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Department of Cardiovascular Sciences, Center for Cardiovascular Regeneration, Houston Methodist Research Institute, Houston, Texas, USADepartment of Ophthalmology, Blanton Eye Institute, Houston Methodist Institute for Academic Medicine, Houston Methodist Research Institute, Houston, Texas, USADepartment of Cardiothoracic Surgeries, Weill Cornell Medical College, Cornell University, New York City, New York, USA
Cardiovascular Biology Research Program, Oklahoma Medical Research Foundation, Oklahoma City, Oklahoma, USADepartment of Cell Biology, University of Oklahoma Health Sciences Center, Oklahoma City, Oklahoma, USA
Metabolic flexibility is the capacity of cells to alter fuel metabolism in response to changes in metabolic demand or nutrient availability. It is critical for maintaining cellular bioenergetics and is involved in the pathogenesis of cardiovascular disease and metabolic disorders. However, the regulation and function of metabolic flexibility in lymphatic endothelial cells (LECs) remain unclear. We have previously shown that glycolysis is the predominant metabolic pathway to generate ATP in LECs and that fibroblast growth factor receptor (FGFR) signaling controls lymphatic vessel formation by promoting glycolysis. Here, we found that chemical inhibition of FGFR activity or knockdown of FGFR1 induces substantial upregulation of fatty acid β-oxidation (FAO) while reducing glycolysis and cellular ATP generation in LECs. Interestingly, such compensatory elevation was not observed in glucose oxidation and glutamine oxidation. Mechanistic studies show that FGFR blockade promotes the expression of carnitine palmitoyltransferase 1A (CPT1A), a rate-limiting enzyme of FAO; this is achieved by dampened extracellular signal–regulated protein kinase activation, which in turn upregulates the expression of the peroxisome proliferator–activated receptor alpha. Metabolic analysis further demonstrates that CPT1A depletion decreases total cellular ATP levels in FGFR1-deficient rather than wildtype LECs. This result suggests that FAO, which makes a negligible contribution to cellular energy under normal conditions, can partially compensate for energy deficiency caused by FGFR inhibition. Consequently, CPT1A silencing potentiates the effect of FGFR1 knockdown on impeding LEC proliferation and migration. Collectively, our study identified a key role of metabolic flexibility in modulating the effect of FGFR signaling on LEC growth.
). In addition to skeletal muscle, metabolic flexibility also occurs in a few other tissues (e.g., adipose tissue) and cell types (e.g., pancreatic β-cells, cardiomyocytes) to help ensure proper cellular and physiological functions (
). Notably, the impairment of metabolic flexibility is strongly associated with obesity, type 2 diabetes mellitus, and heart failure and impacts the pathophysiological processes underlying these diseases (
). Excessive lymphangiogenesis (i.e., lymphatic vessel formation) is involved in inflammatory disease and tumor metastasis, while insufficient lymphatic vessel growth may cause lymphedema, a medical condition that is characterized by swelling because of abnormal accumulation of interstitial fluid in tissues (
). In contrast, the role of fibroblast growth factor (FGF) signaling in lymphatic vessel formation is much less understood. The FGF family of growth factors is composed of 22 members, with FGF2 as a robust mitogen to stimulate lymphangiogenesis both in vitro and in vivo (
). However, during lymphatic vessel formation in vivo, the effect caused by loss of FGFR1 can be compensated by FGFR3, which is upregulated in LECs during lymphatic differentiation and is also required for lymphangiogenesis (
). Knockdown of FGFR1—the most abundant member of the FGFR family—in LECs selectively reduces the expression of the glycolytic enzyme hexokinase 2 (HK2), and consequently suppresses glycolysis and ATP production, which are required for active angiogenic behavior of LECs (
). Recent studies further demonstrate that glutamine metabolism is crucial for EC proliferation and angiogenesis by maintaining tricarboxylic acid cycle anaplerosis, asparagine synthesis, and redox balance (
). Despite the importance of glucose, glutamine, and fatty acids, how metabolism of these different substrates is coordinated to support LEC growth is unknown.
In our current study, we sought to determine the impact of FGFR signaling on utilization of glucose, glutamine, and fatty acids in LECs. During this process, we unexpectedly discovered a balance between glycolysis and FAO, which plays an important role in regulating FGFR-mediated energy generation and angiogenic behavior of cultured LECs.
FGFR inhibition upregulates FAO while reducing glycolysis in LECs
To assess the potential impact of FGFR signaling on glucose oxidation, glutamine oxidation, and FAO, we treated proliferating human dermal LECs (HDLECs) with ASP5878, a highly specific FGFR inhibitor (
), for 2 days. HDLECs were cultured in EC growth medium and kept at subconfluency throughout the experiments to maintain them in a proliferative state. At the end of ASP5878 treatment, we incorporated 5-3H-glucose, 6-14C-glucose, 14C(U)-glutamine, 9,10-3H-palmitic acid, or 1-14C-oleic acid into the culture medium and calculated flux of glycolysis, glucose oxidation, glutamine oxidation, and FAO through the measurement of 3H2O or 14CO2 generation. Consistent with our previous report (
), ASP5878 treatment drastically reduced glycolytic flux (Fig. 1A). FGFR inhibition also impaired glucose oxidation but did not affect glutamine oxidation (Fig. 1, B and C). In contrast, FAO in HDLECs was significantly enhanced after ASP5878 treatment (Fig. 1, D and E). To further confirm these observations that were generated with the FGFR chemical inhibitor, we transfected HDLECs with nontargeting or FGFR1-specific siRNA. Three days following siRNA transfection, we trypsinized control or FGFR1 siRNA-transfected HDLECs and replated them into new culture dishes to obtain subconfluent cells for analyses the next day. Western blotting showed that FGFR1 proteins were effectively depleted by siRNA treatment (Fig. 1, F and G). We further used 5-3H-glucose, 9,10-3H-palmitic acid, and 1-14C-oleic acid to examine the effect of FGFR1 knockdown on flux of glycolysis and FAO. Our data showed that FGFR1 siRNA treatment significantly upregulated FAO while reducing glycolysis (Fig. 1, H–J). Together, our findings suggest the existence of a balance between glycolysis and FAO in LECs and reveal the importance of FGFR signaling in maintaining this balance.
FGFR blockade increases CPT1A expression for promoting FAO in LECs
Next, we sought to elucidate the molecular mechanism by which FGFR signaling inhibition enhanced FAO. Because CPT1A promotes FAO as a rate-limiting step in LECs and plays a critical role in lymphatic vessel formation (
), we examined whether FGFR activity regulates CPT1A expression. Treatment of HDLECs with ASP5878 for 2 days increased CPT1A protein levels while reducing the expression of HK2 (Fig. 2, A–C), which is consistent with our previous report (
). FGFR1 knockdown also led to reduction of HK2 and upregulation of CPT1A expression, albeit to a lesser extent than with the FGFR inhibitor (Fig. 2, D–G). We further confirmed these observations with a second siRNA, which targets a different region in FGFR1 mRNA (Fig. S1, A–C). We then assessed whether CPT1A elevation was required for the effect of FGFR inhibition on FAO. Our data showed that depletion of CPT1A expression by siRNAs prevented ASP5878 or FGFR1 siRNA treatment from increasing FAO flux in HDLECs (Fig. 2, H–J). Collectively, our findings suggest that FGFR signaling regulates FAO by controlling CPT1A expression.
FAO inhibition enhances the effect of FGFR1 knockdown on suppressing energy generation, proliferation, and migration of LECs
We next explored the functional significance of FGFR inhibition–induced FAO upregulation for LECs. Previous studies show that FAO inhibition does not cause energy stress in ECs (
). Consistently, we also found that CPT1A knockdown in HDLECs did not affect total ATP levels (Fig. 3A). However, CPT1A deficiency significantly potentiated the effect of FGFR1 siRNA on suppressing ATP production (Fig. 3A). We then examined the impact of FAO inhibition on LEC proliferation and migration, which are critically involved in lymphatic vessel formation. Our data demonstrated that although knockdown of CPT1A or FGFR1 alone reduced LEC proliferation, their combined depletion resulted in more dramatic impairment (Fig. 3B). We further used an in vitro scratch assay to determine the role of FAO in LEC migration. We found that CPT1A knockdown did not affect migration in wildtype LECs (Fig. 3, C and D), which was consistent with a previous report (
). However, CPT1A depletion aggravated LEC migration defects caused by FGFR1 knockdown (Fig. 3, C and D).
Suppression of glycolysis per se does not lead to upregulation of CPT1A expression in LECs
To test whether CPT1A upregulation was secondary to an impairment of glycolysis caused by FGFR inhibition, we treated HDLECs with 2-deoxy-d-glucose (2DG), a potent inhibitor of glycolysis. Our data showed that 2DG failed to increase CPT1A levels (Fig. S2, A and B). These data suggest that FGFR signaling controls CPT1A expression independent of glycolysis.
FGFR1 depletion upregulates CPT1A expression in LECs through extracellular signal–regulated protein kinase signaling
One major pathway downstream of FGFR activation is RAF–MEK–extracellular signal–regulated protein kinase (ERK) signaling (
). Therefore, we sought to determine whether ERK signaling is involved in FGFR inhibition–induced CPT1A upregulation in LECs. We first examined ERK phosphorylation (Thr202/Tyr204) in HDLECs transfected with control or FGFR1 siRNA. We found that FGFR1 knockdown reduced levels of phosphorylated ERK while elevating CPT1A expression (Fig. 4A and Fig. S3A). Next, to investigate whether FGFR1 deficiency–caused CPT1A increase was due to impaired ERK signaling, we took advantage of RAF1S259A, an RAF1 gain-of-function mutation that can activate ERK in ECs (
). Consistent with the previous studies, transduction of HDLECs with a lentiviral vector expressing RAF1S259A increased ERK phosphorylation (Fig. 4B). Importantly, we found that CPT1A levels were reduced concurrently with ERK activation in these cells (Fig. 4B). Moreover, RAF1S259A expression was sufficient to normalize CPT1A expression in FGFR1 siRNA-transfected HDLECs (Fig. 4C). We also treated HDLECs with 20 μM of U0126, a highly specific MEK inhibitor, for 2 days. We found that inhibition of ERK activation led to CPT1A upregulation, similar to the effect of FGFR1 knockdown (Fig. 4, D and E). Collectively, these data suggest that FGFR-dependent regulation of CPT1A is mediated by ERK signaling.
To understand how FGFR–ERK signaling controls CPT1A expression, we performed quantitative PCR (qPCR) to examine CPT1A mRNA levels. Our data showed that FGFR inhibition by knocking down FGFR1 significantly increased CPT1A mRNA (Fig. 4F and Fig. S3B). Moreover, FGFR1 depletion–induced CPT1A increase could be rescued by RAF1S259A expression in HDLECs (Fig. 4G). Collectively, these results suggest that FGFR–ERK signaling regulates CPT1A expression at the transcriptional level.
FGFR–ERK signaling regulates CPT1A expression and FAO through the peroxisome proliferator–activated receptor alpha
Because our data suggest that FGFR–ERK signaling controls CPT1A transcription, we sought to identify transcriptional regulators that mediate this process. Peroxisome proliferator–activated receptors (PPARs) are a family of nuclear receptor proteins that regulate transcription of genes involved in a variety of cellular processes, including those involved in lipid metabolism (
). These data led us to hypothesize that PPARα and PPARγ may mediate the effect of FGFR–ERK signaling on CPT1A expression. To test this idea, we first used GW6471, a well-characterized PPARα inhibitor (
). We found that treatment of HDLECs with GW6471 suppressed CPT1A upregulation caused by the FGFR inhibitor ASP5878 (Fig. 5, A and B). Similarly, inhibition of PPARα also reduced FGFR1 knockdown–induced elevation of CPT1A expression in HDLECs (Fig. 5, C and D). To exclude potential nonspecific effects of GW6471, we confirmed these observations by using two different PPARα siRNAs. We found that PPARα knockdown in FGFR1-deficient HDLECs normalized CPT1A expression at both mRNA and protein levels (Fig. 5, E and F and Fig. S3C). Consistent with CPT1A expression, FAO increase caused by ASP5878 treatment or FGFR1 knockdown was also rescued by PPARα inhibition (Fig. 5, G and H). In contrast to PPARα, suppression of PPARγ by a specific inhibitor GW9662 (
) failed to prevent FGFR blockade–induced upregulation of CPT1A (Fig. 5I). These data collectively suggest that PPARα, but not PPARγ, mediates FGFR-dependent regulation of CPT1A.
We further examined whether FGFR signaling controls PPARα. We found that FGFR1 knockdown significantly increased mRNA levels of PPARA, which encodes PPARα (Fig. 5J and Fig. S3D). Moreover, FGFR1 depletion–induced PPARA upregulation could be normalized by RAF1S259A expression in HDLECs (Fig. 5K), indicating that PPARα functions downstream of ERK activation. Taken together, our data suggest that FGFR–ERK signaling regulates CPT1A expression and FAO in LECs through PPARα.
In the current study, we discovered that a balance between glycolysis and FAO exists in LECs and plays an important role in FGFR-mediated energy generation, LEC proliferation, and migration. While FGFR inhibition impairs glycolysis by reducing HK2 expression, it enhances FAO by upregulating CPT1A expression through ERK–PPARα signaling (Fig. 6). Such metabolic flexibility allows FAO to partially compensate for energy deficiency caused by FGFR1 knockdown. Functionally, suppression of FAO can exacerbate impaired LEC proliferation and migration caused by FGFR1 depletion. Our findings are reminiscent of the AMP-activated protein kinase (AMPK), which is an extensively studied energy sensor linking glycolysis with FAO. Energy stress because of defective glycolysis activates AMPK, which in turn enhances CPT1 activity and FAO by reducing the generation of a CPT1 inhibitor, malonyl-CoA (
), does not increase CPT1A levels (Fig. S2), indicating that FGFR-mediated regulation of CPT1A expression is independent of AMPK. Therefore, our study reveals a new mechanism (promoting CPT1A expression) that functions in parallel with AMPK (enhancing CPT1 activity). These two mechanisms might collectively contribute to FGFR inhibition–induced FAO upregulation. Our results are also consistent with a recent study, which shows that compared with blood ECs (BECs) in the proliferative state, quiescent BECs, which were obtained through contact inhibition or activating Notch signaling, reduce glycolytic flux but potentiate FAO by increasing CPT1A (
). Although we obtained similar results in wildtype LECs, our data showed that CPT1A silencing decreases ATP levels and migration in FGFR1-deficient LECs (Fig. 3, A, C, and D), suggesting that FGFR suppression makes LECs rely on FAO for acquiring energy. FAO has been demonstrated to regulate EC proliferation by contributing carbons for nucleotide synthesis (
). As such, we found that knockdown of CPT1A is sufficient to impede LEC proliferation, but depletion of both CPT1A and FGFR1 caused a more dramatic effect (Fig. 3B), which likely reflects the outcome of dual inhibition of ATP generation and nucleotide synthesis.
Our data show that ERK activation by RAF1S259A expression can normalize PPARα and CPT1A expression in LECs with FGFR inhibition (Figs. 4, C and G and 5K and Fig. S3C), suggesting that ERK mediates the effect of FGFRs on PPARα and CPT1A expression. These observations are consistent with previous studies on tumor and liver metabolism. For example, using the BRAF inhibitor PLX to suppress mitogen-activated protein kinase in BRAFV600E-driven melanoma upregulates PPARA and CPT1A mRNA (
). Given our findings in the current study, targeting both FGFR signaling and FAO may offer a more effective means of suppressing the formation of tumor-associated lymphatic vessels and therefore may improve current cancer therapies that use FGFR inhibitors.
Cell culture and transfection
HDLECs (HMVEC-dLyNeo-Der Lym Endo EGM-2MV) were purchased from Lonza and cultured in EBM2 basal medium with EGM-2 MV BulletKit. HDLECs were tested negative for mycoplasma by Lonza. Culture medium was changed every other day. Tissue culture plates were coated with 0.1% gelatin (Sigma) for 30 min at 37 °C and washed with Dulbecco's phosphate-buffered saline (Life Technologies) before cell plating. FGFR1 siRNA (Dharmacon; SMARTpool siRNAs with four target sequences: GCCACACUCUGCACCGCUA, CCACAGAAUUGGAGGCUAC, CAAAUGCCCUUCCAGUGGG, and GAAAUUGCAUGCAGUGCCG), FGFR1 siRNA #2 (Qiagen; target sequence: CAGAGATTTACCCATCGGGTA), CPT1A siRNA (Qiagen; target sequence: CTGGATGGGTATGGTCAAGAT), CPT1A siRNA #2 (Qiagen; target sequence: AACGATGTACGCCAAGATCGA), PPARα siRNA (Qiagen; target sequence: CAAGAGAATCTACGAGGCCTA), PPARα siRNA #2 (Qiagen; target sequence: AAGCTTTGGCTTTACGGAATA), and nontargeting siRNA were commercially purchased and transfected by Lipofectamine RNAiMAX (Life Technologies). To assess the effect of FGFR1, CPT1A, and/or PPARα knockdown on metabolic processes and protein expression, HDLECs, transfected with relevant siRNA 2 or 3 days in advance, were replated and collected for indicated analysis approximately 16 h later when the cell confluency reached ∼80%. To assay the effect of PPARα inhibitor on FAO and protein expression after FGFR1 knockdown, HDLECs were transfected with indicated siRNAs 2 days in advance and then treated with GW6471 (10 μM) or GW9662 (20 μM) for another 2 days. HDLECs were treated with ASP5878 (200–400 nM) for 2 days and then assayed the resulting effect on metabolic processes and protein expression. To assay the effect of GW6471 and ASP5878 on FAO and CPT1A expression, HDLECs were treated with both ASP5878 and GW6471 for 2 days before the analyses.
The FGFR inhibitor ASP5878 was purchased from Selleckchem (#S6539). The PPARα antagonist GW6471 was purchased from Cayman Chemical (#11697). The PPARγ antagonist GW9662 was purchased from Sigma (#M6191). 2-DG was ordered from Sigma (#D8375). The MEK inhibitor U0126 was purchased from Sigma (#U120). Luminescence ATP detection assay system (ATPlite) was purchased from PerkinElmer. For Western blot analysis, the following antibodies were used: HK2 (Cell Signaling Technology; #2867, 1:1000), FGFR1 (Cell Signaling Technology; #9740, 1:1000), phospho-ERK (Cell Signaling Technology; #4370, 1:1000), ERK1/2 (Cell Signaling Technology; #4695, 1:2000), CPT1A (Abcam; #128568, 1:5000), GAPDH (Cell Signaling Technology, #5174; 1:5000), α-tubulin (Cell Signaling Technology, #2144; 1:5000), β-actin (Sigma, #A5316; 1:10,000). ImageJ (NIH) was used for densitometry quantification of Western blot bands.
Lentiviral transfer plasmids encoding CMV promoter-driven RAF1S259A (available at addgene) or mCherry (pLVX-mCherry-N1) were cotransfected with psPAX2 and VSV-G into 293T cells. Virus-containing media were collected 1 or 2 days after transfection and used to infect HDLECs in the presence of polybrene (2–4 μg/ml).
Measurement of glycolysis with [5-3H]-glucose
Glycolytic flux was measured as previously described (
). Briefly, subconfluent HDLECs cultured in 12-well plates were incubated with 1 ml per well EBM2 medium (containing appropriate amounts of serum and supplement) with [5-3H]-glucose (PerkinElmer) for 2 to 3 h. Then 0.8 ml per well medium was transferred into glass vials with hanging wells and filter papers soaked with water. After incubation in a cell culture incubator for at least 2 days to reach saturation, filter papers were taken out and the amount of evaporated 3H2O was measured in a scintillation counter.
Measurement of FAO with [9,10-3H]-palmitic acid
FAO flux was measured using [9,10-3H]-palmitic acid essentially as reported (
). Briefly, subconfluent HDLECs cultured in 12-well plates were incubated with 1 ml per well EBM2 medium (containing appropriate amounts of fatty acid–free bovine serum albumin, carnitine, and cold palmitic acid) with [9,10-3H]-palmitic acid (PerkinElmer) for 6 h. Then, 0.8 ml per well medium was transferred into glass vials with hanging wells and filter papers soaked with water. After incubation in a cell culture incubator for at least 2 days to reach saturation, filter papers were taken out, and the amount of evaporated 3H2O was measured in a scintillation counter.
Measurement of FAO with [1-14C]-oleic acid
FAO flux was measured using [1-14C]-oleic acid as reported (
). Briefly, subconfluent HDLECs cultured in 12-well plates were incubated with 1 ml per well EBM2 medium (containing appropriate amounts of fatty acid–free bovine serum albumin and carnitine) with [1-14C]-oleic acid (PerkinElmer) for 6 h. Then, the cells were lysed using 12% perchloric acid (Sigma–Aldrich; #244252), and the wells were covered immediately using filter papers soaked with hyamine hydroxide (PerkinElmer). After incubation in a fume hood for at least 12 h to reach saturation, filter papers were taken out, and the amount of evaporated 14CO2 was measured in a scintillation counter.
Measurement of glucose oxidation with [6-14C]-glucose
Glucose oxidation flux was measured as previously described (
). Briefly, subconfluent HDLECs cultured in 12-well plates were incubated with 1 ml per well EBM2 medium (containing appropriate amounts of serum and supplement) with [6-14C]-glucose (PerkinElmer) for 6 h. Then, the cells were lysed using 12% perchloric acid, and the wells were covered immediately using filter papers soaked with hyamine hydroxide (PerkinElmer). After incubation in a fume hood for at least 12 h to reach saturation, filter papers were taken out, and the amount of evaporated 14CO2 was measured in a scintillation counter.
Measurement of glutamine oxidation with [14C(U)]-glutamine
Glutamine oxidation flux was measured as previously described (
). Briefly, subconfluent HDLECs cultured in 6-well plates were incubated with 2 ml per well EBM2 medium (containing appropriate amounts of serum and supplement) with [14C(U)]-glutamine (PerkinElmer) for 6 h. Then, the cells were lysed using 12% perchloric acid, and the wells were covered immediately using filter papers soaked with hyamine hydroxide (PerkinElmer). After incubation in a fume hood for at least 12 h to reach saturation, filter papers were taken out, and the amount of evaporated 14CO2 was measured in a scintillation counter.
Cells were lysed using radioimmunoprecipitation buffer, and protein concentrations were measured using the bicinchoninic acid assay. Proteins were first loaded and separated by SDS-PAGE and then transferred to a polyvinylidene difluoride membrane. The membrane was blocked with 5% nonfat milk for 45 min at room temperature and then incubated with relevant primary antibodies overnight at 4°. After incubating with horseradish peroxidase–conjugated secondary antibodies, the membrane was visualized with the SuperSignal West Pico PLUS Chemiluminescent Substrate (Thermo Fisher Scientific) using the ChemiDoc MP imaging system (Bio-Rad).
About 80,000 HDLECs were seeded into each well of a 6-well plate and incubated overnight. Cells were transfected with control, FGFR1, CPT1A, or combined FGFR and CPT1A siRNA and cultured for 4 days. Then, cell numbers were counted using a hemocytometer (Hausser Scientific Horsham).
ATP levels were measured according to the manufacturer’s instruction (PerkinElmer; 6016943). Briefly, HDLECs, which were transfected with control, FGFR1, CPT1A, or combined FGFR and CPT1A siRNA 3 days in advance, were replated to a 96-well plate, and ATP production analysis was performed approximately 12 h later. About 50 μl of mammalian cell lysis solution was added to each well, and the plate was shaken for 5 min at 700 rpm. Then, 50 μl of the substrate solution was added to the cells, and the plate was shaken for another 5 min at 700 rpm. After adapting the plate for 10 min, the luminescence was measured using FLUOstar Omega (BMG LABTECH).
Wound-healing migration assay
HDLEC migration was measured in a wound-healing assay, which used ibidi Culture-Inserts (ibidi) to generate the wound. An ibidi Culture-Insert has dimensions 9 × 9 × 5 mm (width × length × height) and is composed of two wells. One or two inserts were placed into one well of 6-well plates. HDLECs, which were transfected with control, FGFR1, CPT1A, or combined FGFR and CPT1A siRNA 3 days in advance, were replated to the wells of the inserts. When cells became fully confluent after attachment, Culture-Inserts were carefully removed by sterile tweezers to start cell migration. For studying the effect of FGFR1 and CPT1A siRNA on migration, the wound-healing process was monitored for approximately 11 h. A Nikon ECLIPSE Ti microscope with an ANDOR camera was used to image cells at the first time point (T0) and the last time point (Tend point). For data analysis, ImageJ was used to measure the wound area at T0 and Tend point. Migration area was obtained by subtracting area at Tend point from area at T0.
Total RNA was extracted from HDLECs using the TRIzol reagent (Thermo Fisher Scientific) or the RNeasy Plus Mini Kit (Qiagen) according to the manufacturer's instructions. The concentrations and quality of RNA were analyzed by Nanophotometer (IMPLEN). Complementary DNA synthesis was performed using the iScript cDNA synthesis kit (Bio-Rad). qPCR was performed with EvaGreen qPCR Master Mix (Bullseye) using the CFX96 Real-Time system (Bio-Rad).
Data represent mean ± SD. The number of biological and technical replicates and independent experiments is indicated in the legends to the figures. Statistical significance between two groups was determined by two-tailed unpaired t test, and statistical significance between multiple groups was calculated using one-way ANOVA with Sidak's multiple comparisons test. p Values are reported to four decimal places. Difference between experimental groups is considered statistically significant if p < 0.05.
The authors declare that they have no conflicts of interest with the contents of this article.
We thank Ms Summer Simeroth for proofreading the article and Drs Courtney Griffin, Sathish Srinivasan, Holly Van Remmen, and Chi Fung Lee for helpful discussions.
P. Y. conceptualization; H. S., J. Z., P. L., and F. H. methodology; H. S., J. Z., P. L., and F. H. formal analysis; H. S., J. Z., P. L., and F. H. investigation; L. F. resources; H. S. and P. Y. writing–original draft; H. S., J. Z., L. F., and P. Y. writing–review and editing; P. Y. supervision; P. Y. funding acquisition.
Funding and additional information
This project was supported by the American Heart Association 16BGIA27790081 and Internal Restorative Medicine to L. F. This work was also supported by the American Heart Association 19CDA34760260, grants from the Oklahoma Center for Adult Stem Cell Research and the Presbyterian Health Foundation, and Oklahoma Medical Research Foundation startup funding to P. Y.