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inhibitor PF-4708671 partly contributes to its glucose
metabolic effects in muscle and liver cells
Michael Shum, Vanessa Houde, Vicky Bellemare, Rafael Moreira, Kerstin
Bellmann, Philippe St-Pierre, Benoit Viollet, Marc Foretz, André Marette
To cite this version:
Michael Shum, Vanessa Houde, Vicky Bellemare, Rafael Moreira, Kerstin Bellmann, et al.. Inhibi-tion of mitochondrial complex 1 by the S6K1 inhibitor PF-4708671 partly contributes to its glucose metabolic effects in muscle and liver cells. Journal of Biological Chemistry, American Society for Biochemistry and Molecular Biology, 2019, 294 (32), pp.12250-12260. �10.1074/jbc.RA119.008488�. �hal-02348777�
Inhibition of mitochondrial complex 1 by the S6K1 inhibitor PF-4708671 partly contributes to its glucose metabolic effects in muscle and liver cells
Michael Shum1, Vanessa P. Houde1, Vicky Bellemare1, Rafael Junges Moreira1, Kerstin Bellmann1,
Philippe St-Pierre1, Benoit Viollet2,3,4, Marc Foretz2,3,4, and André Marette1*
1
Department of Medicine, Quebec Heart and Lung Institute, Québec, G1V 4G5, Université Laval Canada,
2
INSERM, U1016, Institut Cochin, Paris 75014, France, 3CNRS, UMR8104, Paris 75014, France,
4
Université Paris Descartes, Sorbonne Paris Cité, Paris 75014, France
Running title: PF regulate glucose metabolism independently of insulin
*Corresponding author: André Marette, Department of Medicine, Quebec Heart and Lung Institute, Hôpital Laval, Pavillon Marguerite d'Youville, Room Y4308, 2705 Chemin Ste-Foy, Québec, Canada G1V 4G5
Email: andre.marette@criucpq.ulaval.ca
Keywords: PF-4708671, S6K1, glucose uptake, hepatic glucose production, diabetes
Abstract
mTOR complex 1 (mTORC1) and p70 S6 kinase (S6K1) are both involved in the development of obesity-linked insulin resistance. Recently, we showed that the S6K1 inhibitor PF-4708671 (PF) increases insulin sensitivity. However, we also reported that PF can increase glucose metabolism even in the absence of insulin in muscle and hepatic cells. Here, we further explored the potential mechanisms by which PF increases glucose metabolism in muscle and liver cells independently of insulin. Time-course experiments revealed that PF induces AMP-activated protein kinase (AMPK) activation before inhibiting S6K1. However, PF-induced glucose uptake was not prevented in primary muscle cells from AMPK α1/2 double-KO (dKO) mice. Moreover, PF-mediated suppression of hepatic glucose production was maintained in hepatocytes derived from the AMPK α1/2–dKO mice. Remarkably, PF could still reduce glucose production and activate AMPK in hepatocytes from S6K1/2-dKO mice. Mechanistically, bioenergetics experiments revealed that PF reduces mitochondrial complex I activity in both muscle and hepatic cells. The stimulatory effect of PF on glucose uptake was partially reduced by expression
of the Saccharomyces cerevisiae NADH: ubiquinone oxidoreductase in L6 cells. These results indicate that PF-mediated S6K1 inhibition is not required for its effect on insulin-independent glucose metabolism and AMPK activation. We conclude that although PF rapidly activates AMPK, its ability to acutely increase glucose uptake and suppress glucose production does not require AMPK activation. Unexpectedly, PF rapidly inhibits mitochondrial complex I activity, a mechanism that partially underlies PF’s effect on glucose metabolism.
The mechanistic target of rapamycin (mTOR) complex 1 (mTORC1) and its downstream effector p70 ribosomal S6 kinase (S6K1) are important modulators of energy homeostasis (1,2). This pathway regulates cell proliferation, growth and protein synthesis, which are tightly regulated depending on substrate availability (3). Furthermore, the mTORC1/S6K1 pathway also acts as an energy sensor by sensing amino acids, lipids and energy status and triggers a negative feedback loop towards the PI3K/Akt pathway to terminate its activation after insulin or growth factor stimulation (4-7). However, this negative feedback loop is overactivated by nutrient satiation in metabolic diseases such as obesity contributing
to the development of insulin resistance and type 2 diabetes (7-9). Therefore, targeting downstream effectors of mTORC1 such as S6K1 might be a promising strategy to improve the metabolic syndrome.
We have previously shown that pharmacologic inhibition of S6K1 using PF-4708671 (PF) improved glucose tolerance in mice fed high-fat diet by restoring insulin-induced Akt phosphorylation in obese mice (10). Furthermore, in vitro studies using L6 myocytes and FAO hepatoma cells revealed the ability of PF to increase basal muscle glucose uptake and to decrease basal hepatic glucose production (10). These results were obtained in the absence of insulin stimulation suggesting an additional mechanism of this compound to modulate energy homeostasis independently from enhancing insulin signaling. Previously, several studies have shown a role of S6K to regulate AMPK (adenosine 5’ monophosphate activated protein kinase) which is known to be important for energy homeostasis. Indeed, S6K has been shown to phosphorylate AMPK on Ser 491 in neurons, inhibiting its activity (11). In addition, selective S6K1/2 deletion in muscle is associated with an increase in AMPK activity due to an increase of the AMP/ATP ratio (12) suggesting that S6K may participate in the inhibition of AMPK by either a direct and/or indirect mechanisms. However, a previous study using mouse embryonic fibroblasts derived from S6K1/2 KO mice showed that PF activates AMPK independently of S6K1 inhibition (13). AMPK is an important regulator of energy homeostasis by sensing the energy level via the AMP/ATP ratio thus activating catabolic processes to provide energy to the cells. AMPK activation induces glucose uptake, fatty acid oxidation, autophagy and inhibits cell growth and protein synthesis (14-16). However, it remains unknown whether the insulin-independent effect of PF on glucose metabolism are linked to either S6K1 inhibition or AMPK activation in skeletal muscle and liver, the principal sites of glucose disposal.
In this paper, we thus explored the mechanisms behind the changes in glucose metabolism in muscle and hepatic cells upon S6K1 inhibition independently of insulin stimulation. We evaluated the contribution of S6K1, AMPK and
mitochondrial function to PF-4708671-mediated glucose uptake in muscle cells and decreased basal glucose production in hepatocytes. In both muscle cells and hepatocytes, we observed that PF-4708671 increases both AMPK Thr172 and ACC Ser79 phosphorylation reflecting an increase in AMPK activity. We next used primary muscle and liver cells derived from AMPK α1/2 dKO and S6K1/2 dKO mice to determine the role of both AMPK and S6K in the glucose metabolic effect of PF-470867. These experiments showed that neither AMPK or S6K are required for the acute effect of the compound on glucose metabolism. On the other hand, we found that PF-4708671 reduces mitochondrial complex I activity which is partly responsible for its glucose enhancing effect in muscle cells.
Results
PF-4708671 rapidly increased AMPK phosphorylation before inhibiting S6K1
We have previously shown that S6K1 is a key regulator of glucose disposal in both muscle and liver (7,17) and that the S6K1 inhibitor PF-4708671 (PF) increases both dependent and insulin-independent glucose metabolism in muscle and hepatic cells (10). However, the effect of PF on AMPK activation remains unexplored in metabolically relevant cellular models. As shown in Fig. 1A and 1B, dose-response experiments showed that acute (2h) PF treatment (1 to 20 µM) increases phosphorylation of AMPK on Thr172 in both L6 myocytes and FAO hepatocytes. The drug also enhanced the phosphorylation of the AMPK substrate ACC on Ser79. These effects were associated with a dose-dependent inhibition of S6 phosphorylation on Ser 240/44 suggesting that S6K1 inhibition by PF led to AMPK activation. To confirm that S6K1 inhibition is required for AMPK activation, we next performed time-course experiments (Fig. 1C-D). Unexpectedly, AMPK Thr172 and ACC Ser79 phosphorylation were already robustly increased 5 min after PF treatment while S6 phosphorylation was only found to be reduced at least 30 min after PF exposure in L6 cells (Fig. 1C) and FAO cells (Fig. 1D). These results indicate that AMPK activation by PF precedes S6K1 inhibition and further suggest that rapid
AMPK activation may contribute to PF action on glucose metabolism in muscle and liver cells. PF-4708671 increased glucose uptake in muscle cell and decreased hepatic glucose production independently from AMPK
Since AMPK is a key regulator of muscle glucose uptake (18), we next determined whether the induction of glucose uptake by PF was AMPK-dependent. We used differentiated muscle cells from WT mice or mice lacking both AMPKα1 and AMPKα2 catalytic subunits (AMPK α1/2 dKO muscle cells). After 5 and 48 h of treatment with PF, AMPK phosphorylation on Thr172 and ACC phosphorylation on Ser79 were increased in differentiated WT myocytes but, as expected, not in AMPK α1/2 dKO muscle cells (Fig. 2A). In both AMPK WT and dKO muscle cells, PF treatment decreased basal and insulin-stimulated S6 phosphorylation in both WT and AMPK α1/2 dKO muscle cells. PF treatment of both WT and AMPK α1/2 dKO muscle cells for 5 and 48 h increased glucose uptake, compared to vehicle treated cells (Fig. 2B). However, lack of AMPK activation did not reduce the stimulatory effects of PF treatment on muscle glucose uptake. These data demonstrate that AMPK is dispensable for the stimulatory effect of PF on muscle glucose uptake.
To further investigate the role of AMPK in the PF-induced suppression of hepatic glucose production (HGP), we next isolated hepatocytes from WT mice or mice lacking both AMPKα1 and AMPKα2 catalytic subunits (AMPK α1/2 dKO hepatocytes). As expected, western blot analysis confirmed that AMPK α1/2 dKO hepatocytes did not express AMPK (Fig. 2C). In WT hepatocytes, PF increased both AMPK phosphorylation on Thr172 and ACC phosphorylation on Ser79 without (left panel) or with (right panel) glucagon incubation (Fig 2C). In addition, PF dose-dependently inhibited S6 phosphorylation in both AMPK WT and α1/2 dKO hepatocytes. As a positive control, we used metformin which increased AMPK and ACC phosphorylation with or without glucagon (Fig. 2C). Interestingly, metformin did not affect S6 phosphorylation in WT hepatocytes but decreased S6 phosphorylation in AMPK α1/2 double KO hepatocytes (Fig. 2C).
We next determined HGP in primary hepatocytes in basal and glucagon-stimulated conditions (Fig.2 D). As reported previously (19), AMPK is not required for basal or glucagon-induced HGP. Inhibition of S6K1 with PF (20 µM) was found to reduce basal and glucagon-stimulated HGP in AMPK WT and this response was amplified in α1/2 dKO hepatocytes, especially in the glucagon-stimulated state. Similar results were obtained using metformin, which was found to blunt HGP even more in α1/2 dKO than in WT hepatocytes, in agreement with a previous report (19). These results indicate that AMPK is also dispensable for the inhibitory effect of PF on hepatic glucose production.
PF-4708671 decreased basal hepatic production independently from S6K1
To determine whether PF-mediated inhibition of HGP was S6K1-dependent, we next isolated hepatocytes from S6K1/2 double KO mice. As anticipated, S6K1 expression was absent in S6K1/2 dKO hepatocytes and PF decreased S6 phosphorylation in WT hepatic cells (Fig. 3A). In addition, PF increased AMPK and ACC phosphorylation in both S6K1/2 WT and double KO hepatocytes suggesting that S6K1 is dispensable for PF-induced AMPK activation (Fig. 3A). Similarly, metformin strongly increased AMPK and ACC phosphorylation in both WT and S6K1/2 dKO hepatocytes but did not affect S6 phosphorylation in WT hepatocytes (Fig. 3A). In both WT and S6K1/2 dKO hepatocytes, PF (20 µM) decreased glucagon-stimulated HGP (Fig. 3B). Metformin also decreased HGP with no differences between S6K1/2 WT and double KO hepatocytes. Thus, neither AMPK nor S6K is required for the ability of PF to blunt glucose production in hepatocytes.
PF-4708671 decreased mitochondrial respiration and mitochondrial spare respiratory capacity in muscle cells and hepatocytes. Since metformin has been previously shown to control glucose production through regulation of hepatic energy state and inhibition of mitochondrial respiration (19-21) we next investigated whether PF could exert its glucose metabolic effects through a similar mechanism. Using a Seahorse XF 24 system to determine mitochondrial respiratory parameters (Fig. 4A), we next examined the effect of PF on
oxygen consumption in L6 myotubes and FAO hepatocytes. Cells were first treated with PF (1-20 µM) for 400 min to determine the maximal effect of the compound on basal respiration and whether this effect could be sustained over time. In L6 cells, there was a trend for PF to reduce basal mitochondrial respiration (Fig. 4 B-C) which was more apparent in FAO cells, as well as respiration associated with ATP production at 20 µM (Fig. 4 D-E). In addition, PF lowered the maximal respiration and the spare respiratory capacity in both L6 and FAO cells. These results indicate that PF reduced mitochondrial respiratory capacity in both cell types.
PF-4708671 reduced mitochondrial complex 1 activity
We then assessed whether this decrease in maximal respiratory capacity was due to a decrease in mitochondrial complexes I and/or II activity. Cells were permeabilized to allow ADP, pyruvate (Pyr), malate (Mal) and succinate (Succ) to pass through the plasma membrane and to reach mitochondria. L6 cells were acutely treated with different concentrations of PF (i.e. drug added just before the assay) and the plate was added in the Seahorse XF 24 system. Addition of Pyr/Mal in the presence of ADP increased mitochondrial complex I activity which was dose-dependently inhibited by PF (Fig. 5A). Mitochondrial complex II activity was measured by adding the complex I inhibitor rotenone first, followed by succinate. No significant effect of PF treatment was observed on mitochondrial complex II activity (Fig. 5B). Similar results were obtained in FAO cells. Indeed, PF reduced mitochondrial complex I activity in hepatic cells but failed to affect mitochondrial complex II activity (Fig. 5C-D). Taken together, these results show that PF acts mainly on mitochondrial complex 1 to regulate mitochondrial function in both muscle and hepatic cells.
PF-inducing glucose uptake is partially dependent of mitochondrial complex 1 activity. To determine the role of mitochondrial complex I in mediating glucose uptake in L6 myocytes, we stably expressed the Saccharomyces cerevisiae NADH:ubiquinone oxidoreductase (NDI1) in these cells, which can maintain respiration even in the presence of complex-I inhibitors (22-25) . Expression of NDI1 in muscle cells (Fig. 6A) was
found to significantly reduce PF-mediated glucose uptake without interfering with insulin-induced glucose disposal in these cells (Fig. 6B). When taking into account the changes in basal glucose uptake in NDI1-expressing cells in order to calculate the specific effect of PF on this parameter, we conclude that about 45% of the drug effect may be accounted by inhibition of mitochondrial complex 1 activity in muscle cells (Fig. 6C). These results demonstrate that PF increases insulin-independent glucose uptake at least in part by inhibition of mitochondrial complex 1 activity in muscle cells but that other unknown mechanism(s) are also involved.
Discussion
We recently reported that pharmacologic inhibition of S6K1 by PF for only one week improves glucose tolerance in high-fat fed obese mice and increases insulin-induced Akt phosphorylation in liver, muscle and white adipose tissue (10). Interestingly, these studies also revealed that PF enhances basal glucose uptake in myocytes and decreases basal glucose production in hepatocytes suggesting that the S6K1 inhibitor also modulates glucose metabolism through insulin-independent pathways. This is consistent with other studies in C2C12 cells where transfection of a siRNA targeting S6K1 also enhanced basal glucose uptake (26). In the present study we found that PF rapidly modulate glucose metabolism in both L6 myotubes and FAO hepatocytes in association with increased phosphorylation of AMPK and its downstream effector ACC.
However, following acute PF treatment, we observed an increase in both AMPK Thr172 and ACC Ser79 phosphorylation, a well-known substrate of active AMPK. The time-course experiment performed in PF treated cells revealed that AMPK is activated rapidly (Thr172 AMPK and Ser79 ACC phosphorylation), faster than PF-mediating S6K1 inhibition. Despite this rapid increase in AMPK activation, we showed that PF-regulating glucose homeostasis is independent from AMPK. Thus, ruling out a potential role of AMPK on PF-stimulating glucose uptake and decreasing hepatic glucose production. These results are also supported by other studies showing that AMPK deficiency failed to impact hepatic glucose
production and the suppressive effect of metformin on this process (19,27).
PF has been previously suggested to activate AMPK by inhibiting mitochondrial complex I of the respiratory chain independently of S6K1 in mouse embryonic fibroblasts (13). Using muscle cells and hepatocytes, we also observed that PF acutely decreased mitochondrial complex I activity suggesting that the drug modulates glucose metabolism in these cells through inhibition of mitochondrial function. Using S6K1/2 dKO hepatic cells, we further showed that PF inhibits basal or glucagon-stimulated hepatic glucose production independently from S6K. These results suggest that PF action on hepatic glucose production is dispensable from S6K inhibition and thus might be entirely due to a higher energy demand and metabolism rewiring towards replenishing acetyl-CoA pools caused by lower mitochondrial complex I activity.
Glucose uptake can be attributed to glucose transport, glucose oxidation and glycogen synthesis. Our respirometry analysis revealed that PF decreased OXPHOS suggesting that reduction in mitochondrial ATP production might have induced glucose transport for ATP production from glycolysis. Reduction of mitochondrial complex 1 activity explains, in part, the insulin-independent action of PF on glucose metabolism. In L6 muscle cells, the ability of PF to enhance glucose uptake was partially reversed by stable NDI1 expression, indicating that the decrease of mitochondrial complex 1 activity contributes to the stimulatory effect of the compound on muscle glucose uptake. This conclusion is also supported by previous studies showing that inhibition of mitochondrial complex 1 by a very low dose of rotenone or siRNA-mediated silencing of NADH:Ubiquinone Oxidoreductase Subunit A13, a subunit of complex I, can reduce hepatic glucose production and stimulate glucose consumption in muscle cells, independently of AMPK (28). In addition, other studies have reported similar mechanisms. Indeed, berberine has been shown to enhance glucose utilization and glycolysis by reducing mitochondrial glucose oxidation in L6 myotubes. Altogether, our results suggest that the energy demand generated by the inhibition of mitochondrial complex 1 activity is enough to drive
glucose uptake in muscle and to decrease hepatic glucose production in PF-treated cells, independently from AMPK activation. However, the finding that stable expression of NDI1 only partially blunted PF-inducing glucose uptake in muscle cells suggests that other mechanism(s) such as S6K1 inhibition may also contribute to the insulin-independent metabolic effect of the drug. This dual effect of PF-4708671 might be an interesting advantage to treat type 2 diabetes more efficiently. We found that part of the action of PF-4708671 is by inhibiting mitochondrial complex 1, which is also a well-known target for the most prescribed type 2 diabetes drug, metformin (20,29). These results suggest that PF might reproduce many beneficial effects mediated by metformin in addition to those associated with S6K1 inhibition such as direct inhibition of the negative feedback loop regulating insulin signaling. Metformin’s ability to blunt hepatic glucose production is also independent of AMPK and rather mediated by inhibition of mitochondrial complex 1 (20) and mitochondrial glycerol-phosphate dehydrogenase (19,30).
Conclusions
PF-4708671 was characterized as a selective S6K1 inhibitor, but our study and others demonstrate it also has S6K1-independent effects, notably by reducing mitochondrial complex I activity. This inhibition led to a rapid, but transient induction, of AMPK activation and ACC inhibition, followed by S6K1 inhibition in muscle cells and hepatocytes. While S6K1 inhibition by this drug restored insulin signaling in cells and tissues from obese insulin-resistant mice (10), the present data reveal that PF also regulates glucose metabolism independently from AMPK by reducing mitochondrial complex 1 activity. These results suggest that PF might represent an interesting therapeutic tool since it can both reverse S6K1-mediated insulin resistance (10) as well as improving glucose control through inhibition of mitochondrial respiration, thus combining two key mechanisms that are well recognized for the treatment of type 2 diabetes. Experimental procedures
Reagents and antibodies. DMSO was from Sigma-Aldrich (St. Louis, MO, USA). PF-4708671
was from Cayman (Ann Harbor, Michigan, USA). Insulin was from Eli Lilly (Toronto, ON, Canada). The phospho-S6 Ser 240/44, S6, phospho-AMPK Thr172, phospho-AMPK Ser491, AMPK, phospho-ACC Ser79, ACC, phosphor-S6K1 Thr389, S6K1 antibodies were from Cell Signaling Technologies (Danvers, MA, USA). The anti-actin antibody was from Millipore (St. Louis, MO, USA). Antibodies from Cell Signaling Technology were used at 1:1,000 dilution. Actin antibody was used at 1:10,000 dilution.
Animals.
All animal studies were approved by the Paris Descartes University ethics committee (no. CEEA34.BV.157.12) and the Direction Départementale des Services Vétérinaires of the Préfecture de Police de Paris authorization no. 75-886). All mice were maintained in a barrier facility under a 12-h light/12-h dark cycle (8 am-8 pm) with free access to water and standard diet. Liver double-knockout of AMPKα1 and AMPKα2 catalytic subunits was achieved by crossing AMPKα1lox/lox mice with AMPKα2lox/lox
mice and then Alfp-Cre transgenic mice (31,32) on C57BL/6 background, to generate AMPKα1 lox/lox, α2lox/lox (control) and
AMPKα1 lox/lox, α2lox/lox-Alfp-Cre (liver AMPK
α1/2 dKO mice). S6K double knockout or S6K wild type mice were a kind gift of Mario Pende (Inserm, Paris, France). The S6K1/2 dKO mice were generated as previously described (33).
Cell culture.
Rat L6 myoblasts were grown in α-MEM (Invitrogen, Burlington, ON, Canada) supplemented with 10% FBS (vol./vol.). L6 myoblasts were differentiated into myotubes in α-MEM with 2% (vol./vol.) FBS as previously described (34). FAO rat hepatocytes were maintained in RPMI media (Invitrogen) and supplemented with 10% (vol./vol.) FBS. Wild-type (WT) and AMPK α1/2 double knock-out (KO) primary muscle cells were grown as reported previously (35).
NDI1 expression in L6 cells. The pMXS-NDI1 (NADH:ubiquinone oxidoreductase) plasmid was a gift from David Sabatini (Addgene plasmid #72876) and the pMXS-control vector was kindly provided by Kivanc Birsoy (Rockefeller
University, USA). Plasmids were amplified as previously described in Phoenix-Eco cells (ATCC #CRL-3214) (28). Undifferentiated rat L6 myoblasts were infected with the amplified retrovirus and selected with 5 µg/ml of blasticidin to generate L6-pMXS and L6-NDI1 stably transfected myoblasts. Both L6 cell lines were then differentiated into myotubes as described in the previous section creating cell lines that stably overexpress NDI1 or the control vector. NDI1 overexpression was verified by purifying total RNA using the GeneJet RNA purification kit (Thermo Scientific, Burlington, ON, Canada) followed by the RNA reverse transcription with a High Capacity cDNA Reverse Transcription Kit from Applied Biosystems (Burlington, ON, Canada). Real-time PCR was performed using SYBR Green Jump-Start Expression Kit (Sigma-Aldrich, Oakville, ON, Canada). Three sets of primers for S. cerevisiae
NDI1 were used (5’→3’) (IDT, Skokie, IL, USA). NDI1 set 1: F-TCAAAGTACATCTGAGA ACGG, R-GTCTTTGCCAACAATTGCT. NDI1 set 2: F-GCTTAATGACCTTCTAC TTATGGAG, R-GATCTTGCAGATAGAA TCATGGAC. NDI1 set 3: F-CTTTGTTA CCTTCTGCACCA,
R-ACGATGGGCTCAA TAATTGAC. Gene
expression was assessed by the ΔΔCt method and the rat Arbp was used as the reference gene
(F-TAAAGACTGGA GACAAGGTG,
R-GTGTAGTCAGTCTCCA CAGA).
Mouse primary hepatocytes. Mouse primary hepatocytes were isolated from 10-week-old male mice by a modified version of the collagenase method as described previously (19). Wild-type (WT), AMPK α1/2 double KO and S6K1/2 double KO primary hepatocytes were plated in M199 medium supplemented with 100 U/ml penicillin, 100 μg/ml streptomycin, 0.1% bovine serum albumin, 500 nM dexamethasone (dex; Sigma-Aldrich), 100 nM triiodothyronine (Sigma-Aldrich), 10 nM insulin (Actrapid, Novo Nordisk), at a density of 2.5 × 105 cells/well on 6-well plates. After attachment (3–4 hours), hepatocytes were maintained in M199 medium with antibiotics and 100 nM dexamethasone for 16 hours, followed by drug treatment as described below. Vehicle (dimethyl sulfoxide, DMSO 0.02% [vol./vol.]) or PF-4708671 was used at the indicated doses.
Western blotting. Western blots were performed as described (36). Briefly, equal amounts of proteins were separated by SDS-PAGE (9 % [wt/vol.]) and transferred onto nitrocellulose membranes. Membranes were blocked in 5% (wt/vol.) milk diluted in 50 mM Tris pH 7.4, 0.1% (vol./vol.) Tween (TBS-T) and incubated overnight at 4°C with the respective antibodies diluted in 3% (wt/vol.) BSA in TBS-T.
Glucose uptake. L6 cells were serum-deprived for 5 h prior to the experiments, and 100 nmol/l of insulin was used to stimulate the cells during the last hour of deprivation. L6 cells were incubated for 8 min in HEPES-buffered saline containing 10 µmol/l unlabeled 2-DG and 10 µmol/l D-2-deoxy-[3H] glucose (0.5 µCi/ml). The reaction was terminated by washing three times with ice-cold 0.9% NaCl (wt/vol). Cell-associated radioactivity was determined by lysing the cells with 0.05 N NaOH, followed by liquid scintillation counting and normalization to protein concentration. Glucose production. FAO cells were incubated 16 h in serum-free medium, with or without the indicated concentration of insulin, PF-4708671 (10 µmol/l). The cells were washed three times with PBS and incubated with phenol- and glucose-free DMEM medium supplemented with 20 mmol/l sodium L-lactate and 2 mmol/l sodium pyruvate for 5 h with or without the indicated concentration of insulin and PF-4708671 (10 µmol/l). Cell supernatants were collected, and glucose concentration was measured with the Amplex-Red Glucose assay kit (Invitrogen, Burlington, ON, Canada) according to the manufacturer’s instructions. Cells were lysed with 50 mmol/l NaOH and protein concentration was determined
using a BCA protein assay kit for normalization. Primary mouse hepatocytes on 6-well plates (2.5 × 105 cells/well) were maintained in M199 medium containing antibiotics and 100 nM dex for 16 hours prior to the measurement of glucose production. Hepatocytes were washed once with PBS, and glucose production was determined after an 8-hour incubation period in glucose-free DMEM containing lactate/pyruvate (10:1 mM) and 100 nM dex alone or with glucagon 25 nM (Sigma-Aldrich) and with various doses PF-4708671. Glucose concentration values were corrected by protein content.
XF metabolic assay
Oxygen consumption rate (OCR) measurements were made using the Seahorse XF24 Extracellular Flux Analyzer. For mitochondrial metabolic parameters, we used successively different concentration (1-20 µM) of acute PF-4708671,
oligomycin (1 µM), FCCP (2 µM), rotenone/ antimycin (1 µM). For determination of complex I and complex II activity, cells were pre-incubated in mannitol/sucrose (MAS) buffer free of any substrate but containing 4 mM ADP for 30 min followed by the addition of XF PMP permeabilizer agent (1 nM) just before adding the plate in the XF24. Pyruvate/malate followed by rotenone (1 µM), succinate (10 mM) and antimycin (1 µM) were injected to the cells.
Quantification and statistical analyses. One- and two-way ANOVA with Newman–Keuls post hoc was performed with Sigma Plot version 12 (Systat Software, San Jose, CA, USA); p values were considered significant if they were <0.05. SDs are represented in the graphs.
Financial support
This work was funded by a Canadian Institutes of Health Research (CIHR) foundation grant (FDN#143247), a grant from the Cardiometabolic Health, Diabetes and Obesity Research Network (CMDO) to AM and grants from the Région Ile-de-France (CORDDIM), the Société Francophone du Diabète (SFD) and Fondation pour le Recherche Médicale (FRM - DEQ20150934718) to MF and BV. Acknowledgements
We thank Prof. Mario Pende at the Institut Necker-Enfants Malades, Paris, France for kindly providing the S6K1/2 WT and KO mice used in the present study. We also thank Michael Schwab for his helps designing primers used to measure NDI1 expression.
Conflict of interests:
The authors declare that they have no conflicts of interest with the contents of this article
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of PF-4708671 for 2 hours in presence of FBS 10%. Cells were then harvested, processed for SDS-PAGE and western blot using the indicated antibodies. (C) L6 differentiated myocytes and (D) FAO hepatoma cells were treated for the indicated time with 10 µM PF in presence of FBS 10%. Actin levels are presented as loading control. PF, PF-4708671. *p<0.05, **p<0.01, ***p<0.001 vs vehicle treated cells, as indicated. Shown are representative blots and mean +/- SD from 3-5 independent experiments.
(A) WT and AMPK α1/2 KO primary muscle cells in absence or presence of insulin (100 nM, 45 min) and/or PF-4708671 (10 µM, 5 or 48 h) were lysed and immunoblotting using the indicated antibodies. (B) Glucose uptake was assessed in similar conditions. a; p<0.05, vs vehicle treated WT cell, b; p<0.05, vs vehicle treated AMPK α1/2 KO cells, c; p<0.05, vs vehicle treated WT cells with insulin, d; p<0.05, vs vehicle treated AMPK α1/2 KO cells with insulin. White bar: AMPK α1/2 WT Grey bar: AMPK α1/2 KO (C) Hepatocytes were isolated from WT and AMPK α1/2 KO mice and after two days, the cells were treated with PF-4708671 (10 µM), metformin (0.5 mM) and/or glucagon (25 nM) for 8h in the hepatic glucose production media followed by cell lysis for immunoblotting using the indicated antibodies and (D) hepatic glucose production was assessed. PF, PF-4708671. Met, Metformin. White bar: AMPK α1/2 WT Grey bar: AMPK α1/2 KO . Glucose value were corrected for protein content. Shown are the mean +/- SD of 3-4 independent experiments and representative blot from 3 independent experiments. a; p<0.05, vs vehicle treated WT cells without glucagon, b; p<0.05, vs vehicle treated AMPK α1/2 KO cells without glucagon, c; p<0.05, vs vehicle treated WT cells with glucagon, d; p<0.05, vs vehicle treated AMPK α1/2 KO cells with glucagon.
(A) Hepatic glucose production was assessed in primary hepatocytes culture isolated from WT and S6K1 1/2 KO mice. After two days, the cells were treated with PF-4708671 (10 µM), metformin (0.5 mM) and/or glucagon (25 nM) for 8h in the HGP media followed by cell lysis and immunoblotting using the indicated antibodies. PF, PF-4708671. Met, Metformin. (B) Glucose produced by the hepatocytes was measured and values were corrected for protein content. White bar S6K1/2 WT, Grey bar S6K1/2 KO. Shown are the mean +/- SD of at least 4 independent experiments and representative blot from 3 independent experiments. a; p<0.05, vs vehicle treated WT cells without glucagon, b; p<0.05, vs vehicle treated S6K 1/2 KO cells without glucagon, c; p<0.05, vs vehicle treated WT cells with glucagon, d; p<0.05, vs vehicle treated S6K 1/2 KO cells with glucagon.
L6 cells were seeded at 20,000 cells per well in 24-well XF plates and incubated for 24 hours with 10% bovine growth serum (BGS) in MEM at 37ºC/5% CO2 incubator. For differentiation, growth media was replaced with differentiation media (2% BGS in MEM). Well differentiated myocytes were used at day 5 for all assays. (A) Mitochondrial parameters determined from the oxygen consumption trace following PF, oligomycin, FCCP and Rotenone/Antimycin A injection. (B-C) L6 cells were acutely treated with 1-20 μM PF-4708671 by using a port from the seahorse cartridge followed by a Mitostress protocol. (D-E) FAO cells were seeded at 20,000 cells per well in 24-well XF plates and incubated for 3 days with 10% bovine growth serum (BGS) in MEM at 37ºC/5% CO2 incubator. Mitochondrial parameters were assessed with the Mitostress protocol and PF acute is the decrease in basal respiration induced by PF after 400min. *p<0.05, **p<0.01, ***p<0.001 vs vehicle treated cells, as indicated. PF, PF-4708671. Shown are the mean +/- SD of 4-5 independent experiments.
treated with the indicated PF-4708671 concentrations before cell permeabilization with XF plasma membrane permeabilizer (1 nM) from Seahorse Bioscience. Then the plate was inserted into the XF 24-well apparatus. Mitochondrial complex I and II activity were assessed by adding successively Pyruvate/Malate (Pyr/Mal; 10mM/1mM), Rotenone (Rot; 1 uM), Succinate (10 mM) and Antimycin A (AA; 1 uM) in differentiated (A) L6 cells and (C) FAO hepatocytes. (B-D) Mitochondrial complex I activity was calculated from OCR differences between Pyruvate/Malate and Rotenone. Mitochondrial complex II activity was calculated from OCR differences between Succinate and Antimycin A for. *p<0.05, **p<0.01 vs vehicle treated cells, as indicated. Shown are the mean +/- SD of 4-5 independent experiments.
Stable L6-pMXS and L6-NDI1 (NADH:ubiquinone oxidoreductase) transfected myoblasts were differentiated as described in method. NDI1 overexpression was verified by purifying total RNA followed by the RNA reverse transcription. (A) Real-time PCR was then performed and NDI1 expression was corrected to ARBP gene expression. N.D.: Not-detected. (B) Glucose uptake was measured in L6 cell transfected with either control vector pMXS or NDI1 and then differentiated. Cells were treated with 10 μM PF for 5h and/or with insulin (100 nM, 45 min). White bar:
pMXS, Grey bar: NDI1 (C) Changes in glucose uptake were corrected for basal
glucose uptake level in respective conditions and reported in %. White bar: pMXS, Grey bar: NDI1. *** p<0.001 vs respective PF-treated pMXS cells, a, b p<0.001 vs respective basal, PF, PF-4708671. Shown are the mean +/- SD of 4 independent experiments.
