Hierarchical activation of compartmentalized pools of AMPK depends on severity of nutrient or energy stress

Zong, Yue; Zhang, Chen-Song; Li, Mengqi; Wang, Wen; Wang, Zhichao; Hawley, Simon A.; Ma, Teng; Feng, **-Wei; Tian, **ao; Qi, Qu; Wu, Yu-Qing; Zhang, Cixiong; Ye, Zhiyun; Lin, Shu-Yong; Piao, Hai-Long; Hardie, D. Grahame; Lin, Sheng-Cai

doi:10.1038/s41422-019-0163-6

Hierarchical activation of compartmentalized pools of AMPK depends on severity of nutrient or energy stress

Article
Published: 04 April 2019

Volume 29, pages 460–473, (2019)
Cite this article

Download PDF

Cell Research Submit manuscript

Hierarchical activation of compartmentalized pools of AMPK depends on severity of nutrient or energy stress

Download PDF

Yue Zong¹^na1,
Chen-Song Zhang¹^na1,
Mengqi Li¹^na1,
Wen Wang^2,3^na1,
Zhichao Wang^2,3,
Simon A. Hawley⁴,
Teng Ma¹,
**-Wei Feng¹,
**ao Tian¹,
Qu Qi¹,
Yu-Qing Wu¹,
Cixiong Zhang¹,
Zhiyun Ye¹,
Shu-Yong Lin ORCID: orcid.org/0000-0002-4419-1713¹,
Hai-Long Piao^2,3,
D. Grahame Hardie⁴ &
…
Sheng-Cai Lin¹

12k Accesses
99 Citations
60 Altmetric
4 Mentions
Explore all metrics

Abstract

AMPK, a master regulator of metabolic homeostasis, is activated by both AMP-dependent and AMP-independent mechanisms. The conditions under which these different mechanisms operate, and their biological implications are unclear. Here, we show that, depending on the degree of elevation of cellular AMP, distinct compartmentalized pools of AMPK are activated, phosphorylating different sets of targets. Low glucose activates AMPK exclusively through the AMP-independent, AXIN-based pathway in lysosomes to phosphorylate targets such as ACC1 and SREBP1c, exerting early anti-anabolic and pro-catabolic roles. Moderate increases in AMP expand this to activate cytosolic AMPK also in an AXIN-dependent manner. In contrast, high concentrations of AMP, arising from severe nutrient stress, activate all pools of AMPK independently of AXIN. Surprisingly, mitochondrion-localized AMPK is activated to phosphorylate ACC2 and mitochondrial fission factor (MFF) only during severe nutrient stress. Our findings reveal a spatiotemporal basis for hierarchical activation of different pools of AMPK during differing degrees of stress severity.

AMPK: guardian of metabolism and mitochondrial homeostasis

Article 04 October 2017

A metabolite sensor subunit of the Atg1/ULK complex regulates selective autophagy

Article Open access 05 February 2024

AMPK: a key regulator of energy stress and calcium-induced autophagy

Article 16 August 2021

Introduction

The AMP-activated protein kinase (AMPK) is a pivotal sensor for monitoring cellular nutrient supply and energy status, and plays crucial roles in adaptive responses to nutrient availability and falling energy levels.^1,2,3,4,5 AMPK comprises a heterotrimeric complex of a catalytic α subunit and regulatory β and γ subunits. The γ subunit provides binding sites for the regulatory nucleotides AMP, ADP and ATP, whose occupancy depends upon the cellular AMP:ATP and ADP:ATP ratios.^{6,1c, followed by immunoblotting. h, i AMP/ATP and ADP/ATP ratios, acetyl-coA and malonyl-coA levels in livers from mice under starvation or hepatic ischemia. Mice were starved for 16 h or subjected to hepatic ischemia (for 10 min), followed by measurement of AMP/ATP and ADP/ATP ratios by CE-MS (h) or acetyl-coA and malonyl-coA levels by HPLC-MS (i). Results are mean ± SD; ***p < 0.001 by Student’s t-test (h), **p < 0.01, N.S., not significant by ANOVA (i), n = 6. j A schematic diagram showing the three fusion constructs of the β1 subunit (with modifications at the N-terminus) that allow AMPK to locate on lysosomal surface, mitochondrial outer membrane, or in cytosol. k ACC2 can only be phosphorylated by cytosol-localized and the mitochondrion-localized AMPK. AMPKβ-DKO HEK293T cells were infected with HA-tagged lyso-β1 (left panel), cyto-β1 (middle panel), and mito-β1 (right panel), respectively. Cells were then treated with 1 μM A-769662 for 2 h to allow full activation of AMPK, followed by fractionation and immunoblotting for analyzing p-AMPKα, or by immunoprecipitation and immunoblotting for analyzing p-ACC1 and p-ACC2. Experiments in a, c, d, e, g, and k were performed three times, and the others twice. See also Supplementary information, Figs. S3, S4}

Full size image

ACC1 and ACC2 contain very similar AMPK substrate recognition motifs⁵⁴ (consequently phospho-specific antibodies recognize both) but vary in their subcellular locations. We hypothesized that the targets of AMPK may be regulated in a spatiotemporal manner under different stress conditions. To test this, we replaced the endogenous AMPK-β subunits with modified forms that target the complex to specific locations. It is known that N-myristoylation is required for AMPK association with intracellular membranes such as the lysosome¹⁴ and the mitochondrion.²⁴ We generated two fusion constructs with modifications at the N-terminus of the β1 subunit: either adding LAMP2 (for tethering to the lysosomal surface, referred to as lyso-β1) or TOMM20 (for tethering to mitochondrial outer membrane, referred to as mito-β1) and also the β1-G2A mutation (preventing N-myristoylation, referred to as cyto-β1) (Fig. 3j). Before reintroduction of the engineered constructs to HEK293T cells, we knocked out the β subunits (β1 and β2) of AMPK to generate AMPKβ-DKO cells (validation in Supplementary information, Fig. S4a). Lyso-β1, mito-β1 and cyto-β1 were then individually re-introduced to the AMPKβ-DKO cells. The engineered β1 subunits assembled into heterotrimeric AMPK complexes as occurs with the wild-type AMPK-β1, and became appropriately localized as validated by immunostaining (Supplementary information, Fig. S4b, c). We next treated the cells with A-769662, a β1-specific activator,⁵⁵ to cause compartmentalized activation of AMPK (Supplementary information, Fig. S4d). We found that in lyso-β1-expressing cells, only cytosol-localized ACC1 can be phosphorylated in response to A-769662 treatment (Fig. 3k). By contrast, in mito-β1- and cyto-β1-expressing cells, both the cytosol-localized ACC1 and the mitochondrion-localized ACC2 were phosphorylated (Fig. 3k). Therefore, mitochondrion-localized ACC2 seems to be specifically phosphorylated under severe nutrient stress, in which mitochondrial and cytosolic AMPK are also fully activated. In support of this, we found that the mitochondrial fission factor MFF, another mitochondrion-localized AMPK substrate,⁴⁷ was phosphorylated in a similar manner to ACC2 (Fig. 1g).

Roles of AMP in the hierarchical activation of AMPK

We postulated that upon starvation for glucose only, basal AMP might act as a necessary cofactor for AMPK activation via the lysosomal pathway, whereas elevated AMP may bind to additional sites on AMPK. Indeed, when the AMPK-γ1 mutant D317A, which affects the “non-exchangeable” site for AMP (site 4), was re-introduced into HEK293T cells with knockouts of all γ subunits (γ1 to γ3) (AMPKγ-TKO cells, Supplementary information, Fig. S4e), the activation of AMPK upon glucose starvation was severely dampened (Fig. 4a). This is consistent with our previous finding that the AMP-promoted interaction between LKB1 and AMPK is impaired by this mutant.¹⁹ We also introduced the R531G mutant, in which the exchangeable site for AMP (site 3)⁸ is disrupted, into AMPKγ-TKO cells, and found that the phosphorylation of lysosomal AMPK upon glucose starvation remained unaffected. This contrasted to cytosolic and mitochondrial AMPK phosphorylation which was blocked under moderate and high AMP levels (Fig. 4b). Similarly, phosphorylation of ACC2 was blocked in R531G-expressing cells (Fig. 4c). It is not clear why the R531G-containing AMPK complex can respond to glucose starvation, yet fails to be activated by severe starvation (Fig. 4b, c). It is possible that under severe starvation such a mutation perturbs the overall structure of the AMPK complex, thereby blocking its activation by any mode. We also determined whether AMP levels underlie the activation of cytosolic and mitochondrial AMPK by manipulating the generation of AMP by knocking out adenylate kinase 1 (AK1, which catalyzes the conversion of 2 ADP to 1 ATP and 1 AMP) in MEFs (Supplementary information, Fig. S4f). Indeed, we found that knockout of AK1 significantly dampened the activation of mitochondrial- and cytosolic-localized AMPK, and the phosphorylation of ACC2 in conditions of severe nutrient starvation (Fig. 4d; Supplementary information, Fig. S4g).

Discussion

In this study, we have systemically determined how differentially localized AMPK complexes are regulated to affect distinct target phosphorylation in response to different stresses (Fig. 4e). Under conditions of glucose starvation that did not elevate cellular AMP:ATP or ADP:ATP ratios, only lysosomally localized AMPK was activated and exclusively through the lysosomal pathway. However, when AMP was moderately elevated, cytosolic AMPK was also activated in addition to lysosomal AMPK. Activation induced by moderate increases in AMP was not mediated via the lysosomal pathway, because knockout of LAMTOR1 (a subunit of the Ragulator complex) had no effect although activation was still dependent on AXIN1. In support of this, moderately elevated AMP promoted the formation of a complex between AXIN1, LKB1 and AMPK. Interestingly, mitochondrial AMPK was not activated by moderate increases in AMP, but only in response to more severe nutrient stress when there were larger increases in AMP. One possible explanation for this is that the concentrations of ATP are higher in the vicinity of mitochondria, inhibiting the activation of AMPK by AMP. However, when cellular AMP is elevated to higher levels during severe nutrient stress or ischemia, mitochondrial AMPK becomes activated independently of AXIN1, phosphorylating the mitochondrially localized substrates, ACC2 and MFF.

Basal AMP is believed to be maintained at low levels by the freely reversible adenylate kinase reaction (2ADP ↔ ATP + AMP), with the high ATP:ADP ratio in unstressed cells driving the reaction towards ADP and away from AMP. AMP is thought to be constantly bound as a cofactor to site 4 of the AMPK-γ subunit,^Plasmids

Point mutations on AMPK-γ1 (D317A), AMPK-γ2 (R531G), and AMPK-β1 (G2A) were made using a PCR-based site-directed mutagenesis method using PrimeSTAR HS polymerase (Takara). Expression plasmids for various proteins were constructed in the pcDNA3.3 vector for transient transfection (ectopic expression), in pBOBI for lentivirus packaging (stable expression). PCR products were verified by sequencing (Invitrogen, China). The lentivirus-based vector pLL3.7 was used for expression of siRNA in HEK293T and HEK293 cells.

CRISPR/Cas9 knockout cell lines

Genes were deleted in HEK293T cells (human PRKAB1, PRKAB2, PRKAG1, PRKAG2, and PRKAG3) or MEFs (mouse Ak1) using the CRISPR-Cas9 system. Targeting nucleotides were designed using http://crispr.mit.edu. Oligonucleotides of human PRKAG1 were inserted into pX260 vector, human PRKAB1, PRKAB2, PRKAG2, and PRKAG3 into pX330 vector, and mouse AK1 into the lentiCRISPRv2 vector. The individual constructs were then subjected to transfection of HEK293T cells (pX260 and pX330) followed by single-cell sorting into 96-well dishes, or lentivirus packaging (lentiCRISPRv2) using HEK293T cells. In both conditions, cells were transfected with 3 µg of DNA with Lipofectamine 2000 (Invitrogen, Cat. 11668-027) per well of a 6-well plate. Clones were expanded and evaluated for knockout status by sequencing.

Cell culture, transient transfection and lentivirus Infection

HEK293T, HEK293 and MEFs were maintained in Dulbecco’s modified Eagle’s medium (DMEM, Gibco, cat. 11965) supplemented with 10% fetal bovine serum (FBS), 100 IU penicillin, 100 mg/ml streptomycin at 37 °C in a humidified incubator containing 5% CO₂. Polyethylenimine (Polysciences, Inc., Cat. #23966) at a final concentration of 10 μM was used to transfect HEK293T cells. Total DNA for each plate was adjusted to the same amount by using relevant empty vector. Transfected cells were harvested at 24 h after transfection. Lentiviruses for infection of the MEFs were packaged in HEK293T cells using Lipofectamine 2000 transfection. At 30 h post transfection, medium was collected and added to the cells. The cells were incubated for another 24 h. LAMTOR1^F/For AXIN^F/F MEFs were established by introducing SV40 T antigen into primary cultured embryonic cells from a mouse litter. LAMTOR1^−/− or AXIN^−/− MEFs were generated by infecting LAMTOR1^F/F or AXIN^F/F MEFs with adenovirus expressing Cre recombinase for 12 h. The infected cells were then incubated in fresh DMEM for another 8–10 h before further treatments. Cells were verified to be free of mycoplasma contamination and authenticated by STR sequencing.

For glucose starvation, cells were rinsed twice with PBS, and then incubated in glucose-free DMEM (Gibco, cat. 11966) supplemented with 10% FBS and 1 mM sodium pyruvate (Gibco, cat. 11360) for desired periods of time at 37 °C. For starvation for glucose plus glutamine, cells were incubated in DMEM with 5 mM glucose overnight, and then incubated for desired periods of time in fresh DMEM lacking glucose, glutamine, phenol red or pyruvate (Gibco cat. A14430-01), in which 1 mM pyruvate was added, with or without Glutamax™ (Gibco cat. 35050-038) and D-glucose (Gibco cat. A2494001), all without FBS.

Subcellular fractionation

All buffers used for subcellular fractionation contained a protease inhibitor cocktail (Sigma, cat. P8340).

Lysosomes were purified by Lysosome Isolation Kit according to the manufacturer’s instructions with minor modifications. Briefly, cells from sixty 10-cm dishes (60–80% confluence) were collected by direct scra** at room temperature, followed by centrifugation for 5 min at 500 g at 37 °C. Cells were resuspend in 7 ml of 1× Extraction Buffer at room temperature. For isolating lysosomes from mouse livers, approximately 500 mg of frozen liver tissue was directly homogenized in 7 ml of ice-cold 1× Extraction Buffer. The cell/liver homogenates were then homogenized in a 7-ml Dounce homogenizer (Sigma, cat. P0610) for 120 strokes on ice followed by centrifuging for 10 min at 1000 g, 4 °C to yield post-nuclear supernatants (PNS). The PNS were then centrifuged for 20 min at 20,000 g and the pellets were suspended in 1× Extraction Buffer by gentle pipetting to generate Crude Lysosomal Fractions (CLF). The volume of CLF was adjusted to 2.4 ml and then equally divided into six 1.5 ml Eppendorf tubes (400 μl per tube). 253 μl of OptiPrep and 137 μl of 1× OptiPrep Dilution Buffer were added to each CLF and mixed by gentle pipetting. The mixtures were defined as the Diluted OptiPrep Fraction (DOF). Each DOF (0.8 ml) was loaded onto an 11 × 60 mm centrifuge tube at the top of 27% (0.4 ml) and 22.5% (0.5 ml) OptiPrep solution cushions, and then overlaid with 16% (1 ml), 12% (0.9 ml) and 8% (0.3 ml) OptiPrep solutions. The tubes were then centrifuged on a SW60 Ti rotor (Beckman) at 150,000 g for 4 h at 4 °C and the fractions at the top of 12% OptiPrep solution were collected as the crude lysosome fractions. The fractions were diluted with two volumes of PBS, followed by centrifugation at 20,000 g for 20 min. The sediment was the lysosome fraction.

Mitochondria were purified as described previously⁷³ with minor modifications. Briefly, forty 10-cm dishes of regularly cultured, severely starved or AICAR-treated cells (60–80% confluence), or sixty 10-cm dishes of glucose-starved cells were collected by direct scra** at room temperature, followed by centrifugation for 5 min at 500 g at 37 °C. Cells were then resuspended in 20 ml of ice-cold IB_cells-1 buffer (225 mM mannitol, 75 mM sucrose, 0.1 mM EGTA and 30 mM Tris-HCl, pH 7.4). To isolate mitochondria from mouse livers, approximately 500 mg of frozen liver was directly homogenized in 10 ml of IB_liver-1 buffer (225 mM mannitol, 75 mM sucrose, 0.5% BSA, 0.5 mM EGTA and 30 mM Tris-HCl, pH 7.4). Cells or liver homogenates were then homogenized in a 40-ml Dounce homogenizer (Sigma, cat. D9188) for 100 strokes, followed by two rounds of centrifugation for 5 min at 600 g (for cells) or 740 g (for liver tissues) at 4 °C. The supernatants were then collected and centrifuged for 10 min at 7000 g (for cells) or 9000 g (for liver tissues) at 4 °C. The pellets were then washed with 20 ml of ice-cold IB_cells-2 buffer (for cells, 225 mM mannitol, 75 mM sucrose and 30 mM Tris-HCl pH 7.4) or ice-cold IB_liver-2 buffer (for liver tissues, IB_cells-2 buffer supplemented with 0.5% BSA) twice. The suspensions were first centrifuged at 7000 g (for cells) or 10,000 g (for liver tissues) and were centrifuged again at 10,000 g, both for 10 min at 4 °C. Note that for liver tissues, the pellets subjected to the second time of centrifugation were resuspended in 20 ml of ice-cold IB_cells-2. The pellets were then resuspended in 2 ml of ice-cold MRB buffer (250 mM mannitol, 5 mM HEPES pH 7.4 and 0.5 mM EGTA) and were loaded on top of 10 ml of Percoll medium (225 mM mannitol, 25 mM HEPES pH 7.4, 1 mM EGTA and 30% Percoll (v/v)) in 14 × 89-mm centrifuge tubes (Beckman, ref. 344059). The tubes were then centrifuged on a SW41 rotor (Beckman) at 95,000 g for 0.5 h at 4 °C, and the dense band located approximately at the bottom of each tube was collected. The collected fractions were diluted with 10 volumes of MRB buffer, followed by centrifugation at 6300 g for 10 min at 4 °C and washed with 2 ml of MRB buffer, followed by centrifugation at 6300 g for 10 min at 4 °C. The pellets contained purified mitochondria.

Cytosol was purified as described.⁷⁴ Briefly, some 0.15 g of each freshly excised liver tissue or ten 10-cm dishes of cells were homogenized in 800 μl of the homogenization buffer (HB) containing 250 mM sucrose, 3 mM imidazole, pH 7.4. Homogenates were then passed through a 22-G needle attached to a 1-ml syringe for six times, and were then centrifuged at 2000 g for 10 min to yield PNS. PNS samples were then loaded on the top of 11 × 60-mm centrifuge tubes that had been loaded sequentially with 1 ml of 40.6% sucrose (dissolved in HB), 1 ml of 35% sucrose (dissolved in HB), and 1 ml of 25% sucrose (dissolved in HB). Tubes were then centrifuged on an SW60 Ti rotor (Beckman) at 35,000 rpm for 1 h at 4 °C, and the top fractions (about 200 μl) were collected as cytosolic fraction.

Fractionation of nuclei was performed as described previously.¹⁹ Briefly, cells were homogenized in Buffer A (10 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, and 0.15% NP-40) and placed in ice for 15 min. The homogenates were centrifuged at 12,000 g for 1 min at 4 °C. The pellets were washed three times with Buffer A and then resuspended in Buffer B (20 mM HEPES, pH 7.9, 0.4 mM NaCl, 1 mM EDTA, 1 mM EGTA, 0.5% NP-40) and sonicated at 4 °C. Cellular debris was removed by centrifugation at 12,000 g for 30 min at 4 °C, and the supernatant was nuclear fraction.

Protein production

AXIN1, and AMPK complexes were expressed and purified in E. coli as described previously.¹⁹ Briefly, full length AXIN1 was cloned into pET32a vector and transformed into the E. coli strain BL21 (DE3). The transformed cells were induced with 0.1 mM IPTG when the culture attained an optical density of 0.4 at 600 nm (for AMPK, to 1.0 at 600 nm). After growing for 4 h at 16 °C (for AMPK, growing for 16 h), the cells were collected, homogenized in buffer (50 mM sodium phosphate, pH 7.4, 150 mM NaCl, 1% Triton X-100, 5% glycerol) and sonicated. Cell debris was removed by centrifugation. The supernatant was collected and subjected to ultracentrifugation at 150,000 g for 30 min, followed by purification of expressed protein with Nickel affinity gel and washed by 50 mM sodium phosphate, pH 7.4, 150 mM NaCl. The protein was eluted from the affinity resin by 50 mM sodium phosphate, pH 7.4, 150 mM NaCl, 250 mM imidazole, and concentrated to about 3 mg/ml before further purification by gel filtration (Superdex-200, GE Healthcare). The buffer for gel filtration contained 50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 5% glycerol and 1 mM DTT.

In vitro AMPK phosphorylation assay

Bacterially expressed and purified AMPK (800 ng) was incubated with active LKB1/MO25/STRAD complex (200 ng) at 32 °C for 15 min in a kinase buffer containing 5 mM ATP with or without 60 μM AMP and 1 μg of AXIN1. Phosphorylation of AMPK was determined by immunoblotting.

Immunoprecipitation (IP) and immunoblotting

IP of endogenous AMPKα was performed as described previously¹⁹ and was also used for IP ACC1 and ACC2, although with some modifications. Briefly, cells from a single 15-cm dish (grown to 80% confluence) or 50 mg liver (for each lane) were collected and lysed with 500 μl of ice-cold lysis buffer (20 mM Tris-HCl, pH 7.5, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100, 1 mM β-glycerolphosphate, with protease inhibitor cocktail. Note that 150 mM NaCl was used for IP of ACC1 and ACC2, and 50 mM NaCl was used for IP of AMPKα. Lysates were incubated with respective antibodies for 4 h (ACC1 and ACC2) or overnight (AMPKα). Protein aggregates were pre-cleared by centrifugation at 20,000 g for 10 min, and protein A/G beads (1:100) were then added into the lysates and mixed for another 1 (for ACC1 and ACC2) to 3 h (for AMPKα). The beads were washed with 100x volumes of lysis buffer for 3 times at 4 °C and then mixed with an equal volume of 2× SDS sample buffer for immunoblotting.

To analyze the levels of p-AMPKα and p-ACC in MEFs, cells grown to 70–80% confluence in a well of a 6-well dish were lysed with 250 μl of ice-cold lysis buffer. The lysates were then centrifuged at 20,000 g for 10 min at 4 °C and an equal amount of 2× SDS sample buffer was into each supernatant. The levels of p-AMPKα and p-ACC were then analyzed by immunoblotting. To analyze the levels of p-AMPKα and p-ACC in liver, the freshly excised liver tissues were added with ice cold lysis buffer (10 μl/mg liver weight), followed by homogenization and centrifugation as described above. The lysates were then mixed with 2× SDS sample buffer and then subjected to immunoblotting. Levels of total proteins and the levels of phosphorylation of proteins were analyzed on separate gels and representative images were shown. The band intensities on developed films were quantified using Image J software (National Institutes of Health Freeware).

Fluorescence microscopy

HEK293T infected with different fusion constructs of β1 were grown on glass coverslips in 6-well dishes and were cultured to 60–80% confluence. Cells were fixed with 1 ml of 4% formaldehyde (diluted in PBS) at room temperature for 20 min. They were rinsed twice with 1 ml PBS (room temperature) and then permeabilized with 1 ml of 0.1% Triton X-100 (diluted in PBS) for 5 min at 4 °C. Cells were rinsed twice with 1 ml PBS, and were incubated with primary antibodies overnight at 4 °C. The cells were then rinsed three times with 1 ml PBS, and then incubated with Alexa-Fluor 594-conjugated anti mouse secondary antibody (Molecular Probes, cat. A11032; diluted 1:100 in PBS) and Alexa-Fluor 488-conjugated anti-rabbit secondary antibody (Molecular Probes, R37117; diluted 1:100 in PBS) for 8 h at room temperature in the dark. They were washed four times with 1 ml PBS, and then mounted on glass slides using ProLong Diamond Antifade Mountant (Molecular Probes, cat. P36970). Cells were imaged under a Zeiss LSM 780. Samples were excited with an Ar gas laser (Zeiss, laser module LGK 7812) using a 488-nm laser line for Alexa-Fluor 488 dye (green channel), and with a HeNe gas laser (Zeiss, LGK 7512 PF) using a 594-nm laser line for Alexa-Fluor 594 dye (red channel). Confocal microscope images were taken with a 63× oil objective and representative images were shown. The parameters, including ‘PMT voltage’, ‘Offset’, ‘Pinhole’ and ‘Gain’, were kept unchanged between each image taken.

CE-MS-based analysis of AMP, ADP, and ATP

Sample preparation for CE (capillary electrophoresis)-MS was carried out as described previously,^75,76 with some modifications. In general, each measurement required cells to be collected from a 10-cm dish (60–70% confluence) or 100 mg of liver tissue. For analysis of metabolites, cells were rinsed with 20 ml of 5% mannitol solution (dissolved in water) and instantly frozen in liquid nitrogen. Cells were then lysed with 1 ml of methanol containing internal standards 1 [IS1 (Human Metabolome Technologies, H3304-1002, 1:200), used to standardize the metabolite intensity and to adjust the migration time], and were scraped from the dish. For analysis of metabolites in liver, the fleshly excised tissue was freeze-clamped first, then washed in pre-cooled 5% mannitol solution and grinded in 1 ml of methanol with 50 μM IS1. The lysate was then mixed with 1 ml of chloroform and 400 μl of water by 20 s of vortexing. After centrifugation at 15,000 g for 15 min at 4 °C, 450 μl of aqueous phase was collected and was then filtrated through a 5 kDa cutoff filter (Millipore, cat. UFC3LCCNB-HMT) by centrifuging at 10,000 g for 3 h at 4 °C. Simultaneously, the quality control (QC) sample was prepared by combining 100 μl of the aqueous phase from each samples and then filtered. The filtered aqueous phase was then freeze-dried in a vacuum concentrator and then dissolved in water containing internal standards 3 [IS3 (Human Metabolome Technologies, H3304-1104, 1:200), to adjust the migration time]. 20 μl of re-dissolved solution was then loaded into an injection vial with a conical insert for CE-TOF MS (Agilent Technologies 7100, equipped with 6224 mass spectrometer) analysis. For quantification of AMP and ATP, [U-¹³C, ¹⁵N]AMP and [U-¹³C, ¹⁵N]ATP were used to generate standard curves by plotting the ratios of detected labelled AMP or ATP (areas) to the products of IS1 and IS3, against the added concentrations of labelled AMP or ATP. The standard curve of [U-¹³C, ¹⁵N]AMP was also used for quantification of ZMP, because of the structural similarity between them. The amount of AMP, ZMP, and ATP were then estimated according to equations generated from standard curves. The average cell volume, 2263 μm³, was determined by Imaris 7.4.0 software (Bitplane) from the axial image stacks of CDFA-SE labelled MEFs taken under Zeiss LSM780.

HPLC-MS-based analysis of short-chain acyl-CoA

Procedure of preparing samples for HPLC-MS-based acyl-CoA analysis was identical to that described in the CE-MS section, except that cells or liver tissues were rinsed in PBS, and that ¹³C2:0-CoA (150 μg/l) and ¹³C3:0-CoA (200 μg/l) prepared in methanol were added as internal standards. Some 800 μl of the aqueous phase of each sample was collected after centrifugation at 15,000 g for 15 min at 4 °C and was lyophilized in a vacuum concentrator. Samples were then re-dissolved in 20 μl of ice-cold 20% methanol. Measurement of acetyl- and malonyl-CoAs was based on a previous study⁷⁷ using a QTRAP (SCIEX, QTRAP 6500 plus) mass spectrometer interfaced with a UPLC system (Waters, ACQUITY UPLC system). Some 5 μl of each sample were injected onto an Acquity HSS T3 column (2.1 × 50 mm, 1.7 μm, Waters) attached to an Acquity BEH C18 pre-column (2.1 × 5 mm, 1,7 μm, Waters). Mobile phases A and B were water and acetonitrile, respectively (both contains 10 mM ammonium acetate and 0.05% ammonium hydroxide). The column temperature was maintained at 30 °C and autosampler temperature at 8 °C. The gradients were as follows: t = 0 min, 3% B; t = 1 min, 3% B; t = 13 min, 15% B; t = 14.1 min, 100% B; t = 14.7 min, 100% B; t = 14.71 min, 3% B; t = 17 min, 3% B. Mass spectrometry was in positive mode with multiple reactions monitoring mode (MRM), declustering potentials (DP) and collision energies (CE) set at 200 and 38 V, respectively (optimized using ¹³C2:0- and ¹³C3:0-CoA). The following transitions and retention time were used for monitoring each compound: 812.0, 305.3, and 2.2 min for ¹³C2:0-CoA, 810.1, 303.1, and 2.2 min for acetyl-CoA, 856.9, 350.2, and 0.8 min for ¹³C3:0-CoA, and 854.1, 347.1, and 0.8 min for malonyl-CoA. It should be noted that acyl-CoAs were extremely easy to degrade even frozen at −80 °C, so no pause was taken during the whole analytic process and no more than 6 samples were re-dissolved and kept in autosampler at one time.

Statistical analysis

One-way or two-way ANOVA with post hoc analysis was used to compare values among different experimental groups. For experiments with only two groups, a two-tailed Student’s t test was used as specified in the figure legends. For ANOVA, the homogeneity of variance was firstly tested by Levene’s test. If the results were similar, the Tukey’s test was preceded, and if not, the Games-Howell’s test was processed. Similar procedures were followed when the Student’s t test was performed. No samples or animals were excluded from the analysis. Tests were performed with SPSS Statistics 17.0 program, and p < 0.05 was considered statistically significant. Statistical significance is shown as *p < 0.05, **p < 0.01, ***p < 0.001; N.S., not significant.

References

Carling, D., Thornton, C., Woods, A. & Sanders, M. J. AMP-activated protein kinase: new regulation, new roles? Biochem. J. 445, 11–27 (2012).
Article CAS Google Scholar
Steinberg, G. R. & Kemp, B. E. AMPK in Health and Disease. Physiol. Rev. 89, 1025–1078 (2009).
Article CAS Google Scholar
Herzig, S. & Shaw, R. J. AMPK: guardian of metabolism and mitochondrial homeostasis. Nature reviews. Mol. Cell Biol. https://doi.org/10.1038/nrm.2017.95 (2017).
Article Google Scholar
Lin, S. C. & Hardie, D. G. AMPK: sensing glucose as well as cellular energy status. Cell Metab. 27, 299–313 (2017).
Article Google Scholar
Viollet, B. et al. AMPK inhibition in health and disease. Crit. Rev. Biochem. Mol. Biol. 45, 276–295 (2010).
Article CAS Google Scholar
Scott, J. W. et al. CBS domains form energy-sensing modules whose binding of adenosine ligands is disrupted by disease mutations. J. Clin. Invest. 113, 274–284 (2004).
Article CAS Google Scholar
**ao, B. et al. Structural basis for AMP binding to mammalian AMP-activated protein kinase. Nature 449, 496–500 (2007).
Article CAS Google Scholar
Gu, X. et al. Deconvoluting AMP-dependent kinase (AMPK) adenine nucleotide binding and sensing. J. Biol. Chem. 292, 12653–12666 (2017).
Article CAS Google Scholar
Ross, F. A., Jensen, T. E. & Hardie, D. G. Differential regulation by AMP and ADP of AMPK complexes containing different gamma subunit isoforms. Biochem. J. 473, 189–199 (2016).
Article CAS Google Scholar
Hawley, S. A. et al. Calmodulin-dependent protein kinase kinase-beta is an alternative upstream kinase for AMP-activated protein kinase. Cell Metab. 2, 9–19 (2005).
Article CAS Google Scholar
Woods, A. et al. Ca2⁺/calmodulin-dependent protein kinase kinase-beta acts upstream of AMP-activated protein kinase in mammalian cells. Cell Metab. 2, 21–33 (2005).
Article CAS Google Scholar
Hurley, R. L. et al. The Ca²⁺/calmoldulin-dependent protein kinase kinases are AMP-activated protein kinase kinases. J. Biol. Chem. 280, 29060–29066 (2005).
Article CAS Google Scholar
Salt, I. P., Johnson, G., Ashcroft, S. J. H. & Hardie, D. G. AMP-activated protein kinase is activated by low glucose in cell lines derived from pancreatic b cells, and may regulate insulin release. Biochem. J. 335, 533–539 (1998).
Article CAS Google Scholar
Zhang, C. S. et al. Fructose-1,6-bisphosphate and aldolase mediate glucose sensing by AMPK. Nature 548, 112–116 (2017).
Article CAS Google Scholar
Gonzalez, P. S. et al. Mannose impairs tumour growth and enhances chemotherapy. Nature 563, 719–723 (2018).
Article CAS Google Scholar
Nada, S. et al. The novel lipid raft adaptor p18 controls endosome dynamics by anchoring the MEK-ERK pathway to late endosomes. EMBO J. 28, 477–489 (2009).
Article CAS Google Scholar
Bar-Peled, L., Schweitzer, L. D., Zoncu, R. & Sabatini, D. M. Ragulator is a GEF for the rag GTPases that signal amino acid levels to mTORC1. Cell 150, 1196–1208 (2012).
Article CAS Google Scholar
Zhang, C. S. et al. The lysosomal v-ATPase-ragulator complex is a common activator for AMPK and mTORC1, acting as a switch between catabolism and anabolism. Cell Metab. 20, 526–540 (2014).
Article CAS Google Scholar
Zhang, Y. L. et al. AMP as a low-energy charge signal autonomously initiates assembly of AXIN-AMPK-LKB1 complex for AMPK activation. Cell Metab. 18, 546–555 (2013).
Article CAS Google Scholar
Miyamoto, T. et al. Compartmentalized AMPK signaling illuminated by genetically encoded molecular sensors and actuators. Cell Rep. 11, 657–670 (2015).
Article CAS Google Scholar
Salt, I. P. et al. AMP-activated protein kinase—greater AMP dependence, and preferential nuclear localization, of complexes containing the a2 isoform. Biochem. J. 334, 177–187 (1998).
Article CAS Google Scholar
Kazgan, N., Williams, T., Forsberg, L. J. & Brenman, J. E. Identification of a nuclear export signal in the catalytic subunit of AMP-activated protein kinase. Mol. Biol. Cell 21, 3433–3442 (2010).
Article CAS Google Scholar
Vincent, O., Townley, R., Kuchin, S. & Carlson, M. Subcellular localization of the Snf1 kinase is regulated by specific beta subunits and a novel glucose signaling mechanism. Genes Dev. 15, 1104–1114 (2001).
Article CAS Google Scholar
Liang, J. et al. Myristoylation confers noncanonical AMPK functions in autophagy selectivity and mitochondrial surveillance. Nat. Commun. 6, 7926 (2015).
Article CAS Google Scholar
Yi, C. et al. Formation of a Snf1-Mec1-Atg1 module on mitochondria governs energy deprivation-induced autophagy by regulating mitochondrial respiration. Dev. Cell 41, 59–71 e54 (2017).
Article CAS Google Scholar
Tsou, P., Zheng, B., Hsu, C. H., Sasaki, A. T. & Cantley, L. C. A fluorescent reporter of AMPK activity and cellular energy stress. Cell Metab. 13, 476–486 (2011).
Article CAS Google Scholar
Warden, S. M. et al. Post-translational modifications of the beta-1 subunit of AMP-activated protein kinase affect enzyme activity and cellular localization. Biochem. J. 354, 275–283 (2001).
Article CAS Google Scholar
Oakhill, J. S. et al. beta-Subunit myristoylation is the gatekeeper for initiating metabolic stress sensing by AMP-activated protein kinase (AMPK). Proc. Natl Acad. Sci. USA 107, 19237–19241 (2010).
Article CAS Google Scholar
Davies, S. P., Sim, A. T. & Hardie, D. G. Location and function of three sites phosphorylated on rat acetyl-CoA carboxylase by the AMP-activated protein kinase. Eur. J. Biochem 187, 183–190 (1990).
Article CAS Google Scholar
Winder, W. W. et al. Phosphorylation of rat muscle acetyl-CoA carboxylase by AMP-activated protein kinase and protein kinase A. J. Appl. Physiol. 82, 219–225 (1997).
Article CAS Google Scholar
Abu-Elheiga, L. et al. The subcellular localization of acetyl-CoA carboxylase 2. Proc. Natl. Acad. Sci. USA 97, 1444–1449 (2000).
Article CAS Google Scholar
Abu-Elheiga, L., Oh, W., Kordari, P. & Wakil, S. J. Acetyl-CoA carboxylase 2 mutant mice are protected against obesity and diabetes induced by high-fat/high-carbohydrate diets. Proc. Natl. Acad. Sci. USA 100, 10207–10212 (2003).
Article CAS Google Scholar
Wakil, S. J. & Abu-Elheiga, L. A. Fatty acid metabolism: target for metabolic syndrome. J. Lipid Res. 50(Suppl), S138–S143 (2009).
Article Google Scholar
Hardie, D. G., Ross, F. A. & Hawley, S. A. AMPK: a nutrient and energy sensor that maintains energy homeostasis. Nat. Rev. Mol. Cell Biol. 13, 251–262 (2012).
Article CAS Google Scholar
Li, Y. et al. AMPK phosphorylates and inhibits SREBP activity to attenuate hepatic steatosis and atherosclerosis in diet-induced insulin-resistant mice. Cell Metab. 13, 376–388 (2011).
Article CAS Google Scholar
Qian, X. et al. Conversion of PRPS hexamer to monomer by AMPK-mediated phosphorylation inhibits nucleotide synthesis in response to energy stress. Cancer Discov. 8, 94–107 (2018).
Article CAS Google Scholar
Inoki, K., Zhu, T. & Guan, K. L. TSC2 mediates cellular energy response to control cell growth and survival. Cell 115, 577–590 (2003).
Article CAS Google Scholar
Gwinn, D. M. et al. AMPK phosphorylation of raptor mediates a metabolic checkpoint. Mol. Cell 30, 214–226 (2008).
Article CAS Google Scholar
Zoncu, R. et al. mTORC1 senses lysosomal amino acids through an inside-out mechanism that requires the vacuolar H(+)-ATPase. Science 334, 678–683 (2011).
Article CAS Google Scholar
Efeyan, A. et al. Regulation of mTORC1 by the Rag GTPases is necessary for neonatal autophagy and survival. Nature 493, 679–683 (2013).
Article CAS Google Scholar
Pehmoller, C. et al. Genetic disruption of AMPK signaling abolishes both contraction- and insulin-stimulated TBC1D1 phosphorylation and 14-3-3 binding in mouse skeletal muscle. Am. J. Physiol. Endocrinol. Metab. 297, E665–E675 (2009).
Article Google Scholar
Vichaiwong, K. et al. Contraction regulates site-specific phosphorylation of TBC1D1 in skeletal muscle. Biochem. J. 431, 311–320 (2010).
Article CAS Google Scholar
Egan, D. F. et al. Phosphorylation of ULK1 (hATG1) by AMP-activated protein kinase connects energy sensing to mitophagy. Science 331, 456–461 (2011).
Article CAS Google Scholar
Kim, J., Kundu, M., Viollet, B. & Guan, K. L. AMPK and mTOR regulate autophagy through direct phosphorylation of Ulk1. Nat. Cell Biol. 13, 132–141 (2011).
Article CAS Google Scholar
Kim, J. et al. Differential regulation of distinct Vps34 complexes by AMPK in nutrient stress and autophagy. Cell 152, 290–303 (2013).
Article CAS Google Scholar
Youle, R. J. & van der Bliek, A. M. Mitochondrial fission, fusion, and stress. Science 337, 1062–1065 (2012).
Article CAS Google Scholar
Toyama, E. Q. et al. Metabolism. AMP-activated protein kinase mediates mitochondrial fission in response to energy stress. Science 351, 275–281 (2016).
Article CAS Google Scholar
Kim, N. et al. AMPKalpha2 translocates into the nucleus and interacts with hnRNP H: implications in metformin-mediated glucose uptake. Cell. Signal. 26, 1800–1806 (2014).
Article CAS Google Scholar
Wu, D. et al. Glucose-regulated phosphorylation of TET2 by AMPK reveals a pathway linking diabetes to cancer. Nature 559, 637–641 (2018).
Article CAS Google Scholar
Chia, I. V. & Costantini, F. Mouse axin and axin2/conductin proteins are functionally equivalent in vivo. Mol. Cell. Biol. 25, 4371–4376 (2005).
Article CAS Google Scholar
Nusse, R. & Clevers, H. Wnt/beta-catenin signaling, disease, and emerging therapeutic modalities. Cell 169, 985–999 (2017).
Article CAS Google Scholar
Corton, J. M., Gillespie, J. G., Hawley, S. A. & Hardie, D. G. 5-aminoimidazole-4-carboxamide ribonucleoside. A specific method for activating AMP-activated protein kinase in intact cells? Eur. J. Biochem 229, 558–565 (1995).
Article CAS Google Scholar
Faupel, R. P., Seitz, H. J., Tarnowski, W., Thiemann, V. & Weiss, C. The problem of tissue sampling from experimental animals with respect to freezing technique, anoxia, stress and narcosis. A new method for sampling rat liver tissue and the physiological values of glycolytic intermediates and related compounds. Arch. Biochem. Biophys. 148, 509–522 (1972).
Article CAS Google Scholar
Dale, S., Wilson, W. A., Edelman, A. M. & Hardie, D. G. Similar substrate recognition motifs for mammalian AMP-activated protein kinase, higher plant HMG-CoA reductase kinase-A, yeast SNF1, and mammalian calmodulin-dependent protein kinase I. FEBS Lett. 361, 191–195 (1995).
Article CAS Google Scholar
Scott, J. W. et al. Thienopyridone drugs are selective activators of AMP-activated protein kinase beta1-containing complexes. Chem. Biol. 15, 1220–1230 (2008).
Article CAS Google Scholar
Luo, W. et al. Axin contains three separable domains that confer intramolecular, homodimeric, and heterodimeric interactions involved in distinct functions. J. Biol. Chem. 280, 5054–5060 (2005).
Article CAS Google Scholar
Kim, S. E. et al. Wnt stabilization of beta-catenin reveals principles for morphogen receptor-scaffold assemblies. Science 340, 867–870 (2013).
Article CAS Google Scholar
**n, F. J., Wang, J., Zhao, R. Q., Wang, Z. X. & Wu, J. W. Coordinated regulation of AMPK activity by multiple elements in the alpha-subunit. Cell Res. 23, 1237–1240 (2013).
Article CAS Google Scholar
Li, X. et al. Structural basis of AMPK regulation by adenine nucleotides and glycogen. Cell Res. 25, 50–66 (2015).
Article Google Scholar
Ross, F. A., MacKintosh, C. & Hardie, D. G. AMP-activated protein kinase: a cellular energy sensor that comes in 12 flavours. FEBS J. 283, 2987–3001 (2016).
Article CAS Google Scholar
Olivier, S., Foretz, M. & Viollet, B. Promise and challenges for direct small molecule AMPK activators. Biochem. Pharmacol. 153, 147–158 (2018).
Article CAS Google Scholar
Khan, A. S. & Frigo, D. E. A spatiotemporal hypothesis for the regulation, role, and targeting of AMPK in prostate cancer. Nat. Rev. Urol. 14, 164–180 (2017).
Article CAS Google Scholar
Boudaba, N. et al. AMPK re-activation suppresses hepatic steatosis but its downregulation does not promote fatty liver development. EBioMedicine 28, 194–209 (2018).
Article Google Scholar
Cao, Y. et al. Activation of gamma2-AMPK suppresses ribosome biogenesis and protects against myocardial ischemia/reperfusion injury. Circ. Res. 121, 1182–1191 (2017).
Article CAS Google Scholar
Woods, A. et al. Liver-specific activation of AMPK prevents steatosis on a high-fructose diet. Cell Rep. 18, 3043–3051 (2017).
Article CAS Google Scholar
Su, H. et al. VPS34 acetylation controls its lipid kinase activity and the initiation of canonical and non-canonical autophagy. Mol. Cell 67, 907–921 e907 (2017).
Article CAS Google Scholar
Cheong, H., Lindsten, T., Wu, J., Lu, C. & Thompson, C. B. Ammonia-induced autophagy is independent of ULK1/ULK2 kinases. Proc. Natl Acad. Sci. USA 108, 11121–11126 (2011).
Article CAS Google Scholar
Soeters, M. R., Soeters, P. B., Schooneman, M. G., Houten, S. M. & Romijn, J. A. Adaptive reciprocity of lipid and glucose metabolism in human short-term starvation. Am. J. Physiol. Endocrinol. Metab. 303, E1397–E1407 (2012).
Article CAS Google Scholar
Cahill, G. F. Jr. Starvation in man. Clin. Endocrinol. Metab. 5, 397–415 (1976).
Article CAS Google Scholar
Pilegaard, H., Saltin, B. & Neufer, P. D. Effect of short-term fasting and refeeding on transcriptional regulation of metabolic genes in human skeletal muscle. Diabetes 52, 657–662 (2003).
Article CAS Google Scholar
Li, T. Y. et al. ULK1/2 constitute a bifurcate node controlling glucose metabolic fluxes in addition to autophagy. Mol. Cell 62, 359–370 (2016).
Article CAS Google Scholar
Abe, Y. et al. Mouse model of liver ischemia and reperfusion injury: method for studying reactive oxygen and nitrogen metabolites in vivo. Free Radic. Biol. & Med. 46, 1–7 (2009).
Article CAS Google Scholar
Wieckowski, M. R., Giorgi, C., Lebiedzinska, M., Duszynski, J. & Pinton, P. Isolation of mitochondria-associated membranes and mitochondria from animal tissues and cells. Nat. Protoc. 4, 1582–1590 (2009).
Article CAS Google Scholar
Kobayashi, T. et al. Separation and characterization of late endosomal membrane domains. J. Biol. Chem. 277, 32157–32164 (2002).
Article CAS Google Scholar
Zhao, J. et al. Study of polar metabolites in tobacco from different geographical origins by using capillary electrophoresis–mass spectrometry. Metabolomics 10, 805–815 (2014).
Article CAS Google Scholar
Zhao, Y. et al. A metabolomics study delineating geographical location-associated primary metabolic changes in the leaves of growing tobacco plants by GC-MS and CE-MS. Sci. Rep. 5, 16346 (2015).
Article CAS Google Scholar
Wang, S., Wang, Z., Zhou, L., Shi, X. & Xu, G. Comprehensive analysis of short-, medium-, and long-chain acyl-coenzyme a by online two-dimensional liquid chromatography/mass spectrometry. Anal. Chem. 89, 12902–12908 (2017).
Article CAS Google Scholar

Download references

Acknowledgements

ACC1-floxed mouse (Stock No. 030954, The Jackson Laboratory) was a generous gift from Prof. Jay Horton (UT Southwestern Medical Center), and the ACC2^−/− mouse from Prof. Tian Xu (Institute of Developmental Biology and Molecular Medicine, Fudan University). Heterotrimeric AMPK cloned into pET21b was a generous gift from Prof. Dietbert Neumann (Swiss Federal Institute of Technology). We also thank Jiayuan Zhang from **amen University for the artwork of Fig. 4a. This work was supported by grants from the National Key R&D Program of China (2016YFA0502001) and the National Natural Science Foundation of China (#31730058, #31430094, #31601152 and #31690101) through S-C.L., by the Open Research Fund of State Key Laboratory of Cellular Stress Biology, **amen University (SKLCSB2017KF002) through H-L.P., and by Wellcome Trust (204766/Z/16/Z) through D.G.H.

Author information

These authors contributed equally: Yue Zong, Chen-Song Zhang, Mengqi Li, Wen Wang

Authors and Affiliations

State Key Laboratory for Cellular Stress Biology, School of Life Sciences, **amen University, 361102, Fujian, China
Yue Zong, Chen-Song Zhang, Mengqi Li, Teng Ma, **-Wei Feng, **ao Tian, Qu Qi, Yu-Qing Wu, Cixiong Zhang, Zhiyun Ye, Shu-Yong Lin & Sheng-Cai Lin
CAS Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences, 116023, Dalian, China
Wen Wang, Zhichao Wang & Hai-Long Piao
University of Chinese Academy of Sciences, 100049, Bei**g, China
Wen Wang, Zhichao Wang & Hai-Long Piao
Division of Cell Signalling and Immunology, College of Life Sciences, University of Dundee, DD1 5EH, Dundee, Scotland, UK
Simon A. Hawley & D. Grahame Hardie

Authors

Yue Zong
View author publications
You can also search for this author in PubMed Google Scholar
Chen-Song Zhang
View author publications
You can also search for this author in PubMed Google Scholar
Mengqi Li
View author publications
You can also search for this author in PubMed Google Scholar
Wen Wang
View author publications
You can also search for this author in PubMed Google Scholar
Zhichao Wang
View author publications
You can also search for this author in PubMed Google Scholar
Simon A. Hawley
View author publications
You can also search for this author in PubMed Google Scholar
Teng Ma
View author publications
You can also search for this author in PubMed Google Scholar
**-Wei Feng
View author publications
You can also search for this author in PubMed Google Scholar
**ao Tian
View author publications
You can also search for this author in PubMed Google Scholar
Qu Qi
View author publications
You can also search for this author in PubMed Google Scholar
Yu-Qing Wu
View author publications
You can also search for this author in PubMed Google Scholar
Cixiong Zhang
View author publications
You can also search for this author in PubMed Google Scholar
Zhiyun Ye
View author publications
You can also search for this author in PubMed Google Scholar
Shu-Yong Lin
View author publications
You can also search for this author in PubMed Google Scholar
Hai-Long Piao
View author publications
You can also search for this author in PubMed Google Scholar
D. Grahame Hardie
View author publications
You can also search for this author in PubMed Google Scholar
Sheng-Cai Lin
View author publications
You can also search for this author in PubMed Google Scholar

Contributions

Y.Z., C-S.Z., M.L., and S-C.L. conceived the study and designed the experiments. Y.Z., C-S.Z., and M.L. performed the subcellular fractionation, immunoprecipitation, in vitro reconstitution and the associated western blot analyses with assistance from S.A.H, T.M., J.-W.F., X.T., Q.Q., and Y.-Q.W. Y.Z., W.W., Z.W., and C.Z. performed the CE-MS- and HPLC-MS-based analysis of adenylates and CoAs under the guidance of H.-L.P. M.L. performed the confocal imaging acquisition. Z.Y., S-Y.L., H.-L.P., and D.G.H. helped with discussion and interpretation of results. D.G.H. and S-C.L. wrote the manuscript.

Corresponding author

Correspondence to Sheng-Cai Lin.

Ethics declarations

Competing interests

The authors declare no competing interests.

Supplementary information

Fig. S1

Fig. S2

Fig. S3

Fig. S4

Rights and permissions

Reprints and permissions

About this article

Cite this article

Zong, Y., Zhang, CS., Li, M. et al. Hierarchical activation of compartmentalized pools of AMPK depends on severity of nutrient or energy stress. Cell Res 29, 460–473 (2019). https://doi.org/10.1038/s41422-019-0163-6

Download citation

Received: 19 February 2019
Accepted: 15 March 2019
Published: 04 April 2019
Issue Date: June 2019
DOI: https://doi.org/10.1038/s41422-019-0163-6
Springer Nature Singapore Pte Ltd.

This article is cited by

The scaffold protein AXIN1: gene ontology, signal network, and physiological function
- Lu Qiu
- Yixuan Sun
- Yanfeng Gao
Cell Communication and Signaling (2024)
TBC1D23 mediates Golgi-specific LKB1 signaling
- Yingfeng Tu
- Qin Yang
- Da Jia
Nature Communications (2024)
MLKL promotes hepatocarcinogenesis through inhibition of AMPK-mediated autophagy
- **anjun Yu
- Mengyuan Feng
- Qun Zhao
Cell Death & Differentiation (2024)
AMPK targets PDZD8 to trigger carbon source shift from glucose to glutamine
- Mengqi Li
- Yu Wang
- Sheng-Cai Lin
Cell Research (2024)
Rapid metabolic reprogramming mediated by the AMP-activated protein kinase during the lytic cycle of Toxoplasma gondii
- Yaqiong Li
- Zhipeng Niu
- Bang Shen
Nature Communications (2023)

Hierarchical activation of compartmentalized pools of AMPK depends on severity of nutrient or energy stress

Abstract

Similar content being viewed by others

Introduction

Roles of AMP in the hierarchical activation of AMPK

Discussion

Cell culture, transient transfection and lentivirus Infection

Subcellular fractionation

Protein production

In vitro AMPK phosphorylation assay

Immunoprecipitation (IP) and immunoblotting

Fluorescence microscopy

CE-MS-based analysis of AMP, ADP, and ATP

HPLC-MS-based analysis of short-chain acyl-CoA

Statistical analysis

References

Acknowledgements

Author information

Authors and Affiliations

Contributions

Corresponding author

Ethics declarations

Competing interests

Supplementary information

Rights and permissions

About this article

Cite this article

Share this article

This article is cited by

Search

Navigation