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Report
aKlotho Regulates Age-Associated Vascular
Calcification and Lifespan in Zebrafish
Graphical Abstract
Highlights
dZebrafish aklotho mutants display reduced lifespans
dThe aklotho phenotype occurs later in zebrafish than in mice
dZebrafish aklotho mutants display adult-onset vascular
calcification
dCalcification coincides with an increase in osteoclast
differentiation pathways
Authors
Ajeet Pratap Singh, Maria X. Sosa,
Jian Fang, ..., Samuel M. Cadena,
Mark C. Fishman, David J. Glass
Correspondence
david_glass@hms.harvard.edu
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.
Singh et al., 2019, Cell Reports 28, 2767–2776
September 10, 2019 ª2019 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. Glass
1,5,6,
*
1
Zebrafish Group, Chemical Biology and Therapeutics, Novartis Institutes for Biomedical Research, 181 Massachusetts Avenue, Cambridge,
MA 02139, USA
2
Preclinical Safety, Novartis Institutes for Biomedical Research, 250 Massachusetts Avenue, Cambridge, MA 02139, USA
3
DAx/Discovery and Translational Pharmacology, Novartis Institutes for Biomedical Research, 181 Massachusetts Avenue, Cambridge,
MA 02139, USA
4
Harvard Department of Stem Cell and Regenerative Biology, Harvard University, 7 Divinity Ave, Cambridge, MA 02138, USA
5
Age-Related Disorders Group, Chemical Biology and Therapeutics, Novartis Institutes for Biomedical Research, 181 Massachusetts
Avenue, Cambridge, MA 02139, USA
6
Lead Contact
*Correspondence: david_glass@hms.harvard.edu
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 acceler-
ated aging due to hyperphosphatemia and soft tis-
sue 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 scaf-
fold 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 regu-
lates 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 fac-
tor-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 heter-
odimerize 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 funda-
mental differences in physiology between fish and terrestrial ver-
tebrates owing to unique demands of aquatic versus terrestrial
environments. In terms of renal function and mineral and fluid ho-
meostasis, 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 ho-
meostasis. 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 ª2019 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/).
(legend on next page)
2768 Cell Reports 28, 2767–2776, September 10, 2019
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 Early-
Onset Mortality
We targeted the aklotho gene using the CRISPR/Cas9 method
(Irion et al., 2014), generating mutations in two background ze-
brafish strains, T€
u and AB, and identified aklotho alleles carrying
frameshift mutations and early stop codons (Table S1). We also
targeted fgf23 because the function of aKlotho in mammals de-
pends in large part upon its binding to FGF receptors and recruit-
ing 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 morpho-
logically 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 zebra-
fish obtained from breeding parents fgf23 heterozygotes, we ob-
tained 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 sponta-
neous 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, age-
related 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 propor-
tionally 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 integ-
ument 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 (ar-
row in Figure 2B), accompanied by a degeneration and fibrosis of
adjacent skeletal muscles with immune cell infiltration (Fig-
ure 2B). Mineralization was often in a concentric pattern in the
affected areas. The gill arch demonstrated bone overgrowth (hy-
perostosis) 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 strik-
ing 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 bloodflow to the thin-walled capillaries
of the gills (Farrell, 1979; Grimes and Kirby, 2009). To validate
calcification in the BA, we used alizarinred, a stain for calcification
(Walker and Kimmel, 2007). The BA in aklotho
/
mutants is
prominently stained with alizarin red in contrast to wild-type ani-
mals (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 skel-
etal system, there were multifocal areas of hyperostosis and
Figure 1. Zebrafish aklotho and fgf23 Mutants
(A–F) Body condition of aklotho and fgf23 mutant males at 5 mpf: (A) T€
u wild-type strain, (B) aklotho (kl
D5
), and (C) fgf23 (fgf23
ins1
) mutant in T€
u background; (D) AB
wild-type strain, (E) aklotho (kl
D5
), and (F) fgf23 (fgf23
D11
) 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)
in T€
u background and for (I) aklotho (n = 24 wild-type siblings, 21 mutants; p = 0.0001) and (J) fgf23 mutants (n = 60 background contro ls, 68 mutants; p < 0.0001)
in AB background. Log-rank (Mantel-Cox) test for statistical analysis on survival curves in GraphP ad 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.
Cell Reports 28, 2767–2776, September 10, 2019 2769
chondrodysplasia. These changes were prominent in the caudal
region (Figure S2A). Frequently, regions of bone overgrowth
were accompanied by areas of dystrophic calcification, connec-
tive 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 calcifica-
tion in zebrafish is accompanied by an increase in osteoclast ac-
tivity (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
Figure 2. Vascular Calcification and Inflam-
mation in aklotho Mutants
H&E staining on paraffin sections from 5-month-old
wild-type control (T€
u) and aklotho (kl
D5
) males.
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); (D0and D00) alizarin red-
stained 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 kl
D5
(right) hearts. TRAP staining
on (I) whole mount and (J) cryosection of 5-mpf
hearts.
See also Figure S2.
RNA sequencing (RNA-seq) analysis of the
kidney, heart (including BA), and gills at
3 mpf, when mutantsappeared comparable
phenotypically to wild-type siblings, and at
5mpf,whenaklotho
/
mutants became
phenotypically distinct from wild-type sib-
lings (n = 8 animalsper genotype and condi-
tion; 4 malesand 4 females). In the wild-type
siblings, aklotho expression is highest in the
kidneys (Figure 3A[3]). As expected, a sig-
nificant downregulation of aklotho expres-
sion is observed in aklotho
/
mutants at
both 3 and 5 mpf (Figure 3A[1]). In wild-
type 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 indistin-
guishable 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 wide-
spread 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 over-
lap 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 Kolmo-
gorov-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 calcifica-
tion in the BA at 3 mpf. All eight hearts displayed calcification in
2770 Cell Reports 28, 2767–2776, September 10, 2019
(legend on next page)
Cell Reports 28, 2767–2776, September 10, 2019 2771
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 up-
regulation 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 re-
modeling, and inflammation are upregulated in aklotho
/
mutant hearts (Figure 4). spp1 is the most significantly upregu-
lated 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 (Fig-
ure 4D). However, we do not observe an upregulation of the con-
ventional 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 osteo-
blast 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 activ-
ity 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 local-
ized spp1 expression in the BAs of aklotho
/
mutants
(Figure 4H).
DISCUSSION
The aKlotho/FGF23 pathway appears to play a role in the ag-
ing 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 associ-
ated 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 re-
flects the osteogenic cell fate change of vascular smooth mus-
cle. Here, we find that in the absence of aKlotho/FGF23,
vascular tissue in the BA changes its pattern of gene expres-
sion 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 anti-
osteogenic mechanisms, including a local differentiation of os-
teoclasts. 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 se-
vere tumoral calcinosis including ectopic calcifications, indi-
cating the zebrafish model is predictive of the human condition
(Ichikawa et al., 2017).
The BA is an elastic, valveless cardiac outflow tract in tele-
osts that is believed to act as a windkessel to protect the deli-
cate 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; Mizobu-
chi 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.
2772 Cell Reports 28, 2767–2776, September 10, 2019
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 sup-
plementation 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 verte-
brates necessitated the evolution of mechanisms to regulate
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 log
10
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 (kl
D5
) 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.
Cell Reports 28, 2767–2776, September 10, 2019 2773
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; Rimi-
nucci 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:
dKEY RESOURCES TABLE
dLEAD CONTACT AND MATERIALS AVAILABILITY
dEXPERIMENTAL MODEL AND SUBJECT DETAILS
BZebrafish
dMETHOD DETAILS
BLifespan Analysis
BBehavioral Analysis
BHistology
BRNA-Seq Sample Preparation
dQUANTIFICATION AND STATISTICAL ANALYSIS
BRNA-Seq Data Analysis
dDATA 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 Age-
Related 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 RNA-
seq; 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 mi-
croscopy 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. Fremont-
Rahl, 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.; Gener-
ation 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
Collection, G.J.M. and A.P.S.; Behavioral Data Analysis, T.T. and W.T.; Manu-
script 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 Thera-
peutics 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
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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
Chemicals, Peptides, and Recombinant Proteins
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
Critical Commercial Assays
RNAscopeProbe- Dr-spp1 Advanced Cell Diagnostics Cat No. 409501
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
Deposited Data
RNaseq data This study Sequence Read Archive, NCBI. BioProject
Accession: PRJNA556842
Experimental Models: Organisms/Strains
Zebrafish (Danio rerio), T€
u strain N€
usslein-Volhard lab RRID:ZIRC_ZL57
Zebrafish (Danio rerio), AB strain ZIRC RRID:ZIRC_ZL1
klotho
D5
(T€
u strain) This study N/A
klotho
D5
(AB strain) This study N/A
fgf23
D1
(T€
u strain) This study N/A
fgf23
D11
(AB strain) This study N/A
Oligonucleotides
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
(Continued on next page)
Cell Reports 28, 2767–2776.e1–e5, September 10, 2019 e1
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 (david.glass@novartis.com). 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 28C. 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).
T€
u and AB strains of zebrafish were used as wild-type for the study.
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 pu-
rified sgRNA and Cas9 protein (500 ng/ml; PNA Bio – CP01) was co-injected into single-cell stage zebrafish embryos. Injected fertil-
ized embryos were raised to adulthood (F
0
). F
0
adults were crossed to wild-type zebrafish to identify F
1
generation by NGS. Animals
carrying frameshift mutation were identified and propagated for the purpose of this study. For genotyping, DNA was isolated from fin-
clips (Meeker et al., 2007) for PCR and next-generation sequencing (NGS).
sgRNA target for klotho: GGCTGGAGTAATTCGGTTATGG; primers for NGS (forward: CGGCACCGCTGCATATTCAGTGG; and
reverse: CCAGATTTGACTTCAGTACTGAC)
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
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. An-
imals 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 spi-
nal 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 pres-
ence. 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 compro-
mising 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 (0-4C). Samples were fixed in Modified Davidson’s fixative (Fisher Sci-
entific; catalog number 50-292-28) for up to 72 hours. At the time of fixation, abdominal cavity was cut open in order to expose in-
ternal 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 Decal-
cifier (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 5mmona
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 per-
formed 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 (0-4C). 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 4C, and samples were
stored at 80C until processing.
RNA Extraction
All tissues were homogenized using the TissueLyser II (QIAGEN) plus Lysis buffer containing b-mercaptoethanol and stored at
80C. RNA extraction was performed using the automated protocol in the QIAcube workstation utilizing the RNeasy Fibrous Tissue
Cell Reports 28, 2767–2776.e1–e5, September 10, 2019 e3
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 clus-
tering. 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
C
t
values were automatically calculated by the QuantStudio 7 Flex system and outliers among technical triplicates were manually
eliminated. Data were last analyzed using the DDC
t
method: C
t
values were averaged, then subtracted to the average C
t
value of
b-actin (DC
t
), 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 4C. The hearts were washed and then dehydrated through serial methanol incubations and stored at 20C
overnight. Following rehydration, the samples were permeabilized and then incubated in the probe mixture overnight at 40C.
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 sam-
ple). 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 log-
2-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 Sig-
natures 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
e4 Cell Reports 28, 2767–2776.e1–e5, September 10, 2019
to control the False Discovery Rate (FDR). In a separate analysis, for each pathway and comparison, a weighted Kolmogorov-
Smirnov 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.
Cell Reports 28, 2767–2776.e1–e5, September 10, 2019 e5
