Sequestration-Mediated Downregulation of de Novo Purine Biosynthesis by AMPK.

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ACS Chem Biol. Author manuscript; available in PMC 2017 November 07. Published in final edited form as: ACS Chem Biol. 2016 July 15; 11(7): 1917–1924. doi:10.1021/acschembio.6b00039.

Sequestration-Mediated Downregulation of de Novo Purine Biosynthesis by AMPK Danielle L. Schmitt†, Yun-ju Cheng‡, Junyong Park‡, and Songon An*,† †Department

of Chemistry and Biochemistry, University of Maryland Baltimore County (UMBC), 1000 Hilltop Circle, Baltimore, Maryland 21250, United States

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‡Department

of Mathematics and Statistics, University of Maryland Baltimore County (UMBC), 1000 Hilltop Circle, Baltimore, Maryland 21250, United States

Abstract

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Dynamic partitioning of de novo purine biosynthetic enzymes into multienzyme compartments, purinosomes, has been associated with increased flux of de novo purine biosynthesis in human cells. However, we do not know of a mechanism by which de novo purine biosynthesis would be downregulated in cells. We have investigated the functional role of AMP-activated protein kinase (AMPK) in the regulation of de novo purine biosynthesis because of its regulatory action on lipid and carbohydrate biosynthetic pathways. Using pharmacological AMPK activators, we have monitored subcellular localizations of six pathway enzymes tagged with green fluorescent proteins under time-lapse fluorescence single-cell microscopy. We revealed that only one out of six pathway enzymes, formylglycinamidine ribonucleotide synthase (FGAMS), formed spatially distinct cytoplasmic granules after treatment with AMPK activators, indicating the formation of single-enzyme self-assemblies. In addition, subsequent biophysical studies using fluorescence recovery after photobleaching showed that the diffusion kinetics of FGAMS were slower when it localized inside the self-assemblies than within the purinosomes. Importantly, high-performance liquid chromatographic studies revealed that the formation of AMPK-promoted FGAMS selfassembly caused the reduction of purine metabolites in HeLa cells, indicating the downregulation of de novo purine biosynthesis. Collectively, we demonstrate here that the spatial sequestration of FGAMS by AMPK is a mechanism by which de novo purine biosynthesis is downregulated in human cells.

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*

Corresponding Author: Tel.: (+1-410) 455-2514. Fax: (+1-410) 455-1874. [email protected]. Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acschem-bio.6b00039. Figures S1–S6 (PDF) Author Contributions S.A. and D.L.S. designed research. D.L.S. performed experiments. S.A. and D.L.S. analyzed data. Y.C. and J.P. performed statistical analysis. S.A. and D.L.S. wrote the paper.

Notes The authors declare no competing financial interest.

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De novo purine biosynthesis is a 10-step enzymatic process catalyzed by six cytoplasmic enzymes, three of which are multifunctional. In this pathway, phosphoribosylpyrophosphate is converted to inosine monophosphate (IMP) in a process requiring a multitude of cofactors and 4 mol of ATP. In addition, de novo purine biosynthetic enzymes have been shown to compartmentalize into metabolic complexes, namely the purinosomes, in human cancer cells.1,2 Purinosome formation has been functionally correlated with the increased rate of de novo purine biosynthesis,3 followed by an increased pool of the final product, IMP.4 Recently, cellular signaling pathways have been identified to control the activity of de novo purine biosynthesis.5–8 Collectively, purinosome formation appears to be the mechanism for a cell to upregulate de novo purine biosynthesis. However, the basal level activity of de novo purine biosynthesis has been detected in the absence of purinosome assemblies or under conditions in which purinosome formation was not favorable.3,4,9 These data imply that purinosome-negative cells maintain a certain level of metabolic activity of de novo purine biosynthesis. Nevertheless, a downregulation mechanism of de novo purine biosynthesis has been elusive to date.

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We have sought to understand how AMP-activated protein kinase (AMPK) activation metabolically influences de novo purine biosynthesis in live human cells. AMPK is a key energy regulator in the cell, responding to the increased ratio of AMP/ATP for energy homeostasis.10 AMPK consists of three subunits: a kinase domain α and two regulatory subunits β and γ. In addition to AMP as an AMPK activator, the metabolic intermediate of the eighth step of de novo purine biosynthesis, 5-aminoimidazole-4-carboxamide ribonucleotide (ZMP in this article), is characterized as an allosteric activator of AMPK; both bind to the nucleotide-binding pocket in the γ subunit.11,12 Furthermore, pharmacological inhibition of the folate-requiring steps 3 and 9 of de novo purine biosynthesis was shown to temporally increase ZMP levels in cells, subsequently activating AMPK.13–15 However, the influence of AMPK activation on de novo purine biosynthesis, the other way around, has not been elucidated at the molecular level.

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In this work, we reveal that AMPK activators promote the spatial rearrangement of the subcellular localization of one of purine biosynthetic enzymes, formylglycinamidine ribonucleotide synthase (FGAMS), which catalyzes the fourth step of the pathway. The other five enzymes are unaffected by AMPK activation, indicating that the self-assembly of FGAMS is metabolically independent of the purinosome assembly. In addition, we demonstrate that AMPK activation decreases the steady-state levels of IMP, AMP, GMP, and ATP, implying the downregulation of de novo purine biosynthesis. Furthermore, we show

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that the FGAMS self-assembly is biologically different from the purinosome and spatially distinguishable from other cellular bodies within the cytoplasm, including autophagosomes, lysosomes, and other lipid-bound organelles. Given that FGAMS is structurally one of the core elements of the purinosome assembly,2,16 we conclude that the spatial sequestration of FGAMS by AMPK is the mechanism by which de novo purine biosynthesis is downregulated in human cells.

RESULTS AMPK-Activating Small Molecules Induced Clustering of FGAMS-EGFP

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To determine the mechanism of AMPK action on de novo purine biosynthesis at the molecular level, we expressed a reporter protein with an enhanced green fluorescent protein (EGFP) tag in live HeLa cells. Given the reversible compartmentalization property of the EGFP-tagged FGAMS (i.e., FGAMS-EGFP) in live cells,1 we studied the effect of three subgroups of AMPK activators on the subcellular localization of FGAMS-EGFP. First, HeLa cells expressing FGAMS-EGFP were treated with 150–300 μM of cell-permeable 5aminoimidazole-4-carboxamide ribonucleoside (AICAR). Based on the mechanism of AICAR’s action in HeLa cells, exogenously supplied AICAR will contribute to a temporal increase of cellular ZMP level within an hour,11,17 which in turn binds to the γ subunit to allosterically activate AMPK in cultured cell systems.18 We treated HeLa cells with 200 μM AICAR, in which FGAMS-EGFP is distributed diffusively in the cytoplasm. We monitored spatial clustering of FGAMS-EGFP in ~45% of transfected cells after 4 h incubation (Figure 1A and B). Similarly, under immunofluorescence microscopy, we observed the cytosolic clustering of endogenous FGAMS in fixed HeLa cells after the treatment of 200 μM AICAR (Supporting Information Figure S1). Second, we treated HeLa cells with biguanides, metformin (170–430 μM) and phenformin (210–410 μM), which are also known to indirectly activate AMPK through the inhibition of mitochondrial electron transfer organization.19 Likewise, both biguanide AMPK activators were found to induce the clustering of FGAMS-EGFP in ~69% and ~23% of transfected HeLa cells, respectively (Figure 1C–F). Third, we treated HeLa cells with 100 μM A769662, which binds in a pocket between the α and β subunits of AMPK, resulting in allosteric activation.20 After 4 h of A769662 incubation, we monitored that ~21% of HeLa cells displayed FGAMS-EGFP clusters (Figure 1G and H). As negative controls, a purine salvage enzyme, hypoxanthine phosphoribosyltransferase 1 (HPRT1), fused with EGFP (i.e., HPRT1-EGFP) was also subjected to our pharmacological studies. However, HPRT1-EGFP remained diffusive throughout the cytoplasm in the presence of either AICAR (250 μM) or metformin (170–290 μM; Figure 1I–L). HeLa cells expressing EGFP alone also showed no change in its localization for ~5 h in the presence of 300 μM AICAR, indicating no interference by the EGFP tag (Supporting Information Figure S2). Additionally, we confirmed the activation of AMPK by immunostaining the phospho-Thr172 of AMPK with a monoclonal antibody in nontransfected HeLa cells after 4 h of incubation with 200 μM AICAR and 170 μM metformin, respectively (Supporting Information Figure S3). Collectively, we conclude that the clustering of FGAMS-EGFP is promoted by pharmacologically activated AMPK in HeLa cells.


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AMPK Activation-Mediated Clusters of FGAMS-EGFP Did Not Recruit Other Purine Biosynthetic Enzymes We next determined the effect of AICAR treatment on five other purine biosynthetic enzymes.

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First, we evaluated the enzymes involved in the core structure of the purinosome based on previous studies.2,16 Since FGAMS robustly formed clusters in the presence of AICAR, the coclustering of phosphoribosyl pyrophosphate amidotransferase (PPAT) and a trifucntional enzyme possessing the activities of glycinamide ribonucleotide synthetase, glycinamide ribonucleotide transformylase and aminoimidazole ribonucleotide synthase (namely, TrifGART) would indicate the formation of the core structure of purinosomes. However, EGFP-tagged PPAT (i.e., PPAT-GFP), which catalyzes the first step of de novo purine biosynthesis, remained diffusive throughout the cytoplasm in the presence of 200 μM AICAR (Figure 2A and B). GFPS65T-tagged TrifGART (i.e., TrifGART-GFP; steps 2, 3, and 5) did not respond to 200 μM AICAR in HeLa cells, either (Figure 2C and D). It appears that the core structure of the purinosome does not form in the presence of AICAR.

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Second, we further expanded our localization studies to the rest of pathway enzymes. EGFPtagged phosphoribosyl aminoimidazole carboxylase/phosphoribosyl aminoimidazole succinocarboxamide synthase (PAICS-EGFP, steps 6 and 7) was exposed to 200 μM AICAR in HeLa cells for 4 h, but it remained diffusive under our conditions (Figure 2E and F). Adenylosuccinate lyase tagged with EGFP (ASL-EGFP, step 8), which catalyzes the production of ZMP, was not responsive to 200 μM AICAR, either (Figure 2G and H). Even EGFP-tagged aminoimidazolecarboxamide ribonucleotide transformylase/IMP cyclohydrolase (EGFP-ATIC, steps 9 and 10), which consumes ZMP, remained diffusive in the presence of 150–300 μM AICAR (Figure 2I and J) or 172 μM metformin (Supporting Information Figure S4) in HeLa cells, respectively. Thus, other than FGAMS-EGFP, no purine biosynthetic enzymes spatially compartmentalize into the observed cytosolic granules upon AMPK activation.

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Third, we examined whether coexpression of purine enzymes with FGAMS would promote coclustering events. We prepared HeLa cells coexpressing EGFP-ATIC with orange fluorescent protein-tagged FGAMS (FGAMS-OFP). Indeed, 200 μM AICAR and 290 μM metformin promoted the clustering of FGAMS-OFP, but EGFP-ATIC remained diffusive throughout the cytoplasm, respectively (Figure 3). These data indicate that the FGAMS clustering does not recruit other pathway enzymes and also that the clustering event is independent of the stoichiometric expression levels of the transfected pathway enzymes in cells. Collectively, these data support that the self-assembly of FGAMS is promoted in response to AMPK activation in HeLa cells. The Formation of FGAMS Self-Assembly Is Directly Associated with Activated AMPK We have also used small hairpin RNAs to knock down both AMPK α1 and α2 isoforms in HeLa cells (i.e., shRNAAMPK). After dual transfection of shRNAAMPK with a plasmid expressing FGAMS-EGFP, we treated the transfected HeLa cells with 200 μM AICAR for 4 h. We observed that ~15 ± 4% of cells exhibited clusters of FGAMS-EGFP (Figure 4),


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indicating that the knock-down of AMPK impaired the AICAR-induced activation of AMPK. In contrast, ~38 ± 9% of HeLa cells transfected with scrambled shRNAs (i.e., shRNAControl) and FGAMS-EGFP promoted fluorescent clusters in response to 200 μM AICAR (Figure 4), which is in good agreement with our observation presented in Figure 1A,B. Along with our extensive pharmacological studies, these data strongly support that the clustering of FGAMS-EGFP in response to AMPK activators is directly associated with AMPK. Characterization of the FGAMS Self-Assembly versus the Purinosome in HeLa Cells


It appears clear that the observed FGAMS self-assemblies above are not the purinosomes based on our imaging studies. We wonder whether there would be an alternative strategy, which would distinguish the FGAMS self-assembly from the purinosome. First, we quantitatively compared two sets of fluorescent images obtained from the AICAR-treated cluster-positive HeLa cells and the purinosome-positive HeLa cells1 from a purine-deficient medium. Unfortunately, their average sizes were very similar to each other, so that these two assemblies were indistinguishable (Supporting Information Figure S5), 0.46 μm2 versus 0.52 μm2 for FGAMS self-assemblies and purinosomes, respectively. Next, we measured apparent diffusion coefficients (Dapp) of FGAMS-EGFP in respective HeLa cells. To measure the diffusion coefficients of FGAMS-EGFP in the AICAR-treated cluster-positive HeLa cells, transfected HeLa cells were treated with 200 μM AICAR for 4 h prior to fluorescence recovery after photobleaching (FRAP). Independently, we also measured the diffusion coefficients of purinosome-associated FGAMS-EGFP in HeLa cells that were grown under purine-deficient conditions.1 Our FRAP analysis revealed that FGAMS-EGFP involved in the self-assemblies showed a ~2-fold slower diffusion coefficient (Dapp = 0.0017 ± 0.0010 μm2/s; NFRAP = 20) than the diffusion coefficient of FGAMS-EGFP in the purinosome-positive HeLa cells (Dapp = 0.0042 ± 0.0026 μm2/s; NFRAP = 36; Figure 5). Subsequent statistical analysis using three independent procedures (refer to Methods) revealed that all three p values between two groups were less than 0.0001, indicating their apparent diffusion coefficients are significantly different from each other. Due to the different protein composition between the self-assembly and the purinosome within the similar size of clusters, FGAMS-EGFP experiences statistically different molecular environments in two different spatial organizations. FGAMS Self-Assemblies Do Not Associate with Other Cellular Bodies or Organelles

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Since the FGAMS self-assembly is different from the purinosome, we further compared the FGAMS self-assembly with other known cellular bodies and organelles. First, we coimaged the FGAMS self-assemblies with lysosomes in HeLa cells. We used LysoTracker Red to concurrently visualize lysosomes with FGAMS self-assemblies in AICAR-treated HeLa cells. Dual-color imaging revealed that the AICAR-induced FGAMS-EGFP clusters did not colocalize with lysosomes (Supporting Information Figure S6A–C). Second, we investigated if AICAR-induced FGAMS-GFP clusters would associate with mitochondria or any other lipid-bound organelles. Cellular staining of mitochondria and lipid-bound organelles confirmed that the self-assemblies of FGAMS-EGFP did not colocalize with these organelles under our conditions (Supporting Information Figure S6D–I). Third, we sought to determine if the observed FGAMS-EGFP clusters were associated with autophagosomes. ACS Chem Biol. Author manuscript; available in PMC 2017 November 07.

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The activation of AMPK is also known to subsequently activate autophagy in cells.21,22 To visualize endogenous autophagosomes, the membrane-associated protein LC3 was targeted in immunocytochemistry.21 That is, fixed HeLa cells displaying AICAR-induced FGAMSEGFP assemblies were treated with an LC3-specific antibody, with the result showing that LC3 and the FGAMS-GFP self-assemblies did not colocalize (Supporting Information Figure S6J–L). Collectively, the FGAMS self-assembly is an independent cellular granule of the organelles and other cellular bodies, including the purinosome. Metabolic Effect of AICAR Treatment on Purine Nucleotide Pools

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In order to assess if the self-assemblies of FGAMS-EGFP result in any alterations in purine nucleotide pools, high performance liquid chromatography (HPLC) was employed to measure the steady-state levels of purine metabolites. Lawns of HeLa cells were incubated for 4 h with 200 μM AICAR, and subsequently cellular nucleotides were extracted (as described in Methods). Our data revealed that the cellular levels of various nucleotides apparently changed in response to AICAR treatment. According to our peak assignment using commercially available standards of purine intermediates, we monitored the decreased levels of IMP, AMP, GMP, and ATP (Figure 6). Quantitatively, IMP, AMP, GMP, and ATP were decreased ~19 ± 12%, ~44 ± 19%, ~80 ± 5%, and ~13 ± 1%, respectively, after 4 h of AICAR treatment. Subsequent statistical analysis using one-sample t-test with one-tailed rejection region23 showed that the cellular levels of purine metabolites are significantly reduced in the presence of AICAR. These data strongly indicate that pharmacological activation of AMPK downregulates de novo purine biosynthesis. Collectively, we propose that activated AMPK spatially sequesters FGAMS from the other pathway enzymes to downregulate de novo purine biosynthesis in HeLa cells.

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DISCUSSION A metabolic intermediate of purine biosynthesis, ZMP, is a well-characterized allosteric activator of AMPK, which is responsible for maintaining cellular energy homeostasis.10 However, AMPK’s action on de novo purine biosynthesis has not been explored in detail and largely remains elusive. In this work, we investigated the biochemical and cellular relationship between AMPK and de novo purine biosynthesis in the presence of small molecules and shRNAs, respectively. Our inquiry has unveiled an unanticipated mechanism by which de novo purine biosynthesis is downregulated in a cell. We propose that the downregulation of the pathway is achieved through the spatial sequestration of FGAMS into its own self-assemblies from the rest of the pathway enzymes.

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In addition, we demonstrate three different strategies to distinguish the self-assembly of FGAMS from the purinosome. First, examining all the enzyme components involved in de novo purine biosynthesis provides the most compelling evidence for determining whether the purinosome or the FGAMS self-assembly is involved in given cellular mechanisms. Second, measuring the diffusion coefficient of FGAMS-EGFP dictates the nature of the subcellular protein compartments where FGAMS is localized. Due to the slower diffusion kinetics of FGAMS-EGFP inside the self-assembly, we demonstrate FRAP as a biophysical technique to distinguish these two assemblies in live cells. In this regard, we note that, due to


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different transcriptional and translational profiles in different cell lines,24 it was not surprising that the diffusion coefficients of FGAMS-EGFP are not only assembly-dependent but also cell-line-dependent (this work versus ref 2). Third, profiling metabolic changes of purine metabolites indicates whether the observed compartments in single cells are purinosomes or FGAMS self-assemblies. Purinosome formation was associated with the increased level of IMP,4 whereas we showed here that the formation of FGAMS selfassemblies decreased the steady-state levels of purine metabolites of IMP, AMP, GMP, and ATP (Figure 6). HPLC-assisted metabolic profiling of cellular nucleotide pools was sensitive enough to demonstrate the downregulation of de novo purine biosynthesis. Therefore, given that the spatial compartmentalization of FGAMS under fluorescence imaging could indicate either upregulation or downregulation of de novo purine biosynthesis, it appears critical to corroborate the activity of de novo purine biosynthesis with alternative strategies beyond cellular imaging.


It is also important to discuss here the authenticity of the FGAMS self-assembly in cells. Our FRAP measurement showed the mobility of FGAMS-EGFP when it is involved in the self-assembly. It means that this assembly is not formed by insoluble protein aggregates and is different from protein aggresomes,25,26 which previously showed no fluorescence recovery during our FRAP studies.2 In addition, the FGAMS self-assembly is spatially different from membrane-bound organelles, including lysosomes, mitochondria, and autophagosomes, the latter of which might be activated by AMPK under our conditions. Furthermore, the AICAR-induced clustering of endogenous FGAMS was also observed in fixed HeLa cells by immunocytochemistry. Last, self-assembled FGAMS, whose enzymatic activity remained intact, was observed in vitro several decades ago by Buchanan and coworkers,27 although the observation has not been followed up on. Taken together, we conclude that the FGAMS self-assembly is an independent, bona fide phenomenon in the cell.

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Additionally, such self-assemblies have been observed with other metabolic enzymes involved in nucleotide biosynthesis in various organisms. For instance, filament, rod, and/or ring structures were formed by cytidine triphosphate synthase, glutamate dehydrogenase, and/or inosine-5′-monophosphate dehydrogenase 2 in various organisms, including human cells.28–30 Although it is not clear yet what the biological significance of such novel cellular self-assemblies is, their metabolic activities and nonmetabolic moonlighting functions have been progressively elucidated.28–30 Of particular note, this phenomenon appears to be widespread among yeast metabolic enzymes.31–34 Although we do not know whether FGAMS is enzymatically active or inactive inside the self-assemblies under our conditions, the spatial sequestration of FGAMS from the other pathway enzymes metabolically downregulates the activity of de novo purine biosynthesis in HeLa cells. Collectively, the sequential metabolic enzymes in de novo purine biosynthesis are shown to form reversible multienzyme metabolic compartments, purinosomes, to upregulate the pathway in human cells.35 Now, we reveal an AMPK-mediated downregulation mechanism of de novo purine biosynthesis, which agrees with recent studies6,7 showing the functional relationship between a mechanistic target of rapamycin and de novo purine biosynthesis. We


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conclude here that human de novo purine biosynthesis can be turned on and off through the spatiotemporal rearrangement of the enzymes in cells.

METHODS Materials

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Plasmids expressing de novo purine biosynthetic enzymes conjugated with monomeric enhanced green fluorescent protein (EGFP) or monomeric orange fluorescent protein (OFP) were mostly prepared previously.1 Note that EGFP in this article contains three point mutations, A206K, L221K, and F223R, which prevent potential GFP-mediated oligomerization of tagged proteins.36 We also prepared a new plasmid expressing EGFPATIC using the pEGFP-C1 vector (Clontech) to replace the previous clone of GFP-ATIC,1 which was constructed to express GFPS65T-fusion proteins from the pC1-Neo vector (Promega). The cloned plasmid in this work was confirmed by restriction enzyme digestions and DNA sequencing (GeneWiz). The plasmids expressing shRNAAMPK (cat# sc-45312SH) and shRNAControl (cat# sc-108060) were purchased from Santa Cruz. 5-Aminoimidazole-4-carboxamide ribonucleoside (AICAR) and ribonucleotide (ZMP) were obtained from EMD Biosciences. Metformin, phenformin, IMP, AMP, and GMP were obtained from Sigma-Aldrich. A769662 was obtained from Santa Cruz Biotechnology. ATP was obtained from Invitrogen. Cellular staining reagents, including LysoTracker Red DND-99 (cat# L7528), 1,1′-dihexadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (DiIC16(3); cat# D384), and MitroTrackerOrange CMTMRos (cat# M7510), were obtained from Molecular Probes (Life Technologies).

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Cell Culture Human cervix adenocarcinoma cell line, HeLa, was obtained from the American Type Culture Collection. HeLa cells were maintained in either a purine-rich medium (Minimum Essential Medium (MEM, Mediatech cat# 10-010-CV) supplemented with 10% fetal bovine serum (FBS, Atlanta Biological) and 50 ug/mL gentamycin sulfate (Sigma)) or a purinedepleted medium (Roswell Park Memorial Institute 1640 (RPMI 1640, Mediatech Cat# 10-040-CV) supplemented with 10% dialyzed FBS (dFBS, Atlanta Biological) and 50 ug/mL gentamycin sulfate).2 Cells were kept at 37 °C, 95% humidity, and 5% CO2 in a HeraCell CO2 incubator. To assess mycoplasma status, the cells were maintained under the growth media supplemented with Plasmocin Prophylactic (InvivoGen) at a concentration of 5 μg/mL for at least four passages prior to experiments.

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Transfection Cells were plated in an antibiotic-free growth medium to either glass-bottomed 35 mm Petri dishes (MatTek) or eight-well chambers (LabTek). The following day, transfection was carried out using Lipofectamine 2000 or Lipofectamine 3000 (Invitrogen) following the manufacture’s recommendation. After incubation with the lipofectamine–DNA complex for ~5 h, the medium was replaced with a fresh growth medium and incubated overnight at 37 °C, 95% humidity, and 5% CO2. Prior to imaging, the transfected cells were washed for 10 min three times with buffered saline solution (20 mM HEPES (pH 7.4), 135 mM NaCl, 5


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mM KCl, 1 mM MgCl, 2.5 or 5.6 mM glucose, and 1 mg mL−1 bovine serum albumin (Fisher Scientific)) and allowed to incubate in the dark at ambient temperature for at least 1 h. Cells were then imaged in the buffered saline solution. Fluorescence Live-Cell Imaging

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All images were obtained using a Nikon Eclipse Ti fluorescence microscope equipped with a 60× 1.45 numerical aperture oil objective (Nikon CFI PlanApoTIRF). Images were collected using either a Photometric CoolSNAP EZ monochrome CCD camera or photomultipliers mounted to a Nikon Eclipse Ti C2 confocal microscope. For wide-field imaging, EGFP was imaged with a set of Z488/10-HC cleanup, HC TIRF Dichroic, and 525/50-HC emission filter (Chroma Technology Corp). OFP, Cy3, LysoTracker Red, MitoTracker Red, and DiI Stain were imaged with a set of Z561/10-HC cleanup, HC TIRF Dichroic, and 600/50-HC emission filter (Chroma Technology Corp). For confocal imaging, a JDSU argon ion 488 laser line with a 488/561 dichroic mirror and 525/50 emission filter was used for EGFP detection. Data were obtained using NIS-Elements AR 4.13.04 (Nikon). Images were analyzed using the ImageJ processing package (National Institutes of Health). Immunocytochemistry

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Cells were fixed using 3% formaldehyde for 10 min at RT and permeabilized using 0.2% Triton-X in PBS for 5 min at RT. Cells were then washed and incubated with 5% normal donkey serum in PBS (Jackson ImmunoResearch) for 30 min at RT. After washing, cells were incubated with primary antibody (i.e., LC3A/B XP rabbit mAb (1:100, cell signaling), rabbit anti-FGAMS Ab (1:1000, Bethyl Laboratories; cat# A304–218A), rabbit AMPKα (D5A2) mAb (1:100, cell signaling; cat# 5831), and rabbit phospho-Thr172 AMPKα (40H9) mAb (1:100, cell signaling, cat# 2535)) for 30 min at RT, followed by Cy3conjugated Goat anti-Rabbit secondary antibody (1:500, Jackson ImmunoResearch) for 30 min. The fixed cells were then imaged in PBS. Fluorescence Recovery After Photobleaching (FRAP) FRAP was performed using the instrumentation outlined above. For photobleaching, the argon ion 488 nm laser was used at 75% power for 1 s. Before bleaching, at least 10 images were obtained, and postbleaching images were obtained approximately every second for at least 100 s. Individual fluorescence recovery was normalized and corrected for background photobleaching and fitted as we described previously.2 Briefly, fluorescence recovery was individually fitted using the following equation after the degree of background photobleaching was normalized:

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where τ1/2 is a diffusion time constant and t is the time (s). Apparent diffusion coefficients (Dapp) were then calculated using the following equation:


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where re is a measured radius (μm) of a photobleached area. Statistical Analysis of FRAP Data

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We obtained the apparent diffusion coefficients, Dapp1 and Dapp2, of FGAMS-EGFP involved in AICAR-induced fluorescent clusters or purinosomes, respectively, in HeLa cells. Our statistical analysis was carried out with a null hypothesis that the means of the apparent diffusion coefficients, Dapp1 and Dapp2, are the same between the two groups. We examined the equality of the apparent diffusion coefficients using three test procedures: one parametric test and two nonparametric tests. Since the equality of variances are violated based on the Ftest,37 the Welch’s t test38 was conducted as the two-sample t test, which is the most common testing procedure for the comparison of two means values. In addition, two nonparametric tests were applied as well because, with the obtained data showing nonnormal distributions, permutation test and Wilcoxon rank sum test procedures39 would be more appropriate without any specific distributional assumption than Welch’s t test. Cluster Size Analysis

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Cluster size analysis was accomplished using the ImageJ processing software (the National Institutes of Health). Fluorescent images were processed using ImageJ built-in modules. Briefly, raw 12-bit images were scaled according to the resolution of the microscope (i.e., 0.12 μm/pixel) before the Robust Automatic Threshold Selection (RATS) segmentation tool was used to identify fluorescent clusters within a cell by outlining. Default parameters for RATS were used in this analysis (i.e., noise threshold =25, λ factor =3). Once fluorescent clusters were isolated, the Inverse Look-up Table function was used to generate a mask of the original image that only displayed fluorescent clusters. The Analyze Particles module was then applied to this mask in order to attain both the number and area of fluorescent clusters within an image. This process was repeated for all subsequent cell images. The operator then evaluated the original cell images against the particle mask in order to eliminate data in which more than one cluster was counted as a single particle. The data were then analyzed and graphed using Microsoft Excel and Igor Pro (Wavemetrics). Nucleotide Extraction

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Lawns of HeLa cells were harvested using 0.25% Trypsin/2.21 mM EDTA (Corning, cat# MT-25-053-CI), and washed using DPBS (Mediatech, Cat# MT-21-031-CV). Nucleotides were then extracted using 10% perchloric acid followed by neutralization with 1 M potassium hydroxide.40 Last, the cell extracts were filtered through a 0.45 μm syringe filter and either used immediately or kept at −20 to −80 °C.15 High Performance Liquid Chromatography (HPLC) HPLC was performed using a Partisil 10 SAX column (Hichrom) and PerkinElmer Flexar HPLC equipped with a Quaternary LC pump, ultra high performance liquid chromatography


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detector, Peltier LC column oven, and Peltier LC autosampler. The following elution buffers were used to separate nucleotides from cell lysates; i.e., buffer A, 7 mM potassium phosphate dibasic in 7 mM potassium chloride (pH 3.5) and buffer B, 250 mM potassium phosphate dibasic in 500 mM potassium chloride (pH 4.5).40 Nucleotides were eluted using a flow rate of 1.0 mL/min with a 5 min equilibration in 100% buffer A, 30 min gradient to buffer B, and maintenance of 100% buffer B for an additional 40 min. The nucleotides were detected by monitoring absorbance at 260 nm. Retention times of nucleotides were determined using commercially available standards. Data collection and peak analysis were performed using Chromera 4.1 (PerkinElmer), Igor Pro (Wavemetrics), and Microsoft Excel.

Supplementary Material Refer to Web version on PubMed Central for supplementary material.

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Acknowledgments We thank UMBC for providing the Start-up fund to S.A. and also for the SRAIS funding (to S.A.) for the presented study. D.L.S. was partially supported by NIH/NIGMS Training Grant (T32GM066706). We thank R. Karpel (UMBC) for critically reading the manuscript. We also thank G. Balaa, J. Wilhide, C. Kohnhorst, A. Sundaram, J. Ramirez, F. Augustine, D. Jaiswal, M. Kyoung, and E. Green for their assistance on data analysis.

References


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Figure 1.

Effect of AMPK activators on FGAMS-EGFP localization. Transiently expressed FGAMSEGFP in HeLa cells formed clusters after the addition of 150–300 μM AICAR (A and B), 170–430 μM metformin (C and D), 210–410 μM phenformin (E and F), and 100 μM A769662 (G and H) for 4 h. However, transiently expressed HPRT1-EGFP did not form clusters when incubated with 250 μM AICAR (I and J) or 290 μM metformin (K and L) for 4 h. Scale bar, 10 μm.


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Figure 2.

No impact on subcellular localization of other purine biosynthetic enzymes in the presence of AICAR. Transiently expressed PPAT-EGFP (A and B), TrifGART-GFP (C and D), PAICS-EGFP (E and F), ASL-EGFP (G and H), and EGFP-ATIC (I and J) in HeLa cells remained diffusive in the cytoplasm after incubation with 200 μM AICAR for 4 h. Scale bar, 10 μm.


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Author Manuscript Author Manuscript Figure 3.

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AMPK activation on HeLa cells coexpressing FGAMS-OFP and EGFP-ATIC. Treatment of AICAR (200 μM; A–F) or metformin (290 μM; G–L) on dually transfected HeLa cells promoted the clustering of FGAMS-OFP, but EGFP-ATIC remained diffusive. Scale bar, 10 μm.

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Effect of AMPK knock-down on the AICAR-induced clustering efficiency of FGAMSEGFP in HeLa cells. Dually transfected HeLa cells expressing shRNAAMPK or shRNAControl with FGAMS-EGFP were treated with 200 μM AICAR for 4 h. Clustering efficiencies (%) were calculated by dividing the number of cells showing AICAR-induced fluorescent clusters by the total number of cells counted for analysis. The error bars indicate the standard deviations of at least five independent experiments. *Statistical analysis using two-sample t-test reveals p < 0.01, supporting the significant difference between the clustering efficiencies of FGAMS-EGFP in the presence of shRNAAMPK and shRNAControl.


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Diffusion coefficients of FGAMS-EGFP involved in two different granules in HeLa cells. The graph plots the distribution of diffusion coefficients of FGAMS-EGFP involved in the AICAR-induced self-assemblies and the purinosomes in HeLa cells. Statistical analysis using Welch’s t test, a permutation test, and Wilcoxon rank sum test showed p < 0.0001, indicating significant difference between two diffusion coefficients.


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HPLC analysis of purine metabolites in cell lysates. Normalized peak areas of purine metabolites of AMP, IMP, GMP, and ATP from HeLa cells are shown before (gray) and after (black) 4-h incubation with 200 μM AICAR. The error bars indicate the standard deviations of three independent experiments. Statistical analysis using one-sample t-test with one-tailed rejection region23 indicates p < 0.01 for GMP and ATP, p < 0.05 for AMP, and p = 0.06 for IMP. Alternatively, the analysis of variance (ANOVA) indicates p = 0.0141. Note that HPLC chromatograms were corrected for background and the number of cells prior to the peak area integration. The resolved peaks were assigned by commercially available standards.


AMPK Activation via Modulation of De Novo Purine Biosynthesis with an Inhibitor of ATIC Homodimerization.

Some pyrazoles as inhibitors of purine biosynthesis de novo.

Modulation of de novo purine biosynthesis leads to activation of AMPK and results in improved glucose handling and insulin sensitivity.

Quantitative analysis of purine nucleotides indicates that purinosomes increase de novo purine biosynthesis.

Induction of HL-60 leukemia cell differentiation by tetrahydrofolate inhibitors of de novo purine nucleotide biosynthesis.

Stimulation of human purine synthesis de novo by fructose infusion.

Metal stopping reagents facilitate discontinuous activity assays of the de novo purine biosynthesis enzyme PurE.

Regulation of de novo purine biosynthesis in normal and 8-azaguanine-resistant Chinese hamster cells.

G-protein-coupled receptor regulation of de novo purine biosynthesis: a novel druggable mechanism.

De novo purine ribonucleotide biosynthesis in adult Angiostrongylus cantonensis (Nematoda: Metastrongyloidea).

Some regulatory properties of purine biosynthesis de novo in long-term cultures of epithelial-like rat liver cells.

Cloning of three human multifunctional de novo purine biosynthetic genes by functional complementation of yeast mutations.

De Novo Guanine Biosynthesis but Not the Riboswitch-Regulated Purine Salvage Pathway Is Required for Staphylococcus aureus Infection In Vivo.

Evidence in support of biosynthesis de novo of free cytokinins.

De novo biosynthesis of Gastrodin in Escherichia coli.

Sphingolipid De Novo Biosynthesis: A Rheostat of Cardiovascular Homeostasis.

MTHFD1 regulates nuclear de novo thymidylate biosynthesis and genome stability.

S1P Signaling and De Novo Biosynthesis in Blood Pressure Homeostasis.

Endogenous auxin biosynthesis and de novo root organogenesis.

Regulation of de novo purine synthesis in chick liver slices. Role of phosphoribosylpyrophosphate availability and of salvage purine nucleotide synthesis.

Regulation of de novo sphingolipid biosynthesis by the ORMDL proteins and sphingosine kinase-1.

Membrane lipid biosynthesis in Acholeplasma laidlawii B: de novo biosynthesis of saturated fatty acids by growing cells.

The effects of added purines on urate and purine synthesis de novo by isolated chick liver, kidney and lymphoid cells.

Downregulation of de Novo Fatty Acid Synthesis in Subcutaneous Adipose Tissue of Moderately Obese Women.