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Research ArticleCell biologyVascular biology Open Access | 10.1172/JCI150051




SIRT6 PROTECTS VASCULAR SMOOTH MUSCLE CELLS FROM OSTEOGENIC TRANSDIFFERENTIATION
VIA RUNX2 IN CHRONIC KIDNEY DISEASE

WENXIN LI,1 WEIJING FENG,2 XIAOYAN SU,3 DONGLING LUO,1 ZHIBING LI,4 YONGQIAO
ZHOU,4 YONGJUN ZHU,1 MENGBI ZHANG,3 JIE CHEN,5 BAOHUA LIU,6 AND HUI HUANG1

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Li, W. in: JCI | PubMed | Google Scholar

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Feng, W. in: JCI | PubMed | Google Scholar |

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Su, X. in: JCI | PubMed | Google Scholar

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Luo, D. in: JCI | PubMed | Google Scholar

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Li, Z. in: JCI | PubMed | Google Scholar

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Zhou, Y. in: JCI | PubMed | Google Scholar

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Zhu, Y. in: JCI | PubMed | Google Scholar

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Zhang, M. in: JCI | PubMed | Google Scholar

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Chen, J. in: JCI | PubMed | Google Scholar

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Liu, B. in: JCI | PubMed | Google Scholar |

1Department of Cardiology, The Eighth Affiliated Hospital, Sun Yat-sen
University, Shenzhen, China.

2Department of Cardiology, State Key Laboratory of Organ Failure Research,
Guangdong Provincial Key Lab of Shock and Microcirculation, Nanfang Hospital,
Southern Medical University, Guangzhou, China.

3Nephropathy Department, Tungwah Hospital of Sun Yat-sen University, Dongguan,
China.

4Department of Cardiology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

5Department of Radiation Oncology, Sun Yat-sen Memorial Hospital, Sun Yat-sen
University, Guangzhou, China.

6Shenzhen Key Laboratory for Systemic Aging and Intervention, National
Engineering Research Center for Biotechnology-Shenzhen, Shenzhen University
Health Science Center, Shenzhen, China.



Address correspondence to: Hui Huang, Department of Cardiology, The Eighth
Affiliated Hospital, Sun Yat-sen University, No.3025, Shennan Middle Road,
Futian District Shenzhen 518000, China. Email: huangh8@mail.sysu.edu.cn.

Authorship note: WL, WF, XS, and DL are co–first authors.

Find articles by Huang, H. in: JCI | PubMed | Google Scholar

Authorship note: WL, WF, XS, and DL are co–first authors.

Published November 18, 2021 - More info

Published in Volume 132, Issue 1 on January 4, 2022
J Clin Invest. 2022;132(1):e150051. https://doi.org/10.1172/JCI150051.
© 2022 Li et al. This work is licensed under the Creative Commons Attribution
4.0 International License. To view a copy of this license, visit
http://creativecommons.org/licenses/by/4.0/.
Published November 18, 2021 - Version history
Received: March 31, 2021; Accepted: November 12, 2021
View PDF

--------------------------------------------------------------------------------


RELATED ARTICLE:

A MOLECULAR TARGET OF VASCULAR CALCIFICATION IN CHRONIC KIDNEY DISEASE

Mohamed G. Atta
Mohamed G. Atta


Commentary

A MOLECULAR TARGET OF VASCULAR CALCIFICATION IN CHRONIC KIDNEY DISEASE

 * Text
 * PDF

ABSTRACT

Vascular calcification (VC) causes cardiovascular morbidity and mortality in
patients with chronic kidney disease (CKD), particularly those with end-stage
kidney disease (ESKD) on maintenance dialysis treatment. Although many
mechanisms have been proposed, their detailed effects remain incompletely
understood. In this issue of the JCI, Li et al. examined the molecular mechanism
of the protective role of SIRT6 in VC in patients with CKD. Using in vitro and
animal models of CKD, the authors demonstrated that SIRT6 prevents VC by
suppressing the osteogenic transdifferentiation of vascular smooth muscle cells
(VSMCs). Mechanistically, SIRT6 bound and deacetylated the runt-related
transcription factor 2 (Runx2), a key transcription factor for osteogenic
differentiation, promoting its nuclear export for proteasome degradation. These
studies provide a pathway in the pathogenesis of VC and justify investigating
SIRT6 as a potential target in CKD.

AUTHORS

Mohamed G. Atta

×

--------------------------------------------------------------------------------

Abstract

Vascular calcification (VC) is regarded as an important pathological change
lacking effective treatment and associated with high mortality. Sirtuin 6
(SIRT6) is a member of the Sirtuin family, a class III histone deacetylase and a
key epigenetic regulator. SIRT6 has a protective role in patients with chronic
kidney disease (CKD). However, the exact role and molecular mechanism of SIRT6
in VC in patients with CKD remain unclear. Here, we demonstrated that SIRT6 was
markedly downregulated in peripheral blood mononuclear cells (PBMCs) and in the
radial artery tissue of patients with CKD with VC. SIRT6-transgenic (SIRT6-Tg)
mice showed alleviated VC, while vascular smooth muscle cell–specific
(VSMC-specific) SIRT6 knocked-down mice showed severe VC in CKD. SIRT6
suppressed the osteogenic transdifferentiation of VSMCs via regulation of
runt-related transcription factor 2 (Runx2). Coimmunoprecipitation (co-IP) and
immunoprecipitation (IP) assays confirmed that SIRT6 bound to Runx2. Moreover,
Runx2 was deacetylated by SIRT6 and further promoted nuclear export via exportin
1 (XPO1), which in turn caused degradation of Runx2 through the
ubiquitin-proteasome system. These results demonstrated that SIRT6 prevented VC
by suppressing the osteogenic transdifferentiation of VSMCs, and as such
targeting SIRT6 may be an appealing therapeutic target for VC in CKD.

Graphical Abstract

Introduction

Vascular calcification (VC), especially in tunica media, is prevalent in
patients with chronic kidney disease (CKD) (1–3). Previous research has revealed
that VC is a major contributor to major adverse cardiovascular events in CKD and
thus is considered an important pathological change in cardiovascular disease
(4–6). Despite severe clinical consequences, the molecular mechanism underlying
VC remains ill defined and no effective therapeutic strategies are currently
available to prevent or halt the progression of VC in CKD. Recent studies
suggest that VC in CKD is a complex and highly regulated process. Patients with
CKD develop hyperphosphatemia, which subsequently promotes the osteogenic
transdifferentiation of vascular smooth muscle cells (VSMCs) (5, 7, 8).
Phosphate-induced (Pi-induced) remodeling of VSMCs is essential for the
mineralization of vascular tissue, and is highly regulated by osteogenic
transcription factor runt-related transcription factor 2 (Runx2, also known as
core-binding factor subunit α1, CBFA1; refs. 9–12). In this context, it is
tempting to suggest that treatment strategies are needed to block osteogenic
transdifferentiation of VSMCs for preventing or halting the progression of VC.
However, inhibition of osteogenic transdifferentiation of VSMCs has not been
developed, and such approaches are still lacking.

Sirtuins (SIRTs) are conserved NAD+-dependent protein deacetylases that have
beneficial effects against aging and metabolic diseases, and have been
recognized as a potential effective target for cardiovascular disease (CVD)
(13–17). They can maintain endothelial homeostatic functions, delay vascular
aging (18), and protect cardiomyocyte against cardiomyocyte hypertrophy (19). In
addition, SIRTs also show a protective role in CKD (20–22). This moderating
effect may indicate that SIRTs are involved in CVD associated with CKD.
Therefore, further understanding of the functional mechanism of SIRTs to serve
as a therapeutic target for CVD, especially in CKD, is needed.

This study explored the role and underlying molecular mechanism of SIRT6 in VC
induced by CKD. Using clinical samples from patients with CKD, we identified
that SIRT6 was decreased in PBMCs and calcified arteries. We explored the effect
of SIRT6 on VC in CKD and osteogenic transdifferentiation of VSMCs both in vivo
and in vitro. We verified that SIRT6 prevented VC in our models, and elaborated
on the molecular mechanism by which it does so. These findings highlight the
critical role of SIRT6 in VC and indicate that SIRT6 may act as a novel
potential therapeutic target for VC in CKD.

Results

SIRT6 reduction is associated with increased risk of VC in patients with CKD.
The expression levels of the sirtuins family (SIRT1-7) in primary human aortic
smooth muscle cells (HAoSMCs) with different calcification status induced by Pi
were detected (Supplemental Figure 1A; supplemental material available online
with this article; https://doi.org/10.1172/JCI150051DS1). As shown in Figure 1A,
the mRNA expression of SIRT6 was the only significantly downregulated SIRT at
different calcification levels. To investigate the association between SIRT6 and
VC, SIRT6 expression was detected by using the mRNA of PBMCs in 39 patients with
CKD with or without VC and 20 healthy people. Patients with CKD presented lower
SIRT6 expression compared with healthy people (Supplemental Figure 1B). Patients
with VC had significantly lower levels of SIRT6 (3.32 ± 1.47 vs. 6.84 ± 1.96, P
< 0.001; Figure 1B) and higher body mass index (24.94 ± 4.06 vs. 22.02 ± 2.10, P
= 0.02; Table 1) than those without VC. SIRT6 expression was inversely
correlated with VC Agatston scores of thoracic aorta (P < 0.001; Figure 1C).
There were no differences in age, sex distribution proportion, kidney function,
and traditional risk factors between the groups with and without VC (systolic
blood pressure [SBP], diastolic blood pressure [DBP], and lipid profile) (Table
1). Von Kossa assays were performed to verify VC, in addition to
immunofluorescence (IF) staining for SIRT6 in radial arteries from patients
undergoing hemodialysis. In tunica media, SIRT6 expression was detected in more
than 65% of nuclei with no calcification of the arteries, while it exhibited
significantly lower expression (about 30% nuclei) in calcified arteries (Figure
1, D and E). These data indicated that SIRT6 expression decreased in VC among
patients with CKD.

Figure 1

Low level of SIRT6 expression was associated with increased risk of vascular
calcification. (A) The qPCR showed expression of SIRT1-7 in WT VSMCs with
different calcification statuses. SIRT4 was not detected in VSMCs (n = 4 per
group). Data were expressed as mean ± SD, *P < 0.05. (B) SIRT6 mRNA levels in
PBMCs from patients with CKD with (n = 27) or without (n = 12) VC. Data were
expressed as mean ± SD. (C) Correlation between the SIRT6 mRNA level and VC
scores in patients with CKD (n = 39, the Pearson’s correlation coefficient R
value and the P value are shown). (D) von Kossa assay and IF staining for SIRT6
in radial arteries sections from hemodialysis patients with CKD (n = 4 per
group). Scale bars: von Kossa 100 μm; IF 50 μm. (E) The bars showing SIRT6
protein expression (mean ± SD; n = 4 per group; AU) in nuclei of aortic tissues
between patients with CKD with and without VC. Statistical significance was
assessed using 1-way ANOVA followed by Dunnett’s test (A) and 2-tailed t tests
(B and E).

Table 1

Baseline characteristics of the study patients with or without vascular
calcification

SIRT6 impedes vascular calcification in vivo and in vitro. To gain insight of
the role of SIRT6 in VC, we induced VC through 2 CKD models (adenine and
phosphorus diet–induced [AP-induced] mode and 5/6 nephrectomy mode) in WT mice.
CKD status promoted VC in WT mice (Figure 2, A and B; Supplemental Figure 2,
A–C). SIRT6 protein expression in calcified aortas was decreased compared with
healthy controls (Figure 2C and Supplemental Figure 2D). We then generated the
SIRT6-transgenic mice (SIRT6-Tg, for stable overexpression of SIRT6) and
subsequently induced VC through CKD status. SIRT6 expression was enhanced in the
aorta of SIRT6-Tg mice (Supplemental Figure 3). Calcification in aorta was
reduced significantly in SIRT6-Tg mice (Figure 2, A and B and Supplemental
Figure 2A). Of note, SIRT6 protein expression in calcified aortas was also
decreased in SIRT6-Tg mice, similar to WT mice (Figure 2C and Supplemental
Figure 2E). Furthermore, we used adeno-associated viral (serotype 2 gene, AAV2)
to specifically knock down SIRT6 expression in VSMCs. AAV2-sh-SIRT6 successfully
reduced SIRT6 expression in aorta but there was no change in kidney
(Supplemental Figure 4). As expected, SIRT6 reduction in aorta induced severe VC
in CKD status (Supplemental Figure 5, A–C).

Figure 2

SIRT6 attenuated VC. (A) Computed tomography (CT) images showing calcification
in the abdominal aorta. The green arrows and circle indicated the calcification
in abdominal aorta of the WT mouse (n = 12 per group). The bar chart shows the
relative VC Agatston score (fold change) of mouse aortas. Scale bars: 10 mm. (B)
Representative von Kossa staining of abdominal aorta sections (n = 12 per
group). Scale bars: 100 μm. (C) Western blot shows SIRT6 protein in abdominal
aorta was reduced in VC. (D and E) VSMCs were exposed to Pi (3.0 mM) for 7 days
and then stained for mineralization by Alizarin red (D), and the quantitative
analysis of calcium content (E) and ALP (F) were detected (n = 3 per group). (G)
SIRT6 protein expression was reduced in WT and SIRT6-Tg VSMCs in response to Pi
(3.0 mM) treatment (n = 4 per group). (H–J) WT and SIRT6-Tg VSMCs were
pretransfected with siSIRT6 or si-negative control (siNC) and then exposed to Pi
(3.0 mM) for 7 days. VSMCs were stained for mineralization by Alizarin red S
(H), and calcium content (I) and ALP (J) were quantified (n = 3 per group).
Statistical significance was assessed using 1-way ANOVA followed by Dunnett’s
test (A, C–F, I, and J). *P < 0.05. All values are mean ± SD.

To better understand the role of SIRT6 in regulating VC, we constructed
experiments on primary VSMCs in vitro. The VSMCs were identified by smooth
muscle myosin heavy chain and SM22α (Supplemental Figure 6). Treated with Pi
(3.0 mmol/L), SIRT6-Tg VSMCs exhibited lower calcium deposition than WT VSMCs,
as evidenced by Alizarin red staining, calcium content assay, and alkaline
phosphatase (ALP) (Figure 2, D–F). SIRT6 expression decreased upon VSMC
calcification (Figure 2G). Furthermore, in vitro loss-of-function analyses were
performed using small interfering RNA (siRNA) or specific SIRT6 inhibitor
OSS-128167. SIRT6 expression was successfully suppressed (Supplemental Figure 7,
A and B). Silencing of SIRT6 in VSMCs resulted in severe calcium deposition and
increased ALP (Figure 2, H–J and Supplemental Figure 7, C–E), which indicated
that SIRT6 deficiency aggravated VC. Collectively, these data suggested that
SIRT6 played a protective role against VC in vivo and in vitro.

SIRT6 suppresses osteogenic transdifferentiation of VSMCs via downregulation of
Runx2. Osteogenic transdifferentiation of VSMCs serve a critical role in VC, so
we explored the potential role of SIRT6 in this process. SIRT6 reduced the
expression of osteogenic markers osteopontin (OPN) and osteocalcin (OCN) and
maintained the expression of contractile property markers α-smooth muscle actin
(α-SMA) and smooth muscle-22α (SM22α) in vivo (Figure 3, A and B and
Supplemental Figure 2F). As expected, SIRT6 restrained the reduction of SM22α
and α-SMA, and downregulated OPN and OCN in SIRT6-Tg VSMCs when treated with Pi
in vitro (Figure 3C and Supplemental Figure 8A). Conversely, the contractile
markers decreased while osteogenic markers increased in VSMCs when treated with
siSIRT6 and OSS-128167 (Figure 3D; Supplemental Figure 8, B–E and Supplemental
Figure 9A). The same results were observed in AAV2-treated mice. SIRT6
deficiency promoted osteogenic transdifferentiation of VSMCs in CKD mice
(Supplemental Figure 5, D and E). Taken together, these results suggested that
SIRT6 protected against VC by suppressing osteogenic transdifferentiation of
VSMCs.

Figure 3

SIRT6 suppresses osteogenic transdifferentiation of VSMCs via regulation of
Runx2. (A) Expression levels of α-SMA and OPN in abdominal arteries of indicated
groups were determined by IF staining (n = 4 per group). Scale bars: 50 μm. (B)
Western blot analysis of osteogenic and contractile property factors expression
in abdominal arteries (n = 3 per group). (C) Analysis of osteogenic and
contractile property factor expression in WT and SIRT6-Tg VSMCs after Pi (3.0
mM) treatment by Western blot (n = 4 per group). (D) VSMCs were pretransfected
with siSIRT6 or siNC, and then incubated with Pi (3.0 mM) for 7 days, and the
downstream osteogenic markers (OPN, OCN) and contractile property markers
(α-SMA, SM22α) were analyzed by Western blot (n = 4 per group). (E) Runx2
expression was analyzed in WT and SIRT6-Tg VSMCs after Pi (3.0 mM) treatment by
Western blot (n = 4 per group). (F–H) SIRT6-Tg VSMCs were pretransfected with
Runx2 plasmid or vector plasmid, and then exposed to Pi (3.0 mM) for 7 days. The
expression of SIRT6 and Runx2 were analyzed by Western blot (F). VSMCs were
stained for mineralization by Alizarin red S (G), and osteogenic markers (OPN,
OCN) and contractile property markers (α-SMA, SM22α) were analyzed by qPCR (n =
3 per group) (H). Statistical significance was assessed using 1-way ANOVA
followed by Dunnett’s test (H). *P < 0.05. All values are mean ± SD.

Since the osteogenic transdifferentiation of VSMCs was highly regulated by Runx2
(9, 10), we next examined whether SIRT6 regulated VC through Runx2. Runx2
expression was much lower in the SIRT6-Tg group in vivo and in vitro (Figure 3,
B and E and Supplemental Figure 2F). Interestingly, the mRNA expression level of
Runx2 had no marked change between the 2 groups (Supplemental Figure 9D).
Additionally, overexpression of Runx2 removed the protective capacity of SIRT6
(Figure 3, F–H and Supplemental Figure 9, B and C). These results demonstrated
that SIRT6 suppressed osteogenic transdifferentiation of VSMCs via
downregulation of Runx2.

SIRT6 deacetylates Runx2 in osteogenic transdifferentiation of VSMCs. We then
sought to investigate the regulatory role of SIRT6 for Runx2. Quantitative PCR
(qPCR) showed that Runx2 mRNA expression was not significantly changed between
SIRT6-Tg and WT groups (Supplemental Figure 9D), which implied that SIRT6 had
little impact on Runx2 transcription. Since SIRT6 is a NAD+-dependent
deacetylase, we hypothesized that SIRT6 regulated Runx2 through influencing its
acetylation status. As shown in IF staining assays, SIRT6 and Runx2 were
colocalized in the nucleus of SIRT6-Tg VSMCs under Pi treatment (Figure 4A). We
confirmed that SIRT6 physically interacted with Runx2 in co-IP assays (Figure 4,
B and C) and this finding was further verified in human embryonic kidney (HEK)
293T cells transfected with HA-tagged Runx2 and Flag-tagged SIRT6 (Figure 4D).

Figure 4

SIRT6 deacetylates Runx2. (A) Representative IF images showing the
colocalization of SIRT6 and Runx2. Scale bars: 50 μm. (B) Anti-SIRT6 IP followed
by Western blot with anti-Runx2 or anti-SIRT6 antibody in SIRT6-Tg VSMCs after
treatment with Pi (3.0 mM) for 7days. Anti-rabbit IgG IP was used as a negative
control. (C) Anti-Runx2 IP in SIRT6-Tg VSMCs after treatment with Pi (3.0 mM)
for 7days. Western blot was carried out with anti-SIRT6 or anti-Runx2 antibody.
Anti-mouse IgG IP was used as a negative control. (D) The anti-HA IP and anti-
flag IP followed by Western blot with anti-HA or anti-flag antibody in HEK-293T
cells infected with HA-Runx2 plasmid, flag-SIRT6 plasmid, or both. Anti-rabbit
IgG IP was used as a negative control. (E) WT and SIRT6-Tg VSMC lysates were
immunoprecipitated with anti-Runx2 antibody and immunoblotted with
anti-acetylated lysine antibody. (F) HEK-293T cells were infected with HA-Runx2
plasmid, flag-SIRT6 plasmid, or both. The anti-HA IP followed by Western blot
with anti-acetylated lysine antibody and anti-HA antibody. (G) SIRT6-Tg VSMCs
were pretransfected with siSIRT6 or siNC together with Pi (3.0 mM) for 7 days
and OSS-128167 or DMSO were incubated with Pi (3.0 mM) for 7 days. The cell
lysates were immunoprecipitated with anti-Runx2 antibody and immunoblotted with
anti-acetylated lysine antibody and anti-Runx2 antibody. All the above
experimental processing were duplicated 3 times.

We then assessed the acetylation level of Runx2. We found that Runx2 acetylation
level decreased in SIRT6-Tg VSMCs compared with WT VSMCs under Pi treatment
(Figure 4E). Similarly, the acetylation level of Runx2 was decreased in HEK-293T
cells transfected with both Flag-SIRT6 and HA-Runx2 compared with cells
transfected with HA-Runx2 alone (Figure 4F). Conversely, the Runx2 acetylation
level was increased when silencing SIRT6 (Figure 4G). Taken together, these
results suggested that SIRT6 deacetylated Runx2 in osteogenic
transdifferentiation VSMCs.

SIRT6 promotes Runx2 degradation via ubiquitin-proteasome system. Since Runx2
acetylation was responsible for its stabilization (23, 24), we investigated if
SIRT6 could influence Runx2 stabilization. The stability of Runx2 protein was
reduced in SIRT6-Tg VSMCs after treatment with the protein synthesis inhibitor
cycloheximide (CHX) (Figure 5A). Conversely, silencing SIRT6 prolonged the
stability of Runx2 (Figure 5, B and C). In addition, SIRT6 protein stability
didn’t show a significant change under Pi treatment (Supplemental Figure 10). To
explore the manner of Runx2 degradation, the proteasome inhibitor MG132 and the
lysosomal proteases inhibitor leupeptin were applied. As shown, leupeptin had no
impact on Runx2 protein stability, but MG132 dramatically enhanced the protein
stability of Runx2 in SIRT6-Tg VSMCs (Figure 5, D and E). These data indicated
that SIRT6-induced Runx2 reduction was mediated by the proteasome but not the
lysosome. Proteasome protein degradation often correlates with the specificity
of target protein ubiquitin, and protein acetylation and ubiquitination are
involved in the regulation of various cellular functions (25, 26). Therefore, we
investigated the ubiquitination levels of Runx2 in SIRT6-Tg and WT VSMCs under
Pi treatment. The ubiquitination level of Runx2 was upregulated in SIRT6-Tg
VSMCs (Figure 5F). Similar results were observed in HEK-293T cells transfected
with HA-Runx2 alone or together with Flag-SIRT6 (Figure 5G). In contrast,
silencing SIRT6 resulted in a decrease of Runx2 ubiquitination in VSMCs (Figure
5H). Moreover, we further explored smad ubiquitin regulatory factor 1 (Smurf1)
expression and its interaction with Runx2, since Smurf1 is a E3 ubiquitin ligase
reported on degradation of Runx2. The results showed that there was less of a
difference in Smurf1 expression between Pi-treated WT and SIRT6-Tg VSMCs.
Interestingly, the interaction between Smurf1 and Runx2 was weaker in WT VSMCs
than in SIRT6-Tg VSMCs (Supplemental Figure 11). These results further
demonstrated that SIRT6 mediated the ubiquitination of Runx2 in VSMCs. Taken
together, these data indicated that SIRT6 promoted Runx2 ubiquitination and
subsequent proteasome-dependent degradation via Runx2 deacetylation.

Figure 5

SIRT6 promotes Runx2 degradation via the ubiquitin-proteasome system. (A) WT and
SIRT6-Tg VSMCs were treated with Pi (3.0 mM) for 7 days and incubated with the
protein translation inhibitor CHX (0.2 mM) for the indicated times before
harvest, followed by immunoblotting with the anti-Runx2 antibody and anti-GAPDH
anti-body. The curve shows the stability of Runx2 protein. (B and C) SIRT6 was
decreased in primary VSMCs via siRNA (B) or specific inhibitor (C) together with
Pi (3.0 mM) incubation for 7 days. The protein translation inhibitor CHX (0.2
mM) was added for indicated times before harvest, followed by immunoblotting
with the anti-Runx2 antibody and anti-GAPDH antibody. The curve shows the
stability of Runx2 protein. (D and E) SIRT6-Tg VSMCs were incubated with Pi (3.0
mM) together with the leupeptin (1.5 μM) (D) or MG132 (10 μM) (E) for 7 days,
and then the protein translation inhibitor CHX (0.2 mM) was added for the
indicated times before harvest, followed by immunoblotting with the anti-Runx2
antibody and anti-GAPDH antibody. The curve shows the stability of Runx2
protein. (F) WT and SIRT6-Tg VSMC lysates were immunoprecipitated with
anti-Runx2 antibody and immunoblotted with anti-ubiquitin (anti-Ub) antibody.
(G) HEK-293T cells were transfected with His-Ub together with HA-Runx2 plasmid,
flag-SIRT6 plasmid, or both. The anti-HA IP was followed by Western blot with
anti-Ub antibody and anti-HA antibody. (H) SIRT6-Tg VSMCs were pretransfected
with siSIRT6 or siNC together with Pi (3.0 mM) for 7 days, and OSS-128167 or
DMSO were incubated with Pi (3.0 mM) for 7 days. The cell lysates were
immunoprecipitated with anti-Runx2 antibody and immunoblotted with anti-Ub
antibody and anti-Runx2 antibody. Statistical significance was assessed using
2-way ANOVA (A–E). All the above experimental processing was duplicated 3 times.

SIRT6 promotes Runx2 degradation through XPO1-dependent nuclear export. A high
Runx2 expression level was observed in calcified aorta from WT mice, while a low
level was detected in SIRT6-Tg mice (Figure 6A). Interestingly, the nuclear
accumulation of Runx2 was more abundant in WT VSMCs than in SIRT6-Tg VSMCs
(Figure 6A). We explored whether the subcellular localization of Runx2 was
related to SIRT6-mediated degradation. IF staining showed that nuclear
accumulation of Runx2 was less predominant in SIRT6-Tg VSMCs under Pi treatment
(Figure 6B). Similar results were found in immunoblotting analysis (Figure 6C).
Conversely, nuclear accumulation of Runx2 was increased when silencing SIRT6
(Figure 6, D and E). These results demonstrated that SIRT6 modulated Runx2
subcellular localization in Pi-treated VSMCs.

Figure 6

SIRT6 mediates Runx2 nuclear export depending on XPO1. (A) Runx2 IF staining was
performed in abdominal arteries. Scale bar: 50 μm. Statistical significance was
assessed using 2-tailed t tests, *P < 0.05. (B) VSMCs were incubated with Pi for
7 days. IF staining was performed for Runx2. Scale bars: 50 μm. (C) VSMCs were
incubated with Pi for 7 days. Cells were harvested and immunoblotted for the
indicated proteins. (D) SIRT6-Tg VSMCs were incubated with Pi for 7 days after
posttransfection of siSIRT6. Cells were harvested and immunoblotted for the
indicated proteins. (E) SIRT6-Tg VSMCs were incubated with Pi together with
nicotinamide for 7 days. Cells were harvested and immunoblotted for the
indicated proteins. (F) SIRT6-Tg VSMCs were transfected with shRNA targeting
XPO1, XPO4, XPO7, or their vector control, and then incubated with Pi for 7 days
after transfection. Nuclear extracts were immunoblotted for Runx2. (G and H)
SIRT6-Tg VSMCs were incubated with Pi together with Leptomycin A (0.5 nM) for 7
days. Cells were harvested and immunoblotted for the indicated proteins (G). IF
staining was performed for Runx2. Scale bars: 50 μm (H). (I) Anti-XPO1 IP
followed by Western blot with anti-Runx2 or anti-XPO1 antibody in SIRT6-Tg VSMCs
after treatment with Pi for 7 days. Anti-rabbit IgG IP was used as negative
control. (J) Anti-Runx2 IP in SIRT6-Tg VSMCs after treatment with Pi for 7 days.
Western blot was carried out with anti-XPO1 or anti-Runx2 antibody. Anti-mouse
IgG IP was used as negative control. (K) SIRT6-Tg VSMCs were incubated with Pi
together with Leptomycin A for 7 days, and then CHX (0.2 mM) was added for the
indicated times before harvest, followed by immunoblotting for the indicated
proteins. (L) Curve shows the stability of Runx2 and was assessed using 2-way
ANOVA. Pi treatment is 3.0 mM. All the above experimental processing was
duplicated 3 times.

It has been reported that importin β superfamily members exportin-1 (XPO1),
exportin-4 (XPO4), and exportin-7 (XPO7) are related to protein nuclear export
(27). Therefore, we knocked down these genes (Supplemental Figure 9B) to
investigate their potential regulation of this process. Silencing XPO1, but not
the other 2 members, abrogated the SIRT6-induced redistribution of Runx2 (Figure
6, F–H). Furthermore, we examined Runx2–XPO1 interaction by IP and found that
Runx2 directly binds to XPO1 (Figure 6, I and J). Inhibiting XPO1 by leptomycin
A treatment can prolong the stability of Runx2 in SIRT6-Tg VSMCs (Figure 6, K
and L). Taken together, our data suggested that SIRT6-mediated Runx2
deacetylation resulted in redistribution of Runx2 through XPO1.

SIRT6 impedes vascular calcification depending on nuclear export of Runx2. We
performed additional experiments to confirm the nuclear export role of XPO1 in
VC attenuation mediated by SIRT6. As expected, XPO1 inhibitor treatment
significantly increased calcium deposition in both SIRT6-Tg and WT VSMCs (Figure
7, A–C). Similarly, Leptomycin A inhibition of XPO1 reversed the suppressive
role of SIRT6 in osteogenic transdifferentiation of VSMCs (Figure 7, D and E).
Based on these findings, we concluded that XPO1 played a critical role in
SIRT6-mediated VC attenuation.

Figure 7

Nuclear export of Runx2 is a key component of SIRT6 vascular calcification
suppressor function. (A–C) WT and SIRT6-Tg VSMCs were incubated with Pi (3.0 mM)
together with Leptomycin A for 7 days. VSMCs were stained for mineralization by
Alizarin red S (A), and calcium content (B) and ALP (C) were quantified (n = 3
per group). (D and E) The osteogenic markers (OPN, OCN) and the contractile
property markers (α-SMA, SM22α) were analyzed by qPCR for the WT (D) and
SIRT6-Tg VSMCs (E) mouse being incubated with Pi (3.0mM) together with
Leptomycin A for 7 days (n = 3 per group). Statistical significance was assessed
using 1-way ANOVA followed by Dunnett’s test (B–E). *P < 0.05. All values are
mean ± SD.

Discussion

In this study, we elucidated a novel SIRT6/Runx2 pathway in vascular
calcification. For the first time, we found that SIRT6 suppressed VSMC
osteoblastic transdifferentiation and attenuated VC both in vivo and in vitro.
Mechanistically, SIRT6 deacetylated Runx2 and promoted its ubiquitination and
subsequent degradation through the ubiquitin-proteasome system.

There are 7 sirtuins (SIRT1-7) in mammals and each family member has a different
function and subcellular localization. The common molecular targets suggest that
sirtuins might act synergistically. Here, using VSMC calcification in vitro, we
showed that all members of the sirtuin family except SIRT4 are expressed in
VSMCs. It’s known that SIRT1 is implicated in the transcriptional and epigenetic
modifications of cellular and systemic processes. SIRT1 has proved to act in a
protective role against VC (28, 29). However, SIRT1 modulators have not seen
marked results in clinical studies (13). In this study, we found that only SIRT6
not SIRT1 was significantly downregulated at different calcification levels. The
result indicated that SIRT6 played a critical role in VC.

SIRT6 is mainly located in the nucleus, and it is a class IV sirtuin that
exhibits deacetylase and ADP-ribosyltransferase activity. SIRT6 is known to
exert a protective role in atherogenesis and ischemic stroke, and act against
VSMC differentiation in response to the cyclic strain (30–32). SIRT6 plays a
role in a variety of biological processes, and it is responsible for a set of
age-related disorders (33). CKD is one of the most typical age-related metabolic
diseases. However, the association between SIRT6 and VC in CKD remains unknown.
Using 2 canonical CKD models (adenine-induced and 5/6 nephrectomy-induced CKD
mice), we reported that VC was less prominent in SIRT6-Tg mice than the WT
controls. And SIRT6 prevented VC of VSMCs induced by Pi in vitro. VSMC-specific,
SIRT6 knock down of aorta by AAV2 caused severe VC in the WT mouse model. In our
clinical study, a lower expression level of SIRT6 was observed in calcified
radial arteries and PBMCs of patients with CKD with VC. No significant
differences were observed in kidney function or traditional risk factors between
those with or without VC. Thus, these findings indicated that SIRT6 may act as a
protective regulator in vascular calcification and its protective effect was
independent of renal function changes.

Previous studies have demonstrated that the phenotypic transdifferentiation of
VSMCs, from contractile to osteochondrogenic, is a pro-calcifying process and
appears to initiate before mineral deposition (9, 10, 34). During this process,
the osteoblastic features of VSMCs predominate, with decreased expression of
contractile proteins (α-SMA and SM22α) and increased levels of the synthetic
proteins (OPN and OCN). We investigated the effect of SIRT6 on phenotypic
transdifferentiation of VSMCs. SIRT6 can reverse protein expression and mRNA
level of α-SMA and SM22α and reduce protein expression and mRNA transcription of
synthetic proteins such as OPN and OCN during the process of VC. Thus, the
protective role of SIRT6 in VC attenuation was potentially mediated by
inhibiting the phenotypic transdifferentiation of VSMCs. Upregulation of Runx2
expression has been observed in vascular calcification and its core role in
VSMCs osteochondrogenic differentiation has been well documented (35–38).
Posttranslational modifications of Runx2 can influence its stability and
transcriptional activity. Runx2 can be phosphorylated by Erk/MAPK (24) and Akt
(39). In atherosclerotic calcification, AMPKα1 promotes Runx2 SUMOylation,
decreasing its stability (40). PTEN/AKT also modulated Runx2 ubiquitination via
phosphorylating FOXO1/3 in VSMC calcification (41). In addition, enhancing
acetylation of Runx2 promotes its stability and transcriptional activity
(42–44).

We found that protein expression of Runx2 was significantly decreased in a SIRT6
overexpression VC model in vivo and in vitro. Enhancing Runx2 expression via
plasmid reversed the protective effect of SIRT6 in vitro. This indicated Runx2
was regulated by SIRT6. The transcription level of Runx2 was not significantly
affected by SIRT6, so we hypothesized that posttranslational regulation of Runx2
may be involved. Emerging evidence has shown that among the posttranslational
modifications of Runx2 (24, 26, 45–47), SIRT6 is a deacetylase that could
deacetylate the lysine residues of histone and nonhistone substrates, which is
closely related to protein degradation via ubiquitination (48, 49). It has been
reported that acetylation of Runx2 plays an important role in osteogenesis (50).
Here, we found that Runx2 acetylation was reduced in VSMCs with SIRT6
overexpression, and identified physical interaction between Runx2 and SIRT6
proteins via co-IP assay. At the same time, reduction of Runx2 protein in the
SIRT6 overexpression group was attributed to a shorter half-life. Normally, the
acetylation of Runx2 could protect against the ubiquitin-proteasome degradation
process (26). Inhibition of the proteasome via MG132 prevented SIRT6-mediated
downregulation of the Runx2 protein. As expected, the ubiquitinated Runx2 was
increased in SIRT6-Tg VSMCs, and the ubiquitinated Runx2 was almost abrogated in
SIRT6-deficient WT VSMCs. These data indicated that SIRT6 was vital for
ubiquitin-dependent proteolysis of Runx2. As reported in previous studies,
Smurf1-mediated degradation of Runx2, and Runx2 acetylation, inhibited this
interaction. We also found that the combination/interaction of Smurf1 and Runx2
was weaker in WT VSMCs than in SIRT6-Tg. Collectively, these data suggested that
SIRT6 deacetylates Runx2, which was subsequently ubiquitinated, and degraded
through the proteasome.

Runx2 undergoes diverse posttranslational modifications, some of which may
regulate its subcellular distribution, and nuclear-cytoplasmic shuttling of
Runx2 may regulate cell fate (51). However, the subcellular distribution of
Runx2 has not been explored in VC. Our data suggested that nuclear levels of
Runx2 were higher in WT than in SIRT6-Tg VSMCs. There was an increase in the
Runx2 nuclear fraction under SIRT6 deficiency. We explored the Runx2 nuclear
export mechanism and identified XPO1 as the specific transporter, in accordance
with studies that reported that XPO1 regulated Runx2 nuclear-cytoplasmic
shuttling (51, 52). Our in vitro VC models revealed that the XPO1 was vital for
SIRT6-mediated attenuation of VC and we also observed that reduced nuclear
export of Runx2 can prolong its half-life. These findings demonstrated a unique
mechanism of Runx2 degradation, which was mediated through
deacetylation-dependent Runx2 nuclear export.

Our previous study demonstrated alkB homolog 1 (Alkbh1) upregulation on the
progression of VC via activation of the osteogenic protein, bone morphogenetic
protein 2 (BMP2; ref. 53).Runx2 was a major target of BMP2 pathway and BMP2 was
proved to regulate the acetylation and ubiquitination level of Runx2 (26, 43).
Thus, BMP2 and Runx2 cooperatively interact to induce VC. These indicate that
SIRT6 upregulation may play an important role against the BMP2 pathway in VC. In
addition, further studies are required to demonstrate the exact regulatory
effect of Alkbh1/BMP2 pathway on SIRT6 expression in VC. SIRT6 is known for
improving longevity, modulating genome stability and telomere integrity, and
reducing oxidative stress and inflammation (14, 33, 54). It has also been
reported that Runx2 negatively regulates SIRT6 expression at both the
transcriptional and posttranslational levels in breast cancer (55). In our
results, we found that Runx2 did not play a role in regulation of SIRT6
expression in VSMCs (neither in transcription nor posttranslation). SIRT6
expression was significantly associated with disease status of blood vessels,
and SIRT6 expression data from PBMCs can be used as a disease marker for
predicting calcification in patients with CKD. Further studies are needed to
demonstrate the relationship between PBMCs and VSMC calcification.

Collectively, our studies demonstrate for the first time that SIRT6 prevents VC
through posttranslational regulation of Runx2 activity and stability. These
findings suggest that SIRT6 may be an innovative therapeutic strategy for VC.

Methods

CKD patient samples. Peripheral blood samples from patients with CKD and healthy
people were collected from Donghua Hospital of Sun Yat-sen University from
November 2019 to January 2020. Thirty-nine patients with CKD and 20 healthy
people were recruited to this study. PBMCs from peripheral blood were extracted
using Histopaque-1077 (Sigma) gradient. The extract mixture was centrifuged at
400g for 20 minutes and the interface was collected as PBMCs. Clinical and
biochemical parameters were collected from the patient electronic medical
records in the hospital. The radial arteries from patients with CKD undergoing
hemodialysis were collected from The Eighth Affiliated Hospital of Sun Yat-sen
University from November 2019 to January 2020.

Assessment of thoracic aorta calcification score. Patients underwent a chest
multi-detector computed tomography (MDCT) scan with standard
electrocardiographically (ECG) gated protocol to evaluate thoracic aorta
calcification. Agatston scores of images were blind-quantified by 3 independent
investigators with Siemens Syngo CT Workplace software according to standard
criteria (56). The thoracic aorta refers to the section between the ascending
and descending aorta. To measure calcification scores, the CT images were
reconstructed with 1 mm–thick slices. The presence of calcification was defined
as Agatston score in the present study.

Induction of VC in mice. Male mice were used in this study to avoid the
potential interference of changing levels of hormones on VC. WT C57BL/6J mice at
8 weeks and weighing 25 to 30 grams were purchased from the Laboratory Animal
Center of Sun Yat-sen University. Cloned mSirt6 cDNA with CAG promoter was
injected into fertilized eggs to constructed Sirt6-transgenic mice (SIRT6-Tg) of
C57BL/6J background as was previously reported (57). The phenotype of SIRT6-Tg
mice and genotyping identification procedure were identified by One Step Mouse
Genotyping Kit (Vazyme) according to the manufacturer’s instructions. Tail DNA
was used to confirm mice positive for the transgene at 2 to 3 weeks of age. The
following primers were used for genotyping: forward,
5′-GCCGTCTGGTCATTGTCAACCTG-3′; reverse, 5′-AAAGACCCCTAGGAATGCTCGTCAA-3′.
Eight-week-old SIRT6-Tg mice weighing 25 to 30 grams were used for these
experiments. All mice were raised in the Laboratory Animal Center of Sun Yat-sen
University and were maintained in a temperature-controlled room on a 12-hour
light/dark cycle with available access to food and water. WT and SIRT6-Tg mice
were randomly assigned to experimental groups with at least 12 animals in each
group: the control group was fed with standard pellet chow diet (normal diet,
ND) and the CKD model group was fed with special chow containing 0.75% adenine
and high (1.5%) levels of phosphorus (AP diet) or performed a 5/6 nephrectomy
model. After 12 weeks of AP diet or 8 weeks of high phosphorus diet after 5/6
nephrectomy, the animals were analyzed to confirm the vascular calcification of
aorta, and then sacrificed. The aorta was harvested from each animal and was
kept at –80°C for further use. The VC Agatston scores of aortas were analyzed by
3 independent investigators and the score was normalized to the lowest score
(not zero) in SIRT6-Tg group. For VSMC-specific SIRT6 knockdown, the WT mice
were injected in the lateral tail vein with recombinant AAV serotype 2 gene
transfer vectors bearing a VSMC-specific promoter combination (SM22α promoter)
with mouse sh-SIRT6 sequence. After 4 weeks, some of the mice were sacrificed
and aortas and kidneys were collected. Western blot was used to confirm the
efficiency of AAV-sh-SIRT6 in aortas and kidneys. The remaining mice were
treated with AP diet for 12 weeks, or a 5/6 nephrectomy was performed and then
mice were fed with high phosphorus diet for another 8 weeks. Then the mice were
sacrificed and aortas were collected. The detailed protocols were shown in our
previous study (53). The AAV2 was generated by Hanbio.

Cell culture. Primary HAoSMCs were purchased from ATCC and cultured in DMEM
containing 10% FBS supplemented with 100 U/mL penicillin, 100 μg/mL
streptomycin.

Mice VSMCs were isolated from 6-week-old SIRT6-Tg mice and WT C57BL/6J control
mice. Briefly, the adventitia and endothelium were removed from the thoracic
aortic arteries and the remaining tissue was cut into approximately 1 mm2
sections. Aorta segments were placed in cell culture dishes with DMEM containing
10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin in a 37°C incubator
with 5% CO2 for 5 to 7 days. The VSMCs migrated from the explants, and cells
between passages 5 and 8 were used in experiments.

VSMCs calcification induction. To induce calcification, VSMCs at 80% confluence
were incubated in DMEM containing 10% FBS, 100 U/mL penicillin, and 100 μg/mL
streptomycin, with the addition of 3.0 mmol/L sodium phosphate (Pi) (Sigma) and
cultured at 37°C in an incubator containing 5% CO2 for 7 days. The medium and Pi
were refreshed every 2 days. The control VSMCs were treated with DMEM containing
10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin, but without Pi, and
the medium was also refreshed every 2 days.

von Kossa assay. To examine aorta calcification, slides were dehydrated and
rinsed rapidly in double distilled water. The vascular tissue sections were then
incubated with 5% silver nitrate solution and exposed to ultraviolet light for 1
hour until color development was complete. Next, the slides were incubated with
5% sodium thiosulfate and washed with double distilled water. The slides were
photographed by microscopy (Nikon). Calcified nodules were stained brown to
black.

Alizarin red staining. At collection time points, medium was removed and
cultured VSMCs were washed with 4°C PBS 3 times (3 minutes each wash), and then
cell layers were fixed in 4% paraformaldehyde in PBS for 20 minutes. Next, the
paraformaldehyde was removed and the cells were washed in distilled water 3
times (2 minutes each wash). The cells were then exposed to Alizarin red
staining solution (pH 4.2, 1%) for 30 minutes at room temperature, then washed
again with distilled water. Positively stained VSMCs presented a reddish color
to indicate the calcification.

Calcium and ALP quantification. Aortic tissues without adventitia were incubated
with 0.6 mol/L HCl overnight at 37°C. The supernatant of these tissues was then
collected. The cultured VSMCs were washed softly with PBS for 3 times (2 minutes
each wash) and incubated with 0.6 mol/L HCl overnight at 4°C. The supernatant
was collected. Calcium content was determined by using a commercial kit (Biosino
Bio-Technology and Science) according to the manufacturer’s instructions. VSMCs
or aortic tissues were equilibrated with 1% Triton X-100 in 0.9% saline on ice
and the supernatant was collected for ALP quantification assay after
centrifugation in a microfuge at 8000g for 5 minutes. ALP activity was analyzed
using a commercial assay kit (Biosino Bio-Technology and Science). Results are
shown normalized to total protein levels.

Quantitative real-time PCR. Total RNA was extracted from aortic tissue and VSMCs
by using Trizol Reagent (Takara) according to the manufacturer’s instructions.
For mRNA quantification, a PrimeScriptRT Reagent Kit (Takara) was used for RNA
reverse transcription into cDNA. Real-time PCR was performed with SYBR Green
(Takara) and data were collected and analyzed using a LightCycler 96 real-time
system (Roche Diagnostics). Relative quantification was calculated according to
the 2ΔΔCt method, with GAPDH level as a reference. The primer sequences are
listed in Supplemental Table 1.

Transfection and transduction of VSMCs and HEK-293T cells. For siRNA and shRNA
transfection, VSMCs were plated at 5 × 105 cells in 6-well plates. At 50%
confluence, cells transfected with specific siRNA at a final concentration of 10
nmol/L with Lipofectamine 3000 (Invitrogen) according to the manufacturer’s
instructions. After 6 hours of transfection with opti-MEM, the DMEM containing
10% FBS was replaced. The full-length of the target gene cDNA was amplified from
a mouse cDNA library using standard PCR techniques and inserted into pcDNA3.1.
For plasmid transfection, cultured VSMCs or HEK-293T cells were transfected with
specific plasmids by Lipofectamine 3000 regent according to the manufacturer’s
instructions. The relative siRNA and shRNA are listed in Supplemental Table 2.

Immunofluorescence staining and immunohistochemistry. The VSMCs were first
washed with 1× PBS 3 times, and then fixed with 4% paraformaldehyde solution for
20 minutes. Next, the paraformaldehyde was removed and cells were washed in PBS
3 times. Cells were permeabilized using 0.1% Triton-X. After another 3 PBS
washes, cells were incubated with 5% BSA for 30 minutes. Following this, the
primary antibody for rabbit anti-SIRT6 (Abcam) or mouse anti-Runx2 (Abcam) was
incubated overnight at 4°C. FITC-labeled (Sigma) or Alexa Fluor 647–labeled
secondary antibodies (Abcam) were incubated for 1 hour at room temperature. DAPI
(Solarbio) for staining nuclei was incubated for 5 minutes at room temperature
and then cells were washed in PBS 3 times. Imaging was performed using Olympus
IX73fluorescence microscope (Olympus). The antibody details can be found in
Supplemental Table 3.

Radial arteries from hemodialysis patients and mice aortic tissues were
formalin-fixed and further embedded with paraffin. For immunostaining, tissue
sections were deparaffinized in xylene and rehydrated through a graded alcohol
series to distilled water. Antigen retrieval was performed by microwave
irradiation in ethylene diamine tetraacetic acid (EDTA). Then tissue sections
were incubated with 5% normal goat serum in PBS/0.1% Triton X-100 for 1 hour at
room temperature to reduce nonspecific background staining. Sections were then
incubated overnight at 4°C with primary antibody for rabbit anti–α-SMA
(Abclone), rabbit anti-OPN (Proteintech) or rabbit anti-Runx2 (CST). For IF,
binding of primary antibodies was visualized using goat anti-rabbit FITC-labeled
antibody incubated for 1 hour at room temperature. Nuclei were counterstained
with DAPI. Prolong Gold antifade reagent was used to decrease fluorescence
quenching of the slides. For IHC, expression of SIRT6 in WT and SIRT6-Tg mouse
was stained with SIRT6 antibody by universal SP kit (ZSGB-BIO) according to the
manufacturer’s instructions. The images were collected with an Olympus IX73
fluorescence microscope. The primary antibodies are listed in Supplemental Table
3.

Nuclear/cytoplasmic extraction. At collection time points, culture medium was
removed and then VSMCs were washed with 1× PBS for 3 times. The nuclear and
cytoplasmic protein lysate extraction of VSMCs was performed using the Nuclear
Protein Extraction Kit (Solarbio) according to the manufacturer’s
recommendations.

Immunoprecipitation and Western blot analysis. Harvested VSMCs and HEK-293T
cells were lysed with lysis buffer (Beyotime) together with protease and
phosphatase inhibitors on ice for 15 minutes. The lysate was then sonicated on
ice at 10% power for 2 minutes. After centrifugation at 12,000g for 20 minutes
at 4°C, the supernatant was precleared by incubation with protein A+G magnetic
beads (Millipore) and IgG (CST) for 1 hour at 4°C. The samples were then place
in a magnetic separator for 1 minute. The supernatant was incubated with
indicated antibody overnight at 4°C on a rotating platform. Protein A+G magnetic
beads were then added to the supernatants and incubated for 2 hours at room
temperature. The immunocomplexes were washed 3 times with the lysis buffer,
boiled at 95°C for 10 minutes with 2× SDS sample buffer, and analyzed by Western
blot. For Western blot analysis, the cells lysates or tissue pieces were
prepared by adding the lysis buffer on ice for 15 minutes, supplemented with
protease and phosphatase inhibitors, scraping into a 1.5 mL tube, and
centrifuging for 20 minutes at 12,000g at 4°C. The protein content was measured
by enhanced BCA protein assay kit (Beyotime). The proteins were boiled in
loading buffer (Beyotime) at 100°C for 10 minutes. Equal amounts of proteins
were separated on SDS-polyacrylamide gels and transferred to PVDF membranes
(Millipore). The membranes were incubated with the primary antibodies overnight
at 4°C. The membranes were then incubated with secondary anti-rabbit (CWBIO) or
anti-mouse (CWBIO) HRP-conjugated antibody (diluted 1:10,000) for 1 hour at room
temperature. Antibody binding was detected with ECL detection reagent
(Millipore). The relative quantification of immunoblots was analyzed by
grayscale in ImageJ. The antibodies used in this study are listed in
Supplemental Table 3.

Statistics. All data are mean ± SD. Statistical analyses were performed with the
Graphpad Prism v6.00 for Windows (GraphPad Software Inc.). Student’s t test was
used to compare 2 groups and 1-way ANOVA followed by Dunnett’s test was used for
more than 2 groups. VC Agatston scores were nonnormalized parameters, and
logarithmic transformation of VC Agatston scores was used in correlation
analysis (Pearson Correlation Analysis). Statistical significance was accepted
at P less than 0.05.

Study approval. All the related procedures for collection of the samples of
patients with CKD and normal people were performed with the approval from the
internal review and ethics board of Donghua Hospital of Sun Yat-sen University
and The Eighth Affiliated Hospital of Sun Yat-sen University. All participants
signed informed consent before entering this study. Experimental animal
protocols were approved by the Institutional Animal Care and Use Committee of
Sun Yat-sen University.

Author contributions

HH conceived the project. WL, WF, ZL, and BL performed and analyzed in vivo
experiments. WL, WF, and Y Zhou performed the in vitro experiments and analyzed
the data. XS and MZ performed and analyzed the biochemical and biophysical
experiments. DL and Y Zhu were responsible for human clinical and molecular
genetic studies. WL, DL, and JC wrote the paper with input from all authors. The
images were photographed by WL. The order of co–first authors was determined by
their efforts and contributions to the manuscript.

Supplemental material

View Supplemental data

Acknowledgments

The authors would like to thank Long Shuang Huang (The University of Illinois
College of Medicine) for useful suggestions regarding the graphical
representation and the experimental design. The authors would also like to thank
Jessica Tamanini (Shenzhen University and ETediting) for editing the manuscript
prior to submission. This work was supported by National Key Research and
Development Program (grant 2020YFC2004405); the the National Natural Science
Foundation of China (grants 8201101103, 81870506, 91849208, 81670676, and
81422011); Project of Traditional Chinese Medicine in Guangdong Province (grant
20201062); Basic Research Project of Shenzhen Science and Technology Innovation
Committee (grants JCYJ20180306174648342 and JCYJ20190808102005602); Shenzhen
Futian District Public Health Research Project (grant FTWS2019003); and the
Shenzhen Key Medical Discipline Construction Fund (grant SZXK002), all to HH.
This work was also supported by the National Natural Science Foundation of China
(grant 82073408 to JC) and Dongguan Social Science and Technology Development
Project (grant 2018507150461629 to XS).

Footnotes

Conflict of interest: The authors have declared that no conflict of interest
exists.

Copyright: © 2022, Li et al. This is an open access article published under the
terms of the Creative Commons Attribution 4.0 International License.

Reference information: J Clin Invest.
2022;132(1):e150051.https://doi.org/10.1172/JCI150051.

See the related Commentary at A molecular target of vascular calcification in
chronic kidney disease.

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