Longer life from reduced insulin signaling may require glycine N-methyltransferase to produce spermidine

In mammals, Gnmt is a key regulator of the Met cycle and is mostly expressed in the liver (Uhlén et al., 2015). In mice, expression of Gnmt declines with age (Armstrong, Rakoczy, Rojanathammanee, & Brown‐Borg, 2014). To determine the tissue‐specific expression of Gnmt in Drosophila and its response to reduced IIS, we performed Western blot analysis to quantify Gnmt protein expression in flies lacking 3 of the 7 insulin‐like peptides, dilp2‐3,5 mutant flies (Grönke, Clarke, Broughton, Andrews, & Partridge, 2010). In control flies (w^Dah), Gnmt protein was detected in total fly and fat body protein extracts (Figure 1), but not in the thorax or the gut (Figure S1). Gnmt levels were substantially increased in the whole fly and the fat body of dilp2‐3,5 mutant flies (Figure 1), but again were not detected in the thorax or the gut (Figure S1). This suggests that increased expression of Gnmt in the fat body may play a role in modulating the phenotypes induced by reduced IIS, possibly including longevity.

To determine whether increasing the level of Gnmt in the fat body was sufficient to extend lifespan, we over‐expressed Gnmt using a UAS‐Gnmt construct (Obata & Miura, 2015) and two independent, constitutive, fat body drivers, Fat Body‐Gal4 (FB‐Gal4) (Grönke et al.,2003) and Pumpless‐Gal4 (Zinke, Kirchner, Chao, Tetzlaff, & Pankratz, 1999). Both FB‐Gal4 and Pumpless‐Gal4 drivers led to increased Gnmt transcript levels in the fat body (Figure S2a,b) and extended lifespan, by 10% and 8%, respectively, compared to controls (Figure 2a,b). To prevent possible developmental effects of Gnmt expression, UAS‐Gnmt was then expressed using the adult‐specific S106GS GeneSwitch driver line (Osterwalder, Yoon, White, & Keshishian, 2001), which is expressed in fat body and gut. Inducing expression of UAS‐Gnmt with the S106GS driver increased Gnmt levels ~ 3‐fold in the fat body compared to uninduced controls (Figure S2d) and significantly extended lifespan by 9% (Figure 2c, Figure S2e). To determine whether the over‐expression of Gnmt in the gut played a role in the extension of lifespan, we drove expression only in the gut using the constitutive, gut‐specific driver NP1‐Gal4 (Jiang et al., 2009). Using NP1‐Gal4, the gut‐specific expression of Gnmt was comparable to that of fat body‐specific over‐expression (Figure S2c); however, lifespan in these flies was unchanged (Figure S2f). These findings indicate that increased Gnmt protein, specifically in the fat body, can extend lifespan.

To determine whether Gnmt is required for increased lifespan in response to lowered IIS, we measured lifespan of the long‐lived dilp2‐3,5 mutants (Grönke et al., 2010) in the presence and absence of Gnmt. Loss of Gnmt (Gnmt^Mi04290, hereafter referred to as Gnmt^Mi) did not reduce lifespan of wild‐type flies (Figure 3), indicating that Gnmt is not essential for normal lifespan. However, the extended lifespan of dilp2‐3,5 mutants was significantly diminished in the absence of Gnmt (Gnmt^Mi,dilp2‐3,5), with the double mutants living 33% less long than the dilp2‐3,5 mutants alone (Figure 3). Gnmt is thus necessary for the full extension of lifespan of dilp2‐3,5 mutants.

Gnmt is a key enzyme in methionine metabolism, catabolizing SAM into SAH. To determine the molecular mechanism underlying the prolongevity role of Gnmt in response to lowered IIS, we focused on Met and its associated metabolic pathways. These include the transsulfuration pathway (TSP), oxidative and xenobiotic detoxification pathway (glutathione metabolism), and the polyamine pathway (Lu & Mato, 2012; Minois, Carmona‐Gutierrez, & Madeo, 2011) (Figure 4a).

Ubiquitous, or fat body‐specific, over‐expression of Gnmt increases SAM catabolism (Obata & Miura, 2015), possibly converting it into sarcosine and S‐adenosyl homocysteine (SAH). SAH can then be hydrolyzed to homocysteine by SAH hydrolase. Homocysteine is, in turn, the primary source of cysteine for the TSP and the downstream production of glutathione (Kabil et al., 2011). Therefore, increasing Gnmt expression may elevate SAH, cysteine, and glutathione levels and protect against oxidative stress. To test this idea, we investigated whether over‐expression of GNMT in the fat body was sufficient to confer resistance to oxidative stress, induced by hydrogen peroxide (H₂O₂). We found no change in the survival of flies over‐expressing Gnmt using FBGal4, pumplessGal4, or S106GS drivers, suggesting that resistance to oxidative stress is not causal for the longevity of these flies (Figure S3a,b). This is concordant with the finding that oxidative stress resistance may not be causal for the longevity of IIS mutants (Afschar et al., 2016; Slack, Giannakou, Foley, Goss, & Partridge, 2011). Furthermore, S106GS > UAS‐Gnmt flies did not show increased resistance to the xenobiotic dichlorodiphenyltrichloroethane (DDT), suggesting that resistance to xenobiotics is also not causal for lifespan extension in these flies (Figure S3c).

TSP activity leads to the release of hydrogen sulfide (H₂S) gas (Figure 4a), exposure to which can increase the lifespan of C. elegans (Miller & Roth, 2007). More recently, H₂S and TSP activity have been shown to correlate with the effects of dietary restriction, including extension of lifespan, in yeast, worm, fruit fly, and rodent models (Hine et al., 2015). The TSP could, therefore, mediate longevity of Gnmt‐expressing flies, through H₂S production. To test this idea, we analyzed H₂S production capacity as a readout of TSP activity, but could not detect any change in H₂S levels in FBGal4 > UAS‐Gnmt, pumplessGal4 > UAS‐Gnmt, or S106GS > UAS‐Gnmt flies (Figure S3d,e). Together, these findings suggest that neither the TSP nor glutathione production branches of Met metabolism underlie the longevity of Gnmt‐over‐expressing flies.

Met metabolism also branches, through SAM, into the polyamine pathway (Pegg, 2009) (Figure 4a). SAM is decarboxylated by SAM decarboxylase (SamDC) and combines with putrescine to form spermidine, or with spermidine to form spermine via spermidine synthase (SpdS) (Pegg, 2009). In rodents and humans, spermidine levels decrease with age (Das & Kanungo, 1982; Vivó et al., 2001). Furthermore, dietary supplementation with spermidine is sufficient to extend the longevity of yeast, worms, flies, and mice (Eisenberg et al., 2016, 2009). If changes in the polyamine pathway underlie IIS‐mediated lifespan extension, we would expect either the expression or the activity of polyamine synthesis enzymes to be elevated in dilp2‐3,5 mutants. However, expression of the two rate‐limiting enzymes in polyamine synthesis, SamDC and Odc1, was not changed in dilp2‐3,5 mutant flies (Figure S4a).

To determine whether polyamine synthesis enzyme activity underlies Gnmt‐mediated dilp2‐3,5 mutant longevity, we quantified key metabolites of the methionine cycle (Met, SAM, and SAH) and of polyamine synthesis (spermidine, ornithine, and putrescine) (Figure 4a) in wild‐type and dilp2‐3,5 mutant flies in the presence and absence of Gnmt (Gnmt^Mi and Gnmt^Mi,dilp2‐3,5 double mutants). Two metabolites of the Met cycle, Met and SAH, were significantly increased in response to reduced IIS, and the level of SAH, but not Met, was dependent on the presence of Gnmt (Figure 4b). Interestingly, spermidine levels were significantly increased in dilp2‐3,5 mutant flies compared to controls (Figure 4b), but TSP and glutathione pathway metabolites (cysteine, cystathionine, γ‐Glu‐Cys, and glutathione) were unchanged (Figure S4b). Furthermore, the IIS‐mediated increase in spermidine levels required Gnmt (Figure 4b). These findings suggest that reduced IIS activity can modulate methionine metabolism and polyamine synthesis, but not the TSP or glutathione, through Gnmt (Figure 4b). This supports our previous conclusion that the beneficial role of Gnmt in response to reduced IIS does not occur through the TSP or glutathione pathways, but rather through spermidine synthesis.

To confirm the role of Gnmt and spermidine in IIS‐mediated changes to Met metabolism, we treated wild‐type and dilp2‐3,5 mutant flies with spermidine in the presence and absence of Gnmt (Gnmt^Mi and Gnmt^Mi,dilp2‐3,5 double mutants) (1mM) and quantified associated metabolites (Figure 4b). Treatment with spermidine led to restricted changes in metabolism, changing only SAM and putrescine levels in Gnmt^MI and dilp2‐3,5 mutant flies, respectively, and spermidine levels in wild‐type (wDah) and dilp2‐3,5; gnmt double mutant flies (Figure 4b). However, two‐way ANOVA revealed a significant interaction between spermidine treatment and genotype, dilp2‐3,5 mutants showed no increase in spermidine levels, as these mutants maintain elevated spermidine levels, independently of spermidine supplementation (Figure 4b). Interestingly, the loss of elevated spermidine levels in dilp2‐3,5 mutants in the absence of Gnmt was completely abrogated by treatment with spermidine (Figure 4b). Together, these results show that reduced IIS increases spermidine synthesis and that this increase requires Gnmt.

To determine the role of Gnmt in spermidine production and IIS mutant longevity, we assayed the lifespan of wild‐type, Gnmt^Mi and dilp2‐3,5 single mutant, and Gnmt^Mi,dilp2‐3,5 double mutant flies with or without spermidine supplementation. Spermidine increased the lifespan of wild‐type flies, but not of either Gnmt^Mi or dilp2‐3,5 mutants (Figure 4c), suggesting that the elevated spermidine level in these mutants was already sufficient to maximize their lifespan by this mechanism. Gnmt also appeared to exert its effects on lifespan through spermidine, because the reduction of the lifespan of the dilp2‐3,5 flies by the Gnmt^Mi mutant was completely abrogated when the double mutants were fed spermidine (Figure 4c). In accordance with our metabolic analysis (Figure 4b), the lifespan analysis indicated that spermidine regulates processes that play a major role in IIS‐mediated longevity.

Reductions in IIS can induce autophagy, and this induction is essential for lifespan extension in IIS mutants in C. elegans (Meléndez et al., 2003). Spermidine can induce autophagy in yeast, worms, flies, and mice (Eisenberg et al., 2009). To determine whether Gnmt plays a role in autophagy induction in response to reduced IIS, we investigated expression of autophagy associated genes in the fat bodies of wild‐type (wDah), Gnmt^Mi and dilp2‐3,5 single mutants, and Gnmt^Mi,dilp2‐3,5 double mutants fed either spermidine (1mM) or control food. Expression of atg5 and atg8a was induced in the fat bodies of dilp2‐3,5 mutants, but not in Gnmt^Mi or Gnmt^Mi,dilp2‐3,5 double mutant flies compared to controls (Figure 5a). Induction of both atg5 and atg8a is thus directly, or indirectly, regulated through Gnmt, in response to reduced IIS. Furthermore, spermidine treatment led to induction of both atg5 and atg8a in wild‐type flies suggesting, in agreement with previous studies (Eisenberg et al., 2009), that spermidine treatment induces autophagy, but it did so only in the presence of Gnmt (Figure 5a). Spermidine treatment did not increase the level of atg5 and atg8a expression beyond that of control fed dilp2‐3,5 (Figure 5a). However, in dilp2‐3,5 mutants lacking Gnmt, spermidine treatment increased atg5, but not atg8a, expression (Figure 5a). Together, the induction of atg5 and 8a expression suggests that autophagy is induced in response to reduced IIS and that induction is dependent on Gnmt. Furthermore, in the case of atg5, spermidine treatment is sufficient to recapitulate the induction of autophagy seen in IIS mutants, even in the absence of Gnmt.

We then used Western blot analysis to quantify the level of p62, a substrate of autophagy. Increased levels of p62 suggest autophagy is blocked or slowed, while decreased levels suggest increased autophagy (Nagy, Varga, Kovács, Takáts, & Juhasz, 2015). The level of p62 was significantly decreased in the fat body of dilp2‐3,5 mutants, Gnmt^Mi mutants, and Gnmt^Mi,dilp2‐3,5 double mutant flies compared to controls (Figure 5b). Treatment with spermidine significantly reduced p62 levels in the fat body of wild‐type flies, but not in the fat body of dilp2‐3,5 mutants, Gnmt^Mi mutants, or Gnmt^Mi,dilp2‐3,5 double mutant flies, when compared to untreated control flies (Figure 5b). Thus, autophagy in the fat body is activated in response to reduced IIS or spermidine treatment. Surprisingly, p62 levels are also reduced in the fat body of Gnmt^Mi flies and Gnmt^Mi,dilp2‐3,5 double mutant flies. This suggests that Gnmt may also play a role in autophagy activation in response to spermidine treatment.

To determine whether the responses to reduced IIS and spermidine treatment led to functional changes in autophagy, we quantified the level of autophagy using LysoTracker Red (LTR) in the fat bodies of control and dilp^2‐3,5 mutant flies in the presence and absence of Gnmt and with and without spermidine treatment. LTR accumulates inside acidic vesicles allowing direct assessment of autophagic status of a tissue. The fat body of dilp^2‐3,5 mutant flies showed significantly higher numbers of LTR‐stained vesicles per cell compared to controls flies, suggesting increased levels of autophagy (Figure 5b,c). This suggests reducing IIS can increase autophagy in the fat body. Importantly, the observed increase in autophagy in dilp2‐3,5 mutants did not occur in the absence of Gnmt, and Gnmt^Mi mutants show no changes in autophagic status compared to controls (Figure 5b,c). This suggests that, either directly or indirectly, Gnmt modulates autophagy specifically in IIS mutant flies.

The fat body of wild‐type flies fed with spermidine (1mM) showed a significant increase in autophagy compared to untreated control flies, increasing to the level of untreated dilp2‐3,5 mutants (Figure 5b,c). Spermidine treatment did not further enhance autophagy in dilp2‐3,5 mutants (Figure 5b,c). Furthermore, supporting our analysis of atg5 and atg8a expression (Figure 5a), Gnmt^MI flies did not increase the level of autophagy in response to spermidine treatment (Figure 5b,c). However, treatment of dilp2‐3,5 mutants, in the absence of Gnmt, with spermidine increased autophagy activation, but not significantly above the level in controls (Figure 5b,c). Thus, Gnmt plays a role in modulating autophagy in response to reduced IIS, and spermidine may be a component within that response.

Gnmt is an evolutionarily conserved regulator of methionine metabolism, performing the same function in both flies and mice (Obata et al., 2014). To determine whether increased expression of Gnmt in response to reduced IIS is also conserved between mice and flies, we examined Gnmt levels in the functional equivalent of the fly fat body, the liver, of liver‐specific IRS1 KO (Alfp‐Cre::IRS1fl/fl) mice (Essers et al., 2016). We confirmed by Western blot that, as in flies, reducing IIS activity resulted in significantly increased levels of Gnmt in the liver (Figure 6a). To determine whether the increased expression of Gnmt led to conserved metabolic changes, we then quantified the spermidine in the liver of liver‐specific IRS1 knockout mice (Figure 6b). In agreement with our previous analysis in IIS mutant flies, liver‐specific loss of IRS1 increased levels of spermidine (Figure 6b). Together, these results suggest that the response of increasing Gnmt expression is a conserved response to reduced IIS activity and that this response may be mechanistically similar, acting through spermidine.

Flies were maintained in a controlled temperature environment of 25°C, 65% humidity with a 12‐hr day–night light cycle, and fed standard sugar/yeast/agar (SYA) diet (Bass et al., 2007). Larval densities were controlled, and adults were once mated. Female flies were sorted under CO₂ at 10 flies/vial. Lifespan analysis was performed using SYA food or SYA food containing 200μM of RU‐486, 1mM spermidine, or vehicle (EtOH) control food, and survival was scored every 2–3 days.

Food containing RU486 (200 μM) and/or spermidine (1 mM) was prepared every two weeks from a 100 mM (in ETOH) or a 1 M (in ddH2O) stock solution, respectively.

RU486, spermidine, or vehicle control supplementation commenced 48 hr posteclosion. Food vials were changed at 2‐ to 3‐day intervals. Lifespan data used to generate survival curves are summarized in Supporting Information Table S1. All flies were backcrossed for 10 generations into the white Dahomey (w^Dah) background. The genotypes used were w^Dah, dilp2‐3,5 mutant (Grönke et al., 2010), Gnmt^Mi04290 mutant (Venken et al., 2011), and Gnmt^Mi04290 mutant in a dilp2‐3,5 background (Gnmt^Mi04290;dilp2‐3,5). For over‐expression analysis, UAS‐Gnmt (Obata & Miura, 2015) flies were crossed with FB‐GAL4 and pumpless‐GAL4 drivers for constitutive expression of Gnmt in the fat body, and with S106GS‐GAL4 driver for adult‐onset expression.

Liver‐specific knockout of IRS1 mice was generated as described in Essers et al., 2016. Briefly, IRS1^loxP/loxP mice were crossed with Cre recombinase‐expressing mice under the control of the mouse albumin enhancer and promoter and the mouse alpha‐fetoprotein enhancers. All mice were maintained at 22°C under a 12‐hr light/dark cycle with ad libitum access to normal chow [ssniff® R/M‐H phytoestrogen‐poor (9% fat, 34% protein, 57% carbohydrate) Spezialdiäten GmbH, Soest, Germany] and water. Mice were sacrificed at 3–4 months. Livers were dissected and snap‐frozen in liquid nitrogen.

Total protein (30 μg) was loaded and separated on precast TGX gels (Any kD, Bio‐Rad) and transferred to nitrocellulose (0.45 μm) membranes (GE Healthcare). Membranes were incubated (1 hr) in blocking solution (5% nonfat milk in 0.05% TBST) at room temperature and then in primary antibody anti‐Gnmt (for detection of mouse Gnmt (1:1,000)(sc‐68871) or of fly anti‐Gnmt (1:1,000)) (a generous gift from Prof. Miura), anti‐p62 (1:5,000) (a generous gift from Prof. Juhász), and tubulin (1:10,000)(Cell Signaling Technologies) overnight at 4°C. Membranes were washed three times with TBST (0.05%) for 10 min. HRP‐conjugated anti‐mouse (1:10,000 dilution) or anti‐rabbit (1:10,000 dilution) secondary antibodies were incubated for 1 hr at room temperature. Blots were then washed three times with TBST (0.05%), and detection was performed with ECL Prime reagent (Amersham). Blots were imaged in a ChemiDoc^TM imaging station, and protein bands were quantified using Image Lab software (Bio‐Rad).

Total RNA was extracted using TRIzol (Invitrogen) to manufacturers’ guidelines and subsequently treated with DNase I (Ambion). cDNA was synthesized using SuperScript® VILO Master Mix (Invitrogen) cDNA Synthesis Kit, following the manufacturers’ protocol. Quantitative real‐time PCR (qRT–PCR) was performed using SYBR Green probes (Applied Biosystems) on a 7900HT Real‐Time PCR System (Applied Biosystems).

The primers used in the study are listed below:

For DDT and H₂O₂ stress assay, 100 female flies (20 flies/vial) were initially fed either SYA food, RU‐486 (200 μM), or ethanol control food for 7 days. Flies were then transferred into fresh vials containing 0.03% DDT mixed in SYA food for xenobiotic stress assay and 1% agar food containing 5% v/v H₂O₂ for oxidative stress assay, respectively. The number of dead flies was counted four times a day and scored in an Excel sheet.

For quantification of H₂S production, 100 female flies were sorted into vials (20 flies/vial) and fed either SYA food, RU‐486 (200 μM), or ethanol control food. Flies were transferred into fresh vials every second day. H₂S measurements were performed as previously described (Hine et al., 2015). Briefly, 20 aged (10 days) flies were chilled on ice and lysed in 300 μl lysis buffer (5× passive lysis buffer (Promega) made up to 1× lysis buffer with PBS containing cysteine (10 mM) and PLP (10 μM)). 10 μl of lysate was removed for later BCA protein quantification (Pierce) and normalization. Fly lysate was then transferred to a 96‐well plate, and individual wells were tightly covered with lead acetate paper and incubated at 37^oc (3 hr). H2S production was then assayed by densitometry analysis (ImageJ) of lead sulfide darkening and normalized to protein content.

For measurement of metabolites (Sigma), an Acquity UPLC^TM I‐Class System/Xevo^TM TQ‐S (Waters^TM) with MassLynx was used. In brief, 25 snap‐frozen female flies were homogenized in 71 μl of 15 mg/ml dithiothreitol (DTT) and 429 μl 50% methanol (total volume 500 μl) and ~35 mg of frozen mouse tissues were homogenized in 1ml sample buffer (methanol:H₂O:chloroform in 7:2:1 ratio). Homogenates were mixed for 5 min and centrifuged at 9,300 g at 4°C for 15 min. The supernatant was filtered using 0.2‐μm VWR centrifugal filters at maximum speed at 4°C for 5 min. The filtrate was then evaporated using SpeedVac for approx. 2 hr at 30°C. The pellet was reconstituted into 100 μl of running buffer (5mM ammonium formate, 0.15% formic acid aqueous solution, and 100 μg/ml DTT) and filtered again. Each metabolite was run in 1:10 and 1:100 dilutions to meet the standard run range. For absolute quantification of metabolites (Sigma) in positive ESI MRM (multireaction monitoring) mode, an Acquity UPLC^TM I‐Class System/Xevo^TM TQ‐S (Waters^TM) with MassLynx and absolute quantification TargetLynx^TM (Waters^TM) were used. With settings for capillary kv 1.5, desolvation temp. 550°C, desolvation gas flow 800 L/Hr, cone 150 L/Hr, and collision gas flow 0.15 ml/min, a Supelco^TM Discovery^TM HS F5‐3 Column from Sigma 3 µm × 2.1 mm × 100 mm was used at 25°C. Solvent A was 5 mM ammonium formate (Sigma) + 0.15% formic acid (Sigma) and B acetonitrile (VWR). A gradient from 100% A to 0% in 8 min at a flow rate of 0.35 ml/min and an equilibration step from 8.3 to 15 min were used. The following MRM transitions were used as quantifier (M + H⁺)⁺ for putrescine 89.10–30.14 m/z, l‐cysteine hydrochloride 121.92–75.93 m/z, ornithine 133.03–70.09 m/z, spermidine 146.14–72.16 m/z, l‐methionine 150.03–55.99 m/z, Cys‐Gly 178.85–75.93 m/z, l‐cystathionine 222.96–87.90 m/z, y‐Glu‐Cys 250.96–83.94 m/z, l‐glutathione reduced 308.02–75.92 m/z, SAH 385.10–136.38 m/z, and SAM 399.10–250.03 m/z. Compounds were dissolved in 5 mM ammonium formate + 0.15% formic acid + 100 µg/ml DTT. For all compounds, a calibration curve was calculated. Using concentrations from 5 to 75,000 ng/ml (prepared from stock solutions 100 µg/ml), correlation coefficient: r < 0.990; response type: external standard, the peak integrations were corrected manually, if necessary. Quality control standards of each standard were used during sample analysis and showed between 0.5% and 40% deviation, respectively. Blanks after the standards, quality control, and sample batch proved to be sufficient. Absolute value of metabolites was normalized to protein content. Relative fold change value was measured to determine the changes in amount of metabolites.

For LysoTracker staining, fat bodies from female flies (10 days) were dissected in PBS and immediately stained with LysoTracker Red DND‐99 (Invitrogen) dye (1 μM in PBS) for 2 min. Tissues were then washed three times in PBS and mounted in mounting medium (VECTASHIELD H‐1200) containing DAPI. Images were taken using confocal microscope (Leica TCS SP5X) with 40x 1.25 oil objective. Laser power and optical settings were kept constant between images. Images were then analyzed using Imaris 8.0 software. The number of LysoTracker puncti and DAPI‐stained nuclei was quantified according to user manual guidelines and puncti/nucleus calculated. All image analysis and quantification were performed under blinded conditions.

Statistical analysis was performed using GraphPad Prism software 5.0f. Individual statistical tests are mentioned in appropriate figure legends. Lifespan assays and stress survival assays were recorded using Excel, and survival was analyzed using log‐rank test.