been shown to be helpful for the treatment of kidney disease in preclinical models (Doi et al., 2011; Hum et al., 2017; Shi et al., 2016). Taken together, the data suggest that one potential consequence of age-related decline in aKlotho, as has been reported in humans, could be inappropriate osteogenesis often observed in older vascular smooth muscles, along with the decline in cardiac

Figure 3. RNA-Seq Analysis of Kidney, Heart, and Gills in aklotho Mutants (A) aklotho expression in (A1) kidney, (A2) heart, and (A3) gills of wild-type siblings (red) and aklotho mutants (blue); 4 males and 4 females for each set. (B) fgf23 expression in (B1) kidney and (B2) gills of wild-type siblings (red) and aklotho mutants (blue); no expression detected in heart. (C–E1) Volcano plots showing differentially expressed genes at 3 and 5 mpf in (C and C1) kidney, (D and D1) heart, and (E and E1) gills. (F and G) Euler diagram, obtained by R package eulerr, showing the number of (F) upregulated and (G) downregulated genes by aklotho mutation in the kidney, heart, gills, and their overlaps at 3 and 5 mpf. The area of each disjointed shape is proportional to the number of its elements as marked. See also Figures S3 and S4 and Table S2.

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Figure 4. Pathway Enrichment Analysis of Differentially Upregulated Genes (A) Gene set enrichment analysis comparing upregulated pathways by aklotho mutation in the kidney, heart, and gills. Each row is a pathway, annotated on the right-hand side, and each column corresponds to the significance of the enrichment analysis for each tissue and adult stage (m3, 3 mpf; m5, 5 mpf). The colors from white to red represent the negative log10 adjusted p value from low to high. Only pathways that were enriched significantly (adjusted p value < 0.05) in at least one tissue were included. (B–G) Box plots showing select examples of genes upregulated at 5 mpf in aklotho mutant hearts (blue, kl) compared to wild-type (WT) siblings (red). Shown are (B) mmp9 (matrix metallopeptidase 9), (C) mmp13a (matrix metallopeptidase 13a), (D) entpd5a (ectonucleoside triphosphate diphosphohydrolase 5a), (E) spp1 (secreted phosphoprotein 1), (F) ctsk (cathepsin K), and (G) acp5a (acid phosphatase 5a, tartrate resistant). Age: 3 and 5 mpf. (H) RNAscope for spp1: (H1 and H2) WT control; (H3 and H4) aklotho (klD5) mutant hearts stained with DAPI (blue), and spp1 RNAscope probe (red). White lines outline the BA (arrow) and the blood vessel leading to gills. See also Figure S4 and Tables S3 and S4.

elasticity and function (de Carvalho Filho et al., 1996; Fleg and Strait, 2012; Hamczyk et al., 2018; McClelland et al., 2006; Semba et al., 2011a, 2011b; Shaw et al., 2015). Such findings could suggest particular therapeutic readouts of aKlotho supplementation in aged humans where aKlotho levels are low.

In vertebrates, calcium phosphate must be carefully regulated to avoid ectopic precipitation of calcium-phosphate crystals. It is suggested that a shift from a calcium carbonate-based skeleton in invertebrates to a calcium phosphate-based skeleton in vertebrates necessitated the evolution of mechanisms to regulate

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calcium phosphate homeostasis (Kuro-o and Moe, 2017). In mammals, aKlotho is highly expressed in the kidneys, whereas bones are a primary source of FGF23 (Kuro-o et al., 1997; Riminucci et al., 2003). Interestingly, although aklotho is expressed in zebrafish kidneys, fgf23 is expressed in corpuscles of Stannius, a teleost-specific, kidney-associated gland involved in mineral homeostasis (Elizondo et al., 2010; Mangos et al., 2012), and in the gills of adults (this study). The gills play a major role in mineral homeostasis—these are the primary site of calcium uptake in adult fish (Evans et al., 2005; Flik et al., 1985, 1995; Liao et al., 2007). In terrestrial vertebrates, the main source of calcium is food. It is absorbed primarily in the intestines and kidneys, and the bones serve as the major reservoir of the calcium. Taken together, this suggests that although the focus organs may have shifted during the evolutionary transition from aquatic to terrestrial life, the function of the aKlotho/FGF23 pathway in maintaining mineral homeostasis has been conserved. STAR+METHODS Detailed methods are provided in the online version of this paper and include the following: d d d d

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KEY RESOURCES TABLE LEAD CONTACT AND MATERIALS AVAILABILITY EXPERIMENTAL MODEL AND SUBJECT DETAILS B Zebrafish METHOD DETAILS B Lifespan Analysis B Behavioral Analysis B Histology B RNA-Seq Sample Preparation QUANTIFICATION AND STATISTICAL ANALYSIS B RNA-Seq Data Analysis DATA AND CODE AVAILABILITY

SUPPLEMENTAL INFORMATION Supplemental Information can be found online at https://doi.org/10.1016/j. celrep.2019.08.013. ACKNOWLEDGMENTS The study was funded by Novartis. We thank the Zebrafish Group, the AgeRelated Disorders Group, and the Chemical Biology and Therapeutics groups for their enthusiastic support; T. Shavlakadze and A. Jaffe for discussions; T. Scott and P. Stolyar for comments; O. Iartchouk and ASI for help with RNAseq; U. Plikat and NIBR Informatics for the RNA-seq analysis pipeline; G. Zhang, Z. Li, C. Russ, and the NGS facility for NGS; N. Kirkpatrick and the microscopy facility; and K. Maloney, F. Vetrano-Olsen, H. Clark, M.-K. Paulina, L. Ponczek, J. Tobin, V. Afere, A. Elliott, R. Brown, N. Jones-Bolduc, J. FremontRahl, E. Theve, and the laboratory animal services for zebrafish care. AUTHOR CONTRIBUTIONS Experiment Design, A.P.S., M.X.S., S.M.C., M.C.F., and D.J.G.; Experiment Execution, A.P.S., M.X.S., A.H., C.H.F., G.J.M., E.S., G.W., and M.C.; Generation and Characterization of the Knockouts, A.P.S. and M.X.S.; Histology, C.H.F., K.W., and P.C.; Analysis of Histopathology Slides, S.K.S. and A.P.S.; Sample Collection and RNA Preparation, M.X.S., A.P.S., and E.S.; RNA-seq Analysis, M.X.S. and J.F.; qPCR, A.H.; RNAscope, C.H.F.; Behavioral Data

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Collection, G.J.M. and A.P.S.; Behavioral Data Analysis, T.T. and W.T.; Manuscript Preparation, A.P.S. and D.J.G., with inputs from all authors. DECLARATION OF INTERESTS This study was funded by Novartis AG. All authors, except for M.C.F., were employees of Novartis at the time the study was conducted. Some authors, including D.J.G., own Novartis stock. M.C.F. is on the BOD of Semma Therapeutics and Beam Therapeutics, the SAB of Tenaya Therapeutics, and serves as advisor to MPM and Burrage Capital. Received: March 28, 2019 Revised: July 2, 2019 Accepted: July 31, 2019 Published: September 10, 2019 REFERENCES Apschner, A., Huitema, L.F., Ponsioen, B., Peterson-Maduro, J., and SchulteMerker, S. (2014). Zebrafish enpp1 mutants exhibit pathological mineralization, mimicking features of generalized arterial calcification of infancy (GACI) and pseudoxanthoma elasticum (PXE). Dis. Model. Mech. 7, 811–822. Carneiro, M.C., Henriques, C.M., Nabais, J., Ferreira, T., Carvalho, T., and Ferreira, M.G. (2016). Short Telomeres in Key Tissues Initiate Local and Systemic Aging in Zebrafish. PLoS Genet. 12, e1005798. Chen, G., Liu, Y., Goetz, R., Fu, L., Jayaraman, S., Hu, M.C., Moe, O.W., Liang, G., Li, X., and Mohammadi, M. (2018). a-Klotho is a non-enzymatic molecular scaffold for FGF23 hormone signalling. Nature 553, 461–466. Daeschler, E.B., Shubin, N.H., and Jenkins, F.A., Jr. (2006). A Devonian tetrapod-like fish and the evolution of the tetrapod body plan. Nature 440, 757–763. de Carvalho Filho, E.T., de Carvalho, C.A., and de Souza, R.R. (1996). Agerelated changes in elastic fibers of human heart. Gerontology 42, 211–217. Doi, S., Zou, Y., Togao, O., Pastor, J.V., John, G.B., Wang, L., Shiizaki, K., Gotschall, R., Schiavi, S., Yorioka, N., et al. (2011). Klotho inhibits transforming growth factor-beta1 (TGF-beta1) signaling and suppresses renal fibrosis and cancer metastasis in mice. J. Biol. Chem. 286, 8655–8665. Edwards, J.R., and Mundy, G.R. (2011). Advances in osteoclast biology: old findings and new insights from mouse models. Nat. Rev. Rheumatol. 7, 235–243. Elizondo, M.R., Budi, E.H., and Parichy, D.M. (2010). trpm7 regulation of in vivo cation homeostasis and kidney function involves stanniocalcin 1 and fgf23. Endocrinology 151, 5700–5709. Erben, R.G. (2018). a-Klotho’s effects on mineral homeostasis are fibroblast growth factor-23 dependent. Curr. Opin. Nephrol. Hypertens. 27, 229–235. Evans, D.H., Piermarini, P.M., and Choe, K.P. (2005). The multifunctional fish gill: dominant site of gas exchange, osmoregulation, acid-base regulation, and excretion of nitrogenous waste. Physiol. Rev. 85, 97–177. Ewels, P., Magnusson, M., Lundin, S., and Ka¨ller, M. (2016). MultiQC: summarize analysis results for multiple tools and samples in a single report. Bioinformatics 32, 3047–3048. Farrell, A.P. (1979). Wind-Kessel Effect of the Bulbus Arteriosus in Trout. J. Exp. Zool. 209, 169–173. Ferna´ndez, A.F., Sebti, S., Wei, Y., Zou, Z., Shi, M., McMillan, K.L., He, C., Ting, T., Liu, Y., Chiang, W.C., et al. (2018). Disruption of the beclin 1-BCL2 autophagy regulatory complex promotes longevity in mice. Nature 558, 136–140. Fleg, J.L., and Strait, J. (2012). Age-associated changes in cardiovascular structure and function: a fertile milieu for future disease. Heart Fail. Rev. 17, 545–554. Flik, G., Vanrijs, J.H., and Bonga, S.E.W. (1985). Evidence for High-Affinity Ca-2+-Atpase Activity and Atp-Driven Ca-2+-Transport in Membrane Preparations of the Gill Epithelium of the Cichlid Fish Oreochromis-Mossambicus. J. Exp. Biol. 119, 335–347.

Flik, G., Verbost, P., and Bonga, S. (1995). Calcium transport processes in fishesVolume 14 (Academic Press, Inc.).

Love, M.I., Huber, W., and Anders, S. (2014). Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550.

Gerhard, G.S., Kauffman, E.J., Wang, X., Stewart, R., Moore, J.L., Kasales, C.J., Demidenko, E., and Cheng, K.C. (2002). Life spans and senescent phenotypes in two strains of Zebrafish (Danio rerio). Exp. Gerontol. 37, 1055–1068.

Maldanis, L., Carvalho, M., Almeida, M.R., Freitas, F.I., de Andrade, J.A., Nunes, R.S., Rochitte, C.E., Poppi, R.J., Freitas, R.O., Rodrigues, F., et al. (2016). Heart fossilization is possible and informs the evolution of cardiac outflow tract in vertebrates. eLife 5, e14698.

Go, A.S., Chertow, G.M., Fan, D., McCulloch, C.E., and Hsu, C.Y. (2004). Chronic kidney disease and the risks of death, cardiovascular events, and hospitalization. N. Engl. J. Med. 351, 1296–1305. Grimes, A.C., and Kirby, M.L. (2009). The outflow tract of the heart in fishes: anatomy, genes and evolution. J. Fish Biol. 74, 983–1036. Gross-Thebing, T., Paksa, A., and Raz, E. (2014). Simultaneous high-resolution detection of multiple transcripts combined with localization of proteins in whole-mount embryos. BMC Biol. 12, 55. Hamczyk, M.R., del Campo, L., and Andre´s, V. (2018). Aging in the Cardiovascular System: Lessons from Hutchinson-Gilford Progeria Syndrome. Annu. Rev. Physiol. 80, 27–48. Hortells, L., Sosa, C., Guille´n, N., Lucea, S., Milla´n, A´., and Sorribas, V. (2017). Identifying early pathogenic events during vascular calcification in uremic rats. Kidney Int. 92, 1384–1394. Hortells, L., Sur, S., and St Hilaire, C. (2018). Cell Phenotype Transitions in Cardiovascular Calcification. Front. Cardiovasc. Med. 5, 27. Huitema, L.F., Apschner, A., Logister, I., Spoorendonk, K.M., Bussmann, J., Hammond, C.L., and Schulte-Merker, S. (2012). Entpd5 is essential for skeletal mineralization and regulates phosphate homeostasis in zebrafish. Proc. Natl. Acad. Sci. USA 109, 21372–21377. Hum, J.M., O’Bryan, L.M., Tatiparthi, A.K., Cass, T.A., Clinkenbeard, E.L., Cramer, M.S., Bhaskaran, M., Johnson, R.L., Wilson, J.M., Smith, R.C., and White, K.E. (2017). Chronic Hyperphosphatemia and Vascular Calcification Are Reduced by Stable Delivery of Soluble Klotho. J. Am. Soc. Nephrol. 28, 1162–1174. Ichikawa, S., Imel, E.A., Kreiter, M.L., Yu, X., Mackenzie, D.S., Sorenson, A.H., Goetz, R., Mohammadi, M., White, K.E., and Econs, M.J. (2007). A homozygous missense mutation in human KLOTHO causes severe tumoral calcinosis. J. Clin. Invest. 117, 2684–2691. €sslein-Volhard, C. (2014). Precise and efficient Irion, U., Krauss, J., and Nu genome editing in zebrafish using the CRISPR/Cas9 system. Development 141, 4827–4830. Kuro-o, M. (2013). Klotho, phosphate and FGF-23 in ageing and disturbed mineral metabolism. Nat. Rev. Nephrol. 9, 650–660. Kuro-o, M., and Moe, O.W. (2017). FGF23-aKlotho as a paradigm for a kidneybone network. Bone 100, 4–18. Kuro-o, M., Matsumura, Y., Aizawa, H., Kawaguchi, H., Suga, T., Utsugi, T., Ohyama, Y., Kurabayashi, M., Kaname, T., Kume, E., et al. (1997). Mutation of the mouse klotho gene leads to a syndrome resembling ageing. Nature 390, 45–51.

€ppner, H., Reiser, J., and Wolf, M. (2012). Mangos, S., Amaral, A.P., Faul, C., Ju Expression of fgf23 and aklotho in developing embryonic tissues and adult kidney of the zebrafish, Danio rerio. Nephrol. Dial. Transplant. 27, 4314–4322. McClelland, R.L., Chung, H., Detrano, R., Post, W., and Kronmal, R.A. (2006). Distribution of coronary artery calcium by race, gender, and age: results from the Multi-Ethnic Study of Atherosclerosis (MESA). Circulation 113, 30–37. Meeker, N.D., Hutchinson, S.A., Ho, L., and Trede, N.S. (2007). Method for isolation of PCR-ready genomic DNA from zebrafish tissues. Biotechniques 43, 610, 612, 614. Mizobuchi, M., Towler, D., and Slatopolsky, E. (2009). Vascular calcification: the killer of patients with chronic kidney disease. J. Am. Soc. Nephrol. 20, 1453–1464. Novack, D.V., and Teitelbaum, S.L. (2008). The osteoclast: friend or foe? Annu. Rev. Pathol. 3, 457–484. O’Neill, W.C., and Adams, A.L. (2014). Breast arterial calcification in chronic kidney disease: absence of smooth muscle apoptosis and osteogenic transdifferentiation. Kidney Int. 85, 668–676. Page-McCaw, A., Ewald, A.J., and Werb, Z. (2007). Matrix metalloproteinases and the regulation of tissue remodelling. Nat. Rev. Mol. Cell Biol. 8, 221–233. Persy, V., and D’Haese, P. (2009). Vascular calcification and bone disease: the calcification paradox. Trends Mol. Med. 15, 405–416. Pillai, I.C.L., Li, S., Romay, M., Lam, L., Lu, Y., Huang, J., Dillard, N., Zemanova, M., Rubbi, L., Wang, Y., et al. (2017). Cardiac Fibroblasts Adopt Osteogenic Fates and Can Be Targeted to Attenuate Pathological Heart Calcification. Cell Stem Cell 20, 218–232.e215. Riminucci, M., Collins, M.T., Fedarko, N.S., Cherman, N., Corsi, A., White, K.E., Waguespack, S., Gupta, A., Hannon, T., Econs, M.J., et al. (2003). FGF-23 in fibrous dysplasia of bone and its relationship to renal phosphate wasting. J. Clin. Invest. 112, 683–692. Romer, A.S. (1967). Major steps in vertebrate evolution. Science 158, 1629– 1637. Schuierer, S., and Roma, G. (2016). The exon quantification pipeline (EQP): a comprehensive approach to the quantification of gene, exon and junction expression from RNA-seq data. Nucleic Acids Res. 44, e132. Semba, R.D., Cappola, A.R., Sun, K., Bandinelli, S., Dalal, M., Crasto, C., Guralnik, J.M., and Ferrucci, L. (2011a). Plasma klotho and cardiovascular disease in adults. J. Am. Geriatr. Soc. 59, 1596–1601. Semba, R.D., Cappola, A.R., Sun, K., Bandinelli, S., Dalal, M., Crasto, C., Guralnik, J.M., and Ferrucci, L. (2011b). Plasma klotho and mortality risk in older community-dwelling adults. J. Gerontol. A Biol. Sci. Med. Sci. 66, 794–800.

Kurosu, H., Yamamoto, M., Clark, J.D., Pastor, J.V., Nandi, A., Gurnani, P., McGuinness, O.P., Chikuda, H., Yamaguchi, M., Kawaguchi, H., et al. (2005). Suppression of aging in mice by the hormone Klotho. Science 309, 1829–1833.

Shaw, L.J., Giambrone, A.E., Blaha, M.J., Knapper, J.T., Berman, D.S., Bellam, N., Quyyumi, A., Budoff, M.J., Callister, T.Q., and Min, J.K. (2015). Long-Term Prognosis After Coronary Artery Calcification Testing in Asymptomatic Patients: A Cohort Study. Ann. Intern. Med. 163, 14–21.

Kurosu, H., Ogawa, Y., Miyoshi, M., Yamamoto, M., Nandi, A., Rosenblatt, K.P., Baum, M.G., Schiavi, S., Hu, M.C., Moe, O.W., and Kuro-o, M. (2006). Regulation of fibroblast growth factor-23 signaling by klotho. J. Biol. Chem. 281, 6120–6123.

Shen, B., Vardy, K., Hughes, P., Tasdogan, A., Zhao, Z., Yue, R., Crane, G.M., and Morrison, S.J. (2019). Integrin alpha11 is an Osteolectin receptor and is required for the maintenance of adult skeletal bone mass. eLife 8, e42274.

Leopold, J.A. (2013). Vascular calcification: an age-old problem of old age. Circulation 127, 2380–2382. Liao, B.K., Deng, A.N., Chen, S.C., Chou, M.Y., and Hwang, P.P. (2007). Expression and water calcium dependence of calcium transporter isoforms in zebrafish gill mitochondrion-rich cells. BMC Genomics 8, 354. Lindberg, K., Amin, R., Moe, O.W., Hu, M.C., Erben, R.G., O¨stman Wernerson, A., Lanske, B., Olauson, H., and Larsson, T.E. (2014). The kidney is the principal organ mediating klotho effects. J. Am. Soc. Nephrol. 25, 2169–2175.

Shi, M., Flores, B., Gillings, N., Bian, A., Cho, H.J., Yan, S., Liu, Y., Levine, B., Moe, O.W., and Hu, M.C. (2016). aKlotho Mitigates Progression of AKI to CKD through Activation of Autophagy. J. Am. Soc. Nephrol. 27, 2331–2345. Shimada, T., Kakitani, M., Yamazaki, Y., Hasegawa, H., Takeuchi, Y., Fujita, T., Fukumoto, S., Tomizuka, K., and Yamashita, T. (2004). Targeted ablation of Fgf23 demonstrates an essential physiological role of FGF23 in phosphate and vitamin D metabolism. J. Clin. Invest. 113, 561–568. Standal, T., Borset, M., and Sundan, A. (2004). Role of osteopontin in adhesion, migration, cell survival and bone remodeling. Exp. Oncol. 26, 179–184.

Cell Reports 28, 2767–2776, September 10, 2019 2775

Stickens, D., Behonick, D.J., Ortega, N., Heyer, B., Hartenstein, B., Yu, Y., Fosang, A.J., Schorpp-Kistner, M., Angel, P., and Werb, Z. (2004). Altered endochondral bone development in matrix metalloproteinase 13-deficient mice. Development 131, 5883–5895.

Vijayakumar, P., Laize´, V., Cardeira, J., Trindade, M., and Cancela, M.L. (2013). Development of an in vitro cell system from zebrafish suitable to study bone cell differentiation and extracellular matrix mineralization. Zebrafish 10, 500–509.

Sugano, Y., and Lardelli, M. (2011). Identification and expression analysis of the zebrafish orthologue of Klotho. Dev. Genes Evol. 221, 179–186.

Walker, M.B., and Kimmel, C.B. (2007). A two-color acid-free cartilage and bone stain for zebrafish larvae. Biotech. Histochem. 82, 23–28.

Tang, W., Zhang, G., Serluca, F., Li, J., Xiong, X., Coble, M., Tsai, T., Li, Z., Molind, G., Zhu, P., et al. (2018). Genetic Architecture of Collective Behaviors in Zebrafish. bioRxiv. https://doi.org/10.1101/350314. Teitelbaum, S.L., and Ross, F.P. (2003). Genetic regulation of osteoclast development and function. Nat. Rev. Genet. 4, 638–649. Thompson, R.C., Allam, A.H., Lombardi, G.P., Wann, L.S., Sutherland, M.L., Sutherland, J.D., Soliman, M.A., Frohlich, B., Mininberg, D.T., Monge, J.M., et al. (2013). Atherosclerosis across 4000 years of human history: the Horus study of four ancient populations. Lancet 381, 1211–1222. Urakawa, I., Yamazaki, Y., Shimada, T., Iijima, K., Hasegawa, H., Okawa, K., Fujita, T., Fukumoto, S., and Yamashita, T. (2006). Klotho converts canonical FGF receptor into a specific receptor for FGF23. Nature 444, 770–774.

2776 Cell Reports 28, 2767–2776, September 10, 2019

Yang, D.C., Tsai, C.C., Liao, Y.F., Fu, H.C., Tsay, H.J., Huang, T.F., Chen, Y.H., and Hung, S.C. (2011). Twist controls skeletal development and dorsoventral patterning by regulating runx2 in zebrafish. PLoS ONE 6, e27324. Yu, G., Wang, L.G., Han, Y., and He, Q.Y. (2012). clusterProfiler: an R package for comparing biological themes among gene clusters. OMICS 16, 284–287. Yue, R., Shen, B., and Morrison, S.J. (2016). Clec11a/osteolectin is an osteogenic growth factor that promotes the maintenance of the adult skeleton. eLife 5, e18782. Zhu, A., Ibrahim, J.G., and Love, M.I. (2019). Heavy-tailed prior distributions for sequence count data: removing the noise and preserving large differences. Bioinformatics 35, 2084–2092.

STAR+METHODS KEY RESOURCES TABLE

REAGENT or RESOURCE

SOURCE

IDENTIFIER

Biological Samples Zebrafish Kidney

This study

N/A

Zebrafish Gills

This study

N/A

Zebrafish Heart

This study

N/A

Alizarin Red-S

Sigma-Aldrich

Cat No. A5533-25G

Tartrate-resistant acid phosphatase

Sigma-Aldrich

Cat No. 387A-1KT

Modified Davidson’s fixative

Fisher Scientific

Cat No. 50-292-28

Hematoxylin (Gill’s Hematoxylin III)

Poly Scientific R&D Corp

Cat No. s211-32oz

Eosin Y Alcoholic Working Solution

Poly Scientific R&D Corp

Cat No. s2186-32oz Cat No. 409501

Chemicals, Peptides, and Recombinant Proteins

Critical Commercial Assays RNAscope Probe- Dr-spp1

Advanced Cell Diagnostics

RNALater Stabilization Solution

ThermoFisher

Cat No. AM7021

RNeasy Fibrous Tissue Mini kit

QIAGEN

Cat No. 74704

Ambion MEGAshortscript T7 Kit

ThermoFisher

Cat no. AM1354

RNAeasy kit

QIAGEN

Cat No. 74104

Cas9 Protein

PNA Bio

Cat No. CP01

This study

Sequence Read Archive, NCBI. BioProject Accession: PRJNA556842

€ strain Zebrafish (Danio rerio), Tu

€sslein-Volhard lab Nu

RRID:ZIRC_ZL57

Zebrafish (Danio rerio), AB strain

ZIRC

RRID:ZIRC_ZL1

€ strain) klothoD5 (Tu

This study

N/A

Deposited Data RNaseq data Experimental Models: Organisms/Strains

klothoD5 (AB strain)

This study

N/A

€ strain) fgf23D1 (Tu

This study

N/A

fgf23D11 (AB strain)

This study

N/A

T7 universal primer for the DNA template for sgRNA: AAAAGCACCGACTCGGTGCCACTTTTT CAAGTTGATAACGGACTAGCCTTATTTTAACTT GCTATTTCTAGCTCTAAAAC

Integrated DNA Technologies, Inc., USA

N/A

klotho-specific primer for the DNA template for sgRNA: GAAATTAATACGACTCACTATAGGCTG GAGTAATTCGGTTAgttttagagctagaaATAGC

Integrated DNA Technologies, Inc., USA

N/A

Forward primer for genotyping aklotho mutant: CGGCACCGCTGCATATTCAGTGG

Integrated DNA Technologies, Inc., USA

N/A

Reverse primer for genotyping aklotho mutant: CCAGATTTGACTTCAGTACTGAC

Integrated DNA Technologies, Inc., USA

N/A

fgf23-specific primer for the DNA template for sgRNA: GAAATTAATACGACTCACTATAGGT CTGAAGTGGTCTGAAGGGTTTTAGAGCTAGA AATAGC

Integrated DNA Technologies, Inc., USA

N/A

Forward primer for genotyping fgf23 mutant: CCGGCTTTACGCGCTCTGTCAAG

Integrated DNA Technologies, Inc., USA

N/A

Oligonucleotides

(Continued on next page)

Cell Reports 28, 2767–2776.e1–e5, September 10, 2019 e1

Continued REAGENT or RESOURCE

SOURCE

IDENTIFIER

Reverse primer for genotyping fgf23 mutant: CCAGACGGTCTCTGCTTTCTGTT

Integrated DNA Technologies, Inc., USA

N/A

Software and Algorithms Exon Quantification Pipeline

Schuierer and Roma, 2016

N/A

MultiQC

Ewels et. al, 2016

https://github.com/ewels/MultiQC

DESeq2

Love et al., 2014

https://bioconductor.org/packages/release/bioc/ html/DESeq2.html

apeglm

Zhu et al., 2019

https://bioconductor.org/packages/release/bioc/ html/apeglm.html

msigdbr

msigdbr package

https://cran.r-project.org/web/packages/msigdbr/ index.html

ClusterProfiler

Yu et al., 2012

https://bioconductor.org/packages/release/bioc/ html/clusterProfiler.html

Other

LEAD CONTACT AND MATERIALS AVAILABILITY Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, David J. Glass ([email protected]). Zebrafish mutants generated in this study are available upon request, under a Materials Transfer Agreement, to be negotiated with Novartis. EXPERIMENTAL MODEL AND SUBJECT DETAILS Zebrafish All animals were maintained and used for scientific research in accordance with the guidelines of The Institutional Animal Care and Use Committee (IACUC) of the Novartis Institutes for BioMedical Research, Cambridge, USA. Zebrafish were housed in 3L tanks in a recirculating Aquatic Habitats facility (Pentair, USA) on a 14:10 hour light:dark cycle at 28 C. Larvae were fed Zeigler Larval Diet AP100 Z3 to M3 (< 100 microns; Zeigler Bros., Inc, USA) from 5 day post-fertilization to 20 days post-fertilization. Juvenile fish were fed Brine shrimps hatched from Premium Grade Brine Shrimp Eggs (Brine Shrimp Direct, USA) and TetraMin Tropical Flake (Tetra, Germany) twice per day. Adults fish were fed a diet of GEMMA Micro 300 (Skretting France) once per day. Zebrafish were anesthetized using 0.0168% buffered Tricaine-S (MS-222, Syndel). € and AB strains of zebrafish were used as wild-type for the study. Tu Generation of Zebrafish Knockouts Knockouts for klotho and fgf23 were generated by CRISPR/Cas9 method. DNA template for in vitro transcription of CRISPR sgRNA was prepared by PCR using gene-specific primer and T7 universal primer: T7 universal primer: AAAAGCACCGACTCGGTGCCACTTTTTCAAGTTGATAACGGACTAGCCTTATTTTAACTTGCTATTTCTAGCT CTAAAAC klotho-specific primer: GAAATTAATACGACTCACTATAGGCTGGAGTAATTCGGTTAgttttagagctagaaATAGC fgf23-specific primer: GAAATTAATACGACTCACTATAGGTCTGAAGTGGTCTGAAGGGTTTTAGAGCTAGAAATAGC For sgRNA generation, PCR product was purified and in vitro transcription was performed using the Ambion MEGAshortscript T7 Kit (cat no. AM1354). sgRNA was purified using RNAeasy kit (QIAGEN; cat no. 74104), and diluted to 500 ng/ml. Equal volume of purified sgRNA and Cas9 protein (500 ng/ml; PNA Bio – CP01) was co-injected into single-cell stage zebrafish embryos. Injected fertilized embryos were raised to adulthood (F0). F0 adults were crossed to wild-type zebrafish to identify F1 generation by NGS. Animals carrying frameshift mutation were identified and propagated for the purpose of this study. For genotyping, DNA was isolated from finclips (Meeker et al., 2007) for PCR and next-generation sequencing (NGS). sgRNA target for klotho: GGCTGGAGTAATTCGGTTATGG; primers for NGS (forward: CGGCACCGCTGCATATTCAGTGG; and reverse: CCAGATTTGACTTCAGTACTGAC)

e2 Cell Reports 28, 2767–2776.e1–e5, September 10, 2019

sgRNA target for fgf23: AGTCTGAAGTGGTCTGAAGGTGG; primers for NGS (forward: CCGGCTTTACGCGCTCTGTCAAG; and reverse: CCAGACGGTCTCTGCTTTCTGTT) METHOD DETAILS Lifespan Analysis In order to analyze the lifespan of control and mutant lines, survival analysis based on humane end-points was performed using GraphPad Prism 7. Parameters for humane end-points were determined in consultation with institutional veterinarian services. Animals displaying the following signs were euthanized: Signs of tissue degeneration and tumors, inability or unwillingness to swim, inability to maintain balance, abnormal swelling or tumors, severe eye protrusion, gross abnormalities in body shape, posture or spinal deformities that affect animal’s ability to swim or eat. Behavioral Analysis For the behavioral assay performed in the circular arena, six fish of matching size and age were used in each assay. Controls and mutants were assayed in parallel. Experiments were conducted at a similar time of day and monitored by video without human presence. Behavioral rooms had room temperature and light cycles consistent with the main facility. The circular arena consisted of the following features: acrylic tank; outside diameter = 50.8cm; inside diameter = 48.9cm; tank height = 20.3cm height; open top (Plastic Supply, Inc., USA). The tanks were filled to a depth of 4.4cm (
9 L total volume) with zebrafish system water fed directly from the main zebrafish housing unit to ensure all water parameters were identical to the housing conditions. The circular arenas were coated on the outside (I00810, Frosted Glass Finish, Krylon, USA) to prevent the fish from being able to see outside of the arenas without compromising the transparency to infrared light. Underneath the tanks were adjustable infrared panels (940 nm IR LEDs, Shenzhen VICO). Basler Ace 2040-90um Near Infrared (NIR) cameras (Order#-106541, Graftek Imaging, USA) were mounted 58.4cm above the arena to collect a dorsal view of zebrafish. Infrared long-pass filters (Midopt LP780-62, Graftek Imaging, USA) were attached to the lens (Schneider Cinegon 1.9, Graftek Imaging, USA) and were set to an aperture of six. Six fish were transferred directly from home tanks to the behavioral arena by netting. All trials were recorded after 10 minutes of habituation to allow for recovery from any stress due to netting from the home tank. Each trial was a recording of 30 min at 60 frames per second. Arenas were rinsed clean with system water at the end of the day and put through a cabinet washer once a week on a hot water only cycle. For behavioral analysis in the home-tank arena, five size and age-matched fish were placed in a 1.4 L zebrafish tank (Pentair, USA) with 1 L of zebrafish system water. We used a side-mounted camera (acA2000-165u mNIR, Basler). To make the background uniform for tracking, we placed a 25cmX25cm infrared illuminating board on the obverse side of the fish tank to illuminate the fish. An optical filter (LP780-72 filter, MidOPT) was placed on a lens (LM8XC 1.3’’ (4/3’’) 8.5mm, F2.8, KOWA) to permit recording of infrared light. Each trial was a recording of 30 minutes at 60 frames per second. Behavioral data were analyzed as described (Tang et al., 2018). Histology Adult zebrafish were euthanized by exposure to chilled water (0-4 C). Samples were fixed in Modified Davidson’s fixative (Fisher Scientific; catalog number 50-292-28) for up to 72 hours. At the time of fixation, abdominal cavity was cut open in order to expose internal organs for efficient fixation. The gills were flushed gently with the fixative, and an incision was made across the spinal cord posterior to the brain for efficient fixation of the central nervous system. After fixation, Zebrafish were placed in Immuno Cal Decalcifier (Stat Lab-McKinney, TX) for a total of 48 hours with continuous agitation, fresh solution was added after 24 hours. The Zebrafish were then rinsed in running tap water to remove any residual calcium salts, and processed through a graded series of alcohols and xylene to be embedded in paraffin wax in a sagittal orientation. Paraffin embedded blocks were then serially sectioned at 5mm on a rotary microtome, and each individual section was placed on a charged glass slide. Every tenth slide was stained with a hematoxylin and eosin (H&E; Gill’s Hematoxylin III (s211-32oz) and Eosin Y Alcoholic Working Solution (s2186-32oz), Poly Scientific R&D Corp, Bay Shore NY) staining procedure to identify different tissue structures. Slides were then scanned into an Aperio slide Scanner (Leica Biosystems). A board-certified histopathologist analyzed the H&E stained samples. Alizarin red (Alizarin Red-S, Sigma-Aldrich; A5533-25G) staining was performed as described (Walker and Kimmel, 2007). Tartrate-resistant acid phosphatase staining was performed as per the manufacturer’s instructions (Sigma-Aldrich; 387A-1KT). RNA-Seq Sample Preparation Tissue Collection and Dissection Adult zebrafish were euthanized by exposure to chilled water (0-4 C). Gills, heart and kidney were collected from 32 individual fish at two time-points, namely at 3 and 5 mpf for aklotho mutants and wild-type sibling controls (AB background; 8 fish per genotype per time-point - 4 males, 4 females) for a total of 96 samples. Tissues were dissected in cold PBS and immediately stored in RNALater Stabilization Solution (ThermoFisher Cat# AM7021). RNALater was removed after an overnight incubation at 4 C, and samples were stored at 80 C until processing. RNA Extraction All tissues were homogenized using the TissueLyser II (QIAGEN) plus Lysis buffer containing b-mercaptoethanol and stored at 80 C. RNA extraction was performed using the automated protocol in the QIAcube workstation utilizing the RNeasy Fibrous Tissue

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Mini kit (QIAGEN Cat No./ID: 74704). RNA integrity and quality were assessed by Agilent TapeStation using High Sensitivity RNA ScreenTapes. Samples were normalized and 300ng of RNA was used for library prep for each sample. ERCC RNA Spike-in mix was added for quality control. RNASeq Library Prep and Sequencing RNASeq libraries were prepared using the Illumina TruSeq stranded mRNA HS sample preparation kit using an automated pipeline. Magnetic poly-T oligo beads were used to purify poly-A containing mRNA for cDNA synthesis. The library quality was assessed by Agilent TapeStation using High Sensitivity DNA 1000 ScreenTapes and quantitated using Invitrogen Quant-iT PicoGreen dsDNA assay kit. Samples were pooled in equal amount before checking them in an Illumina MiSeq flow cell for quality and to optimize clustering. Four samples failed the library preparation step (two aklotho mutant females, one aklotho mutant male, one wild-type sibling male) and could not be sequenced. Final sequencing was performed on a HiSeq 2500 instrument (76 base pair, paired-end). qPCR for the Quantification of RNA Expression Levels RNA levels were quantified and normalized using the Qubit RNA HS Assay kit (Thermo Fisher). Reverse transcription was performed using the Superscript III First-Strand Synthesis System (Thermo Fisher) following the manufacturer’s instructions (with a 1:1 mix of oligo-dT and random hexamers). qPCR was then performed in triplicates using the Power SYBR Green Mix (Thermo Fisher) on a QuantStudio 7 Flex system following the manufacturer’s instructions. The primers were validated for specificity (melting-curve) and efficiency (dilution curve). The following qPCR primer pairs were used (Vijayakumar et al., 2013; Yang et al., 2011): runx2a (runt-related transcription factor 2): Forward primer: AGCCGACCCACGCCAGTTTGAG Reverse primer: TGGGGTGTAG GTGAATGTTGCTGGATA runx2b: Forward primer: ACGCAAACGGAGGACATACG Reverse primer: CCGGCGCTGGGATCTAC osteopontin/spp1: Forward primer: GAGCCTACACAGACCACGCCAACAG Reverse primer: GGTAGCCCAAACTGTCTCCCCG tnf-a: Forward primer: GCGCTTTTCTGAATCCTACG Reverse primer: TGCCCAGTCTGTCTCCTTCT b-actin: Forward primer: CGAGCAGGAGATGGGAAC Reverse primer: CAACGGAAACGCTCATTGC Ct values were automatically calculated by the QuantStudio 7 Flex system and outliers among technical triplicates were manually eliminated. Data were last analyzed using the DDCt method: Ct values were averaged, then subtracted to the average Ct value of b-actin (DCt), and last the fold change was determined using the formula 2–(DCt sample – DCt reference). RNAscope RNAscope probe targeting spp1 gene (ACDBio; Cat No. 409501) was used to visualize spp1 expression as described (Gross-Thebing et al., 2014). Briefly, Adult zebrafish were euthanized in ice-cold water and decapitated. The hearts were dissected out and fixed in 4% PFA overnight at 4 C. The hearts were washed and then dehydrated through serial methanol incubations and stored at 20 C overnight. Following rehydration, the samples were permeabilized and then incubated in the probe mixture overnight at 40 C. Following incubation, the embryos were washed and fluorescence detection steps were performed as described (Gross-Thebing et al., 2014). Samples were then imaged using a Zeiss Lightsheet microscope (Lightsheet Z.1, Zeiss). QUANTIFICATION AND STATISTICAL ANALYSIS RNA-Seq Data Analysis Alignment and quantification were performed with a Novartis internal pipeline, Exon Quantification Pipeline (EQP) (Schuierer and Roma, 2016), using STAR and the Zebrafish Reference GRCz11. MultiQC package was used to do the pre-alignment QC check (Ewels et al., 2016). Two samples failed the QC checked and were removed from further analysis (one kidney sample, one heart sample). The failed samples correlated with low RIN scores. The average total mapped reads per sample was 23.7 million PE reads. Genes with read counts < 10 were filtered out before Differential gene expression analysis (DGE). DGE was performed with the DESeq2 package (Love et al., 2014), comparing mutants to wild-type using sex as a covariate for each time-point with adjusted p value cutoff = 0.05 and using ‘apeglm’ for LFC shrinkage (Zhu et al., 2019). Adjusted p values were calculated using the Benjamini-Hochberg False Discovery Rate approach to correct for multiple testing. Gene Set Enrichment Analysis Gene set enrichment analysis was performed on the up- or downregulated genes in aklotho mutants (adjusted p value < 0.05 and log2-fold change > 1.5) from each tissue (kidney, heart, and gills) and adult stage (three and five mpf). The curated gene sets, including canonical pathways and hallmark pathways were included in the analysis. The gene sets were downloaded from the Molecular Signatures Database (http://software.broadinstitute.org/gsea/msigdb/) and were converted to zebrafish homologs using the R package msigdbr. For each pathway, the corresponding gene set was compared with an up- or downregulated gene list. The overlapping genes were counted, and a hypergeometric test was performed to measure the statistical significance, e.g., p value, on whether the number of overlaps occurred by chance. Finally, adjusted p values were derived by the Benjamini-Hochberg (BH) procedure

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to control the False Discovery Rate (FDR). In a separate analysis, for each pathway and comparison, a weighted KolmogorovSmirnov test was performed using the gsea function (with the number of permutations to be 1e8) from the R package ClusterProfiler (Yu et al., 2012). Adjusted p values were derived by the BH procedure to control the FDR. DATA AND CODE AVAILABILITY Sequencing data have been deposited to the Sequence Read Archive at NCBI under the BioProject Accession: PRJNA556842.

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