Protein N-terminal Acetylation by the NatA Complex Is Critical for Selective Mitochondrial Degradation

Akinori Eiyama; Koji Okamoto

doi:10.1074/jbc.M115.677468

. 2015 Aug 21;290(41):25034–25044. doi: 10.1074/jbc.M115.677468

Protein N-terminal Acetylation by the NatA Complex Is Critical for Selective Mitochondrial Degradation^*

Akinori Eiyama ^1,¹, Koji Okamoto ^1,²

PMCID: PMC4599008 PMID: 26296886

Background: Mitophagy is a catabolic mechanism that degrades mitochondria selectively, and protein N-terminal acetylation is a major modification of eukaryotic proteins.

Results: Deletion of protein N-terminal acetyltransferase A (NatA) leads to suppression of mitophagy in yeast.

Conclusion: Protein N-terminal acetylation by NatA is crucial for mitophagy.

Significance: This is the first report on the link between mitophagy and protein N-terminal acetylation.

Keywords: acetyltransferase, autophagy, mitochondria, mitophagy, yeast

Abstract

Mitophagy is an evolutionarily conserved autophagy pathway that selectively degrades mitochondria. Although it is well established that this degradation system contributes to mitochondrial quality and quantity control, mechanisms underlying mitophagy remain largely unknown. Here, we report that protein N-terminal acetyltransferase A (NatA), an enzymatic complex composed of the catalytic subunit Ard1 and the adaptor subunit Nat1, is crucial for mitophagy in yeast. NatA is associated with the ribosome via Nat1 and acetylates the second amino acid residues of nascent polypeptides. Mitophagy, but not bulk autophagy, is strongly suppressed in cells lacking Ard1, Nat1, or both proteins. In addition, loss of NatA enzymatic activity causes impairment of mitochondrial degradation, suggesting that protein N-terminal acetylation by NatA is important for mitophagy. Ard1 and Nat1 mutants exhibited defects in induction of Atg32, a protein essential for mitophagy, and formation of mitochondria-specific autophagosomes. Notably, overexpression of Atg32 partially recovered mitophagy in NatA-null cells, implying that this acetyltransferase participates in mitophagy at least in part via Atg32 induction. Together, our data implicate NatA-mediated protein modification as an early regulatory step crucial for efficient mitophagy.

Introduction

Mitochondria are essential organelles that supply most of the energy for a cell. This organelle concomitantly generates reactive oxygen species during respiration. Accumulation of reactive oxygen species eventually causes mitochondrial dysfunction that negatively affects cellular integrity and is thought to induce diverse pathology (1). Additionally, cells need to adjust mitochondrial quantity to maintain a suitable balance between ATP production and consumption (2). Therefore, degradation of dysfunctional and excess mitochondria is critical for cell homeostasis. To solve this problem, cells utilize mitophagy, a catabolic system via autophagy that isolates cytosolic components with double-membrane vesicles called autophagosomes and carries them into digestive compartments such as lysosomes (vacuoles in yeast) for degradation and recycling (3, 4). Mitophagy is a selective pathway that specifically eliminates mitochondria. This process is conserved from yeast to humans and is relevant to cellular physiology (5, 6).

In the budding yeast Saccharomyces cerevisiae, the mitochondria-anchored receptor Atg32 is induced in response to oxidative stress and is localized on the surface of mitochondria (7, 8). Loss of Atg32 disrupts mitophagy, ultimately leading to mitochondrial genome instability (9). Atg32 interacts with Atg8, a ubiquitin-like protein conjugated to the phospholipid phosphatidylethanolamine and required for autophagosome formation, and with Atg11, a selective autophagy-specific scaffold protein required for assembly of core Atg proteins (7, 8). Moreover, phosphorylation of Atg32 stabilizes the Atg32-Atg11 interaction (10 –12). Although the ternary complex is important for formation of mitochondria-specific autophagosomes (mitophagosomes), how Atg32 induction and mitophagosome formation are regulated remains poorly understood.

We show here that formation of mitophagosome is blocked by loss of N-terminal acetyltransferase A (NatA) catalyzing transfer of an acetyl group from acetyl-coenzyme A (acetyl-CoA) to the α-amino group of the N-terminal amino acid residue (13, 14). In addition, Atg32 induction is partially suppressed in cells lacking NatA. Our findings suggest that NatA regulates selective mitochondrial degradation at least in part via mitophagosome formation and Atg32 expression.

Experimental Procedures

Yeast Strains and Growth Conditions

Yeast strains and plasmids used in this study are described in Tables 1 and 2, respectively. Standard genetic and molecular biology methods were used for S. cerevisiae and Escherichia coli strains. Yeast cells were incubated in YPD medium (1% yeast extract, 2% peptone, and 2% dextrose), synthetic medium (0.17% yeast nitrogen base without amino acids and ammonium sulfate, 0.5% ammonium sulfate) with 0.5% casamino acids containing 2% dextrose (SDCA), or 0.1% dextrose plus 3% glycerol (SDGlyCA), supplemented with necessary amino acids. For mitophagy and pexophagy under respiratory conditions, cells grown to mid-log phase in SDCA were transferred to SDGlyCA and incubated at 30 °C. For autophagy and mitophagy under starvation conditions, cells grown to mid-log phase in YPD were transferred to nitrogen-free medium (SD-N; 0.17% yeast nitrogen base without amino acids and ammonium sulfate, 2% dextrose) and incubated at 30 °C. For starvation-induced pexophagy, cells grown overnight in YPD were transferred to oleate medium (0.17% yeast nitrogen base without amino acids ammonium sulfate, 0.5% ammonium sulfate, 1% casamino acids, 0.12% oleate (v/v), 0.2% Tween 40R (v/v), 0.1% yeast extract) at a 1:10 dilution. Pexophagy was induced by transferring cells from oleate medium to SD-N.

TABLE 1.

Yeast strains used in this study

Name	Genotype	Source
KOY76	his3Δ1 leu2Δ0 met15Δ0 ura3Δ0	BY4741 (36)
KOY382	BY4741 his3Δ1::GPD^P-mito-mCherry::CgHIS3 ypt7::kanMX6
KOY611	BY4741 atg32::natNT2 his3Δ1::GPD^P-mito-mCherry::CgHIS3 ypt7::kanMX6
KOY692	BY4741 vph1::VPH1-mCherry::kanMX6 his3Δ1::GPD^P-mito-GFP::CgHIS3 prb1::hphNT1
KOY694	BY4741 atg32::natNT2 vph1::VPH1-mCherry::kanMX6 his3Δ1::GPD^P-mito-GFP::CgHIS3 prb1::hphNT1
KOY1387	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3
KOY1422	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg32::kanMX6
KOY1430	BY4741 agt32::ATG32–3HAn
KOY1555	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg11::natNT2
KOY1601	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::natNT2
KOY1603	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2
KOY1711	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg32::kanMX6 [pRS316-ATG32–3HAn]
KOY1772	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 [pRS316]
KOY1778	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 [pRS316-GFP-ATG8]
KOY1779	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg7::kanMX6 [pRS316-GFP-ATG8]
KOY2085	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat4::natNT2
KOY2087	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 mak3::natNT2
KOY2089	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat5::natNT2
KOY2097	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat3::natNT2
KOY2113	BY4741 pot1::POT1-mCherry::CgHIS3
KOY2188	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 nat1::hphNT1
KOY2230	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::natNT2 [pRS316-GFP-ATG8]
KOY2232	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 [pRS316-GFP-ATG8]
KOY2254	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::NAT1–3HA::hphNT1
KOY2256	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::ARD1–3HA::hphNT1
KOY2326	BY4741 atg32::ATG32–3HAn his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 [pRS315] [pRS316]
KOY2328	BY4741 atg32::ATG32–3HAn his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 [pRS315-ATG32–3HAn] [pRS316-ATG32–3HAn]
KOY2379	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 [pRS316-ARD1–3HA]
KOY2518	BY4741 pot1::POT1-mCherry::CgHIS3 atg36::hphNT1
KOY2632	BY4741 pot1::POT1-mCherry::CgHIS3 nat1::natNT2
KOY2634	BY4741 pot1::POT1-mCherry::CgHIS3 ard1::natNT2
KOY2807	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 [pRS316]
KOY2848	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 [pRS316-ARD1(R125A)-3HA]
KOY2935	BY4741 pep4::kanMX6 prb1::hphNT1 [pRS316-ATG32]
KOY2936	BY4741 pep4::kanMX6 prb1::hphNT1 atg32::ATG32–3HAn [pRS316-ATG32–3HAn]
KOY2988	BY4741 vph1::VPH1-mCherry::kanMX6 his3Δ1::GPD^P-mito-GFP::CgHIS3 prb1::hphNT1 nat1::natNT2
KOY2990	BY4741 vph1::VPH1-mCherry::kanMX6 his3Δ1::GPD^P-mito-GFP::CgHIS3 prb1::hphNT1 ard1::natNT2
KOY2993	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 [pRS316-ARD1(E26Q)-3HA]
KOY3014	BY4741 atg32::ATG32–3HAn his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::natNT2 [pRS316] [pRS315]
KOY3015	BY4741 atg32::ATG32–3HAn his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::natNT2 [pRS316-ATG32–3HAn] [pRS315-ATG32–3HAn]
KOY3137	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::ARD1–3HA::hphNT1 nat1::natNT2
KOY3139	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::NAT1–3HA::hphNT1 ard1::natNT2
KOY3142	BY4741 pep4::kanMX6 prb1::hphNT1 atg32::ATG32–3HAn ard1::natNT2 [pRS316-ATG32–3HAn]
KOY3159	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::NAT1–3HA::hphNT1 [pRS316]
KOY3166	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::NAT1–3HA::hphNT1 ard1::natNT2 [pRS316]
KOY3168	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::NAT1–3HA::hphNT1 ard1::natNT2 [pRS316-ARD1–3HA]
KOY3170	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::NAT1–3HA::hphNT1 ard1::natNT2 [pRS316-ARD1(E26Q)-3HA]
KOY3176	BY4741 his3Δ1::GPD^P-mito-mCherry::CgHIS3 ypt7::kanMX6 nat1::natNT2
KOY3178	BY4741 his3Δ1::GPD^P-mito-mCherry::CgHIS3 ypt7::kanMX6 ard1::natNT2
KOY3229	BY4741 agt32::ATG32–3HAn nat1::natNT2
KOY3230	BY4741 agt32::ATG32–3HAn nat1::natNT2
KOY3231	BY4741 agt32::ATG32–3HAn ard1::natNT2
KOY3232	BY4741 agt32::ATG32–3HAn ard1::natNT2
KOY3239	BY4741 atg32::ATG32–3HAn his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 [pRS316] [pRS315]
KOY3241	BY4741 atg32::ATG32–3HAn his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 [pRS316-ATG32–3HAn] [pRS315-ATG32–3HAn]
KOY3323	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg11::natNT2 [pRS316]
KOY3326	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg11::natNT2 [pRS316-ATG11]
KOY3360	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg19::natNT2
KOY3561	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::GPD^p-ARD1–3HA
KOY3563	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::natNT2 ard1::GPD^p-ARD1–3HA
KOY3696	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg32::kanMX6 [p416TEF-GFP-ATG32]
KOY3698	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 nat1::natNT2 atg32::hphNT1 [p416TEF-GFP-ATG32]
KOY3700	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 ard1::natNT2 atg32::hphNT1 [p416TEF-GFP-ATG32]
KOY3745	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg32::kanMX6 [pRS316-ATG32(V2P)-3HAn]
KOY4028	BY4741 his3Δ1::TEF^P-mito-DHFR-mCherry::CgHIS3 atg11::natNT2 [pRS316-ATG11(A2P)]

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TABLE 2.

Plasmids used in this study

Name	Relevant characteristics	Source
pRS315	CEN LEU2	37
pRS316	CEN URA3	37
pRS315-ATG32–3HAn	CEN LEU2 580-bp 5′-UTR and 744-bp 3′-UTR from ATG32	7
pRS316-ATG32	CEN URA3 580-bp 5′-UTR and 744-bp 3′-UTR from ATG32	11
pRS316-ATG32–3HAn	CEN URA3 580-bp 5′-UTR and 744-bp 3′-UTR from ATG32	7
pRS316-ATG32(V2P)-3HAn	CEN URA3 580-bp 5′-UTR and 744-bp 3′-UTR from ATG32	This study
p416TEF-GFP-ATG32	CEN URA3 TEF^P	7
pRS316-GFP-ATG8	CEN URA3 5′-UTR and 3′-UTR from ATG8	38
pRS316-ARD1–3HA	CEN LEU2 500-bp 5′-UTR from ARD1	This study
pRS316-ARD1(E26Q)-3HA	CEN LEU2 500-bp 5′-UTR from ARD1	This study
pRS316-ARD1(R125A)-3HA	CEN LEU2 500-bp 5′-UTR from ARD1	This study
pRS316-ATG11	CEN URA3 500-bp 5′-UTR and 500-bp 3′-UTR from ATG11	This study
pRS316-ATG11(A2P)	CEN URA3 500-bp 5′-UTR and 500-bp 3′-UTR from ATG11	This study

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Microscopy

Cells were observed using an inverted microscope (Axio Observer. Z1; Carl Zeiss) equipped with differential interference contrast optics, epifluorescence capabilities, a ×100 objective lens (αPlan-APOCHROMAT ×100, NA: 1.46; Carl Zeiss), a monochrome CCD camera (AxioCam MRm; Carl Zeiss), and filter sets for green fluorescent protein (GFP) and mCherry (13 and 20, respectively; Carl Zeiss). Images were captured using acquisition and analysis software (Axio Vision 4.6; Carl Zeiss).

Immunoblotting

Samples corresponding to 0.1 OD₆₀₀ units of cells were separated by SDS-PAGE followed by western blotting and immunodecoration. After treatment with enhanced chemiluminescence reagents, proteins were detected using a luminescent image analyzer (LAS-4000 mini; GE Healthcare). Quantification of the signals was performed using ImageQuant TL (GE Healthcare).

Quantitative RT-PCR

RNA was isolated by Master Pure Yeast RNA purification kit (Epicenter) from 2 OD₆₀₀ units of cells, according to the manufacturer's protocol. Equal amounts of total RNA were mixed with RNA-direct SYBR Green Real Time PCR Master Mix (TOYOBO) and the following primers: ATG32 forward 5′-TGTCACTGCAGCATACGAACAC and reverse 5′-CTGCTCAGTTGAAGAAGGAGATG; ACT1 forward 5′-TATCGTCGGTAGACCAAGACAC and reverse 5′-TCGTCCCAGTTGGTGACAATAC. The Applied Biosystems Step One Plus (Applied Biosystems) was used for quantitative RT-PCR analysis. For each gene, triplicate samples were calculated to average value and then normalized by averaged actin. The value in wild-type cells at the 0-h time point was set to 1. These numbers represent relative expression of ATG32 mRNA.

Immunoprecipitation

Coimmunoprecipitation assays were performed using a vacuolar protease-deficient strain transformed with a plasmid encoding Atg32 (negative control) or Atg32-HA (positive control). 120–140 OD₆₀₀ units of cells grown in SDGlyCA for 24 h were collected by centrifugation, washed once with H₂O, resuspended in TD buffer (0.1 m Tris-SO₄ (pH 9.4), 10 mm DTT), and incubated for 10 min at 30 °C. Cells were collected by centrifugation, resuspended in SP buffer (20 mm potassium phosphate buffer (pH 7.4), 1.2 m sorbitol) containing Zymolyase 100T (120493; Seikagaku), and incubated for 100 min at 30 °C. Spheroplasts were washed with SP buffer and resuspended in SH buffer (0.6 m sorbitol, 20 mm HEPES-KOH (pH 7.4)) containing protease inhibitor mixture (1861284; Thermo Scientific). Whole cell homogenates were subjected to centrifugation (500 × g) at 4 °C for 5 min. Membrane and soluble fractions were separated by centrifugation (17,000 × g) at 4 °C for 5 min. Mitochondria-enriched fractions were then resuspended in lysis buffer (50 mm Tris-HCl (pH 7.5), 100 mm NaCl, 0.1 mm EDTA, 0.4% Triton X-100, and protease inhibitor mixture) at 4 °C for 8 min and subjected to centrifugation (17,000 × g) at 4 °C for 5 min. The supernatant was incubated with 30 μl of anti-HA-agarose conjugate (A2095; Sigma) at 4 °C for 2 h with gentle agitation. The beads were washed twice with wash buffer (50 mm Tris-HCl (pH 7.5), 300 mm NaCl, 0.1 mm EDTA, 0.4% Triton X-100, and protease inhibitor mixture), and once with PBS. Immunoprecipitates were eluted with SDS-sample buffer and analyzed by western blotting.

Results

Mitophagy Is Markedly Suppressed in Cells Lacking NatA

To elucidate the role of NatA in mitophagy, we observed transport of mitochondria to the vacuole, a lytic compartment in yeast. Mitochondria and vacuoles were visualized using a mitochondrial matrix-targeted GFP (mito-GFP) and mCherry fused at the C terminus of Vph1, a membrane-integrated subunit of the vacuolar ATPase (Vph1-mCherry), respectively, in cells lacking Prb1, a vacuole-localized serine protease. When cells are grown in nonfermentable glycerol medium (Gly), mitochondria become active in respiration and are eventually degraded by mitophagy (7). In vacuolar protease-deficient cells, mitochondria transported into the vacuole can be accumulated and observed as degradation intermediates. Cells grown in Gly for 48 h contained mitochondria that were overlapped with the vacuole in a manner dependent on Atg32 (Fig. 1A). We hardly detected mito-GFP signals colocalized with Vph1-mCherry patterns in cells lacking NatA subunits, the catalytic component Ard1 and ribosomal adaptor Nat1 (13, 14), under the same conditions (Fig. 1A). These observations suggest that NatA is important for mitophagy.

FIGURE 1. — **Loss of NatA leads to strong suppression of mitophagy.** A, mitochondrial GFP-expressing and vacuolar Vph1-mCherry-expressing *prb1*Δ, *atg32*Δ *prb1*Δ, *nat1*Δ *prb1*Δ, and *ard1*Δ *prb1*Δ cells grown to mid-log phase in dextrose medium (0 h) were shifted to respiration medium (Gly) for 48 h and observed using fluorescence microscopy. Cells lacking Prb1 are vacuolar protease-deficient and accumulate degradation intermediates within the vacuole. *Scale bar,* 5 μm. *DIC*, differential interference contrast. B, mitochondria-targeted DHFR-mCherry-expressing (mito-DHFR-mCherry, depicted by *arrow*) wild-type, *nat1*Δ, *ard1*Δ, *nat1*Δ *ard1*Δ, and *atg32*Δ cells were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. Generation of free mCherry (depicted by *arrowhead*) indicates transport of the marker to the vacuole. Pgk1 was monitored as a loading control. C, free mCherry in cells under respiratory conditions for 48 h in B was quantified in three experiments. Data represent the averages of all experiments, with *bars* indicating standard deviations. D, mitochondria-targeted DHFR-mCherry-expressing (mito-DHFR-mCherry, depicted by *arrow*) wild-type, *nat3*Δ, *mak3*Δ, *nat4*Δ, and *nat5*Δ cells were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting as in *B. E*, mito-DHFR-mCherry-expressing wild-type, *nat1*Δ, *ard1*Δ, and *atg32*Δ cells grown to mid-log phase in nutrient-rich medium were shifted to nitrogen starvation (−N), collected at the indicated time points, and subjected to western blotting as in *B. F*, free mCherry in cells under starvation conditions for 24 h in E was quantified in three experiments. Data represent the averages of all experiments, with *bars* indicating standard deviations. G, mito-DHFR-mCherry-expressing wild-type, *nat1*Δ, *ard1*Δ, and *atg7*Δ cells were transformed with a plasmid encoding GFP-Atg8, grown to mid-log phase in rich dextrose medium, incubated for the indicated time points in starvation medium (−N), and subjected to western blotting. Generation of free GFP indicates progression of bulk autophagy. Atg7 is an E1 enzyme essential for autophagy. H, mito-DHFR-mCherry-expressing wild-type, *nat1*Δ, *ard1*Δ, and *atg19*Δ cells were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. Precursor form (p) of Ape1 is transported from the cytosol to the vacuole via the Cvt pathway, a selective autophagy-related process, and cleaved to be a mature form (m). Atg19 is an adaptor protein required for the Cvt pathway (35). I, Pot1-mCherry-expressing (a peroxisome marker) wild-type, *nat1*Δ, *ard1*Δ, and *atg36*Δ cells were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. Generation of free mCherry indicated transport of peroxisomes to the vacuole. J, free mCherry in cells under respiratory conditions for 48 h in I was quantified in three experiments. Data represent the averages of all experiments, with *bars* indicating standard deviations. K, Pot1-mCherry (a peroxisome marker) expressing wild-type, *nat1*Δ, *ard1*Δ, and *atg36*Δ cells grown in oleic acid medium were shifted to nitrogen starvation (−N), collected at the indicated time points, and subjected to western blotting. Generation of free mCherry indicated transport of peroxisomes to the vacuole. Atg36 acts as an adaptor essential for pexophagy (16). L, free mCherry in cells under nitrogen starvation for 6 h in K was quantified in three experiments. Data represent the averages of all experiments, with *bars* indicating standard deviations.

We next quantified mitochondrial degradation using mito-dihydrofolate reductase (DHFR)³-mCherry, a reporter located in the matrix of mitochondria (11). Upon mitophagy, this fusion protein is processed to generate free mCherry that is appreciably protease-resistant, thereby indicating degradation of mitochondria. We found that accumulation of free mCherry was greatly decreased in cells lacking Nat1, Ard1, or both (Fig. 1, B and C). By contrast, cells lacking the catalytic subunits of other N-terminal acetyltransferases, NatB–E (13, 14), did not exhibit defects in degradation of mitochondria (Fig. 1D). We have previously demonstrated that mitophagy is induced during nitrogen starvation (15). Starvation-induced mitochondrial degradation was also suppressed in cells lacking Nat1 or Ard1 (Fig. 1, E and F). These results indicate that NatA participates in the regulation of mitophagy.

We further investigated whether NatA affects the other autophagy-related pathways. First, we monitored processing of GFP-Atg8, an indicator of autophagy flux, and confirmed that free GFP was detected in cells lacking Nat1 or Ard1 under nitrogen deprivation (Fig. 1G). Therefore, it is likely that NatA is dispensable for starvation-induced autophagy. Next, we examined the cytoplasm-to-vacuole targeting (Cvt) pathway, a selective autophagy process that mediates transport of vacuolar enzymes such as Ape1, an amino peptidase, from the cytosol to the vacuole. Mature Ape1 transported to the vacuole was detectable in Nat1 or Ard1-null cells under respiratory conditions, indicating that NatA is dispensable for the Cvt pathway (Fig. 1H). Finally, we evaluated selective peroxisome autophagy (pexophagy). Degradation of peroxisomes was monitored using Pot1-mCherry, a marker localized in the peroxisomal matrix, under respiratory conditions that are also able to induce pexophagy (16). We found that accumulation of free mCherry was strongly depressed in cells lacking Nat1 or Ard1 (Fig. 1, I and J). Starvation-induced pexophagy was also inhibited in these mutant cells (Fig. 1, K and L). Together, our results raise the possibility that NatA is a common factor critical for selective elimination of organelles.

Enzymatic Activity of NatA Is Crucial for Mitophagy

To ask whether protein N-terminal acetylation by NatA is important for selective mitochondrial degradation, we investigated a NatA variant lacking enzymatic activity. Previous research unravels the structure of NatA in the fission yeast Schizosaccharomyces pombe, demonstrating that glutamine substitution for Glu-24 of Ard1 leads to loss of enzymatic activities but does not affect substrate binding capacity (17). This critical glutamate is conserved from budding yeast (Glu-26) to higher eukaryotes (Fig. 2A). We thus monitored mitophagy using mito-DHFR-mCherry in cells expressing hemagglutinin (HA)-tagged Ard1^E26Q. Under respiratory conditions, mitochondrial degradation was inhibited to the levels similar to cells lacking Ard1 (Fig. 2, B and C). HA tagging of Nat1 and Ard1 did not significantly affect mitophagy (Fig. 2D). In addition, expression of Nat1 was not severely impaired in cells expressing the Ard1^E26Q mutant (Fig. 2E). Another previous study has reported that human Ard1^R82A, a variant containing a mutation in the acetyl-CoA binding domain, decreases enzymatic activities (18). Because this crucial arginine is also conserved among eukaryotes (Fig. 2F), we introduced a plasmid encoding HA-tagged Ard1^R125A (corresponding to human Ard1^R82A) into ard1Δ cells, and we monitored processing of mito-DHFR-mCherry under respiratory conditions. Cells expressing Ard1^R125A-3HA displayed a reduced accumulation of free mCherry (Fig. 2, G and H). Taken together, we conclude that protein N-terminal acetylation by NatA is critical for mitophagy in yeast.

FIGURE 2. — **Catalytically inactive Ard1 mutant is defective in mitophagy.** A, part of amino acid sequence alignment from Ard1 homologs. The glutamic acid residues critical for enzymatic activity are indicated with *asterisk. B*, mito-DHFR-mCherry-expressing wild-type and *ard1*Δ cells transformed with empty vector, plasmid-encoding Ard1–3HA, or plasmid-encoding Ard1^E26Q (E26Q)-3HA were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. Generation of free mCherry (depicted by *arrowhead*) indicates transport of mitochondria to the vacuole. C, free mCherry in cells under respiratory conditions for 48 h in B was quantified in three experiments. Data represent the averages of all experiments, with *bars* indicating standard deviations. D, wild-type, Nat1–3HA-expressing, Ard1–3HA-expressing, and *atg32*Δ cells containing mito-DHFR-mCherry were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting as in *B. E*, wild-type, Nat1–3HA-expressing, and *atg32*Δ cells containing mito-DHFR-mCherry were transformed with empty vector, and a plasmid encoding Ard1–3HA or Ard1^E26Q (E26Q)-3HA, grown for the indicated time points in glycerol medium (*Gly*), and subjected to western blotting as in *B. F*, part of amino acid sequence alignment including acetyl-CoA binding domains from Ard1 homologs. The arginine residues critical for acetyl-CoA binding are indicated with *asterisk. G*, mito-DHFR-mCherry-expressing wild-type and *ard1*Δ cells transformed with empty vector and a plasmid encoding Ard1–3HA or Ard1^R125A (R125A)-3HA were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting as in *B. H*, free mCherry in cells under respiratory conditions for 48 h in G was quantified in three experiments. Data represent the averages of all experiments, with *bars* indicating standard deviations.

Ribosomal Association of Ard1 Is Important for Efficient Mitophagy

Although Ard1 stably associates with the ribosome via Nat1 in yeast (19, 20), it has also been reported that monomeric Ard1 purified from fission yeast is still catalytically active with altered substrate specificity in vitro (17) and that free Ard1 post-translationally acetylates β- and γ-actin in humans (21). To ask whether monomeric Ard1 dissociated from the ribosome contributes to regulation of mitophagy in S. cerevisiae, we monitored mitochondrial degradation in cells overexpressing Ard1 because the Ard1 levels are drastically reduced in nat1Δ cells (Fig. 3A). We found that overexpression of Ard1 did not lead to recovery of mitophagy in the absence of the ribosome adaptor (Fig. 3, C and D). Hence, it seems likely that NatA-mediated cotranslational acetylation is crucial for mitophagy.

FIGURE 3. — **Ribosome-associated Ard1 is required for efficient mitophagy.** A, Ard1–3HA-expressing wild-type and *nat1*Δ cells were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. B, Nat1–3HA-expressing wild-type and *ard1*Δ cells were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. C, wild-type and *nat1*Δ cells expressing Ard1–3HA under the endogenous promoter or the strong *GPD* promoter were grown for the indicated time points in glycerol medium (*Gly*), and subjected to western blotting. Mitophagy was monitored using mito-DHFR-mCherry (depicted by *arrow*). Generation of free mCherry (depicted by *arrowhead*) indicates transport of mitochondria to the vacuole. D, free mCherry in cells under respiratory conditions for 48 h in C was quantified in three experiments. Data represent the averages of all experiments, with *bars* indicating standard deviations.

Mitophagosome Formation Is Impaired in Cells Lacking NatA

In which stage of mitophagy does NatA participate? It is noteworthy that the Nat1 and Ard1 protein levels declined during respiratory growth and became exceedingly low at the 48-h time point (Fig. 3, A and B). We thus speculated that NatA affects the early phase of mitophagy, in particular mitophagosome formation through which mitochondria are enclosed by the isolation membrane. To test this possibility, we used cells lacking Ypt7, a Rab family GTPase essential for homotypic vacuole fusion. Although autophagosomes are formed normally in this mutant, they do not fuse to the vacuole and accumulate in the cytosol (22). When ypt7Δ cells grown in fermentable medium (Dex) are shifted to respiratory medium (Gly), their mitochondrial shape changes from tubules to fragments (Fig. 4A). Conversely, upon a shift from respiratory to fermentable conditions, mitochondrial tubular networks are reformed by their fusion, but mitophagosomes remain isolated from intact mitochondria. Accordingly, we can detect mitophagosomes as dot-like structures (7). We found mitochondrial dots in cells grown under fermentable conditions for 3 h after respiratory growth (Fig. 4B). By contrast, cells containing mitochondrial dots were few in the absence of Nat1 or Ard1 (Fig. 4C). Hence, these observations suggest that NatA plays a key role in the early step of mitophagosome formation.

FIGURE 4. — **Disruption of NatA causes inhibition of mitophagosome formation.** A, experimental scheme for observation of mitophagosomes in *ypt7*Δ cells. B, mitochondrial mCherry-expressing *ypt7*Δ, *ypt7*Δ *atg32*Δ, *ypt7*Δ *nat1*Δ, and *ypt7*Δ *ard1*Δ cells grown to mid-log phase in dextrose medium (*Dex*) were shifted to respiration medium (*Gly*) for 48 h, transferred to dextrose medium for 3 h, and investigated using fluorescence microscopy. Loss of Ypt7 prevents fusion between mitophagosomes and vacuoles. *Scale bar,* 5 μm. *DIC*, differential interference contrast. C, number of cells containing mitochondrial mCherry dots at stage 3 in B was quantified in three experiments. Date represent the averages of all experiments, with *bars* indicating standard deviations. n >100. **, p < 0.05.

Atg32 Expression Levels Are Low in nat1Δ and ard1Δ Cells

Based on the previous findings that Atg32 acts as a mitophagy receptor to generate mitophagosomes (7, 8), we investigated whether NatA affects Atg32 functions. At first, to examine Atg32 expression in NatA-null cells under respiratory conditions, HA-tagged Atg32 was expressed from the endogenous locus and monitored by western blotting. In light of previous reports that the Atg32 levels are transiently elevated in respiring cells at mid-log phase (7), and that growth of cells lacking NatA is slow in nonfermentable medium (13), we prepared samples from nat1Δ and ard1Δ cells at later time points than wild-type cells and compared their highest amounts of Atg32. Cells lacking Nat1 or Ard1 exhibited a partial reduction in the Atg32 protein levels (Fig. 5, A and B). Additionally, transcriptional levels of ATG32 mRNA in these cells were quantified by real time PCR. We found that ATG32 mRNA expressions in nat1Δ and ard1Δ cells were less than half of those in wild-type cells during respiratory growth (Fig. 5C), raising the possibility that NatA may regulate transcription of ATG32.

FIGURE 5. — **Atg32 induction is partially suppressed in *nat1*Δ and *ard1*Δ cells.** A, Atg32–3HA-expressing wild-type, *nat1*Δ, and *ard1*Δ cells were grown in glycerol medium (*Gly*), collected at the indicated time points, and subjected to western blotting. B, peaks of Atg32–3HA expression in A were quantified in three experiments (wild type, 18 h; *nat1*Δ, 24 h; *ard1*Δ, 24 h). Data represent the averages of all experiments, with *bars* indicating standard deviations. *, p < 0.01. C, mito-DHFR-mCherry-expressing wild-type, *nat1*Δ, and *ard1*Δ cells were grown for the indicated time points in glycerol medium (*Gly*). *ATG32* mRNA expression was analyzed by real time PCR and normalized to *ACT1* mRNA expression. Data represent the averages of all experiments, with *bars* indicating standard deviations. D, Atg32–3HA- and mito-DHFR-mCherry-expressing wild-type, *nat1*Δ, and *ard1*Δ cells transformed with empty vectors or two plasmids encoding Atg32–3HA were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. Generation of free mCherry (depicted by *arrowhead*) indicates transport of mitochondria to the vacuole. E and F, free mCherry in *nat1*Δ and *ard1*Δ cells under respiratory conditions for 48 h in D was quantified in three experiments. Data represent the averages of all experiments, with *bars* indicating standard deviations. G, mito-DHFR-mCherry-expressing *atg32*Δ, *nat1*Δ *atg32*Δ, and *ard1*Δ*atg32*Δ cells were transformed with a plasmid encoding GFP-Atg32 with the constitutive *TEF2* promoter, grown to log phase in glucose medium, and investigated using fluorescence microscopy. *Scale bar*, 5 μm. H, *pep4*Δ *prb1*Δ and *pep4*Δ *prb1*Δ *ard1*Δ cells expressing chromosome- or plasmid-encoded versions of Atg32 or Atg32-HA were grown in glycerol medium and subjected to coimmunoprecipitation using anti-HA antibody-conjugated agarose. Eluted immunoprecipitates (IP) and detergent-solubilized mitochondria-enriched fractions (*input*) were analyzed by western blotting. I, mito-DHFR-mCherry-expressing wild-type and *atg32*Δ cells transformed with an empty vector or a plasmid encoding Atg32–3HA or Atg32^V2P (V2P)-3HA were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. Generation of free mCherry (depicted by *arrowhead*) indicates transport of mitochondria to the vacuole.

To ask whether these reductions of Atg32 expression levels impact mitochondrial degradation, we introduced two low-copy plasmids encoding Atg32–3HA into NatA-null cells and monitored processing of mito-DHFR-mCherry. When grown in nonfermentable medium, Nat1 or Ard1-null cells overexpressing Atg32 displayed partial recovery of mitochondrial degradation (Fig. 5, D–F). These data are consistent with the idea that partial impairment of Atg32 induction may be one of the factors for severe mitophagy defects in cells lacking NatA.

We also confirmed Atg32 localization to mitochondria and interaction with Atg8 and Atg11 in the early stage of mitophagy (7, 11). Fluorescence live cell imaging and coimmunoprecipitation assays were performed using GFP- and HA-tagged Atg32, respectively. GFP-Atg32 was normally localized to mitochondria in cells lacking Nat1 or Ard1 (Fig. 3G), and Atg32–3HA coprecipitated with Atg11 and Atg8 in the absence of Ard1 (Fig. 3H). It is likely that NatA does not regulate organelle targeting and protein-protein interactions of Atg32.

In yeast, NatA acetylates the second amino acids, Ser, Ala, Thr, Val, Gly, and Cys, of nascent polypeptides whose N-terminal methionine residues are removed by methionine peptidase (23 –25). Atg32 contains a valine at the second amino acid position, raising the possibility that Atg32 is acetylated by NatA. We thus substituted proline for Val-2 of Atg32, introduced a plasmid encoding Atg32^V2P-3HA into Atg32-null cells, and monitored processing of mito-DHFR-mCherry under respiratory conditions. A protein containing proline at the second amino acid position is not acetylated by all N-terminal acetyltransferases in accordance with the XPX rule (26). We found that mitophagy is not altered in cells expressing Atg32^V2P-3HA (Fig. 3I). Thus, NatA is unlikely to directly control Atg32.

Atg8 and Atg11 Are Expressed Normally in the Absence of NatA

Finally, we examined whether loss of NatA affects Atg8 and Atg11, two Atg32-interacting proteins essential for mitophagy, under respiratory conditions. The protein levels of Atg8 and Atg11 were not decreased in cells lacking Nat1 or Ard1 (Fig. 6, A and B). Moreover, the phospholipid conjugation of Atg8 occurred properly in NatA-null cells (Fig. 6A). These data are in agreement with the results that NatA is dispensable for the Cvt pathway under respiratory conditions (Fig. 1H). Because Atg11 has alanine at the second amino acid position that can be acetylated by NatA, we introduced a plasmid encoding Atg11^A2P into cells lacking Atg11 and monitored mitophagy under respiratory conditions. Cells expressing Atg11^A2P did not exhibit significant changes in mitochondrial degradation (Fig. 6C). Therefore, it seems likely that NatA acts in mitophagy irrespectively of Atg8 and Atg11.

FIGURE 6. — **Atg8 and Atg11 are not significantly affected in cells lacking NatA.** A, wild-type, *nat1*Δ *ard1*Δ, and *atg7*Δ cells were grown in glycerol medium (*Gly*), collected at the indicated time points, and subjected to western blotting. B, wild-type, *nat1*Δ *ard1*Δ, and *atg11*Δ cells were grown in glycerol medium (*Gly*), collected at the indicated time points, and subjected to western blotting. C, mito-DHFR-mCherry (depicted by *arrow*)-expressing wild-type and *atg11*Δ cells transformed with an empty vector or a plasmid encoding Atg11 or Atg11^A2P (*A2P*) were grown for the indicated time points in glycerol medium (*Gly*) and subjected to western blotting. Generation of free mCherry (depicted by *arrowhead*) indicates transport of mitochondria to the vacuole.

Discussion

In this study, we demonstrated that NatA plays a critical role in selective degradation of mitochondria in yeast. This catabolic process requires NatA catalytic activity and association with the ribosome, indicating that nascent polypeptide N-terminal acetylation of NatA target(s) is a key step for activation of mitophagy. Notably, mitophagosome formation is compromised in nat1Δ and ard1Δ cells, suggesting that NatA participates in the early phase of mitophagy. In addition, Atg32 induction is partially suppressed in the absence of Nat1 or Ard1. In particular, the transcriptional levels of ATG32 are significantly reduced. When we up-regulate Atg32 expression in these mutant cells, mitochondrial degradation is not fully recovered to a wild-type level. We thus speculate that NatA regulates mitophagy through Atg32 and other unknown factors (but not Atg8 and Atg11) (Fig. 7).

FIGURE 7. — **Model for regulation of mitophagy via NatA-mediated protein N-terminal acetylation.** NatA consists of the catalytic Ard1 and the ribosomal adaptor Nat1. This heterodimeric enzyme cotranslationally catalyzes N-terminal acetylation of substrates that are involved in mitophagosome formation and regulation of Atg32 expression at the early stage of mitophagy.

Recently, the Ume6-Sin3-Rpd3 complex, a transcriptional regulator, has been reported to suppress Atg32 expression (27). It remains possible that NatA might acetylate Sin3 and Rpd3 containing a serine and a valine at the second amino acid position, respectively. As N-terminally acetylated residues act as degradation signals (28), Sin3, Rpd3, or both could become stabilized, leading to reduction of Atg32 levels in NatA-null cells.

Bulk autophagy normally occurs in NatA-null cells, indicating that NatA does not act on the core autophagy machinery. Moreover, NatA is unlikely to serve as a common regulator of selective autophagy because it is dispensable for the Cvt pathway. Interestingly, pexophagy is suppressed in NatA-null cells, raising the possibility that NatA regulates selective organelle autophagy via a common mechanism. It has recently been reported that the MAPK signaling pathway, Wsc1-Pkc1-Bck1-Mkk1/2-Slt2, is important for mitophagy and pexophagy in yeast (29). However, these proteins acting in the MAPK pathway have so far not been identified as NatA targets (23). Alternatively, it remains possible that NatA affects distinct molecules specific for either mitophagy or pexophagy.

Does NatA-mediated protein modification affect mitochondrial shaping during mitophagy? Fragmentation is a crucial step for mitochondria to be efficiently surrounded by the isolation membrane in mammalian cells (30 –32). In yeast, it has been reported that Atg11 interacts with Dnm1, a dynamin-related GTPase required for mitochondrial fission, and that mitophagy is suppressed in dnm1Δ cells (33). However, mitochondrial fragmentation is almost normal in NatA-null cells under respiratory conditions (Figs. 1A and 4B). It is therefore unlikely that altered mitochondrial dynamics causes impairment of mitophagy in the absence of NatA. Because more than half of all proteins are acetylated by N-terminal acetyltransferases in yeast, NatA acetylates a lot of mitochondrial proteins, including import receptor, ATP synthase subunits, and ribosomal proteins (23). We thus do not exclude the possibility that changes in multiple mitochondrial functions could lead to mitophagy defects in NatA-null cells. Moreover, a recent study has reported that ERMES, a protein complex tethering the endoplasmic reticulum to mitochondria, is crucial for mitophagosome formation (34). The endoplasmic reticulum-mitochondrial tethering does not seem to be completely disrupted in NatA-null cells, because mitochondrial morphology defects, typical phenotypes of the ERMES mutants, are not seen in cells lacking Ard1 or Nat1 (Fig. 4B).

To our knowledge, this is the first report that protein N-terminal acetylation is linked to mitophagy. To clarify NatA-mediated activation of mitochondrial degradation in more detail, further studies are needed to identify NatA substrate(s) involved in mitophagy, to understand Atg32 induction mechanisms under respiratory conditions, and to investigate whether mitochondrial functions alter in NatA-null cells. These future approaches will elucidate regulation of mitophagy via protein N-terminal acetylation.

Author Contributions

K. O. designed the experiments, conceived and coordinated the study, and wrote the paper. A. E. designed, performed, and analyzed the experiments and wrote the paper. All authors reviewed the results and approved the final version of the manuscript.

Acknowledgments

We are grateful to Toshiya Endo, Hayashi Yamamoto, Kaori Sakakibara, and Ayako Hashimoto for the kind gifts of antibodies, plasmids, and yeast strains. We also thank Noriko Okamoto for valuable comments on this manuscript.

This work was supported in part by Japan Society for the Promotion of Science, Grants-in-aid from JSPS Fellows 261682 (to A. E.) and Scientific Research (B) 25291045 (to K. O.), and by Ministry of Education, Culture, Sports, Science, and Technology Grant-in-aid for Scientific Research on Innovative Areas 15H01536 (to K. O.). The authors declare that they have no conflicts of interest with the contents of this article.

The abbreviations used are:

DHFR: dihydrofolate reductase
Cvt: cytoplasm-to-vacuole targeting.

References

1. Nunnari J., Suomalainen A. (2012) Mitochondria: in sickness and in health. Cell 148, 1145–1159 [DOI] [PMC free article] [PubMed] [Google Scholar]
2. Okamoto K., Kondo-Okamoto N. (2012) Mitochondria and autophagy: critical interplay between the two homeostats. Biochim. Biophys. Acta. 1820, 595–600 [DOI] [PubMed] [Google Scholar]
3. Mizushima N., Yoshimori T., Ohsumi Y. (2011) The role of Atg proteins in autophagosome formation. Annu. Rev. Cell Dev. Biol. 27, 107–132 [DOI] [PubMed] [Google Scholar]
4. Mizushima N. (2011) Autophagy in protein and organelle turnover. Cold Spring Harb. Symp. Quant. Biol. 76, 397–402 [DOI] [PubMed] [Google Scholar]
5. Youle R. J., Narendra D. P. (2011) Mechanisms of mitophagy. Nat. Rev. Mol. Cell Biol. 12, 9–14 [DOI] [PMC free article] [PubMed] [Google Scholar]
6. Okamoto K. (2014) Organellophagy: eliminating cellular building blocks via selective autophagy. J. Cell Biol. 205, 435–445 [DOI] [PMC free article] [PubMed] [Google Scholar]
7. Okamoto K., Kondo-Okamoto N., Ohsumi Y. (2009) Mitochondria-anchored receptor Atg32 mediates degradation of mitochondria via selective autophagy. Dev. Cell 17, 87–97 [DOI] [PubMed] [Google Scholar]
8. Kanki T., Wang K., Cao Y., Baba M., Klionsky D. J. (2009) Atg32 is a mitochondrial protein that confers selectivity during mitophagy. Dev. Cell 17, 98–109 [DOI] [PMC free article] [PubMed] [Google Scholar]
9. Kurihara Y., Kanki T., Aoki Y., Hirota Y., Saigusa T., Uchiumi T., Kang D. (2012) Mitophagy plays an essential role in reducing mitochondrial production of reactive oxygen species and mutation of mitochondrial DNA by maintaining mitochondrial quantity and quality in yeast. J. Biol. Chem. 287, 3265–3272 [DOI] [PMC free article] [PubMed] [Google Scholar]
10. Aoki Y., Kanki T., Hirota Y., Kurihara Y., Saigusa T., Uchiumi T., Kang D. (2011) Phosphorylation of serine 114 on Atg32 mediates mitophagy. Mol. Biol. Cell 22, 3206–3217 [DOI] [PMC free article] [PubMed] [Google Scholar]
11. Kondo-Okamoto N., Noda N. N., Suzuki S. W., Nakatogawa H., Takahashi I., Matsunami M., Hashimoto A., Inagaki F., Ohsumi Y., Okamoto K. (2012) Autophagy-related protein 32 acts as autophagic degron and directly initiates mitophagy. J. Biol. Chem. 287, 10631–10638 [DOI] [PMC free article] [PubMed] [Google Scholar]
12. Kanki T., Kurihara Y., Jin X., Goda T., Ono Y., Aihara M., Hirota Y., Saigusa T., Aoki Y., Uchiumi T., Kang D. (2013) Casein kinase 2 is essential for mitophagy. EMBO Rep. 14, 788–794 [DOI] [PMC free article] [PubMed] [Google Scholar]
13. Starheim K. K., Gevaert K., Arnesen T. (2012) Protein N-terminal acetyltransferases: when the start matters. Trends Biochem. Sci. 37, 152–161 [DOI] [PubMed] [Google Scholar]
14. Arnesen T. (2011) Towards a functional understanding of protein N-terminal acetylation. PLoS Biol. 9, e1001074. [DOI] [PMC free article] [PubMed] [Google Scholar]
15. Eiyama A., Kondo-Okamoto N., Okamoto K. (2013) Mitochondrial degradation during starvation is selective and temporally distinct from bulk autophagy in yeast. FEBS Lett. 587, 1787–1792 [DOI] [PubMed] [Google Scholar]
16. Motley A. M., Nuttall J. M., Hettema E. H. (2012) Pex3-anchored Atg36 tags peroxisomes for degradation in Saccharomyces cerevisiae. EMBO J. 31, 2852–2868 [DOI] [PMC free article] [PubMed] [Google Scholar]
17. Liszczak G., Goldberg J. M., Foyn H., Petersson E. J., Arnesen T., Marmorstein R. (2013) Molecular basis for N-terminal acetylation by the heterodimeric NatA complex. Nat. Struct. Mol. Biol. 20, 1098–1105 [DOI] [PMC free article] [PubMed] [Google Scholar]
18. Asaumi M., Iijima K., Sumioka A., Iijima-Ando K., Kirino Y., Nakaya T., Suzuki T. (2005) Interaction of N-terminal acetyltransferase with the cytoplasmic domain of β-amyloid precursor protein and its effect on Aβ secretion. J. Biochem. 137, 147–155 [DOI] [PubMed] [Google Scholar]
19. Gautschi M., Just S., Mun A., Ross S., Rücknagel P., Dubaquié Y., Ehrenhofer-Murray A., Rospert S. (2003) The yeast Nα-acetyltransferase NatA is quantitatively anchored to the ribosome and interacts with nascent polypeptides. Mol. Cell. Biol. 23, 7403–7414 [DOI] [PMC free article] [PubMed] [Google Scholar]
20. Polevoda B., Brown S., Cardillo T. S., Rigby S., Sherman F. (2008) Yeast N(α)-terminal acetyltransferases are associated with ribosomes. J. Cell. Biochem. 103, 492–508 [DOI] [PubMed] [Google Scholar]
21. Van Damme P., Evjenth R., Foyn H., Demeyer K., De Bock P.-J., Lillehaug J. R., Vandekerckhove J., Arnesen T., Gevaert K. (2011) Proteome-derived peptide libraries allow detailed analysis of the substrate specificities of N(α)-acetyltransferases and point to hNaa10p as the post-translational actin N(α)-acetyltransferase. Mol. Cell. Proteomics 10, M110.004580. [DOI] [PMC free article] [PubMed] [Google Scholar]
22. Kirisako T., Baba M., Ishihara N., Miyazawa K., Ohsumi M., Yoshimori T., Noda T., Ohsumi Y. (1999) Formation process of autophagosome is traced with Apg8/Aut7p in yeast. J. Cell Biol. 147, 435–446 [DOI] [PMC free article] [PubMed] [Google Scholar]
23. Arnesen T., Van Damme P., Polevoda B., Helsens K., Evjenth R., Colaert N., Varhaug J. E., Vandekerckhove J., Lillehaug J. R., Sherman F., Gevaert K. (2009) Proteomics analyses reveal the evolutionary conservation and divergence of N-terminal acetyltransferases from yeast and humans. Proc. Natl. Acad. Sci. U.S.A. 106, 8157–8162 [DOI] [PMC free article] [PubMed] [Google Scholar]
24. Mullen J. R., Kayne P. S., Moerschell R. P., Tsunasawa S., Gribskov M., Colavito-Shepanski M., Grunstein M., Sherman F., Sternglanz R. (1989) Identification and characterization of genes and mutants for an N-terminal acetyltransferase from yeast. EMBO J. 8, 2067–2075 [DOI] [PMC free article] [PubMed] [Google Scholar]
25. Polevoda B., Norbeck J., Takakura H., Blomberg A., Sherman F. (1999) Identification and specificities of N-terminal acetyltransferases from Saccharomyces cerevisiae. EMBO J. 18, 6155–6168 [DOI] [PMC free article] [PubMed] [Google Scholar]
26. Goetze S., Qeli E., Mosimann C., Staes A., Gerrits B., Roschitzki B., Mohanty S., Niederer E. M., Laczko E., Timmerman E., Lange V., Hafen E., Aebersold R., Vandekerckhove J., Basler K., et al. (2009) Identification and functional characterization of N-terminally acetylated proteins in Drosophila melanogaster. PLos Biol. 7, e1000236. [DOI] [PMC free article] [PubMed] [Google Scholar]
27. Aihara M., Jin X., Kurihara Y., Yoshida Y., Matsushima Y., Oku M., Hirota Y., Saigusa T., Aoki Y., Uchiumi T., Yamamoto T., Sakai Y., Kang D., Kanki T. (2014) Tor and the Sin3-Rpd3 complex regulate expression of the mitophagy receptor protein Atg32 in yeast. J. Cell Sci. 127, 3184–3196 [DOI] [PubMed] [Google Scholar]
28. Hwang C.-S., Shemorry A., Varshavsky A. (2010) N-terminal acetylation of cellular proteins creates specific degradation signals. Science. 327, 973–977 [DOI] [PMC free article] [PubMed] [Google Scholar]
29. Mao K., Wang K., Zhao M., Xu T., Klionsky D. J. (2011) Two MAPK-signaling pathways are required for mitophagy in Saccharomyces cerevisiae. J. Cell Biol. 193, 755–767 [DOI] [PMC free article] [PubMed] [Google Scholar]
30. Gomes L. C., Di Benedetto G., Scorrano L. (2011) During autophagy mitochondria elongate, are spared from degradation and sustain cell viability. Nat. Cell Biol. 13, 589–598 [DOI] [PMC free article] [PubMed] [Google Scholar]
31. Rambold A. S., Kostelecky B., Elia N., Lippincott-Schwartz J. (2011) Tubular network formation protects mitochondria from autophagosomal degradation during nutrient starvation. Proc. Natl. Acad. Sci. U.S.A. 108, 10190–10195 [DOI] [PMC free article] [PubMed] [Google Scholar]
32. Twig G., Elorza A., Molina A. J., Mohamed H., Wikstrom J. D., Walzer G., Stiles L., Haigh S. E., Katz S., Las G., Alroy J., Wu M., Py B. F., Yuan J., Deeney J. T., et al. (2008) Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. EMBO J. 27, 433–446 [DOI] [PMC free article] [PubMed] [Google Scholar]
33. Mao K., Wang K., Liu X., Klionsky D. J. (2013) The scaffold protein Atg11 recruits fission machinery to drive selective mitochondria degradation by autophagy. Dev. Cell 26, 9–18 [DOI] [PMC free article] [PubMed] [Google Scholar]
34. Böckler S., Westermann B. (2014) Mitochondrial ER contacts are crucial for mitophagy in yeast. Dev. Cell 28, 450–458 [DOI] [PubMed] [Google Scholar]
35. Kim J., Kamada Y., Stromhaug P. E., Guan J., Hefner-Gravink A., Baba M., Scott S. V., Ohsumi Y., Dunn W. A., Jr., Klionsky D. J. (2001) Cvt9/Gsa9 functions in sequestering selective cytosolic cargo destined for the vacuole. J. Cell Biol. 153, 381–396 [DOI] [PMC free article] [PubMed] [Google Scholar]
36. Brachmann C. B., Davies A., Cost G. J., Caputo E., Li J., Hieter P., Boeke J. D. (1998) Designer deletion strains derived from Saccharomyces cerevisiae S288C: a useful set of strains and plasmids for PCR-mediated gene disruption and other applications. Yeast 14, 115–132 [DOI] [PubMed] [Google Scholar]
37. Sikorski R. S., Hieter P. (1989) A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122, 19–27 [DOI] [PMC free article] [PubMed] [Google Scholar]
38. Suzuki K., Kirisako T., Kamada Y., Mizushima N., Noda T., Ohsumi Y. (2001) The pre-autophagosomal structure organized by concerted functions of APG genes is essential for autophagosome formation. EMBO J. 20, 5971–5981 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B1] 1. Nunnari J., Suomalainen A. (2012) Mitochondria: in sickness and in health. Cell 148, 1145–1159 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B2] 2. Okamoto K., Kondo-Okamoto N. (2012) Mitochondria and autophagy: critical interplay between the two homeostats. Biochim. Biophys. Acta. 1820, 595–600 [DOI] [PubMed] [Google Scholar]

[B3] 3. Mizushima N., Yoshimori T., Ohsumi Y. (2011) The role of Atg proteins in autophagosome formation. Annu. Rev. Cell Dev. Biol. 27, 107–132 [DOI] [PubMed] [Google Scholar]

[B4] 4. Mizushima N. (2011) Autophagy in protein and organelle turnover. Cold Spring Harb. Symp. Quant. Biol. 76, 397–402 [DOI] [PubMed] [Google Scholar]

[B5] 5. Youle R. J., Narendra D. P. (2011) Mechanisms of mitophagy. Nat. Rev. Mol. Cell Biol. 12, 9–14 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B6] 6. Okamoto K. (2014) Organellophagy: eliminating cellular building blocks via selective autophagy. J. Cell Biol. 205, 435–445 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B7] 7. Okamoto K., Kondo-Okamoto N., Ohsumi Y. (2009) Mitochondria-anchored receptor Atg32 mediates degradation of mitochondria via selective autophagy. Dev. Cell 17, 87–97 [DOI] [PubMed] [Google Scholar]

[B8] 8. Kanki T., Wang K., Cao Y., Baba M., Klionsky D. J. (2009) Atg32 is a mitochondrial protein that confers selectivity during mitophagy. Dev. Cell 17, 98–109 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B9] 9. Kurihara Y., Kanki T., Aoki Y., Hirota Y., Saigusa T., Uchiumi T., Kang D. (2012) Mitophagy plays an essential role in reducing mitochondrial production of reactive oxygen species and mutation of mitochondrial DNA by maintaining mitochondrial quantity and quality in yeast. J. Biol. Chem. 287, 3265–3272 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B10] 10. Aoki Y., Kanki T., Hirota Y., Kurihara Y., Saigusa T., Uchiumi T., Kang D. (2011) Phosphorylation of serine 114 on Atg32 mediates mitophagy. Mol. Biol. Cell 22, 3206–3217 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B11] 11. Kondo-Okamoto N., Noda N. N., Suzuki S. W., Nakatogawa H., Takahashi I., Matsunami M., Hashimoto A., Inagaki F., Ohsumi Y., Okamoto K. (2012) Autophagy-related protein 32 acts as autophagic degron and directly initiates mitophagy. J. Biol. Chem. 287, 10631–10638 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B12] 12. Kanki T., Kurihara Y., Jin X., Goda T., Ono Y., Aihara M., Hirota Y., Saigusa T., Aoki Y., Uchiumi T., Kang D. (2013) Casein kinase 2 is essential for mitophagy. EMBO Rep. 14, 788–794 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B13] 13. Starheim K. K., Gevaert K., Arnesen T. (2012) Protein N-terminal acetyltransferases: when the start matters. Trends Biochem. Sci. 37, 152–161 [DOI] [PubMed] [Google Scholar]

[B14] 14. Arnesen T. (2011) Towards a functional understanding of protein N-terminal acetylation. PLoS Biol. 9, e1001074. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B15] 15. Eiyama A., Kondo-Okamoto N., Okamoto K. (2013) Mitochondrial degradation during starvation is selective and temporally distinct from bulk autophagy in yeast. FEBS Lett. 587, 1787–1792 [DOI] [PubMed] [Google Scholar]

[B16] 16. Motley A. M., Nuttall J. M., Hettema E. H. (2012) Pex3-anchored Atg36 tags peroxisomes for degradation in Saccharomyces cerevisiae. EMBO J. 31, 2852–2868 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B17] 17. Liszczak G., Goldberg J. M., Foyn H., Petersson E. J., Arnesen T., Marmorstein R. (2013) Molecular basis for N-terminal acetylation by the heterodimeric NatA complex. Nat. Struct. Mol. Biol. 20, 1098–1105 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B18] 18. Asaumi M., Iijima K., Sumioka A., Iijima-Ando K., Kirino Y., Nakaya T., Suzuki T. (2005) Interaction of N-terminal acetyltransferase with the cytoplasmic domain of β-amyloid precursor protein and its effect on Aβ secretion. J. Biochem. 137, 147–155 [DOI] [PubMed] [Google Scholar]

[B19] 19. Gautschi M., Just S., Mun A., Ross S., Rücknagel P., Dubaquié Y., Ehrenhofer-Murray A., Rospert S. (2003) The yeast Nα-acetyltransferase NatA is quantitatively anchored to the ribosome and interacts with nascent polypeptides. Mol. Cell. Biol. 23, 7403–7414 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B20] 20. Polevoda B., Brown S., Cardillo T. S., Rigby S., Sherman F. (2008) Yeast N(α)-terminal acetyltransferases are associated with ribosomes. J. Cell. Biochem. 103, 492–508 [DOI] [PubMed] [Google Scholar]

[B21] 21. Van Damme P., Evjenth R., Foyn H., Demeyer K., De Bock P.-J., Lillehaug J. R., Vandekerckhove J., Arnesen T., Gevaert K. (2011) Proteome-derived peptide libraries allow detailed analysis of the substrate specificities of N(α)-acetyltransferases and point to hNaa10p as the post-translational actin N(α)-acetyltransferase. Mol. Cell. Proteomics 10, M110.004580. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B22] 22. Kirisako T., Baba M., Ishihara N., Miyazawa K., Ohsumi M., Yoshimori T., Noda T., Ohsumi Y. (1999) Formation process of autophagosome is traced with Apg8/Aut7p in yeast. J. Cell Biol. 147, 435–446 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B23] 23. Arnesen T., Van Damme P., Polevoda B., Helsens K., Evjenth R., Colaert N., Varhaug J. E., Vandekerckhove J., Lillehaug J. R., Sherman F., Gevaert K. (2009) Proteomics analyses reveal the evolutionary conservation and divergence of N-terminal acetyltransferases from yeast and humans. Proc. Natl. Acad. Sci. U.S.A. 106, 8157–8162 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B24] 24. Mullen J. R., Kayne P. S., Moerschell R. P., Tsunasawa S., Gribskov M., Colavito-Shepanski M., Grunstein M., Sherman F., Sternglanz R. (1989) Identification and characterization of genes and mutants for an N-terminal acetyltransferase from yeast. EMBO J. 8, 2067–2075 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B25] 25. Polevoda B., Norbeck J., Takakura H., Blomberg A., Sherman F. (1999) Identification and specificities of N-terminal acetyltransferases from Saccharomyces cerevisiae. EMBO J. 18, 6155–6168 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B26] 26. Goetze S., Qeli E., Mosimann C., Staes A., Gerrits B., Roschitzki B., Mohanty S., Niederer E. M., Laczko E., Timmerman E., Lange V., Hafen E., Aebersold R., Vandekerckhove J., Basler K., et al. (2009) Identification and functional characterization of N-terminally acetylated proteins in Drosophila melanogaster. PLos Biol. 7, e1000236. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B27] 27. Aihara M., Jin X., Kurihara Y., Yoshida Y., Matsushima Y., Oku M., Hirota Y., Saigusa T., Aoki Y., Uchiumi T., Yamamoto T., Sakai Y., Kang D., Kanki T. (2014) Tor and the Sin3-Rpd3 complex regulate expression of the mitophagy receptor protein Atg32 in yeast. J. Cell Sci. 127, 3184–3196 [DOI] [PubMed] [Google Scholar]

[B28] 28. Hwang C.-S., Shemorry A., Varshavsky A. (2010) N-terminal acetylation of cellular proteins creates specific degradation signals. Science. 327, 973–977 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B29] 29. Mao K., Wang K., Zhao M., Xu T., Klionsky D. J. (2011) Two MAPK-signaling pathways are required for mitophagy in Saccharomyces cerevisiae. J. Cell Biol. 193, 755–767 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B30] 30. Gomes L. C., Di Benedetto G., Scorrano L. (2011) During autophagy mitochondria elongate, are spared from degradation and sustain cell viability. Nat. Cell Biol. 13, 589–598 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B31] 31. Rambold A. S., Kostelecky B., Elia N., Lippincott-Schwartz J. (2011) Tubular network formation protects mitochondria from autophagosomal degradation during nutrient starvation. Proc. Natl. Acad. Sci. U.S.A. 108, 10190–10195 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B32] 32. Twig G., Elorza A., Molina A. J., Mohamed H., Wikstrom J. D., Walzer G., Stiles L., Haigh S. E., Katz S., Las G., Alroy J., Wu M., Py B. F., Yuan J., Deeney J. T., et al. (2008) Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. EMBO J. 27, 433–446 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B33] 33. Mao K., Wang K., Liu X., Klionsky D. J. (2013) The scaffold protein Atg11 recruits fission machinery to drive selective mitochondria degradation by autophagy. Dev. Cell 26, 9–18 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B34] 34. Böckler S., Westermann B. (2014) Mitochondrial ER contacts are crucial for mitophagy in yeast. Dev. Cell 28, 450–458 [DOI] [PubMed] [Google Scholar]

[B35] 35. Kim J., Kamada Y., Stromhaug P. E., Guan J., Hefner-Gravink A., Baba M., Scott S. V., Ohsumi Y., Dunn W. A., Jr., Klionsky D. J. (2001) Cvt9/Gsa9 functions in sequestering selective cytosolic cargo destined for the vacuole. J. Cell Biol. 153, 381–396 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B36] 36. Brachmann C. B., Davies A., Cost G. J., Caputo E., Li J., Hieter P., Boeke J. D. (1998) Designer deletion strains derived from Saccharomyces cerevisiae S288C: a useful set of strains and plasmids for PCR-mediated gene disruption and other applications. Yeast 14, 115–132 [DOI] [PubMed] [Google Scholar]

[B37] 37. Sikorski R. S., Hieter P. (1989) A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122, 19–27 [DOI] [PMC free article] [PubMed] [Google Scholar]

[B38] 38. Suzuki K., Kirisako T., Kamada Y., Mizushima N., Noda T., Ohsumi Y. (2001) The pre-autophagosomal structure organized by concerted functions of APG genes is essential for autophagosome formation. EMBO J. 20, 5971–5981 [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

Protein N-terminal Acetylation by the NatA Complex Is Critical for Selective Mitochondrial Degradation^*

Akinori Eiyama

Koji Okamoto

Abstract

Introduction