JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M704409200 on August 7, 2007

J. Biol. Chem., Vol. 282, Issue 41, 30239-30245, October 12, 2007
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
282/41/30239    most recent
M704409200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Garrity, J.
Right arrow Articles by Escalante-Semerena, J. C.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Garrity, J.
Right arrow Articles by Escalante-Semerena, J. C.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

N-Lysine Propionylation Controls the Activity of Propionyl-CoA Synthetase*Formula

Jane Garrity{ddagger}1, Jeffrey G. Gardner{ddagger}, William Hawse§2, Cynthia Wolberger§3, and Jorge C. Escalante-Semerena{ddagger}4

From the {ddagger}Department of Bacteriology, University of Wisconsin-Madison, Madison, Wisconsin 53706 and §Department of Biophysics and Biophysical Chemistry, The Johns Hopkins University School of Medicine, Baltimore, Maryland 21205

Received for publication, May 29, 2007 , and in revised form, July 27, 2007.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Reversible protein acetylation is a ubiquitous means for the rapid control of diverse cellular processes. Acetyltransferase enzymes transfer the acetyl group from acetyl-CoA to lysine residues, while deacetylase enzymes catalyze removal of the acetyl group by hydrolysis or by an NAD+-dependent reaction. Propionyl-coenzyme A (CoA), like acetyl-CoA, is a high energy product of fatty acid metabolism and is produced through a similar chemical reaction. Because acetyl-CoA is the donor molecule for protein acetylation, we investigated whether proteins can be propionylated in vivo, using propionyl-CoA as the donor molecule. We report that the Salmonella enterica propionyl-CoA synthetase enzyme PrpE is propionylated in vivo at lysine 592; propionylation inactivates PrpE. The propionyl-lysine modification is introduced by bacterial Gcn-5-related N-acetyltransferase enzymes and can be removed by bacterial and human Sir2 enzymes (sirtuins). Like the sirtuin deacetylation reaction, sirtuin-catalyzed depropionylation is NAD+-dependent and produces a byproduct, O-propionyl ADP-ribose, analogous to the O-acetyl ADP-ribose sirtuin product of deacetylation. Only a subset of the human sirtuins with deacetylase activity could also depropionylate substrate. The regulation of cellular propionyl-CoA by propionylation of PrpE parallels regulation of acetyl-CoA by acetylation of acetyl-CoA synthetase and raises the possibility that propionylation may serve as a regulatory modification in higher organisms.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Protein acetylation is a ubiquitous means for the rapid control of diverse cellular processes (1, 2). Acetylation occurs at lysine residues, with acetyl-CoA (Ac-CoA)5 serving as the acetyl group donor. In higher organisms, aberrant acetylation of lysine residues in histone tails correlates with diseases such as cancers and developmental disorders and may contribute to modulation of cell life span (3, 4). From bacteria to humans, N-Lys acetylation controls the activity of acetyl-coenzyme A synthetase (AMP-forming; Acs) and potentially that of other members of the AMP-forming family of enzymes (5-7). In Salmonella enterica, Acs is deacetylated by CobB, a member of the Sir2 family of NAD+-dependent deacetylases (a.k.a. sirtuins) (8). Interestingly, strains of S. enterica lacking CobB deacetylase activity cannot grow on propionate because the propionyl-CoA synthetase (encoded by the prpE gene) that activates propionate to propionyl-CoA is not active (5, 9).

Cells generate propionyl-CoA from several different processes, including the catabolism of odd chain fatty acids, the decarboxylation of succinate, the catabolism of amino acids (e.g. threonine), and the activation of propionate (10-12). Propionate is a powerful inhibitor of cell growth that is widely used as a food preservative. Reports in the literature suggest that propionyl-CoA may be responsible for the cytotoxic effects of propionate. Although the direct effects of propionyl-CoA are unclear, it is clear that cells avoid accumulating this metabolite (13-15). The cell maintains low levels of propionyl-CoA by using it as a source of carbon and energy. The predominant pathway for propionate degradation in prokaryotes and eukaryotes is the 2-methylcitric acid cycle, which converts propionate to pyruvate via reactions similar to those of the Krebs cycle (supplemental Fig. S6) (16, 17).

Propionylated lysine residues were recently identified in histones (18, 19); by analogy with histone acetylation, propionyl-CoA is presumably required as a propionyl donor. However, the physiological significance of lysine propionylation was unknown. We report here that the propionyl-CoA synthetase (PrpE) enzyme of S. enterica is reversibly propionylated in vivo and that this modification regulates enzymatic activity. The modification is removed by the S. enterica sirtuin CobB in an NAD+-dependent reaction that mirrors the sirtuin deacetylation reaction. Our results suggest that propionylation/depropionylation may be a conserved regulatory mechanism in higher organisms and that acylation/deacylation systems for the control of acyl:CoA (AMP-forming) ligases may be a general mechanism for maintaining CoA homeostasis in all cells.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Protein Purification
Protein Purity Assessment—Proteins were resolved by 12% SDS-PAGE (20) and visualized with Coomassie Blue (21). Purity was assessed by band densitometry using a Fotodyne imaging system with Foto/Analyst v.5.00 software (Fotodyne Inc) for image acquisition and TotalLab v.2005 software for analysis (Nonlinear Dynamics).

S. enterica PrpEWT and PrpEK592E Proteins—These proteins were purified by chitin purification as described (11).

Pat—GST-H6-Protein Acetyltransferase (Pat) was overproduced as described (22). The Pat used in all experiments except the propionylation and depropionylation time courses was purified by glutathione affinity chromatography as described (22) and was 32% homogeneous. Pat used for time courses was purified by affinity chromatography using a 5-ml HisTrap HP column on anÄKTA FPLC Purifier system (Amersham Biosciences). Cells expressing GST-H6-Pat were harvested by centrifugation at 10,500 x g for 12 min in a Beckman Coulter Avanti J-20 XOI refrigerated centrifuge with a JLA-8.1000 rotor. Cells were resuspended in buffer A (sodium phosphate buffer (20 mM, pH 7.5, at 24 °C), NaCl (0.5 M), imidazole (20 mM)) and broken by sonication using a 550 Sonic Dismembrator (Fisher Scientific) for 3 min (50% duty). After equilibration with buffer A and loading of cell-free extract, the column was washed with 50 ml of buffer A, followed by 40 ml of 8% buffer B (sodium phosphate buffer (20 mM, pH 7.5, at 24 °C), NaCl (0.5 M), imidazole (0.5 M)). A 50-ml linear gradient increased buffer B to 100%. GST-H6-Pat eluted at ~40% buffer B and was 67% pure. GST-H6-Pat was stored in tris(hydroxymethyl)aminomethane buffer (Tris-HCl, 50 mM, pH 7.5, at 25 °C) containing KCl (100 mM) and 50% glycerol (v/v) at 4 °C.

CobB Sirtuin—Synthesis of CobB sirtuin fused to an N-terminal chitin binding tag was directed by plasmid pCOBB31 (supplemental Table S1) in Escherichia coli strain ER2566 (New England Biolabs). Cells were grown at 25 °C in two liters of lysogenic broth (23, 24) supplemented with ampicillin (100 µg/ml) and ZnSO4 (50 µM). Gene expression was induced by addition of isopropyl-beta-D-thiogalactoside to 1 mM at A600 ~ 0.4.

Cells were grown overnight at 25 °C, harvested by centrifugation at 10,500 x g for 12 min as described above, and broken using a chilled French pressure cell (Spectronic Instruments; two passes at 1.3 kPa). Protein was purified and the tag removed using the standard protocol for the IMPACT-CN Protein Fusion and Purification System (New England Biolabs). Cell-free extract was incubated with chitin beads for 1 h at 4 °C prior to column preparation. After tag removal, CobB protein was stored in Tris-HCl buffer (50 mM, pH 7.5, at 25 °C) containing KCl (100 mM) and 50% glycerol (v/v) at -80 °C.

PncA Nicotinamidase—S. enterica pncA was amplified using primers to add 5'-KpnI and 3'-HindIII restriction sites and cloned into plasmid pTEV6 cut with the same enzymes to yield plasmid pPNC2 (supplemental Table S1), which encodes PncA protein with an N-terminal maltose-binding protein-hexahistidine (MBP-H6) tag. Plasmid pPNC2 was moved into E. coli strain BL21({lambda}DE3) by electroporation (25). The resulting strain was grown overnight and subcultured 1:100 (v/v) into 2 liters of lysogenic broth containing ampicillin (100 µg/ml). The culture was grown shaking at 37 °C to A600 ~ 0.7, and MBP-H6-PncA synthesis was induced with isopropyl-1-thio-beta-D-galactopyranoside (1 mM). The culture was grown overnight at 25 °C. Cells were harvested and MBP-H6-PncA purified as described for purification of GST-H6-Pat. PncA eluted at ~30% buffer B. MBP-H6-PncA-containing fractions were pooled and H6-rTEV protease (26) added to reach a 1:50 H6-rTEV protease:MBP-H6-PncA ratio; H6-rTEV protease was purified as described (27). The cleavage reaction mixture was incubated at room temperature for 3 h and dialyzed overnight against two liters of buffer A at 4 °C. Tagless PncA protein (83% pure) was resolved from the reaction mixture using the 5-ml HisTrap HP column, which did not bind tagless PncA. Protein was stored in Tris-HCl (50 mM, pH 7.5, at 25 °C) containing KCl (100 mM) and 20% (v/v) glycerol at -80 °C.

S. enterica Acs Peptide—Peptide consisting of the C-terminal 52 amino acids of S. enterica Acs was synthesized by the Peptide Synthesis Facility of the University of Wisconsin-Madison Biotechnology Center. Peptide was purified by preparative scale HPLC using a Dynamax C18 column (22 x 250 mm). Peptide eluted at 36.5% acetonitrile with a final chromatographic purity of 89%.

Human Proteins—Human SIRT1 protein was a gift from John Denu. Human SIRT2 and SIRT3 were overexpressed in E. coli strain BL21({lambda}DE3) transformed with plasmids SIRT2-pHEX and SIRT3-pQE-80, respectively (6, 28). Cells were grown in 1 liter of lysogenic broth supplemented with ampicillin (100 µg/ml) and ZnSO4 (50 µM) at 37 °C to A600 ~ 0.7. Protein expression was induced with isopropyl-1-thio-beta-D-galactopyranoside (1 mM), and cultures were grown overnight at 20 °C. Cells were harvested by centrifugation and broken by sonication, and hSirT2 and hSirT3 were purified using His-Bind Quick 900 cartridges (Novagen) according to the manufacturer's instructions. Proteins were dialyzed into storage buffer (Tris-HCl (50 mM, pH 7.5, at 25 °C) containing 1 mM dithiothreitol and 20% glycerol (v/v)) and stored at -20 °C. hSirT2 protein was 52% pure, and hSirT3 protein was 49% pure.

Other Proteins—Bacillus subtilis AcuA, Thermotoga maritima Sir2, human SIRT4, and murine SIRT1 proteins were purified as described (29-31).

Enzyme Activity Assays
Acylation/Deacylation Assays—Conditions for protein acylation and deacylation have been described (22). PrpE (62.5 pmol) was incubated at 37 °C with GST-H6-Pat (62.5 pmol) and [1-14C]Ac-CoA (20 µM, 54 mCi/mmol) or [1-14C]Pr-CoA (20 µM, 53 mCi/mmol) (Moravek) in 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES) buffer (50 mM, pH 7.5, at 24 °C) containing tris(2-carboxyethyl)phosphine hydrochloride (200 µM); final volume was 100 µl. After 2 h, GST-H6-Pat protein was removed using GST-Mag®-agarose beads (Novagen). CobB sirtuin (15.6 pmol), NAD+ (1 mM), and PncA (50 pmol, when noted) were added and reactions incubated at 37 °C for an additional 2 h. Reactions were stopped with trichloroacetic acid (final concentration 0.5 M) or gel-loading buffer (Tris-HCl buffer (50 mM, pH 6.8, at 24 °C), SDS (2%, w/v), bromphenol blue (0.1%, w/v), glycerol (10%, v/v), dithiothreitol (100 mM)) (32). Proteins were resolved by 12% SDS-PAGE and visualized after staining with Coomassie Blue. Gels were dried and exposed overnight to a MultiPurpose Phosphor Screen (Packard). Radiolabeled proteins were identified by phosphorimaging using a Cyclone Storage Phosphor System (Packard) and OptiQuant v 04.00 software (Packard).

Propionylation Time Courses—Propionylation reactions were performed as described above with either PrpE (62.5 pmol) and GST-H6-Pat (31.3 pmol) or PrpE (37.5 pmol) and AcuA (18.7 pmol) added/25 µl. Samples (25 µl) were removed at designated times and stopped by addition of gel loading buffer. Proteins were separated by 4-15% SDS-PAGE. Radiolabeled proteins were quantified with respect to a standard curve using a Cyclone Storage Phosphor System (Packard) and OptiQuant v 04.00 software (Packard).

Depropionylation Time Courses—Propionylation reactions were performed as described above with PrpE (62.5 pmol) and GST-H6-Pat (31.3 pmol) added/25 µl. After 2 h of incubation at 37 °C, GST-H6-Pat was removed using GST-Mag®-agarose beads (Novagen). Sirtuin (31.3 pmol), NAD+ (1 mM), and PncA (50 pmol) were added/25 µl. Samples (25 µl) were removed at designated times and stopped by addition of gel loading buffer. Radiolabeled proteins were quantitated as described above.

Propionyl-CoA Synthetase—PrpE (62.5 pmol) was incubated with Pat (62.5 pmol) and 20 µM Pr-CoA, Ac-CoA, or CoA for 6 h at 37 °C. Reaction conditions were as described above. PrpE activity was determined using a coupled assay to link AMP formation to NADH oxidation (11, 33). Reactions (1 ml) contained HEPES (50 mM, pH 7.5, at 25 °C), CoA (1 mM), phosphoenol-pyruvate (3 mM), ATP (1 mM), MgCl2 (200 µM), NADH (260 µM), lactate dehydrogenase (10 units; Sigma), pyruvate kinase (10 units; Sigma), myokinase (10 units; Sigma), and PrpE (62.5 pmol). Reactions were started by addition of sodium propionate (20 µM). Absorbance at 340 nm was measured for 20 min using a PerkinElmer Lambda 40 UV-visible spectrophotometer. Specific activities were calculated using the molar extinction coefficient of NADH (6,220 M-1 cm-1) (34); two moles of NADH were oxidized per mole of AMP produced. Specific activities are reported as µmol of AMP produced/min/mg of protein and are averaged from three independent determinations.

In Vivo Modification of PrpE
The prpE gene was amplified from plasmid pPRP68 (35), adding 5'-NheI and 3'-EcoRI sites, and ligated into plasmid pTYB12 (Novagen) cut with SpeI and EcoRI. The resulting plasmid, pPRP211, was used to produce PrpE with an N-terminal chitin-binding domain in strains JE9221 ({Delta}acs {Delta}cobB/pTARA rpo+/pPRP211 prpE+) and JE9225 ({Delta}acs {Delta}cobB pat/pTARA T7 rpo+/pPRP211 prpE+).

Strains JE9221 and JE91125 were grown at 37 °C in 250 ml of no-carbon essential medium (36) supplemented with propionate (30 mM), glycerol (0.5 mM), MgSO4 (1 mM), L-methionine (0.5 mM), ampicillin (100 µg/ml), chloramphenicol (12.5 µg/ml). L(+)-Arabinose (250 µM) was added at A600 ~ 0.6 to induce expression of T7 polymerase from plasmid pTARA and subsequent PrpE production. PrpE was purified by chitin affinity chromatography and resolved by SDS-PAGE. MALDI-TOF peptide fingerprinting was used to identify the modification.

MALDI-TOF Peptide Fingerprinting
In-gel digest and MALDI-TOF analysis of PrpE proteins were performed at the Mass Spectrometry Facility of the University of Wisconsin-Madison Biotechnology Center. PrpE proteins were excised, destained, and dehydrated and then reduced with dithiothreitol (25 mM) for 30 min at 56 °C, alkylated with iodoacetamide (55 mM) in darkness at room temperature for 30 min, and digested with 0.4 µg of trypsin (Promega Sequence Grade Modified) overnight at 37 °C. All steps were performed in (NH4)HCO3 (25 mM, pH 8). Resulting peptides were extracted with 0.1% trifluoroacetic acid (TFA) followed by acetonitrile/H2O/TFA (70%:25%:5%, v/v). Peptides were dried in a SpeedVac concentrator (Thermo), resuspended in 20 µl of TFA (0.1%, v/v), and solid phase extracted using ZipTip® µC18 pipette tips (Millipore). Peptides were eluted off the C18 column with acetonitrile/H2O/TFA (70%:25%:0.2%, v/v) onto an AnchorChipTM plate (Bruker Daltonics) and recrystallized with 1 µl of matrix (20 mg/ml of {alpha}-cyano-4-hydroxycinnamic acid in acetonitrile/H2O/TFA (70%:25%:0.2%, v/v)). Peptide fingerprint analysis was performed on a Bruker BIFLEX III MALDI-TOF mass spectrometer (Bruker Daltonics). Peptide mapping analysis was performed with the Mascot search engine (Matrix Science).

MS/MS analysis was performed at the Mass Spectrometry Facility of the University of Wisconsin-Madison Biotechnology Center using a 4800 MALDI TOF/TOF mass spectrometer (Applied Biosystems) in positive ion mode. Fragment ions were assigned using the Protein Prospector MS-Product utility (University of California, San Francisco Mass Spectrometry Facility).

OPADPR Production
In situ O-propionyl-ADP-ribose (OPADPR) generation reactions contained (per 125 µl): Acs peptide (62.5 µM), AcuA (62.5 µM), CobB (6.25 µM), PncA (3.13 µM), Pr-CoA (1 mM), NAD+ (1 mM), HEPES (50 mM, pH 7.5, at 25 °C), and tris(2-carboxyethyl)phosphine hydrochloride (200 µM). CobB protein was omitted from the control reaction. Reactions were incubated overnight at 37 °C, stopped by the addition of TFA to 1% (v/v), and filtered through 0.45-µm Spin-X® Centrifuge Tube Filters (Corning Inc.) prior to HPLC analysis.

Reaction components were separated using a Beckman Coulter System Gold HPLC system equipped with an Alltima HP C18 AQ column (4.6 x 150-mm, 5-µm pore size; Alltech). The protocol used was based on those described for the purification of O-acetyl-ADP-ribose (37, 38). The system was run at 25 °C at a flow rate of 1 ml/min and monitored at 260 and 214 nm. Following injection of a 50-µl sample, the column was developed isocratically in solvent A (0.05% TFA/H2O) for 6 min. The gradient was increased linearly to 40% solvent B (0.02% TFA/acetonitrile) over 40 min. Fractions of interest were collected and frozen at -80 °C prior to mass spectrometry analysis.

Enhanced mass spectrometry of fractions of interest was performed using an ABI 3200 Q Trap linear ion trap quadrupole liquid chromatography/MS/MS mass spectrometer (AB Sciex Instruments). Sample was directly infused with an infusion syringe containing 50:50 acetonitrile:H2O and was ionized by electrospray ionization with negative polarity.


Figure 1
View larger version (22K):
[in this window]
[in a new window]

  		FIGURE 1.
PrpE is acetylated and propionylated by Pat and deacetylated and depropionylated by CobB. A, Rows I and II, Coomassie Blue-stained SDS-PAGE gel and phosphorimage of PrpEWT protein after incubation with [1-14C]Ac-CoA (20 µM; specific activity = 54 mCi/mmol) in the presence or absence of Pat and CobB sirtuin enzymes. Rows III and IV, Coomassie Blue-stained SDS-PAGE gel and phosphorimage of PrpEK592 protein incubated with [1-14C]Pr-CoA (20 µM; specific activity = 53 mCi/mmol). Rows V and VI, Coomassie Blue-stained SDS-PAGE gel and phosphorimage of PrpEK592 protein incubated with [1-14C]Ac-CoA. For all lanes, + indicates Pat or CobB was added to the reaction; - indicates heat-inactivated Pat or CobB was used as control. B, propionyl-CoA synthetase is propionylated in vivo. Peptide fingerprinting of PrpE protein isolated from strain JE9221 ({Delta}acs {Delta}cobB/pprpE+). The peak at m/z = 747.4 atomic mass units corresponded to S590GKPrMLR. A much smaller peak is seen at m/z = 733.4 atomic mass units, corresponding to S590GKAcMLR. This result was consistent with in vitro propionylation of PrpE. C, peptide fingerprinting of PrpE isolated from JE9225 ({Delta}acs {Delta}cobB pat/pprpE+). Signals corresponding to acetylated or propionylated peptides were not observed.

 


Figure 2
View larger version (12K):
[in this window]
[in a new window]

  		FIGURE 2.
Bacterial Gcn-5-related N-acetyltransferase enzymes propionylate propionyl-CoA synthetase in vitro. A, time course of the propionylation of PrpE (62.5 pmol) by S. enterica Pat (31.3 pmol). Specific activity = 14.3 ± 2.2 pmol PrpE propionylated µmol-1 min-1. B, time course of the propionylation of PrpE (37.5 pmol) by B. subtilis AcuA (18.8 pmol). Specific activity = 78.4 ± 4.2 pmol PrpE propionylated µmol-1 min-1. Time courses were performed in triplicate. Graphs depict the mean and range of values for each time point.

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Previously published work (9) suggested that the Pat acetyltransferase and CobB deacetylase modulated the activity of PrpE in vivo. When tested in vitro, the S. enterica Pat enzyme acetylated or propionylated PrpE, whereas the CobB sirtuin deacetylated PrpEAc or depropionylated PrpEPr (Fig. 1A, lane 2, rows II and IV). Because both PrpE and Acs contain a P-X4-GK motif in which the lysine residue is critical for function (5, 11), we reasoned that this lysine could be the modified residue, by analogy with Acs (5). Failure of Pat to acetylate a PrpEK592E mutant (Fig. 1A, lane 1, rows II and IV) was consistent with the hypothesis that Lys-592 was indeed the modification site of PrpE. This result was confirmed by MALDI-TOF mass spectrometry peptide fingerprinting studies. The mass spectrum of PrpE incubated with Pat and propionyl-CoA, then cut with trypsin, included a signal at m/z = 747.4 atomic mass units, matching the mass of the S590GKPrMLR peptide (supplemental Fig. S7A). Similarly, mass spectrometry of PrpE incubated with Pat and acetyl-CoA gave a signal matching that of the S590GKAcMLR peptide (supplemental Fig. S7B; m/z = 733.5 atomic mass units). Peptides containing Lys-592 were not present in the mass spectrum of tryptic digest of PrpE after incubation with Pat and CoA (supplemental Fig. S7C). Cleavage after Lys-592 should result in the peptides S590GK (m/z = 291.1 atomic mass units) and M593LR (m/z = 419.2 atomic mass units), as long as Lys-592 was not modified. The above-mentioned peptides were not observed. Activity assays showed that propionylation of PrpE resulted in ~70% loss of specific activity (1.5 + 0.3 µmol AMP min-1 mg-1) relative to that of unmodified PrpE enzyme (5 + 0.9 µmol AMP min-1 mg-1).

To determine whether PrpE was posttranslationally modified in vivo, we isolated PrpE protein from strains grown on propionate. The strains used in these experiments were JE9221 ({Delta}acs {Delta}cobB/pprpE+) and JE9225 ({Delta}acs {Delta}cobB pat/pprpE+) (supplemental Table S1). Our strategy was to overproduce PrpE in the absence of the CobB sirtuin to allow Pat-dependent acylation and subsequent accumulation of PrpE in its acylated form. Both strains grew on propionate, albeit poorly, because the acetate kinase/phosphotransacetylase system was functional (9). We did not expect PrpE to be modified in strain JE9225 ({Delta}acs {Delta}cobB pat/pprpE+) because the pat gene was inactive in this strain; hence PrpE from strain JE9225 was used as a negative control. In agreement with this prediction, strain JE9225 grew much better on propionate (doubling time = 5.5 h) than did strain JE9221 (doubling time = 15.8 h), suggesting that Pat inactivated PrpE in strain JE9221 (supplemental Fig. S8).

The MALDI-TOF mass spectrum of tryptic peptides of PrpE isolated from strain JE9221 ({Delta}acs {Delta}cobB/pprpE+) contained a signal at m/z = 747.4 atomic mass units (Fig. 1B). The MS/MS fragmentation pattern of the m/z = 747.4 peptide matched that expected for S590GKPrMLR (supplemental Table S3) as well as that observed for a synthetic SGKPrMLR peptide (supplemental Fig. S9). The peptide fingerprint of PrpE protein isolated from strain JE9225 ({Delta}acs {Delta}cobB pat/pprpE+) matched that of PrpE treated with CoA and did not contain any signals for modified peptides (Fig. 1C). Although a very small peak was seen at m/z = 747.4, MS/MS fragmentation analysis showed it to be an isotope of the m/z = 745.4 peptide (supplemental Fig. S10). These results demonstrated that Pat was responsible for the propionylation of PrpE in vivo.

We investigated whether other acetyltransferase enzymes could propionylate PrpE. The Gcn-5-related N-acetyltransferase enzyme AcuA from B. subtilis (29) displayed propionylation activity with the specific activity of AcuA (78.4 pmol PrpE propionylated min-1 µmol-1) 5.5 times higher than that of Pat (14.3 pmol PrpE propionylated min-1 µmol-1) (Fig. 2).

Because deletion of the cobB gene encoding the S. enterica sirtuin inactivates PrpE in vivo, we asked whether the CobB sirtuin depropionylates substrates in vitro. As predicted, the S. enterica CobB sirtuin was able to remove the propionyl modification from PrpE (specific activity = 3.1 pmol PrpE depropionylated min-1 µmol-1 CobB) (Fig. 1A, lane 1, row IV, and Fig. 3B). We next asked whether other bacterial and eukaryotic sirtuins had depropionylase activity. The human SIRT2 (hSirT2) and SIRT3 (hSirT3) and the bacterial T. maritima Sir2 (Sir2Tm) proteins depropionylated PrpEPr within 1 h (Fig. 3, C-E). Depropionylase activity was not observed with human (hSirT1) or murine SIRT1 (mSirT1) or human SIRT4 (hSirT4) (Fig. 3A, lanes 3, 4, 7). However, mSirT1 displayed robust deacetylase activity (17.6 + 0.4 pmol PrpE deacetylated min-1 µmol-1 mSirT1), suggesting that the lack of depropionylase activity of mSirT1 arises from specific discrimination between acetyl and propionyl lysine rather than being due simply to absence of any enzymatic activity (data not shown).


Figure 3
View larger version (33K):
[in this window]
[in a new window]

  		FIGURE 3.
Depropionylase activity of sirtuins. A, propionylation/depropionylation assays performed by Pat and no sirtuin (lane 1), CobB (lane 2), hSIRT1 (lane 3), mSIRT1 (lane 4), hSIRT2 (lane 5), Sir2Tm (lane 6), or hSIRT4 (lane 7). B-E, time courses of the depropionylation of PrpEPr by sirtuins. PrpE protein (62.5 pmol) was incubated for 2 h with Pat (31.3 pmol) and [1-14C]Pr-CoA. Pat was removed, and the indicated sirtuin (31.3 pmol) was added. The reaction mixture contained NAD+ (1 mM) and nicotinaminidase (PncA) added to reactions. Time courses were performed in triplicate with the exception of the hSirT2 time course, which was performed in duplicate. Graphs depict the mean and range of values for each time point. B, CobB sirtuin (specific activity = 3.08 ± 0.55 pmol PrpEPr depropionylated µmol-1 min-1). C, Sir2Tm (specific activity = 12.2 ± 2.4 pmol PrpEPr depropionylated µmol-1 min-1). D, hSIRT2 (specific activity = 28.0 ± 9.4 pmol PrpEPr depropionylated µmol-1 min-1). E, hSIRT3 (specific activity = 7.97 ± 0.94 pmol PrpEPr depropionylated µmol-1 min-1).

 
By analogy with the deacetylation reaction, we predicted that sirtuin-mediated depropionylation would produce OPADPR. OPADPR has been shown to be the product of the sirtuin-catalyzed reaction when synthetic propionylated peptides were used as substrates (39).

We developed a system for the in situ generation of OPADPR through iterative propionylation and depropionylation of protein substrate. Reaction mixtures contained AcuABs in lieu of Pat because of the former's higher propionylase activity and because its smaller size allowed us to use higher enzyme concentrations. A peptide consisting of the C-terminal 52 amino acids of AcsSe was used as protein substrate because it could be added at higher concentration than PrpE. Results of a control experiment verified that AcuABs propionylated the 52-amino acid peptide (supplemental Fig. S11). To prevent inhibition of the sirtuin activity by nicotinamide, all reaction mixtures contained PncA nicotinamidase (supplemental Fig. S12). Reaction mixtures were resolved by C18 reverse-phase HPLC. The large NAD+ peak (t = 29.5 min) observed in the chromatogram of control mixture lacking CobB sirtuin was absent in the chromatogram of the mixture containing CobB (Fig. 4A, dashed line). The compound eluting 36.5 min post-injection was OPADPR ((M-H)- ion; m/z = 614.1 atomic mass units) as determined by mass spectrometry (Fig. 4B). Reaction byproducts were identified using authentic standards (nicotinic acid, 5.8 min; ADP-ribose, 3.8 min). To catalyze lysine depropionylation in a manner analogous to the sirtuin deacetylation reaction, the enzyme must be able to accommodate the additional methyl group in the Michaelis complex formed with NAD+ and the propionylated peptide. We therefore modeled the complex based on the structure of NAD+ and an acetylated peptide bound to Sir2Tm (40), a bacterial sirtuin that exhibits depropionylation activity (Fig. 3C). As shown in Fig. 5, the structure can readily accommodate the additional methyl of propionyl-lysine in a hydrophobic pocket in the enzyme active site. A model constructed with no adjustments produced minimal clashes (van der Waals distances of 3.2 Å or greater) that were completely eliminated during energy minimization by minor side-chain rearrangements, with individual atomic shifts of less than 0.2 Å, in the vicinity of the propionyl-lysine.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Taken together, the data presented here support the conclusion that N-Lys propionylation is a physiologically relevant posttranslational modification for the control of protein function. Because propionylase activity was detectable in two bacterial Gcn-5-related N-acetyltransferase enzymes (S. enterica Pat, B. subtilis AcuA), we suggest that members of this family of enzymes in higher organisms may also have propionylase activity. Similarly, robust depropionylase activity is found among sirtuins from bacteria and higher organisms (Fig. 3), suggesting that other members of this enzyme family may also catalyze this activity. However, because this activity was not detected in human or murine SIRT1 or human SIRT4 enzymes, the ability to accommodate the larger propionyl modification in the enzyme active site appears to be enzyme-specific. Consistent with the current understanding of the CobB sirtuin-catalyzed deacetylase reaction, the product of the depropionylase activity of CobB was OPADPR (Fig. 4). It is unclear whether OPADPR plays any physiological role as has been suggested for the related molecule O-acetyl-ADP-ribose (41-43).


Figure 4
View larger version (11K):
[in this window]
[in a new window]

  		FIGURE 4.
ADP-ribose and OPADPR are produced during CobB-mediated depropionylation. A, reverse-phase HPLC chromatograms of reaction mixture containing S. enterica Acs peptide, B. subtilis AcuA, S. enterica CobB, S. enterica PncA, Pr-CoA, and NAD+ (solid line) and of the same mixture lacking CobB (dashed line). Both analyses were performed following overnight incubation at 37 °C. Elution was monitored at 260 nm. ADP-ribose, NAD+, and nicotinic acid peaks were identified by comparison to authentic standards. B, mass spectrum of the products of the reaction mixture containing CobB, collected from 34.5 to 37.0 min. The m/z = 614.2 atomic mass unit signal corresponded to the signal expected for the (M-H)- ion of OPADPR.

 


Figure 5
View larger version (46K):
[in this window]
[in a new window]

  		FIGURE 5.
Model of propionyl-lysine and NAD+ bound in the Sir2Tm active site. A model of propionyl-lysine in the active site of a sirtuin was modeled based on the structure of the ternary complex formed by the bacterial sirtuin Sir2Tm with NAD+ and an acetylated peptide (Protein Data Bank code 2H4F) (40). The acetyl group was replaced with the propionyl group and the structure modeled using the molecular graphics program Quanta (2006 edition). Initial fits were achieved with minimal clashing, but an improved model was obtained from a minimization calculation performed in Quanta using the CHARMM force field. Only minor rearrangements of side chains within the Sir2Tm active site were needed to accommodate the additional methyl in the propionyl group. The propionylated peptide fit into a small hydrophobic pocket comprised of the Leu-100, Leu-159, and Phe-148 side chains and backbone atoms of Val-160 and His-116.

 
From a physiological standpoint, propionylation is a logical means of posttranslational regulation because it allows the cell to rapidly modulate activity of propionyl-CoA synthetase in direct response to potentially deleterious levels of its product, propionyl-CoA. In contrast, control of propionyl-CoA synthetase activity by acetylation would require conversion of propionyl-CoA to acetyl-CoA via rate-limiting steps of the 2-methylcitric acid cycle (supplemental Fig. S6) (16). This strategy would make inactivation of propionyl-CoA synthetase too slow to prevent any imbalance in CoA homeostasis or to prevent other toxic effects caused by high levels of propionyl-CoA (13). The cell therefore senses propionyl-CoA imbalance directly using the same enzyme pair used to control acetyl-CoA homeostasis.

From a broader physiological perspective, all cells must control their pools of acyl-CoAs to avoid depletion of the pool of free CoA and/or synthesis of toxic metabolites (13, 44). This suggests, by analogy with the work reported here and with earlier findings regarding acetyl CoA homeostasis (8), that there might well be other acyltransferase/deacylase systems that cells from all domains of life use to control the activity of acyl-CoA synthetases. For example, a succinyl-CoA:protein succinyltransferase enzyme might use succinyl-CoA to inactivate the succinate:CoA ligase, while a cognate desuccinylase would reactivate succinyl-CoA ligase. Whether these modifications indeed exist in the cell and, like acetylation, are used to regulate other processes is an intriguing possibility under active investigation.


   FOOTNOTES
 
* This work was supported in part by Public Health Service (PHS) Grant GM62203 (to J. C. E.-S.), by Subaward 8412-76121-8 (to J. C. E.-S.) and PHS Grant U54 RR020839. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

Formula The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. S6-S12, supplemental Tables S1-S3, and supplemental references and "Experimental Procedures." Back

1 Supported in part by PHS Biotechnology Training Grant T32 GM08349 and a Howard Hughes Medical Institute (HHMI) predoctoral fellowship. Back

2 Supported by National Science Foundation (NSF) Grant MCB-0220191. Back

3 Supported by HHMI and NSF Grant MCB-0220191. Back

4 To whom correspondence should be addressed: Dept. of Bacteriology, University of Wisconsin, 1550 Linden Dr., Madison, WI 53706. Tel.: 608-262-7379; Fax: 608-265-7909; E-mail: escalante{at}bact.wisc.edu.

5 The abbreviations used are: Ac-CoA, acetyl-coenzyme A; TFA, trifluoroacetic acid; GST-H6-, glutathione S-transferase hexahistidine tag; MBP-H6-, maltose-binding protein hexahistidine tag; H6-rTEV, C-terminal tag, hexahistine-tagged recombinant tobacco etch virus; HPLC, high pressure liquid chromatography; Pat, protein acetyltransferase; MALDI-TOF, matrix-assisted laser desorption ionization time-of-flight; MS, mass spectrometry; Acs, acetyl-coenzyme A synthetase; OPADPR, O-propionyl-ADP-ribose. Back


   ACKNOWLEDGMENTS
 
We thank J. Denu for the gift of plasmids SIRT2-pHEX and SIRT3-pQE-80 and hSirT1 protein.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Walsh, C. T., Garneau-Tsodikova, S., and Gatto, G. J., Jr. (2005) Angew. Chem. Int. Ed. Engl. 44, 7342-7372[CrossRef]
  2. Clark, R. S., Bayir, H., and Jenkins, L. W. (2005) Crit. Care Med. 33, Suppl. 12 S407-S409[CrossRef][Medline] [Order article via Infotrieve]
  3. Guarente, L., and Picard, F. (2005) Cell 120, 473-482[CrossRef][Medline] [Order article via Infotrieve]
  4. Timmermann, S., Lehrmann, H., Polesskaya, A., and Harel-Bellan, A. (2001) Cell Mol. Life Sci. 58, 728-736[CrossRef][Medline] [Order article via Infotrieve]
  5. Starai, V. J., Celic, I., Cole, R. N., Boeke, J. D., and Escalante-Semerena, J. C. (2002) Science 298, 2390-2392[Abstract/Free Full Text]
  6. Hallows, W. C., Lee, S., and Denu, J. M. (2006) Proc. Natl. Acad. Sci. U. S. A. 103, 10230-10235[Abstract/Free Full Text]
  7. Schwer, B., Bunkenborg, J., Verdin, R. O., Andersen, J. S., and Verdin, E. (2006) Proc. Natl. Acad. Sci. U. S. A. 103, 10224-10229[Abstract/Free Full Text]
  8. Sauve, A. A., Wolberger, C., Schramm, V. L., and Boeke, J. D. (2006) Annu. Rev. Biochem. 75, 435-465[CrossRef][Medline] [Order article via Infotrieve]
  9. Starai, V. J., Takahashi, H., Boeke, J. D., and Escalante-Semerena, J. C. (2003) Genetics 163, 545-555[Abstract/Free Full Text]
  10. Sawers, G. (2001) Mol. Microbiol. 39, 1285-1298[CrossRef][Medline] [Order article via Infotrieve]
  11. Horswill, A. R., and Escalante-Semerena, J. C. (2002) Biochemistry 41, 2379-2387[CrossRef][Medline] [Order article via Infotrieve]
  12. Haller, T., Buckel, T., Rétey, J., and Gerlt, J. A. (2000) Biochemistry 39, 4622-4629[CrossRef][Medline] [Order article via Infotrieve]
  13. Horswill, A. R., Dudding, A. R., and Escalante-Semerena, J. C. (2001) J. Biol. Chem. 276, 19094-19101[Abstract/Free Full Text]
  14. Brock, M., and Buckel, W. (2004) Eur. J. Biochem. 271, 3227-3241[Medline] [Order article via Infotrieve]
  15. Stumpf, D. A., McAfee, J., Parks, J. K., and Eguren, L. (1980) Pediatr. Res. 14, 1127-1131[Medline] [Order article via Infotrieve]
  16. Horswill, A. R., and Escalante-Semerena, J. C. (2001) Biochemistry 40, 4703-4713[CrossRef][Medline] [Order article via Infotrieve]
  17. Miyakoshi, S., Uchiyama, H., Someya, T., Satoh, T., and Tabuchi, T. (1987) Agric. Biol. Chem. 51, 2381-2387
  18. Chen, Y., Sprung, R., Tang, Y., Ball, H., Sangras, B., Kim, S., Falck, J. R., Peng, J., Gu, W., and Zhao, Y. (2007) Mol. Cell. Proteomics
  19. Kim, S. C., Sprung, R., Chen, Y., Xu, Y., Ball, H., Pei, J., Cheng, T., Kho, Y., Xiao, H., Xiao, L., Grishin, N. V., White, M., Yang, X. J., and Zhao, Y. (2006) Mol. Cell 23, 607-618[CrossRef][Medline] [Order article via Infotrieve]
  20. Laemmli, U. K. (1970) Nature 227, 680-685[CrossRef][Medline] [Order article via Infotrieve]
  21. Sasse, J. (1991) in Current Protocols in Molecular Biology (Ausubel, F. A., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K., eds) 10.6.1-10.6.25, Wiley Interscience, New York
  22. Starai, V. J., and Escalante-Semerena, J. C. (2004) J. Mol. Biol. 340, 1005-1012[CrossRef][Medline] [Order article via Infotrieve]
  23. Bertani, G. (1951) J. Bacteriol. 62, 293-300[Free Full Text]
  24. Bertani, G. (2004) J. Bacteriol. 186, 595-600[Free Full Text]
  25. Inoue, H., Nojima, H., and Okayama, H. (1990) Gene 96, 23-28[CrossRef][Medline] [Order article via Infotrieve]
  26. Shih, Y. P., Wu, H. C., Hu, S. M., Wang, T. F., and Wang, A. H. (2005) Protein Sci. 14, 936-941[Abstract/Free Full Text]
  27. Brinsmade, S. R., and Escalante-Semerena, J. C. (2007) J. Biol. Chem. 282, 12629-12640[Abstract/Free Full Text]
  28. North, B. J., Marshall, B. L., Borra, M. T., Denu, J. M., and Verdin, E. (2003) Mol. Cell 11, 437-444[CrossRef][Medline] [Order article via Infotrieve]
  29. Gardner, J. G., Grundy, F. J., Henkin, T. M., and Escalante-Semerena, J. C. (2006) J. Bacteriol. 188, 5460-5468[Abstract/Free Full Text]
  30. Smith, J. S., Avalos, J., Celic, I., Muhammad, S., Wolberger, C., and Boeke, J. D. (2002) Methods Enzymol. 353, 282-300[Medline] [Order article via Infotrieve]
  31. Haigis, M. C., Mostoslavsky, R., Haigis, K. M., Fahie, K., Christodoulou, D. C., Murphy, A. J., Valenzuela, D. M., Yancopoulos, G. D., Karow, M., Blander, G., Wolberger, C., Prolla, T. A., Weindruch, R., Alt, F. W., and Guarente, L. (2006) Cell 126, 941-954[CrossRef][Medline] [Order article via Infotrieve]
  32. Sambrook, J., and Russell, D. W. (2001) Molecular Cloning: A Laboratory Manual, 3rd Ed., p. A1.20 Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  33. Williamson, J. R., and Corkey, B. E. (1969) in Methods in Enzymology (Lowenstein, J. M., ed) pp. 434-513, Academic Press, New York
  34. Dawson, R. M. C., Elliott, D. C., Elliott, W. H., and Jones, K. M. (1986) Data for Biochemical Research, 3rd Ed., pp. 122-123, Oxford University Press, Oxford
  35. Horswill, A. R., and Escalante-Semerena, J. C. (1999) Microbiology 145, 1381-1388[Abstract]
  36. Berkowitz, D., Hushon, J. M., Whitfield, H. J., Jr., Roth, J., and Ames, B. N. (1968) J. Bacteriol. 96, 215-220[Abstract/Free Full Text]
  37. Jackson, M. D., and Denu, J. M. (2002) J. Biol. Chem. 277, 18535-18544[Abstract/Free Full Text]
  38. Tanner, K. G., Landry, J., Sternglanz, R., and Denu, J. M. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 14178-14182[Abstract/Free Full Text]
  39. Smith, B. C., and Denu, J. M. (2007) J. Am. Chem. Soc. 129, 5802-5803[CrossRef][Medline] [Order article via Infotrieve]
  40. Hoff, K. G., Avalos, J. L., Sens, K., and Wolberger, C. (2006) Structure 14, 1231-1240[Medline] [Order article via Infotrieve]
  41. Liou, G. G., Tanny, J. C., Kruger, R. G., Walz, T., and Moazed, D. (2005) Cell 121, 515-527[CrossRef][Medline] [Order article via Infotrieve]
  42. Kustatscher, G., Hothorn, M., Pugieux, C., Scheffzek, K., and Ladurner, A. G. (2005) Nat. Struct. Mol. Biol. 12, 624-625[CrossRef][Medline] [Order article via Infotrieve]
  43. Grubisha, O., Rafty, L. A., Takanishi, C. L., Xu, X., Tong, L., Perraud, A. L., Scharenberg, A. M., and Denu, J. M. (2006) J. Biol. Chem. 281, 14057-14065[Abstract/Free Full Text]
  44. Brass, E. P., Tahiliani, A. G., Allen, R. H., and Stabler, S. P. (1990) J. Nutr. 120, 290-297[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Bacteriol.Home page
J. G. Gardner and J. C. Escalante-Semerena
Biochemical and Mutational Analyses of AcuA, the Acetyltransferase Enzyme That Controls the Activity of the Acetyl Coenzyme A Synthetase (AcsA) in Bacillus subtilis
J. Bacteriol., July 15, 2008; 190(14): 5132 - 5136.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Supplemental Data
Right arrow All Versions of this Article:
282/41/30239    most recent
M704409200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Garrity, J.
Right arrow Articles by Escalante-Semerena, J. C.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Garrity, J.
Right arrow Articles by Escalante-Semerena, J. C.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2007 by the American Society for Biochemistry and Molecular Biology.