The Pennsylvania State University

The Graduate School

Eberly College of Science

REGULATION OF MITOCHONDRIAL TRANSLATION AND OXIDATIVE

PHOSPHORYLATION THROUGH REVERSIBLE ACETYLATION

A Dissertation in

Biochemistry, Microbiology and Molecular Biology

by

Hüseyin Çimen

2012 Hüseyin Çimen

Submitted in Partial Fulfillment of the Requirements

for the Degree of

Doctor of Philosophy

August 2012

The Dissertation of Hüseyin Çimen was reviewed and approved* by the following:

Emine C. Koc Assistant Professor of Biochemistry and Molecular Biology Dissertation Co-adviser Co-chair of Committee

Hasan Koc Assistant Professor of Natural Sciences Dissertation Co-adviser Co-chair of Committee

Craig E. Cameron Paul Berg Professor of Biochemistry and Molecular Biology Associate Department Head for Research and Graduate Education

Joseph C. Reese Professor of Biochemistry and Molecular Biology

Teh-hui Kao Professor of Biochemistry and Molecular Biology

Tae-Hee Lee Assistant Professor of Chemistry and the Huck Institute of the Life Sciences

Craig E. Cameron Paul Berg Professor of Biochemistry and Molecular Biology Associate Department Head of the Department of Biochemistry and Molecular Biology

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ABSTRACT

In a eukaryotic cell, mitochondria provide energy in the form of ATP through

oxidative phosphorylation (OXPHOS), which consists of five electron transport chain

complexes embedded in the inner membrane of mitochondria. Human mitochondria have

their own genome and transcription/translation system to synthesize mitochondrially

encoded thirteen proteins of respiratory chain complexes. We investigated how

acetylation of ribosomal proteins regulates translation and energy production in

mitochondria since reversible acetylation of mitochondrial proteins was found to be

critical for maintaining energy homeostasis. We identified mitochondrial ribosomal

protein L10 (MRPL10) as the major acetylated ribosomal protein in mammalian

mitochondria with two-dimensional gel electrophoresis followed by tandem mass

spectrometry and immunoblotting analyses. In addition, we discovered that SIRT3, which

is the main NAD+-dependent deacetylase localized into mitochondria, interacts with the

ribosome and is responsible for the deacetylation of MRPL10. MRPL10 is a member of

L7/L12 stalk, which is essential for translation since this stalk region stimulates the

activity of translation factors. We employed SIRT3 knock-out mice in order to study the

mechanism of reversible acetylation of MRPL10. The acetylation of MRPL10 resulted in

increased MRPL12 binding to the L7/L12 stalk accompanied by enhanced protein

synthesis in our in vitro translation assays. Moreover, HIB1B, a brown adipocyte tissue

cell line stably overexpressing SIRT3, demonstrated reduction in the acetylation of

MRPL10 and decreased MRPL12 binding to the L7/L12 stalk. By using [35S]-methionine

pulse-labeling assays, we revealed that the mitochondrial protein synthesis was

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suppressed in these cells overexpressing SIRT3. Diminished synthesis of mitochondrial-

encoded protein subunits of respiratory chain complexes resulted in reduced activities of

Complex I and IV and total ATP production. Overall, these findings define a possible

mechanism by which SIRT3-dependent reversible acetylation of MRPL10 and binding of

MRPL12 to the ribosome regulates the mitochondrial protein synthesis and, therefore,

modulates the OXPHOS and ATP production. In the next chapter, the discovery of

another novel SIRT3 substrate, the flavoprotein (SdhA) subunit of Complex II, succinate

dehydrogenase, was demonstrated. We identified and assessed the acetylation of the

SdhA subunit, which resulted in reduced activity of Complex II in SIRT3 knock-out

mice. Due to their location in the enzyme, the acetylated lysine residues may induce

conformational changes to the active site of the enzyme in order to regulate its activity.

Next, the advancement in the available expressed sequence tag (ESTs) databases from

different organisms and the improved sensitivity of mass spectrometry-based proteomic

studies encouraged us to reevaluate the protein components of the mammalian

mitochondrial ribosome. In our analyses, we identified three additional members of the

mitochondrial ribosome; CHCHD1, AURKAIP1, and CRIF1. We found that siRNA

mediated knockdown of the newly identified ribosomal proteins to impair mitochondrial

protein synthesis as determined by [35S]-methionine pulse-labeling assays.

Overall, these biochemical and proteomic studies identified novel acetylated

targets, MRPL10 and SdhA, for SIRT3. Given that the components of mitochondrial

translation are crucial in the synthesis of respiratory chain subunits, the newly identified

ribosomal proteins in addition to acetylation of MRPL10 and SdhA provide a more

v

complete picture of mitochondrial translation and regulation of energy production in the

cell.

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TABLE OF CONTENTS

LIST OF FIGURES ..................................................................................................... ix 

LIST OF TABLES ....................................................................................................... xii 

ABBREVIATIONS ..................................................................................................... xiii 

ACKNOWLEDGMENTS ........................................................................................... xv 

Chapter 1 Introduction ................................................................................................ 1 

1.1 Mitochondrion ................................................................................................ 1 1.2 Mitochondrial Genome ................................................................................... 4 1.3 Oxidative Phosphorylation ............................................................................. 7 1.4 Mitochondrial Translation .............................................................................. 8 

1.4.1 Initiation ............................................................................................... 8 1.4.2 Elongation ............................................................................................. 9 1.4.3 Termination .......................................................................................... 10 

1.5 Mitochondrial Ribosome ................................................................................ 12 1.5.1 Small Subunit (28S) of Mammalian Mitochondrial Ribosome ............ 15 1.5.2 Large Subunit (39S) of Mammalian Mitochondrial Ribosome ............ 15 1.5.3 New Mitochondrial Ribosomal Proteins and Additional Functions ..... 17 

1.6 Post-Translational Modifications of Proteins ................................................. 20 1.6.1 Lysine Acetyltransferases (KATs) and Deacetylases (KDACs) .......... 20 1.6.2 Mitochondrial Acetyltransferases and Sirtuins .................................... 23 1.6.3 Post-Translational Modifications of Mitochondrial Ribosomal

Proteins ................................................................................................... 28 1.7 Research Aims ................................................................................................ 29 1.8 References ....................................................................................................... 32 

Chapter 2 Regulation of Mitochondrial Translation by SIRT3 .................................. 39 

2.1 Rationale ......................................................................................................... 40 2.2 Introduction ..................................................................................................... 41 2.3 Materials and Methods ................................................................................... 45 

2.3.1 Mitochondrial Ribosome Preparation and Reverse Phase – High Performance Liquid Chromatography (RP-HPLC). ............................... 45 

2.3.2 Mass Spectrometric Analysis of Bovine Mitochondrial Ribosomal Proteins ................................................................................................... 46 

2.3.3 In Vitro Deacetylation and Translation Assays .................................... 47 2.3.4 Mouse Mitochondrial Ribosome Isolation ........................................... 48 2.3.5 Immunoblotting Assays ........................................................................ 49 2.3.6 Plasmid Constructs ............................................................................... 50 2.3.7 Cell Culture .......................................................................................... 51 

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2.3.8 [35S]-Methionine Pulse-Labeling Assays ............................................. 52 2.3.9 Preparation of Mitochondrial Ribosomes from Cell Lines .................. 52 2.3.10 Complex I and IV Activity and ATP Determination Assays ............. 53 2.3.11 Expression and Purification MRPL10-MRPL12 Complex and

Reconstitution of Hybrid Ribosome ....................................................... 55 2.3.12 Statistical Analysis ............................................................................. 56 

2.4 Results and Discussion .................................................................................. 57 2.4.1 The Mitochondrial Ribosomal Protein MRPL10 is Acetylated and

SIRT3 is Responsible for its Deacetylation ........................................... 57 2.4.2 SIRT3, NAD+-dependent Deacetylase is Associated with 55S

Mitochondrial Ribosome. ....................................................................... 66 2.4.3 Recombinant and Ribosome-Associated Endogenous NAD+-

dependent SIRT3 Deacetylates MRPL10 .............................................. 74 2.4.4 Overexpression of SIRT3 Regulates Mitochondrial Protein

Synthesis in HIB1B Cells ....................................................................... 85 2.4.5 Effect of SIRT3 Overexpression on MRPL12 Binding to the

Ribosome in HIB1B Cells ...................................................................... 87 2.4.6 The Effect of MRPL12 Knockdown and MRPL10 and MRPL12

Overexpression on Mitochondrial Protein Synthesis. ............................ 94 2.4.7 The Role of Acetylated Lysine Residues of MRPL10 on

Mitochondrial Translation and Cell Growth In Vivo. ............................ 98 2.4.8 The Role of MRPL10 Reversible Acetylation in the Composition

of Ribosomal L7/L12 Stalk and Mitochondrial Translation In Vitro. .... 104 2.5 Conclusions and Future Directions ................................................................. 111 2.6 Acknowledgment ............................................................................................ 116 2.7 References ....................................................................................................... 116 

Chapter 3 Regulation of Succinate Dehydrogenase Activity by SIRT3 ..................... 121 

3.1 Rationale ......................................................................................................... 122 3.2 Introduction ..................................................................................................... 123 3.3 Materials and Methods ................................................................................... 125 

3.3.1 Isolation of Mitochondria from Mice Liver and Enrichment of Complex II .............................................................................................. 125 

3.3.2 Two Dimensional-Gel and Immunoblotting Analysis ......................... 126 3.3.3 Mass Spectrometric Identification and Mapping of Acetylation

Sites ........................................................................................................ 127 3.3.4 Cell Culture .......................................................................................... 128 3.3.5 Complex II Enzymatic Activity Assay ................................................. 129 3.3.6 Statistical Analysis ............................................................................... 129 

3.4 Results and Discussion ................................................................................... 130 3.4.1 Succinate Dehydrogenase is Acetylated and SIRT3 is Responsible

for its Deacetylation ............................................................................... 130 3.4.2 Role of Hyperacetylation of SdhA in Complex II Activity .................. 139 

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3.4.3 Role of Increased Level of SIRT3 Expression in Deaceylation of SdhA and Complex II Activity ............................................................... 140 

3.5 Conclusions and Future Directions ................................................................. 146 3.6 Acknowledgment ............................................................................................ 149 3.7 References ....................................................................................................... 150 

Chapter 4 Identification and Analysis of New Mammalian Mitochondrial Ribosomal Proteins: CHCHD1, AURKAIP1, and CRIF1 ................................... 152 

4.1 Rationale ......................................................................................................... 153 4.2 Introduction ..................................................................................................... 154 4.3 Materials and Methods ................................................................................... 156 

4.3.1 Preparation of Bovine Mitochondrial Ribosomal Subunits .................. 156 4.3.2 Identification of Mitochondrial Ribosomal Proteins by Mass

Spectrometry .......................................................................................... 157 4.3.3 Preparation of Crude Ribosomes from Human Cell Lines and

Isolated Mitochondria ............................................................................ 158 4.3.4 RNase A Treatment of Mitochondrial Ribosomes ............................... 160 4.3.5 Immunoblotting Analysis ..................................................................... 160 4.3.6 [35S]-Methionine Labeling of Mitochondrial Translation Products

In Vivo .................................................................................................... 161 4.3.7 Reverse Transcription Polymerase Chain Reaction (RT-PCR) ............ 162 

4.4 Results and Discussion ................................................................................... 164 4.4.1 Identification of 55S Ribosomal Proteins by Tandem Mass

Spectrometry .......................................................................................... 164 4.4.2 Subunit Assignments of Newly Identified Ribosome-associated

Proteins ................................................................................................... 169 4.4.3 Localization of CHCHD1, AURKAIP1, and CRIF1 into the

Mitochondria and Their Roles in Mitochondrial Translation ................ 177 4.5 Conclusions and Future Directions ................................................................. 192 4.6 Acknowledgment ............................................................................................ 195 4.7 References ....................................................................................................... 195 

Chapter 5 Concluding Remarks and Future Directions .............................................. 199 

5.1 References ....................................................................................................... 206 

Appendix A The Effect of GCN5L1 Knockdown in Hep3B and HIB1B Cells. ........ 209 

Appendix B Strong-Cation Exchange (SCX) Chromatography Purification of Mitochondrial MRPL10-MRPL12 Stalk Complex .............................................. 215 

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LIST OF FIGURES

Figure 1.1: Mammalian mitochondrion. ...................................................................... 3 

Figure 1.2: Products of mitochondrial genome and oxidative phosphorylation. ......... 6 

Figure 1.3: Model for different stages of mitochondrial translation. ........................... 11 

Figure 1.4: The 3D cryo-EM reconstruction mitochondrial ribosomes. ...................... 13 

Figure 1.5: Model representing the mitochondrial L7/L12 stalk. ................................ 19 

Figure 1.6: Substrates and biological functions of sirtuins and mitochondrial acetyltransferase(s). .............................................................................................. 26 

Figure 2.1: Detection of acetylated mitochondrial ribosomal proteins. ....................... 59 

Figure 2.2: Purification of MRPL10 and its acetylated forms by reverse phase -HPLC. ................................................................................................................... 62 

Figure 2.3: Alignment of MRPL10 with identified acetylated lysine residues. .......... 64 

Figure 2.4: Structural model for the location of acetylated lysine residues on ribosome and ribosomal L7/L12 stalk region. ...................................................... 65 

Figure 2.5: The association of SIRT3 with mitochondrial 55S ribosomes. ................. 68 

Figure 2.6: Dissociation of SIRT3 from the large subunit of mitochondrial ribosomes by RNase A treatment. ........................................................................ 69 

Figure 2.7: Structural model of the SIRT3 and MRPL10 interactions in the L7/L12 stalk. ......................................................................................................... 72 

Figure 2.8: Deacetylation of MRPL10 by the NAD+-dependent deacetylase, SIRT3. ................................................................................................................... 76 

Figure 2.9: Role of MRPL10 acetylation in SIRT3 knock-out (Sirt3-/-) mice. ............ 77 

Figure 2.10: Role of nicotinamide (NAM) and emetine on mitochondrial protein synthesis. ............................................................................................................... 79 

Figure 2.11: Enhanced MRPL12 binding to the ribosome in SIRT3 knock-out mice. ...................................................................................................................... 81 

Figure 2.12: Role of MRPL10 acetylation in mitochondrial translation in SIRT3 knock-out mice. .................................................................................................... 82 

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Figure 2.13: Identification of two different forms of MRPL12. .................................. 84 

Figure 2.14: Role of SIRT3 over-expression on mitochondrial protein synthesis in HIB1B cells. ......................................................................................................... 86 

Figure 2.15: Effect of SIRT3 over-expression on MRPL10 acetylation and MRPL12 binding to the ribosome in HIB1B cells. .............................................. 88 

Figure 2.16: Complex I activity in HIB1B cells overexpressing SIRT3. .................... 90 

Figure 2.17: Effect of SIRT3 overexpression on Complex IV activity. ...................... 91 

Figure 2.18: ATP production in HIB1B cells overexpressing SIRT3. ........................ 92 

Figure 2.19: Citrate synthase activity in HIB1B cells overexpressing SIRT3. ........... 93 

Figure 2.20: Effect of MRPL12 knockdown and MRPL10 and MRPL12 overexpressions on mitochondrial protein synthesis. ........................................... 96 

Figure 2.21: Effect of MRPL12 knockdown and MRPL10 and MRPL12 overexpression on ribosomal L7/L12 stalk composition and OXPHOS subunits. ................................................................................................................ 97 

Figure 2.22: The effect of MRPL10 LysAla, Gln, and Arg mutants on mitochondrial protein synthesis. ........................................................................... 102 

Figure 2.23: The effect of MRPL10 LysAla, Gln, and Arg mutants on cell growth. .................................................................................................................. 103 

Figure 2.24: Strong-cation exchange (SCX) chromatography purification of mitochondrial MRPL10-MRPL12 stalk complex. ............................................... 106 

Figure 2.25: Poly(U)-directed in vitro translation assays using hybrid ribosomes. ..... 110 

Figure 3.1A-B: Detection of SdhA as a novel SIRT3 substrate in SIRT3 knock-out mice liver mitochondria. ................................................................................. 131 

Figure 3.1C: Identification of SdhA acetylation in SIRT3 knock-out mice liver mitochondria. ........................................................................................................ 132 

Figure 3.2A: The collision-induced dissociation (CID) spectrum of the acetylated peptide. .................................................................................................................. 136 

Figure 3.2B: Primary sequence alignment of acetylated peptides from mice SdhA and its homologs from different species. .............................................................. 137 

Figure 3.2C: Crystal structure model of the chicken SdhA. ........................................ 138 

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Figure 3.3: Regulation of succinate dehydrogenase (Complex II) activity by deacetylation of SdhA. .......................................................................................... 142 

Figure 3.4A-B: Effect of SIRT3 overexpression on SdhA deacetylation in K562 cells. ...................................................................................................................... 144 

Figure 3.4C: Effect of SIRT3 overexpression on Complex II activity in K562 cells. ...................................................................................................................... 145 

Figure 4.1: An experimental scheme for the purification of mitochondrial ribosomes and 28S and 39S subunits. ................................................................... 166 

Figure 4.2: SDS-PAGE analyses of purified bovine 55S ribosomes and 28S and 39S subunits. ......................................................................................................... 167 

Figure 4.3: Mitochondrial ribosomes prepared at different salt and detergent concentrations. ...................................................................................................... 168 

Figure 4.4: Sedimentation of new MRPs with large complexes in human cell lines and mitochondria. ................................................................................................. 179 

Figure 4.5: Distribution of new MRPs in 28S and 39S subunits. ................................ 182 

Figure 4.6: RNA-dependent association of new MRPs with the 55S ribosome. ......... 183 

Figure 4.7A: siRNA mediated knock-down of new MRPs. ........................................ 186 

Figure 4.7B: RT-PCR analysis of mitochondrial- and nuclear-encoded transcripts. .. 187 

Figure 4.8: Effect of new MRPs knock-down on mitochondrial protein synthesis. .... 188 

Figure 4.9: Effect of new MRPs knock-down on OXPHOS subunits. ........................ 189 

Figure 5.1: Model for the role of MRPL10 acetylation on MRPL12 binding to ribosome. .............................................................................................................. 201 

Figure A1.1: The effect of GCN5L1 knockdown on acetylation and OXPHOS subunits in Hep3B cells. ....................................................................................... 212 

Figure A1.2: The effect of GCN5L1 knockdown on mitochondrial protein synthesis in Hep3B cells. ...................................................................................... 213 

Figure A2.1: Strong-cation exchange (SCX) chromatography purification of mitochondrial MRPL10-MRPL12 stalk complex. ............................................... 217 

Figure A2.2: Dialyzed mitochondrial MRPL10-MRPL12 stalk complex. .................. 218 

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LIST OF TABLES

Table 1.1: List of mitochondrial and bacterial ribosomal proteins .............................. 14 

Table 2.1: Peptides detected from tryptic digests of mitochondrial ribosomal protein bands corresponding to acetylation signals by capLC-MS/MS analysis. ................................................................................................................ 60 

Table 2.2: Peptides detected from tryptic digests of mitochondrial ribosome associated bovine SIRT3 by capLC-MS/MS analysis. ......................................... 70 

Table 3.1: Peptides detected from tryptic digests of protein band corresponding to acetylation signal in SIRT3 knock-out mice mitochondria by capLC-MS/MS analysis. ................................................................................................................ 135 

Table 4.1: Peptide sequences of new mitochondrial ribosomal proteins identified from capLC-MS/MS analyses of in-gel tryptic digestions of 28S, 39S, and 55S proteins. ......................................................................................................... 170 

Table 4.2: Relative distribution of new mitochondrial ribosomal proteins using emPAI (Exponentially Modified Protein Abundance Index) values. ................... 172 

Table 4.3: GenBank™ and Swiss-Prot accession numbers of new mitochondrial ribosomal proteins found in various species. ........................................................ 173 

Table 4.4: List of mammalian mitochondrial ribosomal proteins with their bacterial homologs ................................................................................................ 191 

xiii

ABBREVIATIONS

12S rRNA of the small subunit of the mitochondrial ribosome 16S rRNA of the large subunit of the mitochondrial ribosome 28S small subunit of the mitochondrial ribosome 30S small subunit of the bacterial ribosome 39S large subunit of the mitochondrial ribosome 50S large subunit of the bacterial ribosome 55S intact mitochondrial ribosomes 70S intact bacterial ribosomes aa-tRNA aminoacyl tRNA ADP adenosine diphosphate ATP adenosine triphosphate ATP6 ATP synthase subunit 6 ATP8 ATP synthase subunit 8 bp base pair capLC-ESI capillary liquid chromatography nanoelectrospray ionization CID collision-induced dissociation Complex I NADH dehydrogenase Complex II succinate dehydrogenase Complex III ubiquinol cytochrome c oxidoreductase Complex IV cytochrome c oxidase Complex V ATP synthase COXI cytochrome c oxidase subunit 1 COXII cytochrome c oxidase subunit 2 COXIII cytochrome c oxidase subunit 3 CP central protuberance CTD C-terminal domain cyro-EM cyro-electron microscopy Cytb subunit of ubiquinol cytochrome c oxidoreductase DAP3 death associated protein 3 DTT dithiothreitol EF-G mt mitochondrial elongation factor G EF-Tu mt mitochondrial elongation factor Tu E site exit site for tRNA fMet formylated methionine GCN5L1 general control of amino acid synthesis 5-like 1 HSP60 heat shock protein 60 IF1 bacterial initiation factor 1 IF2 bacterial initiation factor 2 IF2mt mitochondrial initiation factor 2 IF3 bacterial initiation factor 3 IF3mt mitochondrial initiation factor 3 IPG immobilized pH gradient kDa kilo daltons

xiv

mgf Mascot generic file mRNA messenger RNA MRP mitochondrial ribosomal protein MS mass spectrometry MS/MS tandem mass spectrometry mtDNA mitochondrial genome NAD+ nicotinamide adenine dinucleotide NAM nicotinamide ND1 NADH dehydrogenase subunit 1 ND2 NADH dehydrogenase subunit 2 ND3 NADH dehydrogenase subunit 3 ND4 NADH dehydrogenase subunit 4 ND4L NADH dehydrogenase subunit 4L ND5 NADH dehydrogenase subunit 5 ND6 NADH dehydrogenase subunit 6 NTD N-terminal domain OXPHOS oxidative phosphorylation PAGE polyacrylamide gel electrophoresis PDB protein data bank PDCD9 programmed cell death protein 9 PMSF phenylmethylsulfonyl fluoride poly U polyphenylalanine P site peptidyl site for tRNA PTC peptidyl transferase site PTM post-translational modification PVDF polyvinylidene fluoride RF release factor ROS reactive oxygen species RP bacterial ribosomal protein rRNA ribosomal RNA SCX strong cation exchange chromatography SD standard deviation SDS sodium dodecyl sulfate SRL sarcin-ricin loop TCA tricarboxylic acid tRNA transfer RNA WT wild-type

xv

ACKNOWLEDGMENTS

I would like to take this opportunity to appreciate the help and support of my

advisor Dr. Emine Koc. This dissertation would not have been possible without her

inspiring suggestions and constant encouragement throughout productive and stimulating

discussions, not to mention her knowledge of every aspect of mitochondrial ribosomes. It

has been an honor to work in her laboratory first as a research associate and then as a

graduate student. I am grateful for the invaluable advice, support, and friendship of my

co-advisor, Dr. Hasan Koc, on both an academic and a personal level. I would like to

thank him for sharing his expertise in mass spectrometry and for making sure we survive

through times of frustration. I am thankful to all of the members of Koc laboratory, Dr.

Miller-Lee, Dr. Han, and Dr. Akpinar for sharing their experience and for providing an

excellent environment to work and I appreciate all of the work done by the undergraduate

students that are not mentioned here. I especially thank Dr. Han for our collaboration in

MRPL10 acetylation project and the undergraduate students, Beril Kumcuoglu and

Nadiah Abu who did astounding job in new MRPs project. I would also like to

acknowledge my committee members, Dr. Cameron, Dr. Reese, Dr. Kao, and Dr. Lee for

all the intellectual input and guidance throughout my graduate life. I need to express my

gratitude to the members of Cameron laboratory for their friendship and support during

my PhD. I am also grateful to the friends I have made at Penn State and Marshall

University. Finally, I would like to acknowledge my family for their unconditional love

and support in addition to their patience and understanding that helped me endure the

tough times.

Chapter 1

Introduction

1.1 Mitochondrion

Mitochondria (Greek mitos: thread; chondros: granule) were called bioblasts by

Richard Altmann in 1894 (1) and the aliens with permanent residence in our cells by

Erich Gnaiger (2). Mitochondria are composed of two compartmental spaces. First is the

matrix surrounded by the inner membrane impermeable to most small molecules,

including protons, and second is the intermembrane space enclosed by the permeable

outer membrane. In the 1950s, mitochondria were found to be the site of oxidative

phosphorylation (OXPHOS) in eukaryotes and they were called the powerhouse of the

cells (Fig. 1.1). They are responsible for providing more than 90 % of the total cellular

energy extracted from dietary calories. Dietary carbohydrates are degraded through

glycolysis to pyruvate, which is transported into mitochondria and converted to acetyl-

CoA by pyruvate dehydrogenase. The tricarboxylic acid cycle (TCA) oxidizes acetyl-

CoA by transferring hydrogens to the electron carriers, NAD+ and FAD, which drive

OXPHOS (Fig. 1.1). OXPHOS establishes an electrochemical gradient (Δψ) across the

inner mitochondrial membrane with complexes I, III, and IV pumping protons into the

inter membrane space (Fig. 1.2). This process provides the proton motive force which

drives the translocation of protons through the complex V, ATP synthase and this rotary

machine produces adenosine triphosphate (ATP) from adenosine diphosphate (ADP) and

2

phosphate (Pi) (3,4). Overall, mitochondrial bioenergetics involves the intermediates,

acetyl-CoA, ATP, and NAD+ to monitor the availability of nutrition and the redox status

for maintenance of cellular homeostasis (4). However, mitochondria are also source of

reactive oxygen species (ROS) generated during the transfer of electrons through the

OXPHOS complexes. In addition, protons in the intermembrane space are channeled

through the uncoupling proteins (UCP) located in the inner membrane to generate heat in

brown adipocytes (5). The other critical physiological functions of mitochondria involve

apoptosis, calcium storage to respond to calcium signaling, ketone body formation, heme

biosynthesis, and amino acid degradation including some reactions of the urea cycle

(Fig. 1.1). Given this variety of roles in cellular physiology, any mitochondrial

abnormality in these processes could contribute to the mitochondrial diseases such as

type 2 diabetes, neurodegeneration as in Parkinson’s disease, aging, and cancer (3,6,7).

Unfortunately, there is no cure available for mitochondrial disorders, only treatments to

relieve symptoms (8).

3

Figure 1.1: Mammalian mitochondrion.

Mitochondrion has two compartmental spaces, intermembrane space and matrix,

enclosed by inner and outer membranes. Pyruvate enters the mitochondrion through

pyruvate dehydrogenase (PDH) to generate acetyl-CoA. Oxidation of acetyl-CoA by the

tricarboxylic acid cycle (TCA) provides the reduced NADH to drive generation of ATP

which is translocated into cytosol by adenine nucleotide translocator (ANT) and voltage

dependent anion channel (VDAC). Respiratory chain complexes (I, II, III, IV, and V)

also generates reactive oxygen species (ROS) in addition to ATP synthesis. Uncoupling

proteins (UCP) generate heat by uncoupling the proton gradient (Δψ). Additional

mitochondrial metabolic functions include apoptosis with the release of cyctochrome c

(cyt c) into the cytosol and storage of cellular calcium for signaling.

4

1.2 Mitochondrial Genome

Human mitochondria contain several hundreds or thousands of mitochondrial

genome copies depending on the cell type. Mitochondrial DNA (mtDNA) is a 16,569

base pair (bp) circular molecule and is maternally inherited (Fig. 1.2) (9). It encodes 22

mitochondrial tRNAs, 13 mRNAs, and 2 rRNAs, a total of 37 genes without any intron.

(9-11). MtDNA contains a 1.1 kb non-coding region called the D-loop which is involved

in the regulation of transcription and replication of the molecule, and the double stranded

structure is formed by its heavy and light strands (12). The Light strand provides genes

for only one mRNA, ND6, and eight tRNAs while the rest of the 37 genes are encoded by

the heavy strand (10,13,14). Mitochondrially-encoded mRNAs are translated by

mitochondrial ribosomes to synthesize 7 proteins (ND1, 2, 3, 4, 4L, 5, and 6) of Complex

I among 43 subunits in total, 1 protein (Cytb) of Complex III among 11 subunits in total,

3 proteins (COXI, II, and III) of Complex IV among 13 subunits in total, and 2 proteins

(ATPase 6 and 8) of Complex V among 16 subunits in total (Fig. 1.2). Interestingly, none

of the subunits of Complex II are encoded by the mitochondrial genome and there is no

proton translocation through this complex, which may explain why mitochondrial DNA

did not preserve any gene for this complex throughout evolution. The mitochondrially-

encoded proteins are all essential for forming the hydrophobic core and the assembly of

the OXPHOS complexes with the nuclear-encoded subunits. OXPHOS deficiency can be

caused by mutations in any member of protein synthesis machinery including tRNA,

rRNA, and proteins. Many mutations in mtDNA are linked to deafness, metabolic and

5

neurodegenerative diseases, and result in myopathies, encephalopathy, diabetes, and

cancer (15-17).

6

Figure 1.2: Products of mitochondrial genome and oxidative phosphorylation.

Mitochondrion has a circular DNA molecule and its own

transcription/translation machinery for the synthesis of essential subunits of respiratory

chain complexes (I, III, IV, V). Double stranded mtDNA, 16.5 kb, encodes ND1, 2, 3,

4, 4L, 5, and 6 subunits of Complex I (blue), Cytb subunit of Complex III (pink),

COXI, II, and III subunits of Complex IV (sky blue) and ATP6 and 8 subunits of

Complex V (red), which are translated by mitochondrial ribosomes (55S).

7

1.3 Oxidative Phosphorylation

Most of the OXPHOS subunits are encoded by the nuclear DNA, translated by

cytosolic ribosomes, and then translocated into the mitochondria where they are

assembled along with the mitochondrial encoded subunits into functional complexes. The

TCA cycle and fatty acid oxidation are the biochemical pathways providing reduced

forms of co-factors, NADH (nicotinamide adenine dinucleotide) and FADH2 (flavin

adenine dinucleotide) to drive oxidative phosphorylation (6). Five different enzyme

complexes located on the inner membrane are responsible for the synthesis of ATP

(Fig. 1.2). Oxidative phosphorylation is initiated with the transfer of electrons from

NADH or succinate to ubiquinone (also called coenzyme Q or Q) by Complex I (NADH

dehydrogenase) or Complex II (succinate dehydrogenase), respectively (18,19). Then, the

reduced ubiquinone (also called ubiquinol or QH2) carries these electrons to Complex III

(cytochrome c reductase) which mediates the transfer to cytochrome c (cyt c) (20).

Complex IV (cytochrome c oxidase) generates H2O with the electrons from reduced cyt c

and oxygen (Fig. 1.2) (21,22). These processes are accompanied by proton translocation

into the intermembrane space through complexes I, III, and IV to generate an

electrochemical gradient (Δψ). In the final step, the proton-motive force drives protons

back into the matrix to provide energy for ATP synthesis through Complex V (ATP

synthase) (23).

Complexes I and III are involved in the generation of reactive oxygen species

(ROS) which are hydrogen peroxide and oxygen radicals. These species are associated

with mitochondrial dysfunction resulting in neurodegeneration since they target proteins,

8

lipids, and nucleic acids (24,25). In addition, a mutation in Complex I subunit 4

converting highly conserved arginine to a histidine at codon 340 was implicated in the

atrophy of the optic nerve in adults (Leber's hereditary optic neuropathy) (26). Moreover,

the T8993C/G mutations of mtDNA affect ATPase6 subunit of Complex V and result in

Leigh syndrome in addition to the phenotype of ataxia and pigment retinopathy (27,28).

1.4 Mitochondrial Translation

Translation of mitochondrial mRNAs involves different stages similar to those in

bacteria (29). In mitochondria, mRNAs do not contain 5’-untranslated region (UTR) and

3’-UTRs other than a short polyA tail, and do not require a Shine-Dalgarno sequence as

needed in prokaryotes, or a 5’-cap as in eukaryotic cells for initiation of translation

(9,14). However, mitochondrial ribosomes need formyl-methionyl tRNA (fMet-tRNA)

for initiation as in bacteria (30). The three different stages of mitochondrial translation,

initiation, elongation, and termination are explained below (Fig. 1.3).

1.4.1 Initiation

There are two mitochondrial initiation factors, IF2mt and IF3mt involved in protein

synthesis in addition to fMet-tRNA and mRNA. Mitochondrial translation system lacks

IF1 which is essential in the bacterial translation initiation. IF2mt was demonstrated to

perform the role of IF1 with the additional 37 amino acid residue between domains V and

VI (31-33). The first step in translation initiation is the dissociation of intact ribosomes

9

(55S) by IF3mt providing small subunit (28S) for binding of IF2mt-GTP and mRNA which

is fed into the protein rich mRNA entrance gate on the small subunit (31). This complex

promotes the binding of fMet-tRNA to the 5’ start codon and then association of

ribosomal large subunit (39S) releases the initiation factors after the hydrolysis of GTP to

GDP. At this step, the 55S initiation complex is formed and enters into the elongation

phase of protein synthesis (Fig. 1.3) (34).

1.4.2 Elongation

The polypeptide chain elongation phase of mitochondrial protein synthesis is

highly conserved between bacteria and mitochondria compared to the initiation and

termination phases in bacteria (34-36). It is initiated with the binding of the ternary

complex which is formed through the binding of EF-Tumt and aminoacyl-tRNA (aa-

tRNA) to the A-site of the 55S complex provided during the initiation step. Just after the

recognition of codon-anticodon interaction, GTP is hydrolyzed to GDP by EF-Tumt and it

is released from the complex (34). Peptide bond formation is catalyzed by the ribosome

itself followed by the translocation of deacetylated tRNA from the P-site to the E-site.

The energy from GTP hydrolysis on EF-Gmt drives the peptidyl-tRNA from the A-site to

the P-site, which is analogous to the bacterial protein synthesis (34,37). It was also

reported that the L7/L12 stalk proteins are involved in the translocation process by

promoting GTPase activity of elongation factors (38,39). Sequential repeats of the

elongation phase generate growing polypeptide chain through the peptide exit tunnel on

the ribosomal large subunit.

10

1.4.3 Termination

The ribosome stalls when either UAA or UGA stop codon enters into the A-site.

These codons are recognized by release factor, RF1amt, which causes the release of

synthesized polypeptide by the hydrolysis of the peptidyl-tRNA bond by the peptidyl

transferase center with the energy from GTP (34,40). In bacterial protein synthesis, RF1

and RF2, which are small GTPases, recognize UAA or UAG and UAA or UGA as stop

codons to terminate translation, respectively (41,42). A third release factor, RF3,

catalyzes the release of RF1 and RF2 at the end of the termination process in order to

recycle the ribosomes in dissociated forms to keep them available for next initiation

phase.

11

Figure 1.3: Model for different stages of mitochondrial translation.

Mitochondrial translation starts with IF3mt binding which provides dissociated large

subunit (39S) and small subunit (28S) for the initiation phase. The mitochondrial initiation

factors, IF2mt and IF3mt, are shown in green and pink, respectively. The mitochondrial

elongation factors, EF-Tumt and EF-Gmt, are shown in sky blue and red, respectively. The

release factor, RFmt, is shown in purple. The initiator tRNA (fMet-tRNA) and aminoacyl-

tRNA (aa-tRNA) are shown in light and dark green, respectively. GTP is designated with

orange and GDP with yellow color.

12

1.5 Mitochondrial Ribosome

In eukaryotic cells, there are two types of ribosomes, cytoplasmic ribosomes

present in the cytosol and mitochondrial ribosomes located within mitochondria. The

mitochondrial translation machinery synthesizes only thirteen proteins of the oxidative

phosphorylation complexes and the synthesis of the rest of the cellular proteins are all

carried out by cytoplasmic ribosomes. Mammalian mitochondrial ribosomes have low

sedimentation coefficients (55S) and they are formed by association of small subunit

(28S) and large subunit (39S) complexes (Fig. 1.4) (43). These ribosomes are large

ribonucleoprotein complexes with a molecular mass of ~2.6x106 Da. They are closely

related to bacterial ribosomes compared to the cytosolic ribosomes, in terms of conserved

proteins in some functional domains on ribosome and susceptibility to similar antibiotics.

However, their protein to RNA ratio is totally reversed (67 % protein, 33 % RNA in

mitochondrial ribosome) (44). Mitochondrial ribosomes are composed of twenty-nine

proteins with 12S rRNA on the small subunit and forty-eight proteins with 16S rRNA on

the large subunit (45-47). These proteins are listed in Table 1.1 and this list is updated

with our recent findings in Chapter 4 (46).

13

Figure 1.4: The 3D cryo-EM reconstruction mitochondrial ribosomes.

The mitochondrial ribosome contains nuclear-encoded mitochondrial ribosomal

proteins and mitochondrial-encoded rRNAs (12S, 16S). The small subunit is 28S and

shown in yellow and green. The large subunit is 39S and shown in sky blue and dark

blue. The regions conserved in bacterial and mitochondrial ribosomes are shown in

yellow and sky blue. The green and dark blue regions are the mitochondrial specific

proteins which have no homolog in bacterial ribosomes. Details of functional domains

shown in the figure are described in text. SRL: sarcin-ricin loop, CP: central

protuberance and PTC: the peptidyl transferase site in the large subunit. Platform is

located on the opposite side of shoulder in the mRNA binding path. Courtesy of Dr. R.

K. Agrawal.

14

28S and 39S: small and large subunit of mitochondrial ribosome 30S and 50S: small and large subunit of bacterial ribosome New Class: mitochondria-specific ribosomal proteins *: List includes the newly identified ribosomal proteins in Chapter 4.

Table 1.1: List of mitochondrial and bacterial ribosomal proteins.*

28S 30S New Class 39S 50S New Class Missing S1 MRPS22 MRPL1 L1 MRPL37 MRPS2 S2 MRPS23 MRPL2 L2 MRPL38 Missing S3 MRPS24 MRPL3 L3 MRPL39 Missing S4 MRPS25 MRPL4 L4 MRPL40 MRPS5 S5 MRPS26 Missing L5 MRPL41 MRPS6 S6 MRPS27 Missing L6 MRPL42 MRPS7 S7 MRPS28 MRPL12 L7/L12 MRPL43 Missing S8 MRPS29 MRPL9 L9 MRPL44 MRPS9 S9 MRPS30 MRPL10 L10 MRPL45 MRPS10 S10 MRPS31 MRPL11 L11 MRPL46 MRPS11 S11 MRPS32 MRPL13 L13 MRPL48 MRPS12 S12 MRPS33 MRPL14 L14 MRPL49 Missing S13 MRPS34 MRPL15 L15 MRPL50 MRPS14 S14 MRPS35 MRPL16 L16 MRPL51 MRPS15 S15 MRPS36 MRPL17 L17 MRPL52 MRPS16 S16 MRPS37 (CHCHD1) MRPL18 L18 MRPL53 MRPS17 S17 MRPS38 (AURKAIP1) MRPL19 L19 MRPL54 MRPS18-1 S18 MRPS39 (PTCD3) MRPL20 L20 MRPL55 MRPS18-2 S18 MRPL21 L21 MRPL56 MRPS18-3 S18 MRPL22 L22 MRPL57 (RP_63) Missing S19 MRPL23 L23 MRPL58 (ICT1) Missing S20 MRPL24 L24 MRPL59 (CRIF1) MRPS21 S21 Missing L25 MRPL60 (C7orf30) MRPL27 L27 MRPL28 L28 MRPL47 L29 MRPL30 L30 Missing L31 MRPL32 L32 MRPL33 L33 MRPL34 L34 MRPL35 L35 MRPL36 L36

15

1.5.1 Small Subunit (28S) of Mammalian Mitochondrial Ribosome

Analysis of the small subunit of mitochondrial ribosomal proteins (MRP) resulted

in identification of a total of twenty-nine proteins currently listed in Table 1.1 (updated in

Chapter 4) (48-50). Fourteen of them have homologs in the bacterial ribosome while the

other fifteen proteins are classified as mitochondria-specific ribosomal proteins. The

small subunit is structurally divided into three functional segments, the platform, the

head, and the body. First, the platform lines the region in between the head and body

parts and it is involved in the formation of the P-site on the ribosome and the mRNA

binding to the ribosome (50). The platform is formed by MRPS2, MRPS5, MRPS6,

MRPS21, and MRPS21 which interact with the central domain of 12S rRNA in addition

to one of the three MRPS18 variants (47,50). Next, the head region, composed of

MRPS2, MRPS7, MRPS9, MRPS10, and MRPS14, provides their interaction with the 3’

domain of the 12S rRNA (47,50). Proteins located in the platform and head regions have

essential functions in mRNA and tRNA binding to the bacterial ribosome (37,51-56).

Last, the body segment of the small subunit involves MRPS5, MRPS12, MRPS16,

MRPS17 interacting with the 5’ domain of the 12S rRNA (47).

1.5.2 Large Subunit (39S) of Mammalian Mitochondrial Ribosome

The large subunit of the mitochondrial ribosome consists of 16S rRNA and forty-

eight proteins, twenty-eight of which have homologs in the bacterial ribosome

(Table 1.1). The rest of them, MRPL37 to MRPL56, are mitochondria-specific ribosomal

proteins. There are three major structural units extending from the body, L1 stalk, L7/L12

16

stalk, and the central protuberance (CP), which are identified as larger than their bacterial

counterparts (11). The CP region has MRPL18 which is homologous to RPL18 in

bacteria while this region misses other proteins homologous to RPL5 and RPL25 (57).

The L1 stalk proteins such as MRPL1, MRPL9, and MRPL28, have a higher molecular

mass than their corresponding partners in the bacterial ribosome. This might be a

compensation mechanism for the truncated 16S rRNA helices in mitochondrial ribosome

(58). The other functionally important region is the peptide exit tunnel which is

surrounded by MRPL22, MRPL23, and MRPL24. This region does not have a homolog

of RPL29 which is present in bacterial ribosomes. The functional role of the RPL29

might be replaced by other interacting proteins (11,59). The most important and relevant

functional region on the ribosome is the L7/L12 stalk, which is formed by RPL10,

RPL11, and multiple copies of RPL7/L12 dimers in bacteria. RPL7 is the N-terminally

acetylated form of RPL12. Since N-terminal acetylation of mitochondrial protein L12 is

not reported yet, MRPL12 designates the mitochondrial counterpart of L7/L12. In

mitochondria, the evolutionary conserved L7/L12 stalk of the large subunit is composed

of MRPL10, MRPL11, and MRPL12 (Fig. 1.5). The members of this stalk are all

essential and play significant roles during different stages of protein synthesis by

interacting with several different translation factors, IF2, EF-Tu, EF-G, and RF3

(39,60,61). The RPL7/L12 exists in different numbers of dimers on the ribosome. The

length of the L10 helix 8 is an important player in the determination of the total number

of RPL7/L12 dimers (62). In Escherichia coli, there are four copies of L7/L12 as two

dimers bound to the C-terminal domain of L10 (63). In addition, Thermotoga maritima

has six copies of RPL7/L12 forming three dimers localized on RPL10 which has

17

additional amino acid residues at its helix 8 (64). However, there is no information

pertaining to the number of MRPL12 present on the mitochondrial ribosome. The

transient binding of MRPL12 to the ribosome makes it distinct from other ribosomal

proteins in terms of the regulation of ribosomal function. This flexible binding involves

only protein-protein interactions with MRPL10 without any rRNA interaction (65).

1.5.3 New Mitochondrial Ribosomal Proteins and Additional Functions

Mitochondria-specific ribosomal proteins present a new class of proteins which

do not have homologous counterparts in bacterial ribosome. These mitochondria-specific

ribosomal proteins are, MRPS29 (DAP3, death associated protein 3), MRPS30 (PDCD9,

programmed cell death protein 9), MRPL37, and MRPL41, all of which are known to be

involved in apoptosis (48,66-69). Furthermore, MRPL41 plays an important role in cell

cycle arrest by increasing the levels of p21 and p27 under serum starvation (66,70). On

the other hand, a member of peptidyl-tRNA hydrolase family protein, immature colon

carcinoma transcript-1 (ICT1), has been demonstrated to cleave the peptidyl-tRNA and

prevent them from stalling on mRNA lacking a stop codon in mitochondria (59).

In addition, several mitochondrial ribosomal proteins, such as MRPS23,

MRPL11, and MRPL28, were found to be expressed differentially in cervical cancer and

pancreatic tumor cells (71,72). Targeting MRPL28 and MRPL12 in this type of cells

leads to reduced mitochondrial activity but elevated glycolysis (73). Therefore, it is

plausible to suggest that the regulation of mitochondrial protein synthesis in addition to

18

these newly acquired roles might be important in mitochondrial function and energy

production.

19

Figure 1.5: Model representing the mitochondrial L7/L12 stalk.

The L7/L12 stalk comprises L10 (green) having a globular N-terminal domain

involved in binding to L11 (yellow) and providing C-terminal α-helix domain for the

binding of multiple copies of L7/L12 dimers (pink). The L7/L12 N-terminal domain

(NTD) interacting with L10 and the C-terminal domain (CTD) which binds to GTPases

are also labeled. Sarcin-ricin loop (SRL) of 23S rRNA provides the interaction sites for

translation factors. Protein Data Bank entries of 1MMS (T. maritima L11), 1RQU (E.

coli L7/L12 dimer), and 1S72 (Haloarcula marismortui 50S, where proteins and rRNA

are in aqua and gray, respectively) are used in this figure (61). Courtesy of Dr. Koc.

20

1.6 Post-Translational Modifications of Proteins

Post-translational modifications (PTMs) are covalent, mostly reversible,

alterations to one or more amino acids of a protein target after its synthesis. These

modifications include acetylation, phosphorylation, methylation, glycosylation,

ubiquitination, nitrosylation, and proteolysis that determine activity state, localization,

turnover, and interactions with other cellular molecules such as proteins, nucleic acids,

lipids, and cofactors (74). Mass spectrometry-based methods provide highly sensitive

detection and mapping of PTMs using MS/MS spectra of peptides from in-gel or in–

solution digestion with proteases, most common being trypsin.

Acetylation at ε-amino group of lysine residues can be detected by its

characteristic mass shift of +42 Da from unmodified forms. Acetylation of lysine residue

blocks its cleavage by trypsin due to charge neutralization. Therefore, the acetylated

peptides are detected in late fractions during reverse-phase chromatography separation

and identified as identified as a longer peptide with a miss-cleavage. Protein

phosphorylation on serine, threonine, or tyrosine residues is similarly detected by 80 Da

increase in mass, which is diagnostic for the addition of HPO3. Site of phosphorylation

can be mapped using the mass shift in the corresponding fragment ion peak in tandem

mass spectrum of the phosphorylated peptide.

1.6.1 Lysine Acetyltransferases (KATs) and Deacetylases (KDACs)

Acetylation of proteins involves the transfer of acetyl group from acetyl-CoA to

the ε-amino group on target lysine, which results in the charge neutralization. It is one of

21

the best characterized histone tail modifications regulated by histone acetyltransferases

(HATs) and histone deacetylases (HDACs) (75). Lysine acetylation on histones is

associated with transcription activation by opening of chromatin structure and

recruitment of bromodomain containing transcriptional factors, such as SWI/SNF and the

SAGA (76,77). There are three main HATs families: GNATs (general control non-

derepressible 5 (Gcn5)-related N-acetyltransferases), p300/CBP (adenoviral E1A-

associated protein of 300 kDa/CREB-binding protein), and MYST proteins. Members of

the GNAT family include HAT1 (histone acetyltransferase 1), Gcn5, PCAF, Elp3, Hat1,

Hpa2, and Nut1 (78). MYST proteins are founded by MOZ (monocytic leukemia zinc

finger protein), YBF2/SAS3, SAS2 (something about silencing 2), and Tip60 (HIV Tat-

interactive protein of 60 kDa). The members of the MYST family suggest a relation of

histone acetylation to leukemogenesis, gene silencing, and HIV biology (79,80). HATs

mostly form multisubunit complexes which are important in regulating acetyltransferase

activity by targeting them to specific chromosomal regions and in modulating the

substrate specificity of HATs. They are under a dynamic relationship with HDACs. There

are four classes of HDACs in humans; class I (HDAC1, -2, -3, and -8), class II (HDAC4,

-5, -6, -7, -9, and -10), class III (sirtuin [silent information regulator 2 (Sir2)-related

protein] SIRT1-7), class IV (HDAC11) (75,81). NAD+ requirement of sirtuins as the

cofactor makes them distinct from other HDACs, which share sequence homology and

are dependent on Zn2+ for deacetylase activity. They show variation in their cellular

localization (82). NuRD (nucleosome remodeling deacetylase complex), an HDAC1/2

complex, was demonstrated to be involved in cancer metastasis (83). HDAC3 complexes

were linked to histone demethylation and cell-cycle regulation (84). Hda6 (histone

22

deacetylase 6), which was identified as a class I HDAC in Arabidopsis thaliana, was

implicated in heterochromatin formation by interacting with different chromatin

modifiers (85).

HATs and HDACs are broadly referred to as KATs (lysine acetyltransferases) and

KDACs (lysine deacetylases) after the identification of non-histone proteins as targets of

acetylation and deacetylation (86). P53, NF-κB (nuclear factor-κB), STAT3, tubulin, c-

Myc, HIF1α (hypoxia-inducible factor), HMG (high mobility group) proteins, and Ku70

are among these non-histone targets which are involved in many biological processes

such as cell proliferation, survival, and apoptosis (86-88). Acetylation of nonhistone

proteins has been demonstrated to modulate protein functions by altering their stability,

cellular localization, and interactions with nucleotide or other proteins (86).

The members of class III KDACs or sirtuins (SIRT1-7) are homologous to the

yeast Sir2 (silent information regulator 2). They deacetylate the target acetylated lysine

residues by consuming NAD+ as a cofactor and then they release NAM (nicotinamide)

and O-Acetyl-ADPR (O-acetyl-ADP-ribose). SIRT1, SIRT6, and SIRT7 are located in

nucleus, SIRT2 in the cytoplasm, and SIRT3, SIRT4, and SIRT5 are primarily

mitochondrial. SIRT1, SIRT3, and SIRT5 deacetylate the target proteins in an NAD+-

dependent manner; however, SIRT6 functions as an NAD+-dependent ADP-

ribosyltransferase. SIRT2 and SIRT4 have both NAD+-dependent deacetylase and ADP-

ribosyltransferase activity (89). The enzymatic activity of SIRT7 has not yet been

determined. Its involvement in the activation of RNA polymerase I transcription was

demonstrated (90). They control the metabolic response to oxidative stress by targeting a

variety of proteins in different cellular compartments. In mammalian cells, it was shown

23

that calorie restriction activates SIRT1, which increases the life span of the cell by

reducing stress-induced apoptosis (91). SIRT1 deacetylates PGC1α and HIF1α to

modulate the balance between oxidative and glycolytic metabolism under low energy

status of the cell. Low glycolysis rate results in the activation of SIRT1 due to elevated

NAD+ levels and, therefore, deacetylation of PGC1α promotes the mitochondrial

biogenesis and oxidative metabolism (92). It also mediates the response to the oxidative

stress by deacetylating HSF (heat shock factor). Ku70, which is a DNA repair factor

induced by calorie restriction, is deacetylated by SIRT1 to promote cell survival (93).

SIRT2 deacetylates α-tubulin and perturbs cell motility by effecting microtubule

dynamics (94,95). SIRT6 is an ADP-ribosyltransferase which plays a role in the

regulation of genomic stability by functioning in base excision repair of single stranded

DNA breaks (96,97). SIRT6 along with SIRT1 inhibit the transcriptional activity of NF-

κB to protect pancreatic β cells from inflammatory response (98,99). Overall, the NAD+-

dependent reactions of sirtuins connect energy metabolism and cellular survival

pathways. The targets described above implicate them in a wide range of diverse cellular

processes; such as glucose homeostasis, cellular growth, and stress resistance.

1.6.2 Mitochondrial Acetyltransferases and Sirtuins

In recent years, a tremendous amount of work has been reported about the role of

acetylation modifying the ε-amino group of lysine residues of proteins in regulation of

energy metabolism (100-106). Almost ~30 % of mitochondrial proteins have been

identified as acetylated by high-throughput proteomic approaches (87,107). Majority of

24

post-translationally modified proteins are involved in mitochondrial metabolic pathways,

including oxidative phosphorylation, fatty acid oxidation, and the urea cycle

(100,103,108-110).

In a recent report, GCN5L1 has been identified as a component of the

mitochondrial acetyltransferase system. It was demonstrated to acetylate respiratory chain

subunits and inhibit their enzymatic activities (Fig. 1.6) (111). On the other hand, three

members of class III histone deacetylases, SIRT3, SIRT4 and SIRT5, have been localized

into mitochondria (102,103,112,113). Their activity depends on the availability of NAD+

which is a cosubstrate in the deacetylation and ADP-ribosylation of target proteins (114).

This reaction generates nicotinamide (NAM) and 2`-O-Acetyl-ADP-ribose resulting in

deacetylated lysine residues.

Acetyl-CoA synthetase 2 (AceCS2) is one of the first substrates identified for

SIRT3 in mitochondria. It is activated by SIRT3 deacetylation to enhance acetyl-coA

production (109,115,116). Respiratory chain complexes in mitochondria were found to be

regulated by reversible acetylation (Fig. 1.6). Acetylation of the NDUFA9 subunit of

Complex I inhibits its activity and SIRT3 reverses this inhibition by deacetylation. We

also discovered that Complex II activity is regulated in a similar manner by reversible

acetylation of succinate dehydrogenase flavoprotein subunit, SdhA (see Chapter 3) (100).

SIRT3 knock-out mice displays no obvious phenotypes except lower basal ATP levels

(107,108). Overexpression of SIRT3 in brown adipocytes elevates the consumption of

oxygen, which indicates the increased activity of oxidative phosphorylation, and results

in reduced ROS production in these cells (117). Overall, these findings demonstrate the

important role of SIRT3 in direct regulation of oxidative phosphorylation and

25

maintenance of basal ATP levels (100,108). In beta cell mitochondria, SIRT4 is involved

in repressing the activity of glutamate dehydrogenase by ADP-ribosylation and therefore

reducing the insulin secretion in response to amino acids (113). In addition, SIRT5

deacetylates carbamoyl phosphate synthetase 1 (CPS1) which leads to activation of

enzymatic function in ammonia detoxification and disposal during the urea cycle. This

highlights the importance of regulation of the urea cycle during sirtuin-mediated

mitochondrial adaptation under conditions of energy limitation (Fig. 1.6) (103).

Mitochondrial sirtuins appear to work together to regulate the mitochondrial adaptation;

SIRT3 and SIRT4 may stimulate production of ammonia and SIRT5 activates the urea

cycle to process this ammonia (118). In addition, energy production from fat might be

involved in reduction of ROS generation since electrons are translocated to the

respiratory chain Complex II by FADH2. In this way, overall ROS generation per se

might be decreased since one of the ROS sources, the Complex I, is bypassed (119). In

conclusion, mitochondrial sirtuins mediate the metabolic adaptation to funnel fuel

sources, fatty acids and aminoacids, to be used as energy sources during energy

limitation.

26

Figure 1.6: Substrates and biological functions of sirtuins and mitochondrial

acetyltransferase(s).

SIRT1, SIRT6, and SIRT7 reside in nucleus. SIRT1 deactylates PGC1α (PPAR

gamma coactivator 1-alpha) and HIF1α (hypoxia-inducible factor 1-alpha) to modulate the

balance between oxidative and glycolytic metabolism under low energy status of the cell. It

also targets HSF (heat shock factor) and Ku70 (DNA repair factor) to mediate the oxidative

stress and DNA damage response, respectively. SIRT1 along with SIRT6 inhibit the NF-κB

activity to regulate inflammatory response. SIRT7 was demonstrated to be a positive

regulator of RNA polymerase I transcription. SIRT2 functions as α-tubulin deacetylase in

the cytoplasm.

27

SIRT3, SIRT4, and SIRT5 are localized in mitochondria. SIRT3 targets

respiratory chain Complexes I (NDUFA9) and II (SdhA) and activates oxidative

phosphorylation. Recently identified mitochondrial acetyltransferase, GCN5L1, reverses

the effects of SIRT3 on Complexes I and II. In addition, SIRT3 deacetylates MRPL10 in

the ribosome and down regulates protein synthesis. SIRT4 ADP-ribosylates glutamate

dehydrogenase (GDH) and controls insulin secretion in pancreatic β-cells. Carbamoyl

phosphate synthetase (CPS1), which catalyzes the first and rate limiting step of the urea

cycle, is activated by SIRT5.

28

1.6.3 Post-Translational Modifications of Mitochondrial Ribosomal Proteins

Large scale mass spectrometry based analyses of post-translational modifications

(PTMs), especially phosphorylation and acetylation of ribosomal proteins have led to the

discovery of regulation of ribosomal activity by these modifications (120-125).

Mitochondrial ribosomes are responsible for the synthesis of thirteen essential subunits of

respiratory chain complexes that oxidize acetyl-CoA and NADH to generate ATP for

cellular needs. In order to maintain mitochondrial and cellular energy homeostasis, these

metabolites in addition to NAD+ might be important for the modulatory enzymes to sense

the energy and redox state of the mitochondria and cell. These modulators are the

kinases, acetyl transferases, and deacetylases/ADP-ribosyl transferases located in

mitochondria (105,111,122,125,126).

PTMs of ribosomal proteins might lead to conformational changes resulting in

protein-protein and protein-RNA reorganization (126-128). These modified ribosomal

proteins are mostly found in functionally important parts of the ribosome, L7/L12 stalk,

sarcin-ricin loop, and peptide exit tunnel, peptidyl transferase center, which are highly

conserved between bacterial and mitochondrial ribosomes as described earlier. MRPS7,

MRPS11, and variants of MRPS18 were found to be phosphorylated at residues

important for mRNA-binding during the initiation phase of protein synthesis (121).

MRPS16 is another phosphorylated protein involved in subunit association, and has been

found to be lethal when it has a mutation causing immature synthesis (121,129). MRPL2

and MRPL27 are positioned in the region forming the peptidyl transferase center and

their phosphorylation is implicated in regulation of mitochondrial protein synthesis

29

(121,128). Moreover, MRPS29, which is a mitochondria-specific ribosomal protein and

has a role in apoptosis, has been studied in detail for its phosphorylated residues and how

these residues regulate its function in mitochondrial protein synthesis and apoptosis

(120). The sarcin-ricin loop region is involved in promoting GTPase activity of

translation factors along with the ribosomal L7/L12 stalk proteins. Some of the proteins

in this region, MRPL3, MRPL13, and MRPL19, have been identified as phosphorylated

as well. The other member of this stalk, MRPL10 is acetylated which is involved in the

regulation of protein synthesis (see Chapter 2) (124).

The mitochondrial translation machinery and ribosomal proteins are recognized as

essential components of mitochondrial dysfunction in apoptosis and cancer due to their

roles in mitochondrial energy homeostasis. Defects in MRPS16 and MRPS22 have been

associated with severe respiratory dysfunction because of deficiency in oxidative

complexes. (129,130). Furthermore, aberrant expression of certain ribosomal proteins,

MRPS23, MRPL11, MRPS29, and MRPS30 are implicated in several tumors, cervical

and breast cancer (71,72,131,132).

1.7 Research Aims

As described in the previous sections, mitochondrial ribosomes are responsible

for the synthesis of essential proteins involved in energy generation. The regulation of

mitochondrial translation by PTMs of ribosomal proteins would also be involved in

mitochondrial function/dysfunction. To test this hypothesis, the role of ribosomal protein

phosphorylation and acetylation in the regulation of mitochondrial translation and

30

apoptosis has been investigated (120,133,134). Phosphorylation of mitochondrial

ribosomal proteins was demonstrated to be critical for induction of apoptosis and

regulation of translation (133). Steady-state levels of phosphorylation and acetylation of

mitochondrial ribosomal proteins were identified using mass spectrometry-based

proteomics (133,134). We applied capillary liquid chromatography - nanoelectrospray

ionization - tandem mass spectrometry (capLC-MS/MS) based proteomics to identify

acetylated mitochondrial ribosomal proteins involved in translation.

By performing two-dimensional gel electrophoresis followed with mass

spectrometry analyses of mitochondrial ribosomal proteins, we identified MRPL10 as the

major acetylated protein and mapped the acetylated lysine residues which might induce

conformational changes to the protein and might affect its interaction with other

ribosomal proteins. MRPL10 is a member of one of the highly conserved functional sites

in the mitochondrial ribosome, L7/L12 stalk. This stalk is composed of MRPL10,

MRPL11, and multiple dimers of MRPL12. It is essential for protein synthesis in all

organisms due to interactions with elongation and initiation factors during various stages

of translation. In bacteria, acetylation of L12 was demonstrated to stabilize the interaction

between stalk proteins due to increased hydrophobicity by acetylation (135). The

acetylated lysine residues of MRPL10 are located near the pivotal point of the L10 where

it interacts with MRPL12 dimers in the crystal structure model of the stalk (Fig. 5.1). In

addition, in our preliminary analyses of SIRT3 knock-out mice, we detected changes in

the MRPL12 amounts compared to other ribosomal proteins. Based on the evidence

above, we hypothesize that reversible acetylation of MRPL10 is important in the

modulation of MRPL12 binding to ribosomes and regulation of the mitochondrial protein

31

synthesis. Next, the interaction of SIRT3 with the mitochondrial ribosome was revealed

by immunoblotting analyses of sucrose gradient fractions containing separated ribosomal

subunits. Since SIRT3 is involved in the modulation of mitochondrial functions in

response to energetically changing environment by monitoring the altered energy and

redox state, we investigated the effect of SIRT3 in MRPL10 acetylation and

mitochondrial translation. In SIRT3 knock-out mice, elevated acetylation of MRPL10

resulted in improved stability of MRPL12 binding to the L7/L12 stalk accompanied with

enhanced mitochondrial translation. On the other hand, HIB1B, a brown adipocyte tissue

cell line stably overexpressing SIRT3, displayed reduction in the acetylation status of

MRPL10 which resulted in destabilization of MRPL12 on the ribosome. Moreover, the

activities of Complex I and Complex IV and total ATP production were assayed in

SIRT3 overexpressing cells. Overall, we present a regulatory role of MRPL10 acetylation

modulating composition of the L7/L12 stalk and mitochondrial translation in response to

cellular energy needs monitored by SIRT3.

In our analysis of liver mitochondrial proteins from SIRT3 knock-out mice, we

detected acetylation of additional target proteins. The report about the regulation of

Complex I activity and oxidative phosphorylation by SIRT3 encouraged us to perform

more functional studies with SIRT3 knock-out mice liver mitochondria. We revealed the

flavoprotein (SdhA) subunit of Complex II, succinate dehydrogenase as an additional

substrate for SIRT3. Deacetylation of the SdhA subunit resulted in increased Complex II

activity in wild type mice. By employing mass spectrometry-based proteomics, we

mapped the acetylated residues which are possibly involved in the entry of substrate to

the active site of the enzyme in order to regulate its enzymatic activity. This study

32

identifies SdhA as a novel substrate of the NAD+-dependent deacetylase, SIRT3,

involved in regulation of Complex II activity through reversible acetylation.

Mitochondrial ribosomes are more closely related to bacterial ribosomes than

eukaryotic cytosolic ribosomes, which supports the evolutionary perspective of

endosymbiosis theory. Most of the mammalian mitochondrial ribosomal proteins are

homologous to their bacterial counterparts in terms of their structure and function.

However, the amount of protein on the ribosome relative to ribosomal RNA is totally

reversed and mitochondrial ribosome acquired additional proteins. The improved

sensitivity of mass spectrometry-based proteomic tools and increased availability of

expressed sequence tags from different organisms in databases prompted us to reevaluate

the protein components of mitochondrial ribosomes. In Chapter 5, we report three

additional members of the mitochondrial ribosome; CHCHD1, AURKAIP1, and CRIF1,

and their functional implication on mitochondrial protein synthesis by employing siRNA

mediated knockdown in cell lines.

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Chapter 2

Regulation of Mitochondrial Translation by SIRT3

This chapter of dissertation was reproduced in part with permission from Yongjie

Yang*, Huseyin Cimen*, Min-Joon Han*, Tong Shi, Jian-Hong Deng, Hasan Koc,

Orsolya M. Palacios, Laura Montier, Yidong Bai, Qiang Tong, and Emine C. Koc. (2010)

NAD+-dependent deacetylase SIRT3 regulates mitochondrial protein synthesis by

deacetylation of the ribosomal protein MRPL10. J. Biol. Chem.; 285 (10):7417-7429.

Copyright © 2010 by The American Society for Biochemistry and Molecular Biology,

Inc.

* These authors equally contributed to this work.

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2.1 Rationale

Protein lysine acetylation has been identified as a widespread post-translational

modification in almost 30 % of mitochondrial proteins and implicated to be involved in

regulatory mechanisms for energy metabolism, stress responses, and longevity (1-4).

Since most of ATP to be used for cellular energy is produced by oxidative

phosphorylation (OXPHOS) in mitochondria, it is essential to uncover the regulatory

mechanisms of its activity. Phosphorylation of mitochondrial ribosomes and

mitochondrial proteins is implicated in the regulation of protein synthesis and

mitochondrial function. However, there is still limited knowledge on the role of

acetylation in mitochondria. In this study, we identified mitochondrial ribosomal protein

L10 (MRPL10) as the major acetylated ribosomal protein and its NAD+-dependent

deacetylation by SIRT3 associated with the ribosome by employing mass spectrometry-

based proteomic tools and immunoblotting analyses. The acetylated lysine residues were

mapped by using tandem mass spectrometry and the importance of these residues in the

acetylation of MRPL10 was confirmed by site-directed mutagenesis studies. Moreover,

we have demonstrated the increased in vitro translational activity of bovine ribosomes

treated with nicotinamide (NAM), which is an inhibitor of SIRT3. In addition, the

mitochondrial ribosomes isolated from SIRT3 knock-out (Sirt3-/-) mice liver displayed

enhanced translational activity in our in vitro translation assays. To determine the

regulatory mechanism of mitochondrial ribosome acetylation, we analyzed the

mitochondrial ribosomes of SIRT3 knock-out mice and mouse brown adipocyte (HIB1B)

cells stably overexpressing SIRT3 for the changes in ribosomal proteins. The level of

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MRPL12 bound to ribosomes was found to be modulated by MRPL10 acetylation

compared to other ribosomal proteins. Therefore, we hypothesize regulation of MRPL12

binding to ribosomes by MRPL10 acetylation as a plausible mechanism to modulate

mitochondrial protein synthesis. In order to elucidate the role of acetylated lysine

residues mapped on MRPL10 in MRPL12 binding to the ribosome and translation, we

employed hybrid ribosomes constituted with bacterial ribosome without L7/L12 stalk and

mitochondrial stalk proteins, MRPL10 lysine mutants and MRPL12, in in vitro

translation activity assays. In addition, we compared the [35S]-methionine labeled

mitochondrial translation products from the cells transfected with MRPL10 lysine

mutants and their effect on cellular growth rates. Together, the data from these methods

serve as a framework for more directed future studies focusing on additional acetylated

lysine residues on MRPL10 which might be involved in MRPL12 binding to ribosomes.

Overall our findings suggest SIRT3-dependent reversible acetylation of MRPL10

might regulate the mitochondrial protein synthesis by modulating MRPL12 binding to the

ribosomes.

2.2 Introduction

OXPHOS provides over 90 % of the energy in the form of ATP to meet cellular

energy needs in mammalian mitochondria. Proteomic surveys of mitochondria acetylome

have revealed reversible acetylation of almost 30 % of the total mitochondrial proteome,

which are implicated in regulation of mitochondrial energy metabolism, signaling, and

apoptosis (1-6). Recently, it was discovered that GCN5L1 [GCN5(general control of

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amino acid synthesis 5)-like 1; also known as Bloc1s1], is one of the essential

components of the mitochondrial acetyltransferase system involved in acetylation of

electron transport chain proteins (7). Members of the class III histone deacetylases

(sirtuins), SIRT3, SIRT4, and SIRT5, have been found to reside in the mitochondria

(3,8,9). Sirtuins are homologs of yeast SIR2 (silent mating type information regulation 2)

and use NAD+ as a co-substrate, which suggests that they are nutrient sensitive

modulators regulating especially mitochondrial metabolic enzymes in response to the

[NADH]/[NAD+] ratio (10-12). Both SIRT3 and SIRT4 are required in an NAD+-

dependent manner for the maintenance of cell survival after genotoxic stress (13,14).

Genetic variations in the human Sirt3 gene have also been linked to longevity (13,14).

Also, SIRT3 expression in adipose tissue increases upon caloric restriction and cold

exposure (1,15). Mitochondrial acetyl-CoA synthetase 2 (ACS2) and glutamate

dehydrogenase (GDH) are the two key metabolic enzymes regulated by SIRT3 mediated

deacetylation (2,3,16).

Our previous findings about the importance of phosphorylation in mitochondrial

ribosomal proteins regulating mitochondrial protein synthesis served as a framework to

examine reversible acetylation of mitochondrial ribosomal proteins involved in regulation

of mitochondrial translation which is responsible for the synthesis of thirteen subunits

essential for the complexes in OXPHOS and for ATP production (17).

Mitochondria have their own genome encoding 2 rRNAs, 22 tRNAs, and 13

mRNAs that are utilized to synthesize essential subunits of the electron transport chain

complexes by its own translation machinery (18). The mitochondrial ribosome is similar

to the bacterial ribosome both structurally and functionally, but different in terms of

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having fewer rRNA and more proteins compared to the bacterial ribosome. The

mitochondrial ribosome (55S) is composed of a small subunit (28S) and a large subunit

(39S) containing 12S rRNA and 16S rRNA, respectively (19). Previously, our lab has

identified 77 mammalian mitochondrial ribosomal proteins, of which 29 are in the small

subunit and 48 are in the large subunit (20-24). Approximately half of these proteins have

homologs in bacterial ribosomes, while the remainders represent new classes of

ribosomal proteins (22,24). The functional core of the mitochondrial ribosome, which is

essential for proteins synthesis, is conserved according to cryo-EM reconstruction studies

(25). Notably, some mitochondrial ribosomal proteins have been mapped to regions

associated with disorders of mitochondrial energy metabolism (26). Changes in

expression levels and mutations in these ribosomal proteins affect mitochondrial protein

synthesis, cell growth, and apoptosis (27-31). Some of the ribosomal proteins with

bacterial homologs such as MRPS12, MRPS16, and MRPL12, have been shown to be

essential in mitochondrial protein synthesis (27,32-34). Particularly, the L7/L12 stalk

region, which consists of L10, L11, and multiple copies of L7/L12, is highly conserved

between species and is essential since it, along with the sarcin-ricin loop (SRL), forms the

GTPase active center that interacts with the translation initiation and elongation factors,

such as IF-2, EF-Tu, EF-G, and RF-3 (35-38). Previous studies also showed that when

L7/L12 was depleted, GTPase activation of both EF-Tu and EF-G was impaired (39).

Multiple L7/L12 dimers interact with L10 by protein-protein interactions. In Escherichia

coli and Bacillus subtilis, there are four copies of L7/L12 bound to the C-terminus of L10

as two dimers (40,41). On the other hand, in Thermotoga maritima and Thermus

thermophilus, six copies of L7/L12 as three dimers were found to associate with L10,

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which has additional residues at its C-terminus (42). However, the exact number of

MRPL12 needed to form the complete stalk in the mammalian mitochondria is not yet

known. In bacteria, it is known that post-translational modifications of L7/L12 by

phosphorylation and acetylation at the N-termini stabilize the interactions between L10

and L7/L12 (42,43). Moreover, changes in the L7/L12 content of ribosomes and L7/L12-

dependent translation activity in starvation and stationary phase of bacterial growth has

been reported (44-46). Similar to its bacterial homolog, the mitochondrial L7/L12

(MRPL12) is required for cell cycle progression and growth in Drosophila melanogaster

and accumulated during the G1-phase of growth stimulated cells (47-49). Changes in

mRNA expression levels and different isoforms of mitochondrial L7/L12 stalk proteins

were also reported to be critical for regulation of oxidative phosphorylation in cancer and

caloric restriction (50-52). In addition to its essential role in protein synthesis, free pools

of MRPL12 also couple mitochondrial translation to transcription by directly interacting

with the mitochondrial RNA polymerase to stimulate transcription in mitochondria

(32,53).

In this study, we provide evidence that mitochondrial protein synthesis is

regulated by reversible acetylation of MRPL10, and that its deacetylation by ribosome

associated SIRT3 in an NAD+-dependent manner modulates MRPL12 binding to

ribosome and thus the recruitment of translational factors during protein synthesis.

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2.3 Materials and Methods

2.3.1 Mitochondrial Ribosome Preparation and Reverse Phase – High Performance

Liquid Chromatography (RP-HPLC).

Preparation of mitochondrial ribosomes from bovine liver was adapted from

previously described methods (19, 21, 35, 36). To preserve the phosphorylation and

acetylation status of ribosomal proteins, phosphatase inhibitors (2 mM imizadole, 1 mM

Na3VO4, 1.15 mM Na2MoO4.2H2O, 1 mM NaF, 4 mM Na2C4H4O6.2H2O) and

deacetylase inhibitor (1 mM sodium butyrate) were added during the homogenization

process. Crude ribosomes prepared at 0.2, 0.4, and 1.6 % Triton X-100 concentrations

were loaded onto 10–30 % sucrose gradients and fractionated to isolate mitochondrial

55S ribosomes (16, 18, 34). To analyze rRNA dependent association of SIRT3 with

ribosomes, 20 A260 units of crude ribosome preparation obtained from bovine liver were

incubated in the absence and presence of RNase A and loaded onto another 10-30 %

linear sucrose gradient. After centrifugation, equal volumes of gradient fractions were

separated on 12 % SDS- polyacrylamide gels and immunoblotted with indicated

antibodies. Sirtuin inhibitor, 10 mM nicotinamide (NAM) was also used to treat pure

ribosomes to monitor acetylation changes and its effect on ribosomal activity in in vitro

translation assays with additional reaction with 10 µg/mL emetine as a cytoplasmic

translation inhibitor. For two-dimensional gel analysis, purified ribosomes were

sedimented by ultracentrifugation at 40,000 rpm for 5 h in a Beckman Type 40 rotor.

Acetone pellets of ribosome preparations (1.8 A260 units) were resuspended in two-

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dimensional lysis buffer consisting of 9.8 M urea, 2 % (w/v) Nonidet P-40, 2 %

ampholytes, pI 3–10 and 8–10, and 100 mM dithiothreitol (DTT). The samples were

loaded on non-equilibrium pH gradient electrophoresis tube gels and equilibrated in

buffer containing 60 mM Tris-HCl, pH 6.8, 2 % SDS, 100 mM DTT, and 10 % glycerol.

The 14 % second dimension gel was stained with Coomassie Blue, and the protein spots

corresponding to the acetylated ribosomal proteins were excised based on their locations

determined by immunoblotting analysis using anti-acetyl Lys antibody.

Approximately 5 A260 units of purified 55S ribosomes from bovine mitochondria

were incubated in 67 % glacial acetic acid for 16 h at 4oC to precipitate the rRNA. After

centrifuging at 18,000 g for 15 min, supernatant containing ribosomal proteins was

dialyzed in 6 % glacial acetic acid for 16 h at 4oC. HPLC analysis of ribosomal proteins

was conducted using a Shimadzu Model SCL-10Avp equipped with an SCL-10A diode

array detector (Shimadzu). The separation was performed on a 300 Å pore RP4 (5μm)

column (250 x 4.6 mm) (Eprogen Inc.). Solvent A was 0.1 % trifluoroacetic acid and

solvent B consisted of 0.1 % trifluoroacetic acid in acetonitrile. The gradient ranged from

5 % to 35 % B in 100 min, from 35 % to 55 % 100 min. The flow rate was 1.0 ml/min

and the column effluent was monitored at 215 nm.

2.3.2 Mass Spectrometric Analysis of Bovine Mitochondrial Ribosomal Proteins

In order to identify acetylated ribosomal proteins and the other ribosome-

associated proteins, in-gel and in-solution tryptic digestions were carried out using

ribosomes prepared at different Triton X-100 concentrations and analyzed by capillary

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liquid chromatography - nanoelectrospray ionization - tandem mass spectrometry

(capLC/MS/MS) (31,54,55). Tandem MS spectra obtained by fragmenting a peptide by

collision-induced dissociation (CID) were acquired using the capLC-MS/MS system that

consisted of a Surveyor HPLC pump, a Surveyor Micro AS autosampler, and an LTQ

linear ion trap mass spectrometer (ThermoFinnigan). The raw CID tandem MS spectra

were converted to Mascot generic files (.mgf) using the extract msn software

(ThermoFinnigan). Both, the .mgf and .raw files were submitted to site-licensed Mascot

(version 2.2) and Sequest search engines, respectively, to search against in-house

generated sequences of 55S proteins, all known human and bovine mitochondrial

proteins, and proteins in the Swiss-Prot database. The variable modifications were

methionine oxidation (+16 Da), acetylation of lysine residues (+42 Da) and

phosphorylation (+80 Da) of Ser, Thr, and Tyr residues. Up to 2 missed cleavages were

allowed for the protease of choice. Peptide mass tolerance and fragment mass tolerance

were set to 3 and 2 Da, respectively. Tandem MS spectra were manually evaluated at the

raw data level with the consideration of overall data quality, signal-to-noise of matched

peaks, and the presence of dominant peaks that did not match to any theoretical m/z

value.

2.3.3 In Vitro Deacetylation and Translation Assays

Deacetylation of ribosomal proteins was performed in the presence of NAD+ as a

substrate for the deacetylase using previously described methods (12,56). For this

reaction, 0.1 or 0.2 A260 units of sucrose gradient-purified or crude 55 S ribosomes,

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respectively, prepared in 0.2 % Triton X-100, were incubated with 3 mM NAD+ for 30

min at 37 °C in the presence and absence of 0.25 µg of recombinant mouse SIRT3.

Acetylated/deactylated ribosome samples were analyzed by immunoblotting using anti-

acetyl Lys antibody.

Poly(U)-directed in vitro translation assays were performed in 100 µl reactions

containing 50 mM Tris-HCl, pH 7.8, 1 mM DTT, 0.1 mM spermine, 40 mM KCl, 7.5

mM MgCl2, 2.5 mM phosphoenolpyruvate, 0.18 U pyruvate kinase, 0.5 mM GTP, 50 U

RNasin Plus, 12.5 µg/mL poly(U), 20 pmol [14C]-Phe-tRNA, 0.15 µM EF-Tumt, 1 µg EF-

Gmt, and varying amounts of mitochondrial ribosomes obtained from mitochondria. The

EF-Tumt and EF-Gmt were prepared from the recombinant proteins. The reaction mixtures

were incubated at 37°C for 15 min and terminated by the addition of cold 5 %

trichloroacetic acid followed by incubation at 90°C for 10 min. The in vitro translated

[14C] labeled-poly(Phe) was collected on nitrocellulose filter membranes and quantified

using a liquid scintillation counter.

2.3.4 Mouse Mitochondrial Ribosome Isolation

Mice in which the Sirt3 gene was targeted by gene trapping were obtained from

the Texas Institute for Genomic Medicine (Houston, TX) and liver tissues obtained from

Sirt3+/+, Sirt3+/-, and Sirt3-/- mice were kindly provided by Qiang Tong of Baylor College

of Medicine (57,58). The frozen liver tissues were resuspended in an isotonic

mitochondrial buffer (MB) (210 mM mannitol, 70 mM sucrose, 1 mM EDTA, 10 mM

HEPES-KOH, pH 7.5), supplemented with protease inhibitors (1 mM

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phenylmethylsulfonyl fluoride, 50 µg/ml leupeptin), and then homogenized in a Dounce

homogenizer on ice. The suspension was centrifuged at 400 g at 4 °C. This procedure was

repeated twice, and supernatants were centrifuged at 10,000 g at 4°C for 10 min to pellet

mitochondria. Mitochondria were lysed in a buffer containing 0.26 M sucrose, 20 mM

Tris-HCl, pH 7.6, 40 mM KCl, 20 mM MgCl2, 0.8 mM EDTA, 0.05 mM spermine, 0.05

mM spermidine, 6 mM β-mercaptoethanol, and 1.6 % Triton X-100, and the lysates were

loaded onto 34 % sucrose cushions and centrifuged at 100,000 g at 4°C for 16 h. The

crude ribosome pellets were resuspended in 20 mM Tris-HCl, pH 7.6, 40 mM KCl, 20

mM MgCl2 and 1 mM DTT.

2.3.5 Immunoblotting Assays

Protein samples for either one-dimensional or two-dimensional analysis were

loaded onto SDS-polyacrylamide gels and transferred to a polyvinylidene difluoride

(PVDF) membrane. The blot was probed with mouse monoclonal MRPS29 and HSP60

antibodies at 1:5000 dilution (BD-Transduction Laboratories), rabbit polyclonal MRPL10

antibodies against mouse and human at 1:3000 dilution and rabbit polyclonal human

MRPL41 and MRPL47 antibodies at 1:3000 (Covance), rabbit polyclonal anti-SIRT3

antiserum (against C-terminal domain of murine SIRT3) at 1:3000, mouse monoclonal

anti-FLAG M2-peroxidase at 1:3000 (Sigma-Aldrich), mouse monoclonal anti-acetyl Lys

antibody at 1:1000 (Cell Signaling Technology Inc.), and rabbit monoclonal His-tag

antibody at 1:5000 dilution (Rockland Immunochemicals). Oxidative phosphorylation

human antibody mixture (MitoProfile ® Total OXPHOS Antibody Cocktail,

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Mitosciences Inc.) was used at a 1:3000 dilution. The secondary antibody was

ImmunoPure antibody goat anti-mouse IgG (Pierce) at a 1:5000 dilution; goat antirabbit

IgG at a 1:1000 dilution; or Affinipure rabbit anti-mouse IgG, rabbit anti-goat IgG, or

goat anti-rabbit IgG (Jackson ImmunoResearch), all at a 1:10,000 dilution, followed by

development with the SuperSignal West Pico Chemiluminescent Substrate (Pierce)

according to the protocol provided by the manufacturer.

2.3.6 Plasmid Constructs

The human full-length MRPL10 coding sequence was amplified by reverse

transcription-PCR, using cDNA library of HeLa and the primer pair 5_-

AAACGGGGTACCATATGGGCTCCAAGGCTGTTACCCGC-3_ and 5_-

GCGGGTACCCTCGAGTTACTAATGGTGATGGTGATGATGCGAGTCCGGAACA

GTGTCAGG- 3_. For transient expression, the DNA fragments from MRPL10 was

digested with KpnI/XhoI, and then inserted into the pcDNA3.1-MycHis vector

(Invitrogen). MRPL10 mutants were created by site-directed mutagenesis (QuikChange_

II site-directed mutagenesis kits, Stratagene) at K124, K162, and K196. The primers for

K124A (AAG to GCG) forward (5_-cacaagatcctgatgGCggtcttccccaaccag) and reverse

(5_-ctggttggggaagaccGCcatcaggatcttgtg); K124Q (AAG to CAG) forward (5_-

cacaagatcctgatgCAggtcttccccaaccag) and reverse (5_-

ctggttggggaagaccTGcatcaggatcttgtg); K124R (AAG to AGG) forward (5_-

cacaagatcctgatgAGggtcttccccaaccag) and reverse (5_-

ctggttggggaagaccCTcatcaggatcttgtg); K162A (AAG to GCG) forward (5_-

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gaagagcccaaggtcGCggagatggtacgg) and reverse (5_-ccgtaccatctccGCgaccttgggctcttc);

K162Q (AAG to CAG) forward (5_-gaagagcccaaggtcCAggagatggtacgg) and reverse (5_-

ccgtaccatctccTGgaccttgggctcttc); K162R (AAG to AGG) forward (5_-

gaagagcccaaggtcAGggagatggtacgg) and reverse (5_-ccgtaccatctccCTgaccttgggctcttc);

K196A (AAG to GCG) forward (5_-ggctttatcaactactccGCgctccccagcctgccc) and reverse

(5_-gggcaggctggggagcGCggagtagttgataaagcc); K196Q (AAG to CAG) forward (5_-

ggctttatcaactactccCAgctccccagcctgccc) and reverse (5_-

gggcaggctggggagcTGggagtagttgataaagcc); K196R (AAG to AGG) forward (5_-

ggctttatcaactactccAGgctccccagcctgccc) and reverse (5_-

gggcaggctggggagcCTggagtagttgataaagcc) were utilized for the PCR amplification,

respectively. Mutated sequences are capitalized.

2.3.7 Cell Culture

The human embryonic kidney 293T (HEK293T) cells were cultured in DMEM

(Hyclone) supplemented with 10 % bovine calf serum (Hyclone, Logan, Utah), 100

IU/ml penicillin and 100 μg/ml streptomycin, at 37 oC and 5 % CO2 in a humidified

atmosphere. Mouse brown preadipocyte (HIB1B) cells with retroviral stable expression

of murine SIRT3 were previously described and were kindly provided by Qiang Tong of

Baylor college of Medicine (15,58). HIB1B cells were grown in the same complete

medium with puromycin (4 μg/mL).

HEK293T cells (4 X 105) were grown to about 80 % confluence in antibiotic-free

DMEM supplemented with 10 % FBS on 6 well plate. 4 μg of overexpression vector

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constructs and shRNA against MRPL12 (Open Biosystems Inc.) was used for

transfection experiments by using Lipofectamine 2000 (Invitrogen). After 2-4 days of

incubation with culture media, the transfected cells were lysed in cell lysis buffer (50 mM

Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 0.1 % SDS, and 0.5 % NP-40)

supplemented with a protease inhibitor cocktail (Sigma-Aldrich).

2.3.8 [35S]-Methionine Pulse-Labeling Assays

In order to measure mitochondrial protein synthesis activity using [35S]-

methionine, cells were cultured with dialyzed serum (25 mM Tris, pH 7.4, 137 mM

NaCl, 10 mM KCl) and minimum essential DMEM medium lacking methionine,

glutamine, and cysteine, after corresponding treatment. Cells were incubated with

emetine containing medium for 5 min to arrest cytoplasmic protein synthesis and added

0.2 mCi/ml of [35S]-methionine containing medium to label mitochondrially-encoded

thirteen proteins. After 2-4 h incubation, whole cell lysates were separated by SDS-

polyacrylamide gel which was dried and autoradiographed (58-60).

2.3.9 Preparation of Mitochondrial Ribosomes from Cell Lines

Approximately 4 X 107 HIB1B cells were collected and resuspended in lysis

buffer containing 50 mM Tris-HCl, pH 7.6, 0.26 M sucrose, 60 mM KCl, 20 mM MgCl2,

0.8 mM EDTA, 2 mM DTT, 0.05 mM spermine, 0.05 mM spermidine, proteinase

inhibitor cocktail (Sigma-Aldrich) by using 20 G needle. Cells were lysed by Dounce-

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homogenizer with addition of 1.6 % Triton X-100. Cell lysates were loaded onto 34 %

sucrose cushion buffer (50 mM Tris-HCl, pH 7.6, 60 mM KCl, 20 mM MgCl2, 6 mM

BME and 34 % sucrose) and centrifuged in a type 40 rotor (Beckman Coulter Inc.) at

40,000 rpm for 16 h to collect the crude ribosomes. Pellets were resuspended in 50 µL of

Base Buffer (50 mM Tris-HCl, pH 7.6, 60 mM KCl, 20 mM MgCl2, 1 mM DTT, and

protease inhibitor cocktail (Sigma-Aldrich). Ribosome suspensions were stored at -80 0C

for further analyses.

2.3.10 Complex I and IV Activity and ATP Determination Assays

Mitochondrial electron transport chain complex activity was measured by using

whole cell lysates prepared from HIB1B cells overexpressing SIRT3. Cells were

sonicated in a specific buffer for each enzyme activity and treated with n-dodecyl--

maltoside (1.67 μg/uL). The activity of Complex I was determined by monitoring the

reduction of 2,6-dichloroindophenolate (DCIP) at 600 nm as described (61). In brief, the

assay was performed in reaction buffer (25 mM potassium phosphate, pH 7.2, 5 mM

MgCl2, 2.5 mg/ml BSA, 2 mM KCN, 0.13 mM NADH, 60 μM DCIP, 65 μM

decylubiquinone, and 2 μg/mL antimycin A. The reaction was initiated by adding 100 μg

of cell lysate and reduction of DCIP was monitored for 10 min and another 10 min with

the addition of Complex I inhibitor, rotenone (2 μg/mL) to record rotenone insensitive

activity.

Complex IV (cytochrome c oxidase) activity was determined by measuring the

oxidation of ferrocytochrome c at 550 nm. The assay was performed in the presence of 10

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mM Tris-HCl, pH 7.0, 120 mM MgCl2, 25 mM sucrose, 11 μM ferrocytochrome c, and

varying amounts of cell lysate. Ferrocytochrome c was prepared by reducing it in 0.5 mM

DTT (62). The reaction was initiated with the addition of ferrocytochrome c to the

mixture and monitored for 3 min.

Freshly prepared HIB1B cell lysates in boiling water for 5 min were used to

measure cellular ATP concentration with ATP determination kit (Molecular Probe).

Chemiluminescent detection of ATP content was recorded as a function firefly luciferase

and luciferin by using Junitor LB 9509 Luminometer (Berthold Technologies). Protein

concentration of cell lysates was determined by BCA assay (Pierce) and RLU (relative

luminescent unit) was normalized to protein concentration.

Citrate synthase activity was determined by measuring the increase in absorbance

due to reduction of DTNB [5,5'-dithiobis-(2-nitrobenzoic acid)] at 412 nm. In the

presence of oxaloacetate, citrate synthase hydrolyzes thioester of acetyl-CoA resulting in

the formation of CoA with a thiol group. This thiol groups reacts with DTNB to form

TNB (5-thio-2-nitobenzoic acid), which is a yellow product observed by

spectrophotometry. About 100 µg of cell lysate was used for each measurement. The

assay was performed in the presence of 10 mM Tris-HCl, pH 7.0, 10 mM DTNB, 30 mM

acetyl-CoA, and varying amounts of cell lysate. The reaction was initiated with the

addition of 10 mM oxaloacetate to the mixture and monitored for 3 min.

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2.3.11 Expression and Purification MRPL10-MRPL12 Complex and Reconstitution

of Hybrid Ribosome

Human MRPL10 and MRPL12 (GenBank AB051618 and X79865) were

amplified from cDNA library of HeLa cells using the primers; MRPL10-NT1: Forward

5’-AAACGGGGTACCATATGGGCTCCAAGGCTGTTACCCGC-3’, Reverse 5’-

GCGGGTACCCTCGAGTTACTAATGGTGATGGTGATGATGCGAGTCCGGAACA

GTGTCAGG-3’; MRPL12-NT1: Forward 5’-

AAACGGCCATGGGTGCACCCCTGGATAACGCC-3’, Reverse 5’-

GCGGGTCTCGAGTTACTCCAGAACCACGGTGCC-3’. Corresponding MRPL10

mutants were created by site-directed mutagenesis (QuikChange_ II site-directed

mutagenesis kits, Stratagene) at K124, K162, and K196 by using the primers indicated in

plasmid constructs section. After cloning into pETDuet™-1 expression vector using NdeI-

XhoI for MRPL10 and NcoI-SalI for MRPL12, proteins were over-expressed by induction

with IPTG (200 μM) for 16 h at 18 °C. The cell lysate prepared with sonication was

centrifuged first 3,000 g for 15 min and then at 10,000 g for 30 min at 4 C.

Overexpressed complex proteins purified with Ni-NTA affinity column at 250 mM

imidazole, were dialyzed by using Spectra/Por 1 ® (SP, Spectrum Laboratories Inc.)

dialysis tubing in dialysis buffer (50 mM Tris, pH 8.0, 200 mM KCl, 1 mM DTT, and 10

% glycerol) to remove imidazole (63). The dialyzed sample was then applied to a strong

cation exchange gravity column (SCX, SP Sepharose Fast Flow column, GE Healthcare)

and eluted with step gradient salt of KCl; a linear gradient 50 mM –150 mM for 30 min,

400 mM – 500 mM for 10 min, and 500 mM – 1 M for 10 min. The fractions containing

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the MRPL10-MRPL12 complex were collected, combined, and dialyzed for further

analysis.

For the reconstitution of the hybrid ribosome, the L7/L12 stalk was removed from

bacterial 70S ribosomes by incubating in high salt extraction buffer (50 mM Tris-HCl, pH

7.6, 1 M NH4Cl, 20 mM MgCl2, 10 mM -mercaptoethanol) at 30 °C for 5 min followed

with addition of 1 mL of pre-warmed ethanol at 30 °C twice. Stripped ribosomes (70S-str)

were recollected by ultracentrifugation at 40,000 rpm for 16 h at 4 °C and ribosome pellet

was resuspended in reconstitution buffer (20 mM Tris-HCl, pH 7.5, 60 mM NH4Cl, 10

mM MgCl2, and 2 mM DTT). Stripped ribosomes (35 pmol) were incubated with purified

MRPL10-MRPL12 complex (70 pmol) and MRPL12 (280 pmol) at 37 °C for 15 min

before using in in vitro translation assay.

2.3.12 Statistical Analysis

Results are expressed as means ± s.d. of at least three independent experiments.

Statistical difference between test groups was analyzed by one-way ANOVA test.

Statistical significance was defined at P<0.05.

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2.4 Results and Discussion

2.4.1 The Mitochondrial Ribosomal Protein MRPL10 is Acetylated and SIRT3 is

Responsible for its Deacetylation

A combination of two-dimensional gel separation and capLC-MS/MS analyses

has been successfully used by our laboratory for the identification of ribosomal proteins

and their post-translational modifications (17,31,54,64,65). Acetylation of several

components of the translational machinery, either at N-terminal amino groups or at -

amino groups of Lys residues in bacteria, has been reported previously (66). To

determine the acetylated proteins of mammalian mitochondrial ribosomes, 55S ribosomes

were purified from the bovine liver using previously described methods (67,68).

Ribosomal proteins were then separated by one or two-dimensional gel electrophoresis,

and acetylated ribosomal proteins were identified by immunoblotting analysis with anti-

acetyl-Lys antibody, which detects protein only when it has been post-translationally

modified by acetylation on the -amino groups of Lys residues (Fig. 2.1). Protein bands

corresponding to acetylated proteins detected in the gel were excised, digested with

trypsin, and analyzed by capLC-MS/MS for identification as described in Materials and

Methods. The mass spectrometric analyses of one and two-dimensional gel spots revealed

the presence of two mitochondrial ribosomal proteins, MRPL10 (29.3 kDa) and MRPL19

(33.3 kDa) (Fig. 2.1, Table 2.1). After the cleavage of mitochondrial localization signal

sequence, their mature forms have very similar pIs and molecular masses and so they

58

were detected in the same protein band (Fig. 2.1). However, the analyses with mass

spectrometry didn’t reveal any acetylated peptides from MRPL19 in the same protein

band giving the acetylation signal (Table 2.1). Furthermore, we didn’t detect any

significant acetylation of MRPL19 when it was immunoprecipitated using anti-Flag

antibody from HEK293T cells transiently transfected with pcDNA3-Flag vector

containing its coding sequence compared to MRPL10 which was found to be clearly

acetylated in parallel experiment (58). For this reason, MRPL10 has been designated as

the major acetylated protein of the 55S ribosome.

59

Figure 2.1: Detection of acetylated mitochondrial ribosomal proteins.

Approximately 1.8 and 0.3 A260 units of purified bovine 55S ribosomes were

separated on two dimensional (2D) non-equilibrium pH gradient electrophoresis (2D-

NEPHGE) gel (courtesy of Dr. E. Koc) and one dimensional (1D) SDS-polyacrylamide

gel followed by detection of acetylated ribosomal proteins with anti-acetyl Lys antibody.

60

Peptide Sequence m/z Mr Score

(experimental) MRPL10 QKLoMAVTEYIAPKPVVNPR 725.1 2172.3 39QacKLoMAVTEYIAPKPVVNPR 738.8 2213.3 22LoMAVTEYIAPKPVVNPR 958.5 1915.1 72oMIAVCQNVAoMSAEDK 822.5 1642.9 96VFPNQILKPFLEDSK 888.4 1774.8 69YQNLLPLFVGHNLLLVSEEPK 1213.1 2424.1 96VKEoMVRILK 567.7 1132.3 21VacKEoMVRILK 588.5 1173.5 15ILacKpSVPFLPLLGGCIDDTILSR 858.0 2571.0 47QGFINYpSacKLPpSLALAQGELVGGLpTLLTAR 1104.6 3310.9 54LPSLALAQGELVGGLTLLTAR 1048.2 2094.3 145THSLLQHHPLQLTALLDQYAR 1229.3 2455.8 78QQLEGDPVVPASAQPDPPNPVQDS 829.8 2486.3 83 MRPL19 FLSPEFIPPR 602.4 1202.8 79ILHIPEFYVGSILR 829.4 1656.8 68LDDSLLYLR 555.2 1108.4 87DALPEYSTFDMNMKPVAQEPSR 843.5 2527.3 80WSQPWLEFDoMMR 822.1 1642.2 62IEAAIWNEIEASK 737.7 1473.5 91

Collision-induced dissociation spectra of peptides were searched by MASCOT as

described. Modifications of peptides by acetylation (ac), phosphorylation (p), and

oxidation (o) were shown for each peptide. Courtesy of Dr. H. Koc.

Table 2.1: Peptides detected from tryptic digests of mitochondrial ribosomal protein

bands corresponding to acetylation signals by capLC-MS/MS analysis.

61

For further confirmation of MRPL10 acetylation and mapping of the acetylated

Lys residues, mitochondrial ribosomal proteins isolated from bovine 55S ribosomes were

separated on reverse phase – high performance liquid chromatography (RT-HPLC) and

tryptic digests of the band(s) corresponding to the acetylated MRPL10 was analyzed by

capLC-MS/MS (Fig. 2.2 and Table 2.1). Acetylated and non-acetylated forms of

MRPL10 were well separated on the C4-reverse phase column, and the acetylated form

of MRPL10 was eluted in the later fraction as expected due to the neutralization of

positive charges on Lys residues by more hydrophobic acetyl groups. In the

immunoblotting analysis of HPLC fractions by anti-MRPL10 and anti-acetyl-Lys

antibodies, several different forms of MRPL10 were observed. This observation might be

because of differential modification(s) of MRPL10 by post-translational modifications

such as acetylation and phosphorylation (Fig. 2.2). Depending on the relative signal

intensities of acetylated and non-acetylated MRPL10, acetylated MRPL10 can be

estimated to be 30 % of the total MRPL10 found in the dissociated 39S subunits and 55S

ribosomes (Fig. 2.2). However, this estimation is based on recognition of all these

different forms of MRPL10 by anti-MRPL10 antibody equally (Fig. 2.2).

62

Figure 2.2: Purification of MRPL10 and its acetylated forms by reverse phase -

HPLC.

Approximately, 5 A260 units of purified 55S ribosome preparation obtained from

bovine liver were incubated in the presence of glacial acetic acid as described in

Materials and Methods. After centrifugation, supernatant containing the ribosomal

proteins was separated with RP-HPLC using 5-55 % acetonitrile gradient. Equal volumes

of gradient fractions were separated on 12 % SDS-polyacrylamide gels and analyzed with

anti-acetyl Lys and anti-MRPL10 antibodies. Acetylated protein bands corresponding to

MRPL10 protein were shown by arrows.

63

In addition, we also performed capLC-MS/MS analysis to map the acetylated Lys

residues from the tryptic digests of acetylated MRPL10 fractions. This analysis enabled

us to map several highly conserved acetylated Lys residues (Lys 46, 124, 162, 169, and

196) in bovine 55S ribosomes (Table 2.1). The majority of the Lys residues are highly

conserved in human, bovine, and mouse MRPL10 proteins (Fig. 2.3). In order to

investigate the contribution of highly conserved Lys residues in the acetylation of

MRPL10, wild type, double (Lys162Ala and Lys196Ala), or triple (Lys124Ala,

Lys162Ala, and Lys196Ala) mutants of MRPL10 were stably expressed in HeLa cells.

After the enrichment of the proteins using His-tag affinity chromatography, (58). The

acetylation status of MRPL10 gradually declined in both double and triple Lys to Ala

mutants compared to wild type (58).

64

Figure 2.3: Alignment of MRPL10 with identified acetylated lysine residues.

Primary sequence alignment of MRPL10 homologs from different species, human

(NP660298), bovine (XP592952), mouse (NP080430), archaea (Thermotoga maritima

NP228266), and bacteria (Escherichia coli AAC43083) was created using the

CLUSTALW program in Biology Workbench and is displayed in BOXSHADE.

(*) denotes the acetylated lysine residues detected in the capLC-MS/MS analysis.

*

*

*

*

*

65

Figure 2.4: Structural model for the location of acetylated lysine residues on

ribosome and ribosomal L7/L12 stalk region.

Crystal structures of bacterial ribosome (E. coli Protein databank entry 2AW4)

and archaeal L7/L12 stalk (T. maritima PDB: 1ZAX) were used to illustrate the location

of acetylated lysine residues (colored in red). Mitochondrial homologs of L7/L12 stalk

proteins L11 and L10 in complex with three L7/12 N-terminal-domain dimers were

shaded in yellow, green, and orange, respectively. The 50S ribosomal rRNA and

ribosomal proteins were colored in blue and pink, respectively. Pymol software from

DeLano Scientific LLC was used to generate structural models (69). Courtesy of Dr. E.

Koc.

66

2.4.2 SIRT3, NAD+-dependent Deacetylase is Associated with 55S Mitochondrial

Ribosome.

Three members of the sirtuin family NAD+-dependent deacetylases, SIRT3,

SIRT4, and SIRT5, have been identified and localized in the mitochondria (6-8). They

have been implicated in several mitochondrial metabolic pathways. SIRT3 targets

Complex I subunit, NDUFA9 and Complex II SdhA; SIRT4 targets glutamate

dehydrogenase; SIRT5 targets carbamoyl phosphate synthetase 1 (57,70-72). Next, we

investigated whether any of the deacetylases mentioned above are associated with the

ribosome. Crude mitochondrial ribosomes were isolated at different salt and non-ionic

detergent concentrations to preserve their interactions with associated proteins and were

then fractionated on 10-30 % linear sucrose gradients (Fig. 2.5). Immunoblotting analyses

of the sucrose gradient fractions antibodies against acetyl-Lys, MRPL41, and SIRT3

revealed that SIRT3 containing fractions (12-18) overlapped with the acetylated-

MRPL10 and another large subunit ribosomal protein MRPL41 (Fig. 2.5). In order to

confirm the association of SIRT3 with mitochondrial ribosomes, ribosome preparations

were treated with RNase A and loaded on 10-30 % gradients (Fig. 2.6). RNase A

treatment only digests rRNAs of 55S ribosomes. Proteins physically associated with the

rRNA were spread through the gradient and no significant amounts of SIRT3 or MRPL10

were retained in the gradients (Fig. 2.6).

In our analysis, we have found SIRT3 association with the mitochondrial

ribosome only at low detergent and ionic conditions, implying a possibly transient

interaction. In addition to the immunoblotting analysis, in-gel proteolytic digestion and

67

mass spectrometric analysis of the protein band detected with the anti-SIRT3 antibody

confirmed the association of SIRT3 with the ribosome (Table 2.2). Since the other

mitochondrial sirtuins, SIRT4 and SIRT5, were not detected in our analyses of ribosome

preparations, we concluded that SIRT3 is the mitochondrial deacetylase associated with

the ribosome. This finding is supported with a recent report stating that SIRT3 is the

major mitochondrial deacetylase in mice where no mitochondrial hyperacetylation is

detectable when the two other mitochondrial sirtuins, SIRT4 and SIRT5, are deleted (16).

68

Figure 2.5: The association of SIRT3 with mitochondrial 55S ribosomes.

Crude mitochondrial ribosomes isolated from bovine liver were layered onto 10-

30 % linear sucrose gradients in order to sediment 55S ribosomes. Immunoblotting

analyses of corresponding fractions separated on 12 % SDS-polyacrylamide gel were

performed using antibodies against acetylated MRPL10, SIRT3, and MRPL41 to

demonstrate the co-sedimentation of SIRT3 with the 55S ribosome.

69

Figure 2.6: Dissociation of SIRT3 from the large subunit of mitochondrial ribosomes

by RNase A treatment.

Approximately, 20 A260 units of crude ribosome preparation obtained from bovine

liver were incubated in the absence (A) and presence of RNase A (B) and loaded onto

two 10-30 % linear sucrose gradients. After centrifugation, equal volumes of gradient

fractions were separated on 12 % SDS-polyacrylamide gels and PVDF blots were probed

with SIRT3, MRPL10, acetyl-Lys and MRPL41 antibodies.

70

Courtesy of Dr. H. Koc.

Table 2.2: Peptides detected from tryptic digests of mitochondrial ribosome

associated bovine SIRT3 by capLC-MS/MS analysis.

Peptide Sequence m/z Mr Score

(experimental)

KFLLQDIAELIK 717.5 1433.0 101 LYTQNIDGLER 662.8 1323.5 73 LVEAHGSLASATCTVCR 860.2 1718.3 56 DVAQLGDVVHGVK 669.5 1337.1 63 LVELLGWTDDIQDLIQR 1015.7 2029.3 104

71

The interaction of SIRT3 with MRPL10 was investigated further with GST-pull-

down and co-immunoprecipitation assays to map the domains mediating this binding

(58). The first 148 amino acid (aa) residues of the MRPL10 N-terminal domain and

residues aa 156-171 on SIRT3 are found to be important for their association. These

domains were also evaluated using a structural model based on the coordinates of human

SIRT3 and the Thermotoga maritima L10-L7/L12 complex (Fig. 2.7) (73). As illustrated

in Fig. 2.7, the region within the first 148 residues of MRPL10 (marked in pink) required

for SIRT3 binding is probably the most accessible region for this interaction since

MRPL10 also interacts with MRPL11 and MRPL12 to form the mitochondrial L7/L12

stalk. The C-terminal helix 8 of Thermotoga L10 binds three L7/L12 dimers (two

L7/L12 dimers in E. coli) and the N-terminal globular domain interacts with the N-

terminal domain of L11 and SIRT3 (Fig. 2.7). Regions of MRPL10 and SIRT3

responsible for binding to each other, and their interactions with the rest of the ribosome,

are in agreement with the idea of a transient interaction between SIRT3 and the

mitochondrial ribosome. Moreover, acetylated Lys residues in MRPL10 are located near

the SIRT3 binding domain and reversible acetylation/deacetylation of these residues

could be involved in the regulation of ribosomal function by NAD+-dependent SIRT3

(Fig. 2.7). The L7/L12 stalk is known to be essential as well as the most highly regulated

and flexible region of the large subunit due to its interaction with elongation, initiation

and release factors during different stages of translation (37,74,75). For this reason, it is

plausible to suggest that the mitochondrial translational machinery might be regulated by

reversible acetylation of the L7/L12 stalk protein, MRPL10.

72

K169

K196

K162

L7/12

L10 NTD

L10 CTD

K124L10

A B

DI

DII

DIII

39S Subunit

L1 stalk

L7/12 stalkL11 L10 PTC

SRL

SIRT3

CP

Figure 2.7: Structural model of the SIRT3 and MRPL10 interactions in the L7/L12

stalk.

A. Crystal structure model of the human SIRT3 was generated (Protein Data Bank

entry 3GLU) where MRP-L10 interaction site and acetyl-CoA synthetase 2 peptide at the

active site are represented in green and red, respectively. B. Structure of the L10-L7/L12

complex from T.maritima (1ZAX) was used to model L7/L12 stalk in mitochondria. In

the model, MRPL10 was colored in green, pink to represent SIRT3 binding site, and red

for the conserved Lys residues found to be acetylated in bovine MRPL10 (shown by

asterisks in Fig. 2.3). The three L7/L12 dimers (DI, DII, DIII) were colored in yellow.

C

73

C. Models of the human SIRT3 and T. maritima L10-L7/L12 complex were used

to represent their possible interactions with 55S mitochondrial ribosomes using

coordinates from the E. coli 50S subunit (2AW4). The 50S ribosomal rRNAs, L10, L11,

and SIRT3 were colored in blue, green, yellow, and pink, respectively. The other

functional regions such as peptidyl transferase center (PTC), central protuberance (CP),

sarcin-ricin loop (SRL), L1, and L7/L12 stalks of the large subunit and ribosomal

proteins (salmon) were labeled in the model. The structural model was generated using

PyMol software (DeLano Scientific) (69). Courtesy of Dr. E. Koc.

74

2.4.3 Recombinant and Ribosome-Associated Endogenous NAD+-dependent SIRT3

Deacetylates MRPL10

Given that the ribosomal protein MRPL10 is acetylated and interacts directly with

the NAD+-dependent deacetylase SIRT3, we also investigated the effects of SIRT3 by

performing in vitro deacetylation assays using purified bovine mitochondrial ribosomes

containing acetylated MRPL10 as the SIRT3 substrate. The NAD+-dependent

deacetylation of the ribosome associated MRPL10 by recombinant SIRT3 was detected

by immunoblotting analysis with anti-acetyl-Lys antibody (Fig. 2.8). In the presence of

NAD+, the acetylation level of the protein band containing MRPL10 significantly

decreased in the absence of SIRT3, presumably due to the catalytic activity of the

ribosome-associated endogenous SIRT3. This observation also confirmed the presence of

the ribosome-associated SIRT3 in the mitochondrial ribosome preparations. Furthermore,

the addition of recombinant SIRT3 to the in vitro deacetylation assay reaction further

decreased the acetylation level of MRPL10. In contrast, acetylated HSP70, which is

approximately 75 kDa and cosedimented with the ribosome, was neither deacetylated by

an endogenous deacetylase nor by the recombinant SIRT3 in vitro (Fig. 2.8).

We also assessed the acetylation level of MRPL10 in a SIRT3 knock-out (Sirt3-/-)

mouse to evaluate whether SIRT3 was the major deacetylase responsible for the

deacetylation of MRPL10. Analysis of mitochondrial ribosomes isolated from SIRT3

knock-out (Sirt3-/-), wild-type (Sirt3+/+), and heterozygote (Sirt3+/-) mice revealed that the

MRPL10 in the Sirt3-/- mice was more heavily acetylated than in either of the other two

more strains (Fig. 2.9). Immunoblottings were also probed with mouse anti-MRPL10 and

75

anti-MRPS29 antibodies to ensure equal loading of mitochondrial ribosomal proteins

(Fig. 2.9). The increase in acetylation of glutamate dehydrogenase (GDH) confirms the

hyperacetylation state of Sirt3-/- mice (Fig. 2.9)(16).

Overall, the data we obtained strongly suggest that the mitochondrial SIRT3 is

associated with the mitochondrial ribosome and also involved in specific deacetylation of

MRPL10. Moreover, in the absence of SIRT3, the amount of MRPL10 acetylation was

increased compared to the MRPL10 isolated from wild type and heterozygote mice

expressing SIRT3.

We also performed poly(U)-directed in vitro translation assays to monitor

mitochondrial ribosome activity as a function of the acetylation status of MRPL10. This

assay is one of the primary techniques in assessing the interactions of mitochondrial

elongation factors with the L7/L12 stalk of the ribosome (76,77). To test the role of

reversible acetylation in mitochondrial ribosome activity using this in vitro assay, we first

isolated mitochondrial ribosomes from SIRT3 knock-out (Sirt3-/-), wild-type (Sirt3+/+)

and heterozygote (Sirt3+/-) mouse liver mitochondria and performed poly(U)-directed

poly(phenylalanine) synthesis in the presence of [14C]-tRNAPhe and the mammalian

mitochondrial elongation factors EF-Tumt and EF-G1mt (77). To ensure that equal

amounts of mitochondrial ribosomes were used in each in vitro translation assay,

ribosomes were quantified using A260 measurements and immunoblotting analysis was

performed with anti-MRPS29 antibody as an equal loading control (Fig. 2.9). In these

assays, mitochondrial ribosomes isolated from Sirt3-/- mice had a 50-60% higher

translational activity compared to those from Sirt3+/+ or Sirt3+/- mice (Fig. 2.9).

76

Figure 2.8: Deacetylation of MRPL10 by the NAD+-dependent deacetylase, SIRT3.

In vitro deacetylation reactions using approximately 0.1 A260 units of 55S bovine

mitochondrial ribosomes were performed in the presence of 3 mM NAD+ and 0.2 μg of

recombinant SIRT3 as labeled. Immunoblotting analyses using anti-acetyl Lys antibody

to detect acetylated MRPL10 indicate the specific deacetylation of MRPL10, but not the

acetylated HSP70 sedimented with ribosomes, by endogenous and recombinant SIRT3 in

the presence of NAD+.

77

Figure 2.9: Role of MRPL10 acetylation in SIRT3 knock-out (Sirt3-/-) mice.

Immunoblotting analyses with mitochondrial ribosomes prepared from Sirt3-/-,

Sirt3-/+, and Sirt3+/+ mice liver using anti-acetyl Lys antibody indicate the elevated

acetylation of ribosomal protein MRPL10 as well as glutamate dehydrogenase (GDH)

which was used as a control for hyperacetylation in absence of SIRT3. In order to ensure

equal loading, immunoblots were probed with anti-MRPL10 and anti-MRPS29

antibodies. Acetylated ribosomes promote mitochondrial protein synthesis in vitro.

Mitochondrial ribosomes (0.05-0.1 A260 units) isolated from Sirt3+/+, Sirt3+/-, and Sirt3-/-

mice liver mitochondria were used in the poly(U)-directed in vitro translation assays. One

way ANOVA test was used to compare the significance of values from three separate

experiments. Asterisk denotes p<0.05.

78

Since nicotinamide (NAM) is a general inhibitor for sirtuins, including SIRT3,

mitochondrial ribosomes prepared in the presence or absence of nicotinamide and

cytoplasmic translation inhibitor emetine were also examined by poly(U)-dependent

translation assays (Fig. 2.10 ). In these assays, we first verified the increased acetylation

of mitochondrial ribosomes prepared in the presence of nicotinamide and tested the

translation activity of these ribosomes. In agreement with the increased activity obtained

in Sirt3-/- mice, the translation activity of mitochondrial ribosomes from nicotinamide

treated mitochondria was significantly higher and this activity was not inhibited by

cytoplasmic translation inhibitor emetine (Fig. 2.10). These results from SIRT3 knock-

out (Sirt3-/-) mice and NAM-treated bovine mitochondria indicate that ribosomes

containing acetylated MRPL10 are more active in protein synthesis in vitro.

79

Figure 2.10: Role of nicotinamide (NAM) and emetine on mitochondrial protein

synthesis.

A. Immunoblotting analysis of mitochondrial ribosomes (0.2 A260) prepared in

the absence (Con) and presence of 10 mM nicotinamide (NAM) from bovine liver

mitochondria probed with anti-acetyl Lys, MRPL10 and MRPS29 antibodies. Relative

change in MRPL10 acetylation was graphed. B. Mitochondrial ribosomes (0.1-0.2 A260

units) isolated from control and NAM treated bovine liver mitochondria were used in the

poly(U)-directed in vitro translation assays described in Materials and Method. In vitro

translation assays using NAM treated ribosomes were also repeated by adding 1 mM

emetine (Dr. M.J. Han contributed to this experiment). One way ANOVA test was used

to compare the significance of values from three separate experiments. Asterisk denotes

p<0.05.

*

*

80

We have also demonstrated the increased expression of mitochondrially encoded

cytochrome c oxidase subunit II (Complex IV, COII) compared to the nuclear-encoded

components of the oxidative phosphorylation complexes in SIRT3 knock-out mice

mitochondria (Fig. 2.11).

In SIRT3 knock-out mice, MRPL12 signal, indicated with an arrow, was slightly

increased compared to the other mitochondrial ribosomal proteins (Fig. 2.11). The

ribosomal protein MRPL10 is located in the L7/12 stalk of the ribosome and responsible

for binding of multiple copies of MRPL12. The components of this stalk are universally

conserved since they are essential for translocation of tRNA catalyzed by EF-G-

dependent GTP hydrolysis (78,79). Therefore, MRPL12 antibody was specifically chosen

to monitor changes in MRPL12. Mitochondrial ribosomes enriched from SIRT3 knock-

out mice by sedimentation through a 34% sucrose cushion and were evaluated for the

mitochondrial ribosome content using immunoblotting assays (Fig. 2.12). MRPL12 band

corresponding to lower molecular mass in the mitochondrial lysate was significantly

increased compared to MRPL10 and MRPL47. The MRPL10 and MRPL47 bands

displayed fewer signals in SIRT3 knock-out ribosome sample compared to wild type,

possibly due to less loading (Fig. 2.12). The in vitro translation assays were also repeated

and stimulation of mitochondrial translation in knock-out mice was confirmed with

increased acetylation of MRPL10 (Fig. 2.12). This observation implies that the

acetylation of MRPL10 in SIRT3 knock-out mice may stabilize MRPL12 binding to the

ribosome possibly contributing to elevated mitochondrial translation activity.

81

Figure 2.11: Enhanced MRPL12 binding to the ribosome in SIRT3 knock-out mice.

Mitochondrial lysates from wild-type (Sirt3+/+) and knock-out (Sirt3-/-) were

separated on 12 % SDS-polyacrylamide gel and probed with OXPHOS antibody cocktail

to examine the effect of enhanced MRPL10 acetylation on the steady-state levels of

complex subunits. Mitochondrially encoded subunit of Complex IV, COII in SIRT3

knock-out mice is enhanced while no difference is observed in other subunits. Protein

blots were probed with anti-acetyl Lys antibody to show increased acetylation of

mitochondrial proteins and MRPL10 (indicated by arrows) in the absence of SIRT3 and

with HSP60, MRPL47, and MRPL10 antibodies to ensure equal protein loading.

MRPL12 signal displays an increase in its shorter form (indicated by an arrow).

82

Figure 2.12: Role of MRPL10 acetylation in mitochondrial translation in SIRT3

knock-out mice.

Mitochondrial ribosomes were enriched from wild-type (Sirt3+/+) and SIRT3

knock-out (Sirt3-/-) mice liver mitochondria to evaluate changes in ribosomal protein

content by immunoblotting analyses. Approximately, 0.05 A260 units of ribosomes were

separated on 12 % SDS-polyacrylamide gel for the comparison. Immunoblots were

developed with antibodies as indicated in the figure. Arrows indicate the MRPL10

acetylation and ribosome bound MRPL12 (shorter form). Poly(U)-directed in vitro

translation assays were also conducted to demonstrate enhanced protein synthesis in

SIRT3 knock-out mice. Mitochondrial ribosomes (0.1 A260 units) enriched from wild-type

(Sirt3+/+) and SIRT3 knock-out (Sirt3-/-) mice liver mitochondria were used in in vitro

translation assays. The values are the mean SD for three separate measurements. One

way ANOVA test was used to compare the significance of values. Asterisk denotes

p<0.05.

*

83

The two different forms of MRPL12 in mitochondria were characterized by two-

dimensional gel electrophoresis separation of mitochondrial proteins followed by

immunoblotting with MRPL12 antibody and identification of corresponding protein

bands with capLC-MS/MS analysis (Fig. 2.13).

84

Figure 2.13: Identification of two different forms of MRPL12.

Mitochondrial ribosome were separated on two dimensional gels and protein

bands displaying MRPL12 signal after immunoblotting analysis were labeled with arrows

on Coomassie Blue-stained gel. Primary sequences of MRPL12 from bovine, mouse,

human and L12 from bacteria (E. coli) were aligned using CLUSTALW program in

Biology Workbench and is displayed in BOXSHADE. The N-terminal sequences of the

long and short form of MRPL12 are shown by arrows, and the calculated molecular

weights are 17.8 kDa and 16.5 kDa, respectively.

85

2.4.4 Overexpression of SIRT3 Regulates Mitochondrial Protein Synthesis in HIB1B

Cells

The next goal is to examine potential mechanisms to elucidate the role of SIRT3-

dependent deacetylation of MRPL10 and/or ribosomal proteins in protein synthesis. For

this purpose, we employed brown adipocyte cells, HIB1B, overexpressing SIRT3 to label

mitochondrially encoded proteins by [35S]-methionine. The stable expression of the full-

length murine SIRT3 clone decreased the de novo synthesis of mitochondrially encoded

proteins by about 40 % in agreement with our SIRT3 knock-out mice data (Fig. 2.14)

In bacteria, multiple copies of L7/L12 form a very flexible stalk along with L10 to

recruit elongation factors during translation (37,80,81). Previous reports showed that the

bacterial homolog of MRPL12 plays a role in the recruitment of several different

GTPases, such as IF2, EF-Tu, RF-G, and RF3, that function in translation (35,37,79,82).

In order to understand the mechanism involved in regulation of translation by acetylation

of MRPL10, we focused our efforts on MRPL12 which binds to the ribosome by

interacting only with MRPL10 involving only protein-protein interaction. Therefore, it is

plausible to suggest that reversible acetylation of MRPL10 might change the L7/L12

stalk composition to modulate mitochondrial translation in response to changes in SIRT3

activity. Interestingly, the observed increase in MRPL12 incorporation into the

mitochondrial ribosome in SIRT3 knock-out mice without changes in other ribosomal

proteins supports this hypothesis (Fig. 2.12).

86

Figure 2.14: Role of SIRT3 over-expression on mitochondrial protein synthesis in

HIB1B cells.

Control HIB1B cells stably expressing the empty vector (Con) and full-length (1-

334 aa) murine SIRT3 (SIRT3) were exposed to [35S]-methionine in the presence of a

cytosolic translation inhibitor, emetine. A representative electrophoretic pattern of the de

novo synthesized translational products was presented. ND1, -2, -3, -4, -4L, -5, and -6 are

subunits of Complex I; Cytb is subunit of Complex III; COI, -II, and -III are subunits of

Complex IV; ATP6 and ATP8 are subunits of the Complex V. Coomassie Blue staining

of the same gel was performed to ensure equal protein loading in the gel. The combined

intensities of mitochondrially encoded proteins from each lane were used as the overall

quantitation of the mitochondrial protein synthesis (Dr. M. J. Han contributed to this

experiment). One way ANOVA test was used to compare the significance of

measurements from three experiments. Asterisk denotes p<0.05.

*

87

2.4.5 Effect of SIRT3 Overexpression on MRPL12 Binding to the Ribosome in

HIB1B Cells

To examine the effect of SIRT3 over-expression on MRPL12 binding to the

mitochondrial ribosome, we performed immunoblotting analyses to check the expression

of MRPL12 and incorporation of MRPL12 into the mitochondrial ribosome in control

and SIRT3 over-expressing HIB1B cells. We isolated mitochondria from each cell line

and checked expression level of MRPL12. Flag-tagged ectopically expressed SIRT3 was

confirmed using an anti-Flag antibody. Immunoblotting analyses with anti-MRPL10,

anti-MRPL47, anti-SdhA, and anti-HSP60 antibodies were performed to ensure equal

loading amounts (Fig. 2.15). However, no significant change was detected in expression

level of MPRL12 in mitochondrial lysates between HIB1B and SIRT3 over-expressing

cells. To investigate the possible flexible binding of MRPL12 on the mitochondrial

ribosome, immunoblotting analyses were performed using ribosomes enriched from

HIB1B and SIRT3 over-expressing cells. These results showed a significant reduction in

MRPL12 binding on the ribosome (Fig. 2.15). Furthermore, immunoblotting with anti-

acetyl Lys antibody showed reduced acetylation of MRPL10 in SIRT3 over-expressing

cells. Probing with anti-MRPL10 and anti-MRPL47 antibodies was also performed to

ensure equal loading amounts (Fig. 2.15).

88

Figure 2.15: Effect of SIRT3 over-expression on MRPL10 acetylation and MRPL12

binding to the ribosome in HIB1B cells.

A. Approximately, 20 g of mitochondrial lysates from control and SIRT3

expressing cells were separated on 12 % SDS-polyacrylamide gel, and immunoblotting

analyses were performed with corresponding antibodies (Dr. M. J. Han contributed to this

experiment.). B. Over-expression of the Flag-tagged SIRT3 decreased acetylation of

MRPL10 which reduced MRPL12 binding on mitochondrial ribosomes in HIB1B cells.

Approximately, 0.05 A260 of mitochondrial ribosomes, which were enriched from control

and SIRT3 over-expressing cells by centrifuging through 34 % cushion buffer, was

separated on 12 % SDS-polyacrylamide gel, and immunoblotting analyses were performed

using antibodies as indicated.

89

The changes in the translational activity of mitochondrial ribosomes which

synthesize thirteen essential proteins of the electron transport chain complexes might

result in changes of complex activities and ATP production. As shown in Fig. 2.16 and

Fig. 2.17, reduction in the activities of Complexes I and IV was in agreement with the

decline in protein synthesis in SIRT3 over-expressing cells. Not only was the activity of

Complex I and IV reduced in SIRT3 over-expression cells, but also total ATP production

(Fig. 2.18). These effects appeared to be specific, since citrate synthase activity, a key

enzyme of the Krebs cycle and indicator of mitochondrial function, was comparable in

control and SIRT3 over-expressing cells (Fig. 2.19). We observed over 40 % of reduction

in 13 mitochondrially-encoded proteins in SIRT3 over-expressing HIB1B cells; however,

the production of ATP was reduced by only about 20-30 % compared to control cells

(Fig. 2.18). This discrepancy could be explained by direct regulation of OXPHOS

complexes by SIRT3-dependent deacetylation. We and other groups reported that

activities of Complexes I and II are stimulated by SIRT3-deacetylation of their nuclear

encoded subunits (57,72). This implies that the activities of OXPHOS complexes and

ATP production could be regulated by both acetylation and translation of their

components in a SIRT3-dependent manner to maintain basal ATP level (57,72,83).

90

Figure 2.16: Complex I activity in HIB1B cells overexpressing SIRT3.

Whole cell lysates were prepared from HIB1B control (Con) and SIRT3 over-

expressing (SIRT3) cells. Cells were lysed by n-dodecyl--maltoside and sonication in

ice cold water. The activity of Complex I was determined by monitoring the reduction of

2,6-dichloroindophenolate (DCIP) at 600 nm as preciously described in Materials and

Methods. The reaction was initiated by adding 100 μg of cell lysate, and the reduction of

DCIP was monitored for 10 min at 600 nm. After adding Complex I inhibitor, rotenone,

samples were monitored for an additional 10 min. Only the rotenone sensitive Complex I

activity was presented. The results were expressed as the percent of control. The mean ±

SD was calculated from three independent experiments. Value of ٭P < 0.05 was

considered statistically significant. One way ANOVA test was used to compare the

significance of values.

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Figure 2.17: Effect of SIRT3 overexpression on Complex IV activity.

Complex IV activity was determined by measuring the oxidation of

ferrocytochrome c at 550 nm and reported as percent of relative change in OD550. About

100 µg of cell lysate was used for each measurement. One way ANOVA test was used to

compare the significance of values. Asterisk denotes for ٭P < 0.05.

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Figure 2.18: ATP production in HIB1B cells overexpressing SIRT3.

ATP production as a function of luciferase activation measured at 560 nm using

2-10 µg of cell lysates. Relative luminescent unit (RLU) was normalized by the protein

concentration of each lysate. The results were expressed as the percent of control. The

mean ± SD was calculated from three independent experiments (Courtesy of Dr. M. J.

Han). One way ANOVA test was used to compare the significance of values. Value of ٭P

< 0.05 was considered statistically significant.

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Figure 2.19: Citrate synthase activity in HIB1B cells overexpressing SIRT3.

Citrate synthase activity was determined by measuring the increase in absorbance

due to reduction of DTNB [5,5'-dithiobis-(2-nitrobenzoic acid)] at 412 nm, coupled to the

reduction of CoA by citrate synthase in the presence of oxaloacetate. About 100 µg of

cell lysate was used for each measurement.

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2.4.6 The Effect of MRPL12 Knockdown and MRPL10 and MRPL12

Overexpression on Mitochondrial Protein Synthesis.

The composition of the L7/L12 stalk and the nature of MRPL10 and multiple

copies of MRPL12 interactions are unknown in mitochondria. In bacteria, multiple copies

of L12 bind to L10 via protein-protein interactions to form the L7/L12 stalk. The

interaction between L10 and L12 is sensitive to ionic conditions. We employed shRNA

mediated knockdown of MRPL12 (shL12) in HEK293T cells in order to verify the effect

of MRPL12 knockdown on mitochondrial translation by pulse labeling of de novo

synthesized mitochondrially-encoded thirteen proteins in the presence of cytoplasmic

translation inhibitor, emetine, as described in Materials and Methods (Fig. 2.20). In

addition, MRPL10 and MRPL12 were overexpressed (L10 and L12, respectively) to

compare the effect of changing MRPL12 levels on mitochondrial ribosomal activity

(Fig. 2.20). The knockdown of MRPL12 reduced the total pulse labeling of thirteen

proteins compared to control cells (Con), while MRPL10 and MRPL12 overexpression

resulted in an elevated level of [35S]-methionine incorporation into mitochondrially-

encoded proteins. This finding suggests the involvement of MRPL12 in mitochondrial

translation activity. Immunoblotting analysis of these samples revealed the reduction in

MRPL12 expression level; especially the ribosome bound shorter form, in knockdown

cells resulting in lower ribosomal activity (Fig. 2.21). It was recently reported that the

free and longer form of MRPL12, which is ribosome unbound version, is involved in the

regulation of mitochondrial transcription by directly interacting with mitochondrial RNA

polymerase (POLRMT) (53). It is possible that the knock down of MRPL12 results in

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decreased mitochondrial translation activity in addition to decreased POLRMT activity.

HSP60 was used to ensure equal loading of mitochondrial proteins and MRPS29 and

MRPL47 for mitochondrial ribosomal proteins. His-tag immunoblotting was performed

to detect overexpressed MRPL10 and MRPL12. Immunoblotting assay with MRPL12

antibody did not display increase in MRPL12 possibly due to limited recognition by the

antibody. In addition, the steady-state levels of OXPHOS subunits, Complex IV, the

mitochondrial-encoded subunit, did not reveal much difference between the lysates

prepared from these cell lines (Fig. 2.21). We quantitated the immunoblotting signals of

Complex IV and Complex III subunits in MRPL12 knock-down and control cells. Even

they showed fewer signals in control sample, relative signal intensities were similar when

compared to the signal intensities of Complex IV and Complex III (Fig. 2.21).

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Figure 2.20: Effect of MRPL12 knockdown and MRPL10 and MRPL12

overexpressions on mitochondrial protein synthesis.

HEK293T cells transfected with MRPL12 shRNA (shL12) construct and

MRPL10 (L10) and MRPL12 (L12) overexpression constructs were exposed to [35S]-

methionine in the presence of a cytosolic translation inhibitor, emetine, for 2 h at 37oC.

Approximately, 40 μg of cell lysate from each sample were separated on 14 % SDS-

polyacrylamide gel. Electrophoretic pattern of the de novo synthesized translational

products was presented. ND1, -2, -3, -4, -4L, -5, and -6 are subunits of Complex I; Cytb

is subunit of Complex III; COI, -II, and -III are subunits of Complex IV; ATP6 and ATP8

are subunits of the Complex V. Coomassie blue staining of the same gel was performed

to ensure equal protein loading in the gel. The combined intensities from each lane were

used as the overall quantitation of the mitochondrial protein synthesis compared to

control. One way ANOVA test was used to compare the significance of values from three

experiments. Values of ٭P < 0.05 were considered statistically significant.

*

* *

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Figure 2.21: Effect of MRPL12 knockdown and MRPL10 and MRPL12

overexpression on ribosomal L7/L12 stalk composition and OXPHOS subunits.

Cell lysates from HEK293T cells containing MRPL12 knockdown (shL12) and

MRPL10 (L10) and MRPL12 (L12) overexpression were analyzed with immunoblotting

assay using His-tag and MRPL2 antibodies. Reduction of MRPL12 and overexpression

of MRPL10 and MRPL12, were demonstrated. MRPS29 and MRPL47 antibodies were

used as controls for other ribosomal proteins and HSP60 antibody for mitochondrial

proteins. OXPHOS antibody probing shows the steady-state levels of complex subunits,

where relative quantitation of Complex IV to Complex III is given as a graph.

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2.4.7 The Role of Acetylated Lysine Residues of MRPL10 on Mitochondrial

Translation and Cell Growth In Vivo.

MRPL10 is one of the essential components of the mitochondrial L7/L12 stalk

since it possibly provides the binding sites for multiple copies of MRPL12 via protein-

protein interactions. In E. coli, the L7/L12-stalk is a pentameric complex in which there

are four copies of L12 bound to L10 per ribosome (40). On the other hand, in T. maritima

the L7/L12 stalk is formed by a copy of L10 and six copies of L12 as three dimers per

ribosome (37). Bacterial L10 has a globular N-terminal and a flexible C-terminal α-helix

domain involved in binding to L11 and binding of L12 dimers, respectively. In the

mammalian mitochondrial ribosome, however, the composition of the L7/L12 stalk and

the nature of the MRPL10 and MRPL12 interaction(s), in particular the role of MRPL10

acetylation in the composition of the stalk and interaction of proteins forming the stalk is

not known. Previously, expression of two different forms of MRPL12 was reported in

human mitochondria (48). We have also confirmed the presence of two different forms of

MRPL12 in bovine mitochondria (Fig. 2.13). Interestingly, we detected changes in the

expression of one of these two different forms of MRPL12 in the ribosomes of SIRT3

knock-out mice and HIB1B cells overexpressing SIRT3, as a function of MRPL10

acetylation/deacetylation (Fig. 2.11 and Fig. 2.15). Based on these results, the acetylated

Lys residues of MRPL10 might be involved in modulating the number of MRPL12

dimers comprising the stalk and be important for regulation of mammalian mitochondrial

protein synthesis and cell growth.

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To determine the modulation of MRPL12 binding to MRPL10 by reversible

acetylation, we constructed expression vectors (control without insert) including

MRPL10 wild-type (WT) and its triple lysine residues at positions Lys124, Lys169, and

Lys196 which were mutated to Gln to mimic hyperacetylation and Ala or Arg to mimic

hypoacetylation, respectively, as described in Materials and Methods. The triple Gln

mutation of MRPL10 was expected to demonstrate the effect of acetylation since it would

not introduce the charge found on Lys residues and this would possibly provide a

stronger MRPL10-MRPL12 interaction. Conversely, Arg mutations of MRPL10 would

provide the charge on its side chain and this was expected to exert the effect of

nonacetylated Lys residues on MRPL12 binding. Introducing additional positive charges

to MRPL10 may interfere with MRPL12 binding to the ribosome. Moreover, Ala does

not have a long side chain as Lys does so the replacement of Lys with Ala in MRPL10

would introduce a structural and/or conformational change to the surface where MRPL12

binds. The triple Ala mutant of MRPL10 might also display reduced binding of MRPL12

to the mitochondrial ribosome due to the disruption of protein-protein interactions.

HEK293T cells transfected with corresponding constructs were grown for four

more days to overexpress MRPL10 constructs. Then, these cells were treated with

emetine, cytoplasmic translation inhibitor, followed by the incubation with [35S]-

methionine to specifically label mitochondrial protein synthesis products (Fig. 2.22).

Overexpression of MRPL10 (WT) and Gln mutant of MRPL10 resulted in elevated

mitochondrial translation, implying a dominant negative effect of these forms of

MRPL10 in cells (Fig. 2.22). However, Arg mutant of MRPL10 displayed a reduced

level of ribosomal activity compared to control cells. Overexpression of MRPL10

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proteins were confirmed by immunoblotting analysis using a His-tag antibody. The same

samples were also immunoblotted using MRPL12, MRPS29, and MRPL47 antibodies to

reveal the changes in ribosomal proteins and HSP60 antibody to verify mitochondrial

protein content in these cells (Fig. 2.22). Immunoblotting assays with the cell lysates did

not confirm that these changes were caused by alterations in the MRPL12 amounts since

they did not correlate with the ribosomal activity. However, ribosome bound version of

MRPL12 was not detected here. Immunoblotting with mitochondrial ribosomes by using

MRPL12 antibody might reveal the effect of MRPL10 constructs on MRPL12 binding to

ribosomes. Cell proliferation assay using WST1 reagent was performed to compare the

growth rates of cells overexpressing MRPL10 constructs. The calculated proliferation

rates of cells were graphed relative to control cells transfected with empty expression

vector (Fig. 2.23). The cells overexpressing wild type MRPL10 and its Gln mutant

displayed a significant increase in the growth rate. Conversely, proliferation rate of cells

overexpressing Ala mutant of MRPL10 was reduced, but not in cells overexpressing Arg

mutant. Overexpression of MRPL10 constructs might have some effect on cell growth;

however, their incorporation level into mitochondrial ribosomes affecting translation

would be reduced if they impair the mitochondrial protein synthesis. Here, incorporation

of MRPL10 mutants into mitochondrial ribosomes would be enforced to exert the effect

of Lys mutations on mitochondrial protein synthesis and therefore cell growth. This

effect could possibly be enhanced in cells where the endogenous MRPL10 was targeted

to be depleted or knockdown. Overall, these findings suggest the possible importance of

acetylated lysine residues in MRPL10 and their roles in the formation and/or composition

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of the mitochondrial L7/L12 and protein synthesis activity which affects overall cell

growth.

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Figure 2.22: The effect of MRPL10 LysAla, Gln, and Arg mutants on mitochondrial protein synthesis.

HEK293T cells were transfected with pcDNA 3.1+ (NC or CON), pcDNA-MRPL10 wild type (WT) and its lysine mutants (A: Ala, Q: Gln, and R: Arg) to overexpress MRPL10. [35S]-methionine labeling of mitochondrial-encoded proteins was performed in the presence of a cytosolic translation inhibitor, emetine to comparemitochondria translation activity between samples. Approximately, 40 μg of lysates were separated on SDS-polyacrylamide gels and electrophoretic pattern of the de novosynthesized translational products was presented. ND1, -2, -3, -4, -4L, -5, and -6 are subunits of Complex I; Cytb is subunit of Complex III; COI, -II, and -III are subunits of Complex IV; ATP6 and ATP8 are subunits of the Complex V. Coomassie blue stainingof the same gel was performed to ensure equal protein loading in the gel. The combinedintensities from each lane were used as the overall quantitation of the mitochondrial protein synthesis relative to control. One way ANOVA test was used to compare thesignificance of measurements from three experiments. Asterisks denote p<0.05.As control, His-tag antibody for overexpression of MRPL10 constructs, MRPS29 and MRPL47 antibodies for ribosomal proteins and HSP60 antibody for mitochondrialproteins were examined. MRPL12 immunoblotting analysis indicates the change in itsexpression level in cell lysates.

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Figure 2.23: The effect of MRPL10 LysAla, Gln, and Arg mutants on cell growth.

The cell proliferation assay using reagent WST1 was employed to quantitate cell

growth of HEK293T cells overexpressing MRPL10 wild type (WT) and its lysine

mutants (Ala: A, Q: Gln, and R: Arg). After transfection of cells with control (CON) or

corresponding expression vectors, 5 x 103 cells were grown 3 more days. Then, WST1

assay reagent was added and cells were incubated at 37oC following the method provided

by manufacturer. Growth rate of cells were graphed as relative ratio to control cells from

three separate plates. One way ANOVA test was used to compare the significance of

values and the value of *P < 0.05 was considered statistically significant.

*

*

*

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2.4.8 The Role of MRPL10 Reversible Acetylation in the Composition of Ribosomal

L7/L12 Stalk and Mitochondrial Translation In Vitro.

In order to elucidate the mechanism involved in the regulation of mitochondrial

L7/L12 stalk composition by reversible acetylation, we examined the MRPL10 and

MRPL12 interaction(s) in vitro. In our previous studies, MRPL10 and MRPL12 complex

formation and co-purification of the complex was achieved from bacteria co-expressing

MRPL10 and MRPL12 genes cloned into a bacterial expression vector, pET-Duet-1 (63).

We identified the contaminating proteins, such as groEL, HSP70, EF-P, and ribosomal

protein L4, co-purified with our MRPL10-MRPL12 complex by nickel-nitrilotriacetic

acid (Ni-NTA) affinity chromatography (63). Similarly, we have utilized this approach to

test the lysine mutant constructs of MRPL10 to examine how these mutations affect the

stoichiometry of MRPL10-MRPL12. Point mutations of MRPL10 were created by site-

directed mutagenesis at Lys124, Lys162, and Lys196 by generating LysAla in addition

to LysArg and LysGln to mimic hypoacetylation and hyperacetylation, respectively

as described in Materials and Methods. petDUET® overexpression system (Invitrogen

Inc.) having one of these MRPL10 constructs including His-tag at its C-terminal end and

MRPL12 was co-expressed in E. coli. By employing Ni-NTA affinity chromatography

followed by strong-cation exchange (SCX) chromatography, we attempted to purify His-

tagged MRPL10 under native conditions to maintain its interaction with MRPL12

(Fig. 2.24). For SCX-chromatography, we optimized the elution conditions with

application of step-gradient of salt; 1inear 50 mM to 150 mM KCl to remove non-

retained proteins for the first 30 min, then linear 400 mM to 500 mM KCl in 10 min to

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elute the stalk proteins which was followed by a linear increase of KCl concentration to 1

M in another 5 min to elute the retained proteins. Fractions were monitored by UV

detection at 260 nm and then analyzed by SDS-polyacrylamide gel electrophoresis

(Fig. 2.24). The chromatogram shows two separate peaks for wild type MRPL10 –

MRPL12 construct in agreement with our previous purification analysis (blue line in

Fig. 2.24), confirming the additional stable complex formation with chaperones (63).

However, the chromatography profiles for MRPL10 lysine mutants did not overlap with

that of the wild type sample possibly because of different amounts of MRPL12 bound to

MRPL10 lysine mutants, which confirms the importance of these lysine residues in

maintaining the MRPL10-MRPL12 interactions (Fig. 2.24). For this reason, we

performed immunoblotting assays using the His-tag antibody to ensure equal amounts of

MRPL10 were provided in in vitro translation assays using hybrid ribosomes (Fig. 2.25).

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Figure 2.24: Strong-cation exchange (SCX) chromatography purification of

mitochondrial MRPL10-MRPL12 stalk complex.

Ni-NTA chromatography purified wild type and mutant MRPL10-MRPL12

complexes (Ini) were applied on SCX column to remove contaminants (FT: flow-

through, E1: first fraction) where fractions were monitored with a UV detector at 260 nm

to trace the proteins during application of step gradient of salt, KCl (dashed line). In the

elution profile, stalk complexes containing MRPL10 wild type (WT) were labeled with

blue solid line and lysine mutants, Ala (A), Gln (Q), Arg (R) were marked in black, red,

and green, respectively. Fractions were analyzed on SDS-polyacrylamide gels to locate

MRPL10-MRPL12 complexes.

*: Fractions were combined and dialyzed to be used in activity assays.

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The MRPL10-MRPL12 complexes purified from SCX-column were used in the

reconstitution of hybrid ribosomes in order to assess their roles in poly(U)-directed

poly(Phe) synthesis in the presence of [14C]-tRNAPhe and the mammalian mitochondrial

elongation factors EF-Tumt and EF-G1mt (77). SCX-column fractions containing

MRPL10-MRPL12 complexes were combined and analyzed on the same SDS-

polyacrylamide gel to examine the degree of MRPL12 binding to MRPL10 lysine

mutants. Wild type MRPL10 displays the highest amount of MRPL12 retained during

purification followed by Q mutant mimicking hyperacetylation status and by the A

mutant. The amount of MRPL12 bound to MRPL10 was reduced in the R mutant,

hypoacetylated state of the protein (Fig. 2.25). This finding suggests that

acetylation/deacetylation of these lysine residues might be critical in terms of modulating

the L7/L12 stalk composition to regulate mitochondrial translational activity. In order to

examine this possibility, bacterial ribosomes lacking L10 and L12 (stripped ribosomes,

70S-str) were prepared by sedimenting ribosomes in the presence of 1 M NH4Cl and 50

% ethanol at 37oC to selectively remove these proteins (39,63). After incubation of the

purified mitochondrial MRPL10-MRPL12 complexes with the stripped ribosomes, we

recovered hybrid bacterial ribosome containing the mitochondrial L7/L12 stalk by

ultracentrifugation through a 30 % sucrose cushion as described in Materials and

Methods (63). In order to determine the activity of these hybrid ribosomes, polyU-

directed in vitro translation assays were performed as described previously (37,76,77). As

shown in Fig. 2.25, we used the activity of the reconstituted ribosome with recombinant

bacterial L10 and L12 to compare the activities of ribosomes reconstituted with the

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corresponding MRPL10-MRPL12 complexes. While bacterial stalk proteins recovered

the translational activity of ribosomes, mitochondrial stalk complexes did not restore the

poly-(Phe) synthesis activity which was similar to the inactive stripped ribosome level

(Fig. 2.25). It is possible that impurities and improperly folded MRPL10-MRPL12

complex were probably the factors affecting the activity of reconstituted ribosomes.

Purification of MRPL10 is challenging because of its hydrophobicity. Providing

MRPL12 in the same expression system improved its solubility and recovery at the

affinity chromatography purification step. However, proteins co-purified with MRPL10-

MRPL12 complex, such as groEL, HSP70, and EF-P, were probably reducing the

exposure of His-tag on MRPL10 to Ni-NTA resin, which decreased the amount of

purified MRPL10. These chaperon proteins might be associated with MRPL10-MRPL12

complex due to improper folding of overexpressed protein (s), which might also affect

their activity in in vitro translation assays. Removal of the co-purified proteins by SCX-

chromatography possible exposed hydrophobic regions on MRPL10 and its mutants to

induce protein aggregation. In addition, dialysis of protein preparations to remove

imidazole and salt after each Ni-NTA- and SCX-chromatography steps, respectively,

caused precipitation of MRPL10-MRPL12 complex.

These experiments will be repeated with highly purified stalk complex proteins to

demonstrate the effect of the acetylated lysine residues on MRPL12 binding and

ribosomal activity. To improve purification of MRPL10-MRPL12 by using Ni-NTA

affinity chromatography, bacterial cell lysate will be prepared in the presence of

additional ATP and magnesium salts since the association of contaminating proteins

might be reduced in this way (84). Another approach was the immunoprecipitation of

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MRPL10 by using His-tag antibody. However, immunoprecipitation of the complex

using the bacterial lysate containing overexpressed MRPL10-MRPL12 complex or the

Ni-NTA chromatography purified complex, did not enrich MRPL10-MRPL12 complex

due to the lack of binding specificity of the MRPL10-MRPL12 complex to the His-tag

antibody (data not shown). In addition, extended time of incubations with antibody and

solutions might perturb the activity of the MRPL10.

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Figure 2.25: Poly(U)-directed in vitro translation assays using hybrid ribosomes.

Stripped ribosomes (35 pmol of 70S-str) were reconstituted with bacterial L10 (70

pmol of bacL10) and L12 (280 pmol of bacL12) or corresponding mitochondrial L10 (70

pmol of MRPL10) and L12 (280 pmol of MRPL12) stalk proteins purified from SCX

column at 37oC for 15 min. The poly(U)-directed in vitro translation assay was carried

out at 37oC for 30 min and stopped by cold 5 % trichloroacetic acid addition on ice for 5

min. After incubation at 90oC for 10 min, samples were filtered through the nitrocellulose

membrane and the amount of [14C]-labeled-poly(Phe) was quantitated using liquid

scintillation counter. Graph was expressed as the relative percent of synthesized

poly(Phe) compared to the control bacterial ribosome (70S).

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2.5 Conclusions and Future Directions

Mitochondria are the essential organelle for eukaryotic cells since they generate

most of the ATP through oxidative phosphorylation (OXPHOS) for cellular metabolism.

Slower metabolic rates with reduction in OXPHOS and ATP production is correlated

with lower caloric intake resulting in an extended lifespan (186). SIRT3, mitochondrial

NAD+-dependent deacetylase, is implicated in those effects by deacetylating

mitochondrial proteins (108,109,113,117,143,172). Similarly, SIRT3 may be involved in

coordinating the activity of mitochondrial protein synthesis machinery through MRPL10

deacetylation in response to the changes in the [NADH]/[NAD+] ratio in the mammalian

mitochondria. Our findings suggest that mitochondrial protein synthesis is regulated by

NAD+-dependent activity of SIRT3 on MRPL10 acetylation level in bovine ribosomes.

This hypothesis is supported by employing SIRT3 knock-out mice. Liver mitochondria

isolated from SIRT3 knock-out mice liver displays hyperacetylated MRPL10 and/or

ribosomes which results in more protein synthesis, leading to an increase in the

expression of mitochondrially-encoded components of the oxidative phosphorylation.

Conversely, HIB1B cells overexpressing SIRT3 demonstrates deacetylation of MRPL10

and/or ribosomes which may impair the synthesis of mitochondrially-encoded proteins,

and therefore, reduces the rate of oxidative phosphorylation by lowering the expression

of essential subunits of respiratory chain complexes. The analysis of mitochondrial

ribosomes from SIRT3 knock-out mice and HIB1B cells overexpressing SIRT3 displays

that changes in MRPL12 binding to the ribosomes is an important factor involved in the

acetylation mediated regulation of protein synthesis in ribosomes.

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Multiple copies of MRPL12 dimers interact with MRPL10 to form the

mitochondrial L7/L12 stalk. This stalk region is evolutionary conserved and essential for

the recruitment of elongation factors EF-Tu and EF-G during protein synthesis (187,188).

In E. coli, the L7/L12-stalk is composed of four copies of L12 bound to L10 per ribosome

(156). On the other hand, in T. maritima the L7/L12 stalk contains a copy of L10 and to

six copies of L12 as three dimers per ribosome (61). Bacterial L10 has a globular N-

terminal and a flexible C-terminal α-helix domain involved in binding to L11 and binding

of L12 dimers, respectively. In the mammalian mitochondrial ribosome; however, the

composition of the L7/L12 stalk and the nature of the MRPL10 and MRPL12

interaction(s), in particular the role of MRPL10 acetylation on the composition of the

stalk and interaction of proteins forming the stalk is not known. Acetylation of L12 at the

N-terminal end was shown to increase the interaction and stability between the stalk

proteins in E. coli due to increased hydrophobicity by acetylation (135). Intriguingly,

acetylation of the bacterial ribosomal L10 is also reported and the acetylated lysine

residues in both bacterial and mitochondrial L10 are mapped to the N-terminal globular

domain of L10 towards the elongation factor binding cavity under the L7/L12 stalk

region. The bacterial and mitochondrial L12 homologs have a highly conserved lysine

rich CTDs, therefore, packing this lysine-rich domain into a lysine-rich L10 surface

would not be possible due to charge repulsion. In this scenario, having a neutralized L10

surface by acetylation would stabilize the lysine-rich CTD of L12 between the N- and C-

terminal domains of L10 and L11, respectively (Fig. 5.1). Based on these results and our

findings in SIRT3 knock-out mice and HIB1B cells overexpressing SIRT3, it is feasible

to postulate that the SIRT3-dependent reversible acetylation of MRPL10 regulates

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mitochondrial protein synthesis by modulating protein-protein, specifically MRPL10-

MRPL12, interactions in the L7/L12 stalk region of the ribosome.

In order to confirm the importance of mitochondrial L7/L12 stalk proteins,

MRPL10 and MRPL12, in mitochondrial translation, shRNA mediated knockdown of

MRPL12 and overexpression of MRPL10 and MRPL12 was performed in HEK293T

cells. Reduction of the de novo synthesized mitochondrial-encoded thirteen proteins in

pulse labeling assay confirms the importance of MRPL12 in ribosomal activity.

Conversely, the cells overexpressing MRPL10 and MRPL12 displays elevated labeling of

synthesized products, which highlights the implication of these L7/L12 stalk proteins in

protein synthesis in mitochondria. To determine the modulation of MRPL12 binding to

MRPL10 by reversible acetylation and its effect on mitochondrial translation, Gln and

Arg mutants at selected three acetylated residues, Lys124, Lys169, and Lys196,

mimicking hyperacetylation and hypoacetylation, respectively, were employed in in vivo

studies. The triple Gln mutation of MRPL10 is expected to demonstrate the effect of

acetylation since it would not introduce the charge found on Lys residues and this would

possibly provide a stronger MRPL10-MRPL12 interaction. Conversely, Arg mutant

provides the charge on its side chain and is anticipated to reduce MRPL12 binding to

ribosome and thus mitochondrial translation similar to the effect of nonacetylated Lys

residues. The triple Ala mutant of MRPL10 might also display reduced binding of

MRPL12 to the mitochondrial ribosome due to the disruption of protein-protein

interactions since Ala does not have a long side chain as Lys does. However, the pulse

labeling and cell growth rate assays with the cells overexpressing MRPL10 mutants did

not reveal significant effect on mitochondrial protein synthesis and cell proliferation

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compared to the cells containing wild type MRPL10. One possibility of not seeing a

significant change between the cells might be the different import rate into mitochondria

and also different incorporation level of MRPL10 mutants into mitochondrial ribosomes.

It might be challenging to get the same level of incorporation of MRPL10 mutants into

the mitochondrial ribosomes since they would incorporate endogenous MRPL10 to

overcome the defect in protein synthesis caused by the MRPL10 constructs. However,

depletion of endogenous MRPL10 in these cells might enforce mitochondrial ribosomes

to harbor MRPL10 constructs and improve the effect of Lys mutations on MRPL12

binding to ribosomes, mitochondrial protein synthesis, and therefore cell growth in vivo.

Furthermore, there may be additional acetylated lysine residues on MRPL10 and their

mutants might also be required to demonstrate the significant effect of lysine residues

involved in the modulation of MRPL10 interaction with MRPL12 on the stalk. On the

other hand, the in vitro approach employing hybrid ribosomes to assess the effect of

acetylated lysine residues on MRPL10 in MRPL12 binding to the ribosomes and protein

synthesis was not conclusive. The MRPL10-MRPL12 complexes used in the

reconstitution of hybrid ribosomes contain impurities and possibly improperly folded

MRPL10 constructs, which might affect the activity of hybrid ribosomes in poly(U)-

directed poly(Phe) synthesis assays. The experiments to improve the purification of

MRPL10-MRPL12 complex and activity assays are under progress to determine the

effect of acetylated Lys residues and their mutants in MRPL12 binding and protein

synthesis. Instead of the reconstitution of hybrid ribosomes with purified MRPL10-

MRPL12 complexes, bacterial ribosomes which incorporate these constructs during their

overexpression might also be used in poly(U)-directed in vitro translation assays. As

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described in our in vivo approach, different incorporation levels of the MRPL10

constructs into bacterial ribosomes as opposed to wild type MRPL10 and RPL10 might

result in ambiguous results with reduced effect of the MRPL10 mutants in protein

synthesis. Future studies may involve the deletion of bacterial RPL10 (and possibly

RPL12) with recombination techniques and the expression of MRPL10-MRPL12

complexes (189). In this system, the stability and activity of MRPL10 constructs would

be improved since they are incorporated into ribosomes in the bacterial cells.

Overall, our preliminary in vivo and in vitro studies demonstrated that acetylated

Lys residues on MRPL10 might possibly modulate MRPL12 binding to ribosomes

resulting in the regulation of mitochondrial translation and cell growth. These methods

have laid out the initial framework to uncover the significant role of the MRPL10

acetylation in MRPL12 binding to the ribosome. Reversible acetylation of MRPL10 may

be one of the key regulatory mechanisms that modulates MRPL12 binding to the

ribosomal stalk and the synthesis of mitochondrially-encoded essential proteins of

respiratory chain complexes. This regulation is possibly mediated by SIRT3, ribosome

associated NAD+-dependent deacetylase, which monitors the changes in mitochondrial

metabolism through the [NADH]/[NAD+] ratio. Increased levels of NAD+ by caloric

restriction will stimulate SIRT3 and it will perform its protective functions in

mitochondria by deacetylating its target proteins. Most of the metabolic enzymes will be

activated to restore the NADH levels for ATP production by OXPHOS. On the other

hand, deacetylation of MRPL10 and decreased MRPL12 on the ribosomes reduce the

mitochondrial translation rate and additional synthesis of respiratory chain components

due to low NADH levels. Conversely, when the NADH level is high, acetylation of

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MRPL10 triggers the binding of MRPL12 to enhance protein synthesis rate to support the

OXPHOS and ATP production rates.

2.6 Acknowledgment

We would like to thank Dr. Qiang Tong for providing SIRT3 knockout mice liver

mitochondria and HIB1B cells overexpressing SIRT3.

2.7 References

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Chapter 3

Regulation of Succinate Dehydrogenase Activity by SIRT3

This chapter of dissertation was reproduced with permission from Huseyin

Cimen, Min-Joon Han, Yongjie Yang, Qiang Tong, Hasan Koc, and Emine C. Koc.

(2010) Regulation of Succinate Dehydrogenase Activity by SIRT3 in Mammalian

Mitochondria. Biochemistry; 49 (2):304–311. Copyright © 2010 by American Chemical

Society.

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3.1 Rationale

A member of the sirtuin family of NAD+-dependent deacetylases, SIRT3 was

identified as one of the major mitochondrial deacetylase located in mammalian

mitochondria responsible for deacetylation of several metabolic enzymes and

components of oxidative phosphorylation. Regulation of protein deacetylation by SIRT3

is important for mitochondrial metabolism, cell survival and longevity. In this study, we

identified one of the Complex II subunits, succinate dehydrogenase flavoprotein (SdhA)

subunit, as a novel SIRT3 substrate in SIRT3 knock-out mice. Several acetylated Lys

residues were mapped by tandem mass spectrometry and we determined the role of

acetylation on Complex II activity in SIRT3 knock-out mice. In agreement with SIRT3

dependent activation of Complex I, we observed that deacetylation of SdhA subunit

increased the Complex II activity in wild type mice. In addition, we treated K562 cell

lines with nicotinamide and kaempferol to inhibit deacetylase activity of SIRT3 and

stimulate SIRT3 expression, respectively. Stimulation of SIRT3 expression decreased the

acetylation of SdhA subunit and increased Complex II activity in kaempferol treated cells

compared to control and nicotinamide treated cells. Evaluation of acetylated residues in

the SdhA crystal structure from porcine and chicken suggest that acetylation of

hydrophilic surface of SdhA may control the substrate entry to the active site of the

protein and regulate the enzyme activity. Our findings constitute the first evidence for the

regulation of Complex II activity by the reversible acetylation of the SdhA subunit as a

novel substrate of the NAD+- dependent deacetylase, SIRT3.

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3.2 Introduction

Reversible acetylation of mitochondrial proteins is critical for regulation of many

biological processes, including oxidative phosphorylation and the Krebs cycle (1-7).

Flavoprotein of the succinate dehydrogenase complex (Complex II SdhA subunit) was

identified as one of the acetylated proteins of mice liver mitochondria in two independent

high throughput mapping of acetylated proteins by tandem mass spectrometry (2,7).

Complex II or succinate dehydrogenase (SDH) is found as an inner membrane-bound

enzyme complex and it is the only enzyme that participates in both the Krebs cycle and

oxidative phosphorylation in mitochondria. It has four different protein subunits;

hydrophilic subunits SdhA and SdhB facing the matrix side of the inner membrane and

hydrophobic subunits, SdhC and SdhD, tethering the complex in the phospholipid

membrane. SdhA is a 70 kDa large flavoprotein subunit containing covalently bound

FAD and substrate binding site for the entry point of electrons to Complex II. SDH plays

such an important role in the mitochondria, that severe deficiency of this enzyme is

incompatible with life. However, point or milder mutations in the C-terminal domain of

SdhA lead to Leigh syndrome and various neurodegenerative disorders (8). Mutations of

the other SDH subunits containing Fe-S cofactors have been associated with generation

of reactive oxygen species causing tumor formation (9).

Post-translational modifications of SdhA by phosphorylation at tyrosine residues

and acetylation at lysine residues were previously reported (2,7,10). Interestingly, six

acetylated lysine residues in SdhA were mapped in the LC-MS/MS analysis of well-fed

rat mitochondria in two independent studies (2,7). However, neither enzymes responsible

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for reversible acetylation and phosphorylation nor their regulatory roles of these post-

translational modifications on SdhA or Complex II activity are known. Several members

of the class III histone deacetylases (sirtuins) SIRT3, SIRT4, and SIRT5 have been found

to reside in mitochondria (6,11,12). Sirtuins use NAD+ as a cosubstrate, and both SIRT3

and SIRT4 are required to maintain cell survival after genotoxic stress in a NAD+-

dependent manner, and genetic variations in the human SIRT3 gene have been linked to

longevity (13,14). We have previously shown that SIRT3 expression in adipose tissue is

increased by caloric restriction and cold exposure (2,15). Mitochondrial acetyl-CoA

synthetase 2 and glutamate dehydrogenase (GDH) are the two key metabolic enzymes

regulated through deacetylation by SIRT3 (1,6,16). Thus, SIRT3 was determined to be

the major deacetylase that modulates mitochondrial function in response to

[NADH]/[NAD+] ratio by regulating the activity of key metabolic enzymes (6,12,16,17).

In addition to metabolic enzymes, nuclear encoded subunits of the electron

transport chain complexes and ribosomes responsible for the synthesis of 13 essential

proteins of the oxidative phosphorylation were found to be regulated by reversible

acetylation (2). In our recent studies we demonstrated that the mitochondrial ribosomal

protein MRPL10 is acetylated and its deacetylation by the NAD+-dependent deacetylase

SIRT3 regulates mitochondrial protein synthesis (18). Additionally, Complex I subunit

NDUFA9 is also determined to be a SIRT3 substrate and acetylation/deacetylation of this

protein is proposed to regulate and maintain basal ATP levels in mammalian

mitochondria (17). However, contribution of Complex II acetylation was overlooked on

oxidative phosphorylation and ATP production in the same study (17).

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Here, we confirmed that one of the subunits of Complex II, SdhA, is indeed a

highly acetylated protein and it is a novel SIRT3 substrate as shown in SIRT3 knock-out

mice using various proteomic techniques. We have also determined the SIRT3-dependent

activation of Complex II in wild-type mice and in cells over-expressing SIRT3. Our

results reported in this study suggest a more global role for SIRT3 in regulating oxidative

phosphorylation by reversible acetylation of the Complex II subunit SdhA, and therefore,

ATP production in mammalian mitochondria.

3.3 Materials and Methods

3.3.1 Isolation of Mitochondria from Mice Liver and Enrichment of Complex II

SIRT3 knock-out mice were obtained from the Texas Institute for Genomic

Medicine (Houston, TX, USA). Briefly, these mice were produced by generating

embryonic stem (ES) cells (Omnibank no. OST341297) bearing a retroviral promoter trap

that functionally inactivates one allele of the Sirt3 gene, as described previously (19).

Liver tissue obtained from SIRT3 knock-out (Sirt3-/-), wild-type (Sirt3+/+), and

heterozygote (Sirt3+/-) mice was resuspended in an isotonic mitochondrial buffer (MB)

(210 mM mannitol, 70 mM sucrose, 1 mM EDTA, 10 mM HEPES-KOH pH 7.5),

supplemented with protease inhibitors (1 mM phenylmethylsulfonyl fluoride, 50 µg/ml

leupeptin), and then homogenized in a Dounce homogenizer (Wheaton) on ice. The

suspension was centrifuged at 400 x g on a microcentrifuge (ThermoForma) at 4°C. This

126

procedure was repeated twice, and supernatants were centrifuged at 10,000 g at 4°C for

10 min to pellet mitochondria. After the mitochondrial pellets were lysed in a buffer

containing 0.26 M sucrose, 20 mM Tris-HCl, pH 7.6, 40 mM KCl, 20 mM MgCl2, 0.8

mM EDTA, 0.05 mM spermine, 0.05 mM spermidine, 6 mM β-mercaptoethanol, and 1.6

% Triton X-100, mitochondrial lysates were loaded on to 34 % sucrose cushions and

centrifuged at 100,000 g at 4°C for 16 h. The cushion layers enriched for acetylated

proteins were precipitated with acetone.

3.3.2 Two Dimensional-Gel and Immunoblotting Analysis

Acetone precipitated protein pellets were resuspended in Destreak rehydration

buffer (Amersham Biosciences Inc.) and loaded onto the IPG strips ( pI 3-10) (Bio-Rad

Laboratories, Inc.). IPG strips were rehydrated overnight and run on the Ettan IPGphor

(Amersham Biosciences Inc) according to the manufacturer’s protocols. After running the

first dimension the strips were equilibrated in 6 M urea, 0.375 M Tris-HCl pH 8.8, 2 %

SDS, 20 % glycerol, and 2 % (w/v) DTT for 10 min. The strips then were equilibrated in

the equilibration buffer containing 2.5 % (w/v) iodoacetamide and loaded onto the second

dimension SDS-polyacrylamide gel. The gels were either stained with Coomassie Blue or

transferred to a PVDF membrane to be probed with acetyl Lys antibody at a 1:3000

dilution or SIRT3 antibody at a 1:1000 dilution (Cell Signaling Technology Inc.), a

monoclonal SdhA (Complex II subunit 70 kDa Fp) antibody at a 1:5000 dilution

(MitoSciences Inc.) or β-Actin antibody at a 1:5000 dilution (Abcam Inc.). The

secondary antibody was ImmunoPure antibody goat anti-mouse IgG (Pierce

127

Biochemicals Inc.) at a 1:5000 dilution or goat anti-rabbit IgG at a 1:1000 dilution or

Affinipure rabbit anti-mouse IgG, Rabbit anti-goat IgG, or goat anti-rabbit IgG (Jackson

Immuno Research) each at 1:10,000 dilution, followed by development with the

SuperSignal West Pico Chemiluminescent Substrate (Pierce Biochemicals Inc.) according

to the protocol provided by the manufacturer.

3.3.3 Mass Spectrometric Identification and Mapping of Acetylation Sites

SDS-polyacrylamide gel protein bands or 2D-gel spots corresponding to

acetylated proteins were excised and digested in-gel with trypsin prior to liquid

chromatography tandem mass spectrometry (capLC-MS/MS) analysis. The capLC-

MS/MS analysis performed by an LTQ mass spectrometer equipped with a nano-

electrospray ionization source and Surveyor MS Pump Plus HPLC system and Surveyor

Micro AS autosampler (ThermoFisher Co.). The in-gel tryptic digests (3-5 µL) were

injected and loaded onto a peptide trap (MiChrom peptide CapTrap, C8 like resin, 0.3

mm x 1 mm, 5µm) over 3 min at 10 µL/min for on-line desalting and concentration. The

peptide trap was then placed in line with the analytical column, a PicoFrit column (0.075

mm x 150 mm) packed in-house with Supelco's Wide Bore C18 (5 µm, 300 Å) resin. The

column was eluted at 250 nL/min using a gradient that consisted of 0.1 % formic acid

(Solvent A) and 0.1 % formic acid in acetonitrile (Solvent B). The peptides were eluted

by ramping the solvent B to 40 % over 30 min. Tandem MS spectra were acquired for

ions above a predetermined intensity threshold using the automated data-dependent

acquisition. The spectra were processed and searched against the protein sequence

128

database Swiss-Prot using a locally maintained Mascot 2.2 (Matrix Science) and

Proteome Discoverer 1.0 (ThermoFisher) search engines to identify proteins and

modifications. Mass tolerance was 3 amu and 2 amu for precursor and product ions,

respectively. Up to 2 missed cleavages were allowed for digestion by trypsin and

methionine oxidation (+16 Da) and lysine acetylation (+42 Da) were considered as

variable modifications.

3.3.4 Cell Culture

Approximately, 7 x 107 K562 (human chronic myelogenous leukemia cell line)

cells was grown in RPMI 1640 medium (Mediatech Inc.) supplemented with 10 % (v/v)

bovine calf serum (Hyclone) and 100 IU/ml penicillin and 100 μg/ml streptomycin, at

37oC and 5 % CO2 in a humidified atmosphere. Cells were treated with nicotinamide

(Calbiochem) or kaempferol (Sigma, St. Louis, MO) for 16 or 48 h at 10 mM or 50 µM

final concentrations, respectively. For immunoblotting, cell pellets were lysed in a buffer

containing 50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 0.5 %

NP-40, 0.1 % SDS, supplemented with protease inhibitor cocktail (Sigma-Aldrich).

After incubation on ice for 10 min, soluble protein fraction was collected by

centrifugation at 14,000 g at 4°C for 15 min.

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3.3.5 Complex II Enzymatic Activity Assay

Mitochondria and K562 cell pellets prepared as indicated above were lysed in a

buffer containing 300 mM mannitol, 20 mM sodium phosphate, pH 7.2, 10 mM KCl, 5

mM MgCl2, and 2 mg/ml dodecyl-β-D-maltoside. Preincubation of varying amounts of

mitochondrial and K562 cell lysates in a buffer containing 300 mM mannitol, 20 mM

sodium phosphate, pH 7.2, 10 mM KCl, 5 mM MgCl2, 50 mM sodium succinate, 40 mM

sodium azide, prior to the addition of 50 µM 2,6-dichloroindophenolate in order to fully

activate succinate dehydrogenase. Complex II enzymatic activity was recorded by

monitoring the reduction of 2,6-dichloroindophenolate at 600 nm. The rate is calculated

by dividing the absorbance difference between two linear points by the time point

difference [Rate = (absorbance 1 – absorbance 2) / (time 2 – time 1) (20).

3.3.6 Statistical Analysis

Results are expressed as means ± SD of at least three separate experiments.

Statistical difference between test groups was analyzed by one-way ANOVA test and

significance was defined at P<0.05.

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3.4 Results and Discussion

3.4.1 Succinate Dehydrogenase is Acetylated and SIRT3 is Responsible for its

Deacetylation

We have recently identified acetylated and phosphorylated protein(s) of

mitochondrial ribosomes using a combination of immunoblotting and capillary LC-

MS/MS analysis and identified NAD+-dependent SIRT3 as the deacetylase responsible

for deacetylation of MRPL10 (18,21,22). Using a similar strategy, we identified

acetylated proteins specifically deacetylated by SIRT3 in wild type and SIRT3 knock-out

mice liver mitochondria to determine SIRT3 substrates. For this purpose, mitochondria

were isolated from SIRT3 knock-out (Sirt3-/-), wild-type (Sirt3+/+), and heterozygote

(Sirt3+/-) mouse liver mitochondria. Acetylated proteins in mitochondrial lysates were

detected by immunoblotting performed with acetyl-Lys antibody, which revealed two

major protein bands at around ~70 kDa and ~55 kDa with an increased level of

acetylation in SIRT3 knock-out mice mitochondrial lysate as indicated by arrows

(Fig. 3.1A). Our findings suggest that these two proteins are potential substrates of

NAD+-dependent SIRT3 since they were highly acetylated in the absence of SIRT3

expression in knock-out mice (Fig. 3.1A). The lack of SIRT3 expression in the whole

liver or liver mitochondria from the SIRT3 knock-out mice was confirmed by

immunoblotting analysis (data not shown) (23).

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Figure 3.1A-B: Detection of SdhA as a novel SIRT3 substrate in SIRT3 knock-out

mice liver mitochondria.

A) Increased acetylation of mitochondrial proteins was detected in mitochondrial

lysates by immunoblotting with acetyl Lys (N-acetyl K) antibody. As a control for equal

loading, protein blot was developed with Hsp60 antibody. B) Approximately, 2 mg

of Sirt3−/− mice liver mitochondrial lysate were layered on 34 % sucrose cushion and

fractioned into five separate layers (the top is 1 and bottom is 5). Equal volumes of each

fraction were separated on SDS-polyacrylamide gel and acetylated proteins in each

fraction were detected by immunoblotting analysis. The total Sirt3−/− mice liver

mitochondrial lysate (ML) layered on the cushion was also analyzed to locate the

acetylated proteins in the fractions. Arrows show the location of SIRT3 substrates

glutamate dehydrogenase (GDH) and the flavoprotein subunit of succinate

dehydrogenase (SdhA).

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Figure 3.1C: Identification of SdhA acetylation in SIRT3 knock-out mice liver

mitochondria.

C) Approximately, 50 μl of fraction 3 from Sirt3+/+ or Sirt3−/− mice liver

mitochondria was separated on 2D-polyacrylamide gels and acetylated proteins were

detected with acetyl Lys (N-acetyl K) antibody. The acetylated 2D-gel spots

corresponding to the Coomassie Blue-stained gel spots were digested in-gel and

identified by mass spectrometry. The protein identification determined by mass

spectrometry was confirmed by immunoblotting using SdhA antibody.

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To identify the proteins in these bands and simplify the protein content for 2D-gel

separation, mitochondrial lysate obtained from SIRT3 knock-out mice was fractionated

on a 34 % sucrose cushion containing nonionic detergent Triton-X100. Immunoblotting

analysis of fractions showed that the two major acetylated proteins at ~70 and ~55 kDa

were in fractions 3 and 4, respectively, implying the presence of these proteins in large

protein complexes (Fig. 3.1B). For the identification of 70 kDa proteins, 2D-gel

electrophoresis was performed using fraction 3, and protein blot was probed with acetyl-

Lys antibody (Fig. 3.1C). Protein bands corresponding to acetylated protein detected in

2D-gel were excised, digested in-gel with trypsin, and analyzed by cap LC-MS/MS for

identification. The mass spectrometric analyses of the 2D-gel spots revealed the presence

of the flavoprotein subunit of succinate dehydrogenase (SdhA) in ~70 kDa protein spot

(Table 3.1). The other protein was identified as glutamate dehydrogenase (GDH) using

the same approach. Acetylation of glutamate dehydrogenase and role of SIRT3 on its

deacetylation was previously reported (16). Therefore, we focused our efforts in

determining the acetylation and deacetylation of SdhA in mitochondria obtained from

SIRT3 knock-out and wild type mice. In order to confirm the deacetylation of SdhA by

SIRT3, immunoblotting and Coomassie Blue-stained gels of protein lysates were

compared (Fig. 3.1C). Even though the SdhA signals obtained by its specific antibody in

both SIRT3 knock-out and wild type fractions were comparable, the acetylation signal

significantly increased in mitochondrial fraction from SIRT3 knock-out mice (Fig. 3.1C).

This observation supports the possibility that deacetylation of SdhA is due to the

expression of endogenous SIRT3 in wild type mice mitochondria while the absence of

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SIRT3 expression in knock-out mice causes hyper acetylation of the SdhA subunit

(Fig. 3.1C).

In addition to confirming the acetylation of the SdhA subunit by immunoblotting,

one of the acetylated tryptic peptides was also identified with a Mascot score of 74 in the

capLC-MS/MS analysis of the 2D-gel spots that was previously detected. The collision-

induced dissociation (CID) spectrum of the acetylated peptide, AFGGQSLacKFGK, is

given in Fig. 3.2A. In high throughput analysis of acetylated proteins from well-fed rat

liver mitochondria, several other acetylated lysines were identified (1-7). Alignment of

these acetylated peptides with the conserved regions in several other mammalian and

chicken mitochondrial forms and Eshcerichia coli SdhA shows that the acetylated lysines

are highly conserved in these proteins (Fig. 3.2B) (24). To demonstrate the location of

acetylated lysines in the SdhA subunit, we modeled Complex II structure using the

coordinates of the chicken mitochondrial Complex II (Fig. 3.2C). In this structure,

conserved acetylated lysine residues (K179, K485, K498, and K538) in the mouse

sequence are labeled as red surfaces in the SdhA subunit (Fig. 3.2B and C). All these

residues are located on the hydrophilic surface of the subunit supporting the reversible

acetylation of these residues by changes in [NADH]/[NAD+] ratios.

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Courtesy of Dr. H. Koc.

Table 3.1: Peptides detected from tryptic digests of protein band corresponding to

acetylation signal in SIRT3 knock-out mice mitochondria by capLC-MS/MS

analysis.

Peptide Sequence m/z Mascot Score ISQLYGDLK 1036.6 38 HTLSYVDIK 1075.6 41 WHFYDTVK 1095.6 48 NTVIATGGYGR 555.4 83 SMQNHAAVFR 589.2 66

AFGGQSLKacFGK 592.2 74 KHTLSYVDIK 602.8 57 AKNTVIATGGYGR 655.2 82 GEGGILINSQGER 666.2 84 VTLEYRPVIDK 667.3 63 TGHSLLHTLYGR 678.2 88 ANAGEESVMNLDK 698.2 98 VDEYDYSKPIQGQQK 899.8 82 VRVDEYDYSKPIQGQQK 1027.4 102 NTVIATGGYGRTYFSCTSAHTSTGDGTAMVTR 1102.1 41

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Figure 3.2A: The collision-induced dissociation (CID) spectrum of the acetylated

peptide.

A) The CID spectrum of acetylated peptide detected in capLC-MS/MS analysis of

2D-gel spot of SdhA from SIRT3 knock-out mice mitochondria. Courtesy of Dr. H. Koc.

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Figure 3.2B: Primary sequence alignment of acetylated peptides from mice SdhA

and its homologs from different species.

B) Acetylated peptides of mouse SdhA were aligned with corresponding residues

from human, bovine, pig, chicken, and E. coli. (*) denotes the acetylated Lys residues

detected in the capLC-MS/MS analysis. The alignment was created with CLUSTALW

program in Biology Workbench and displayed in BOXSHADE.

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Figure 3.2C: Crystal structure model of the chicken SdhA.

C) Crystal Structure of chicken SdhA (Protein Databank entry 1YQ3) was used to

represent the four subunits SdhA (green), SdhB (cyan), SdhC (pink), and SdhD (yellow).

The conserved Lys residues found to be acetylated in mouse SdhA (shown by asterisks in

B) are colored in red. Courtesy of Dr. E. Koc.

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3.4.2 Role of Hyperacetylation of SdhA in Complex II Activity

In order to determine the effect of acetylation on oxidation of succinate to

fumarate by Complex II activity, we measured the oxidation of 2,6-

dichloroindophenolate (DCIP) in mitochondrial suspensions obtained from SIRT3 knock-

out and wild type mice. First, mitochondrial suspensions obtained from these mice were

separated on a 12 % SDS-polyacrylamide gel and evaluated for the SdhA, Hsp60, and

acetylation levels by immunoblotting with specific antibodies. Although the same amount

of SdhA and Hsp60 were loaded in the gels, the degree of acetylation was much higher in

mitochondrial suspension from SIRT3 knock-out mice compared to wild type mice

(Fig. 3.3A). After confirming the presence of equal amounts of SdhA in these samples,

we performed the Complex II activity assays at several different amounts of

mitochondrial suspensions obtained from SIRT3 knock-out and wild type mice

(Fig. 3.3B). In these assays, the activity of complex II was followed by the transfer of

electrons from succinate to DCIP at 600 nm (Fig. 3.3B). As plotted in Fig. 3.3B, the rate

of reactions was measured as changes in the absorbance at 600 nm over time as a

function of the amount of mitochondrial suspension used in the assays. At 15 µg of

mitochondria suspension, the difference between the rate of Complex II activity from

SIRT3 knock-out mice and wild type mice was about ~30 % (Fig. 3.3B). To demonstrate

the linearity of the percent inhibition detection by the assay, different amounts of

mitochondrial lysate were used; however, the percent inhibition did not change

significantly above 15 µg of mitochondrial suspension. Here, reduction of DCIP was

directly related to the SdhA activity since electrons from succinate is first transferred to

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the enzyme bound cofactor, FAD, in the SdhA subunit. For this reason, decrease in

Complex II activity can be attributed to increased acetylation of SdhA in mitochondria

from SIRT3 knock-out mice (Fig. 3.3B).

3.4.3 Role of Increased Level of SIRT3 Expression in Deaceylation of SdhA and

Complex II Activity

The significant increase in acetylation of several proteins in SIRT3 knock-out

mice mitochondria (Fig. 3.1A and Fig. 3.3A) prompted us to determine the effect of

SIRT3 overexpression. Stimulation of sirtuins, class III histone deacetylases, by several

polyphenolic compounds such as resveratrol and kaempferol has been suggested recently

(25-27). Specifically, kaempferol treatment of the chronic myelogenous leukemia, K562,

cell line has been shown to increase the level of SIRT3 expression in these cell lines (27).

Moreover, nicotinamide is a general sirtuin inhibitor and has been shown to inhibit

SIRT3-dependent deacetylation of GDH and NDUFA9 (17,28). To demonstrate the effect

of SIRT3 expression on Complex II activity, we treated K562 cells with 50 µM

kaempferol or 10 mM nicotinamide for either 16 or 48 h and monitored the changes in

acetylation and expression of SIRT3 by immunoblotting analysis using whole cell lysates

(Fig. 3.4A). Nicotinamide treatment did not further increase the acetylation of proteins

possibly due to their hyperacetylated state caused by glucose rich media used to grow

these cells. Reprobing of the membranes was performed with SdhA and Hsp60 antibodies

to ensure equal amounts of protein loading in the SDS-polyacrylamide gels. Consistent

with the increased expression of SIRT3 in kaempferol-treated cells, the overall

141

acetylation level of proteins decreased compared to that in the control and nicotinamide-

treated cells (Fig. 3.4).

142

Figure 3.3: Regulation of succinate dehydrogenase (Complex II) activity by

deacetylation of SdhA.

Hyperacetylation of SdhA decreases Complex II activity in SIRT3 knock-out

mice. A. Equal amounts of lysates obtained from Sirt3+/+ and Sirt3−/− mice liver

mitochondria were separated on 12 % SDS-polyacrylamide gel and probed with acetyl

Lys (N-acetyl K), SdhA, and Hsp60 antibodies. B. Complex II activity was measured as

the rate of DCIP reduction, monitored at 600 nm using different amounts of

mitochondrial lysates from Sirt3+/+ and Sirt3−/− mice liver mitochondria. One way

ANOVA test was used to compare the significance of values from three separate

experiments. Asterisks denote p<0.05.

143

In addition to the detection of overall changes in acetylation of proteins in K562

cells, we fractionated the cell lysates treated with kaempferol and nicotinamide along

with untreated cells on 34 % sucrose cushion containing 1.6 % Triton-X100 to enrich for

SdhA protein. Similar to the pattern obtained in fractionation of mice liver mitochondria,

SdhA remained associated and sedimented with the rest of the Complex II subunits in

fractionation of kaempferol- and nicotinamide-treated cells as confirmed by

immunoblotting analyses (Fig. 3.4B). Especially in the nicotinamide treated and the

control cells, acetylated protein signal (shown by arrows) overlapped with the SdhA

signal in the reprobing of the membranes with the specific SdhA antibody. On the other

hand, acetylation of SdhA was significantly reduced in kaempferol-treated cells, despite

the strong SdhA signal obtained with the Sdh antibody in the reprobing. Interestingly, the

acetylation signal coming from the lower band was also affected by kaempferol and

nicotinamide treatments (Fig. 3.4B).

Again, to determine the role of SdhA acetylation on Complex II activity, we

performed Complex II enzyme activity assays using whole cell lysates obtained from

nicotinamide-and kaempferol-treated K562 cells, which revealed that the Complex II was

about ~20 % more active in kaempferol-treated cells compared to the Complex II activity

from nicotinamide-treated cells (Fig. 3.4C). The Complex II activity in control cells was

similar to the activity of nicotinamide-treated cells, possibly due to their comparable

acetylation status (Fig. 3.4C).

144

Figure 3.4A-B: Effect of SIRT3 overexpression on SdhA deacetylation in K562 cells.

A. Immunoblotting analysis of K562 cell lysates obtained from control (Cont),

nicotinamide- (Nam), and kaempferol- (Kaem) treated cells. Approximately, 20 μg of

control and treated K562 cell lysates from each sample were loaded onto 12 % SDS-

polyacrylamide gels and immunoblotting was performed as described above. Actin and

Hsp60 blots were shown to ensure equal loading of protein amounts (Dr. M. J. Han

contributed this experiment). B. Equal amounts (about 2 mg) of control and treated K562

cell lysates were layered on a 34 % sucrose cushion and fractionated into eight 1 mL

aliquots after high speed centrifugation. Equal volumes of each fraction (1–8) were

acetone precipitated and loaded on 12 % SDS-polyacrylamide gels for immunoblotting

analysis. Arrows show the location of acetylated protein overlapping with the SdhA

signal.

145

Figure 3.4C: Effect of SIRT3 overexpression on Complex II activity in K562 cells.

C. Complex II activity was monitored using different amounts of kaempferol- and

nicotinamide-treated K562 cell lysates. The analysis was done in triplicate and values

shown are the mean ±SD. One way ANOVA test was used to compare the significance of

the values. Asterisks denote p<0.05.

146

3.5 Conclusions and Future Directions

Mitochondria are required for the production of more than 90 % of the ATP

required for survival of eukaryotic cells in oxidative phosphorylation. Regulation of

oxidative phosphorylation and Krebs cycle components by post-translational

modifications has already been established (2,7,29,30). ADP/ATP and [NADH]/[NAD+]

ratios are important for regulation of these pathways either by post-translational

modifications such as phosphorylation and acetylation or by allosteric regulation.

Regulation of mitochondrial function by phosphorylation is known for a long time;

however, the recent progress in identification of mitochondria specific NAD+-dependent

sirtuins such as SIRT3, SIRT4, and SIRT5, revealed the importance of [NADH]/[NAD+]

ratio in regulation of protein and enzyme function in post-translational modifications by

reversible acetylation (14,28). One of the best characterized mitochondrial NAD+-

dependent deacetylase, SIRT3, has been known to regulate activities of several metabolic

enzymes and the Complex I subunit NDUFA9 by deacetylation (17). Moreover, we have

discovered its pivotal role in the regulation of mitochondrially encoded proteins of

oxidative phosphorylation by mitochondrial protein synthesis through specific

deacetylation of a ribosomal protein MRPL10 (18).

In this study, comparison of acetylated proteins in wild type and SIRT3 knock-out

mice mitochondria has led us to a novel substrate for SIRT3, the flavoprotein of succinate

dehydrogenase complex (SdhA), along with a known substrate, glutamate

dehydrogenase. SdhA is one of the hydrophilic subunits of the succinate dehydrogenase

involved in both the Krebs cycle and oxidative phosphorylation in mammalian

147

mitochondria. Previously, in two independent high throughput surveys of the acetylated

proteins of the rat liver, several acetylated peptides were mapped from SdhA (2,7), while

it was reported as an unacetylated protein in a comprehensive study of SIRT3-dependent

deacetylation of Complex I subunit NDUFA9 (17). However, the role of acetylation in

the enzyme activity and the deacetylase responsible for this modification were not

determined previously.

We believe that the data presented here convincingly clarify the discrepancy

reported in the literature and demonstrate that SIRT3 is indeed the major mitochondrial

deacetylase controlling oxidative phosphorylation by reversible lysine acetylation

(16,17). In the comparison of 2D-gel immunoblotting of SIRT3−/− and SIRT3+/+ mice

liver mitochondria, SdhA was found to be hyperacetylated in the absence of SIRT3;

however, it is possible that the degree of acetylation in wild-type mice is regulated by

availability of acetyl-coA and/or NADH levels in the mitochondria. For this reason, we

have not observed complete deacetylation of SdhA in the wild-type mice liver

mitochondrial lysates (Fig. 3.1C and 3.3A). More importantly, we have shown the effect

of hyperacetylation on Complex II activity in SIRT3−/− liver mitochondria (Fig. 3.3B).

Interestingly, the Complex II activity in SIRT3 knock-out mice was ~ 30 % lower than

that of the wild-type, possibly due to incomplete deacetylation of SdhA in the wild-type

mice (Fig. 3.3B). Previously, none of the Complex II subunit proteins was reported as

acetylated proteins for the immunocaptured Complex II components in SIRT3 knock-out

mice (17). This discrepancy could be due to the sample preparation used by Ahn et al. as

they determined the acetylation of Complex II components after immunocapturing of the

complex (17). In addition to changes in SdhA acetylation and Complex II activity in

148

SIRT3−/− and SIRT3+/+ mice mitochondria, we have shown a decrease in SdhA activity

while a slight increased level of acetylation was observed in cells treated with a general

deacetylase inhibitor, nicotinamide. In contrast, kaempferol treatment of the same cell

line caused an increase in the level of SIRT3 expression and significant deacetylation of

SdhA accompanied by 20 % increase in Complex II activity possibly due to SIRT3-

dependent deacetylation of SdhA. Surprisingly, the changes in acetylation of SdhA did

not completely inhibit the Complex II activity. As proposed previously, it is likely that

only a minor proportion of the protein is acetylated or acetylation only partially regulates

the enzyme activity even though mitochondrial protein hyperacetylation is dramatic in

SIRT3 knock-out mice (16). Additionally, conserved acetylated lysine residues in

mammalian SdhA are located on the surface of the protein, away from the active site of

the enzyme. Therefore, it is feasible to expect that acetylation of the positively charged

residues on the surface of the enzyme might either slightly change the affinity of the

enzyme for its negatively charged substrate, succinate, or induce conformational changes

to reduce the activity of the enzyme (Fig. 3.2C).

Regulation of Complex II activity by reversible acetylation of SdhA subunit

relates how oxidative phosphorylation and Krebs cycle components are regulated by

metabolite levels in mammalian mitochondria. In the case of high levels of reduced

cofactors such as NADH and FADH2 present in the mitochondria, there is no need for

further oxidation of acetyl-CoA in the Krebs cycle for generation of these cofactors to

support oxidative phosphorylation. Thus, it would be reasonable to suggest that

acetylation of SdhA just slows the Krebs cycle, as this process will also cause

accumulation of acetyl-CoA in the mitochondria. On the other hand, when NAD+ level

149

increases in the mitochondria, SIRT3 and other NAD+-dependent deacetylases will be

activated and deacetylate SdhA and other acetylated components of the Krebs cycle. In

agreement with stimulation of catalytic activities of metabolic enzymes such as glutamate

dehydrogenase and acetyl-CoA synthetase 2 by deacetylation, deacetylation of SdhA also

stimulates Complex II or succinate dehydrogenase activity to promote the Krebs cycle for

the generation of reduced NADH and FADH2, as they are the electron donors for ATP

synthesis in oxidative phosphorylation. Another potential regulation of Complex II

activity is by phosphorylation of the SdhA subunit as it was found to be phosphorylated

by Fgr tyrosine kinase in vitro (10). Given its importance in oxidative phosphorylation, it

could be suggested that this enzyme can be regulated through cooperation or interplay

between these two different post-translational modifications at varying metabolite levels.

Moreover, in the case of complete inhibition of the complex, succinate accumulation

resulting from the decreased SdhA activity may cause deleterious effects in the cell due

to the absence of additional mitochondrial metabolic enzymes that can metabolize

succinate (8,9).

3.6 Acknowledgment

We thank Dr. Qiang Tong for providing SIRT3 knockout mice liver

mitochondria.

150

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Chapter 4

Identification and Analysis of New Mammalian Mitochondrial Ribosomal

Proteins: CHCHD1, AURKAIP1, and CRIF1

This chapter of dissertation was reproduced with permission from Emine C.

Koc, Huseyin Cimen, Beril Kumcuoglu, Nadiah Abu, Gurler Akpinar, Md. Haque,

Linda Spremulli, Hasan Koc. Identification and Characterization of CHCHD1,

AURKAIP1, and CRIF1 as New Members of the Mammalian Mitochondrial

Ribosome. Nucleic Acids Research (Manuscript submitted.) Copyright © 2012 by

Oxford University Press.

153

4.1 Rationale

Mitochondrially encoded components of the oxidative phosphorylation

(OXPHOS) complexes are all essential and synthesized by 55S ribosomes in

mammalian mitochondria. In the initial identification of mitochondrial ribosomal

proteins (MRPs), it was challenging to identify MRPs with no clear homologs in

bacteria due to the limited availability of expressed sequence tag (ESTs) from

different organisms in the databases. With the improvement in genome sequencing

and increased sensitivity of mass spectrometry (MS)-based technologies, we have

identified five additional proteins as new MRPs; MRPS37 (Coiled-coil-helix-coiled-

coil-helix domain containing protein 1-CHCHD1), MRPS38 (Aurora kinase A

interacting protein 1-AURKAIP1), MRPS39 (Pentatricopeptide repeat-containing

protein 3-PTCD3), MRPL58 (Immature colon carcinoma transcript 1 protein-ICT1),

and MRPL59 (CR-6 interacting factor 1-CRIF1), and a ribosome-associated assembly

factor, C7orf30, in highly purified 55S ribosomes. The human PTCD3 and ICT1 and

yeast homologs of CHCHD1 and AURKAIP1 had been observed to be associated

with mitochondrial ribosomes previously. Here, we also report that CHCHD1 and

AURKAIP1 are bona fide small subunit proteins and CRIF1 is a bona fide large

subunit protein of the 55S ribosome in mammalian mitochondria and demonstrate that

they have essential roles in mitochondrial protein synthesis in vivo.

154

4.2 Introduction

Mammalian mitochondria are responsible for providing over 90 % of the

energy used by cells in the form of ATP through oxidative phosphorylation

(OXPHOS). ATP production by this process depends on electron transport chain

complexes and the ATP synthase, components of which are encoded by both the

nuclear and mitochondrial genomes. In mammals mitochondrial DNA encodes the

information for only 13 essential proteins required for OXPHOS in addition to 22

tRNAs and 2 rRNAs. The 13 proteins encoded within the mitochondrial genome are

synthesized on 55S ribosomes using the specialized translational system within the

organelle.

Mammalian mitochondrial ribosomes (55S) consist of small (28S) and large

(39S) subunits (1,2). The 55S ribosome is composed of ~80 mitochondrial ribosomal

proteins (MRPs) and all of these proteins are encoded by nuclear genes and imported

into mitochondria where they are assembled into the ribosome with mitochondrially

transcribed rRNAs (3-6). About a decade ago, almost all of the proteins present in

these ribosomes were identified by using various proteomic techniques. This initial

analysis indicated that the small subunit of the ribosome contains 29 proteins while

the large subunit has about 50 proteins (5-11). About half of these proteins in

mammalian mitochondrial ribosomes have homologs in bacterial ribosomes and play

a role either in the assembly and structure of ribosomes or in the initiation, elongation,

or termination phases of mitochondrial translation. The functions and locations of the

mitochondrial-specific proteins are not known; however, they may replace some of

the functions of bacterial ribosomal proteins that are not present in the mitochondrial

ribosome. They may also provide additional function(s) critical for protein synthesis

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or its regulation in mammalian mitochondria (5,6). Although it is not possible to

determine the exact locations of these mitochondrial-specific proteins without x-ray

structural information, cryo-EM reconstruction studies indicated that they are

distributed on the exterior surface of the ribosome (12,13). It is clear that some of the

mitochondrial-specific ribosomal proteins are located in functionally important

regions of the ribosome particularly at the mRNA entrance gate in the small subunit,

and at the polypeptide exit tunnel and the central protuberance region of the large

subunit creating specific structures on the mitochondrial ribosome (12,13).

In many instances mitochondrial-specific ribosomal proteins were identified

when searching for their additional functions due to lack of homology with known

ribosomal proteins in bacteria and eukaryotic 80S ribosomes. For example,

mitochondrial-specific MRPs such as MRPS29 (DAP3-death associated protein 3),

MRPS30 (PDCD9-programmed cell death protein 9), MRPL37, and MRPL41 have

been reported to be involved in apoptosis (5,6,14-17). While major mutations of

MRPs causing functional changes in mitochondrial translation are shown to be lethal,

aberrantly expressed MRPs are also related to many different tumors including breast,

gliomas, squamous cell carcinoma, and osteosarcoma (6,18-21). Therefore, a

complete list of mitochondrial ribosomal proteins will be fundamental for our

understanding of the mitochondrial translational machinery and its contribution to

mitochondrial energy production and biogenesis in health and disease.

In the present study, we reevaluated protein components of the mammalian

mitochondrial ribosome using mass spectrometry (MS)-based proteomics and

identified five potential new mitochondrial ribosomal proteins: coiled-coil-helix-

coiled-coil-helix domain containing protein 1 (CHCHD1), aurora kinase A interacting

protein 1 (AURKAIP1), pentatricopeptide repeat-containing protein 3 (PTCD3),

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immature colon carcinoma transcript 1 protein (ICT1), CR-6 interacting factor 1

(CRIF1 also known as growth arrest and DNA-damage-inducible proteins-interacting

protein 1 (Gadd45GIP1)), and a ribosome-associated assembly factor C7orf30. The

new mitochondrial ribosomal proteins; CHCHD1, AURKAIP1, and PTCD3 were

assigned to the small subunit and ICT1 and CRIF1 were assigned to the large subunit

of the 55S ribosome.

4.3 Materials and Methods

4.3.1 Preparation of Bovine Mitochondrial Ribosomal Subunits

Mitochondrial ribosomes from bovine liver were prepared using a previously

described method at high and low ionic strength and at several different detergent

concentrations (Fig. 4.1) (5,6,22,23). The high ionic strength and detergent

concentrations used the standard conditions of 300 mM KCl and 1.6 % Triton X-100

(10). For the preparation of mitochondrial ribosomes at low salt and detergent

conditions, mitochondrial lysates were prepared in a buffer containing 50 mM Tris-

HCl, pH 7.6, 40 mM KCl, 20 mM MgCl2, 6 mM -mercaptoethanol, 0.2 % Triton X-

100, and 1 mM phenylmethylsulfonyl fluoride (PMSF). The mitochondrial lysate was

layered onto a 34 % cushion solution (40 mM KCl, 20 mM MgCl2, 50 mM Tris-HCl,

pH 7.6, 6 mM -mercaptoethanol, and 34 % (w/w) sucrose). Samples were

centrifuged at 35,000 rpm for 16 h at 4 ºC in a Beckman type-50.2 rotor. The pellet

was collected as crude mitochondrial ribosomes and resuspended in a buffer prepared

with 40 mM KCl, 20 mM MgCl2, 20 mM Tris-HCl pH 7.6, 1 mM dithiothreitol

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(DTT), and protease inhibitor cocktail from Sigma-Aldrich. Samples were then

subjected to centrifugation through a 10-30 % linear sucrose gradient in buffer

containing 40 mM KCl, 20 mM MgCl2, 50 mM Tris-HCl pH 7.6, 1 mM DTT.

Fractions containing 55S ribosomes were combined and the ribosomes were collected

by centrifugation at 40,000 rpm for 16 h (Fig. 4.2A). The concentration of Mg2+ in the

preparations was reduced to 2 mM by dialysis in order to dissociate ribosomes into

28S and 39S subunits and the samples were separated again on a linear 10-30 %

sucrose gradient containing 2 mM Mg2+. Highly purified 28S and 39S subunits were

collected by centrifugation at 40,000 rpm for 16 h (Fig. 4.2B).

4.3.2 Identification of Mitochondrial Ribosomal Proteins by Mass Spectrometry

To identify the proteins of mammalian mitochondrial ribosomes, purified

ribosomal 28S and 39S subunits and 55S samples were separated on SDS-

polyacrylamide gel and corresponding protein bands were excised into at least thirty

equal gel pieces which were processed by performing in-gel tryptic digestion using

methods previously established in our laboratory (Fig. 4.2B) (24-26). Tryptic digests

were analyzed by capillary liquid chromatography-nanoelectrospray ionization-

tandem mass spectrometry (capLC-MS/MS). Extracted tryptic peptides (3-5 μL) were

injected and loaded into a peptide trap (Michrom peptide CapTrap, C8 like resin, 0.3

× 1 mm, 5 μm) over 3 min at 10 μL/min for online desalting and concentration. With

the use of the six-port switching valve, the peptide trap was then placed in-line with

the analytical column, a PicoFrit column (0.075 × 150 mm) packed in-house with

Wide Bore C18 reverse phase resin (Supelco Co., 5 μm, 300 Å). Tandem MS spectra

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of tryptic peptides were obtained by collision-induced dissociation (CID) in an LTQ

linear ion trap mass spectrometer (ThermoFinnigan) system including a Surveyor

HPLC pump and a Surveyor Micro AS autosampler.

MS/MS spectra were processed by Xcalibur 2.0 and searched against

nonredundant protein and Swiss-Prot and UnitProtKB databases using the Mascot

server. Additionally, the search was repeated using a bovine protein sequence

database generated in-house. Protein information obtained from the database searches

and the scores of mitochondrial ribosomal proteins which were observed in multiple

bands were compared. To increase the data quality, proteins with a Mascot score

lower than 45 were excluded from the list.

Ribosomal proteins were assigned to subunits according to their abundance in

28S and 39S fractions. Exponentially modified Protein Abundance Index (emPAI)

score was calculated to compare the protein abundance in each sample using a

previously reported method (27). Briefly, the ratio of observed unique parent ion

number in the analysis to the observable peptide number from in silico digestion is

used as PAI in the formula: emPAI = 10^(PAI) – 1.

4.3.3 Preparation of Crude Ribosomes from Human Cell Lines and Isolated

Mitochondria

HeLa cells were grown in Dulbecco’s Modified Eagle’s Medium (DMEM)

media (Cellgro, Mediatech Inc.), supplemented with 10 % (v/v) bovine calf serum

(Hyclone Laboratories) and 100 IU/ml penicillin and 100 μg/ml streptomycin, at 37oC

and 5 % CO2 in a humidified atmosphere. For the whole cell lysate preparation,

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approximately 4 x 107 HeLa cells were combined and lysed in 2 mL of buffer

containing 50 mM Tris-HCl, pH 7.6, 0.26 M sucrose, 60 mM KCl, 20 mM MgCl2, 0.8

mM EDTA, 2 mM DTT, 0.05 mM spermine, 0.05 mM spermidine, 1.6 % Triton X-

100, and protease inhibitor cocktail from Sigma-Aldrich using a Dounce homogenizer

(Wheaton). In order to isolate mitochondria, approximately 2 x 107 HeLa cells were

resuspended in 1 mL of an isotonic mitochondrial buffer (MB) (210 mM mannitol, 70

mM sucrose, 1 mM EDTA, 10 mM HEPES-KOH, pH 7.5), supplemented with

protease inhibitors (1 mM PMSF and protease cocktail from Sigma-Aldrich), and then

homogenized in a Dounce homogenizer on ice. The suspension was centrifuged at

400 x g on a microcentrifuge (ThermoForma) at 4°C. The pellet was resuspended in

another 1 mL of MB and the 400 x g centrifugation was repeated. Supernatants were

combined and centrifuged at 10,000 x g at 4°C for 10 min to pellet mitochondria. The

mitochondrial pellets were lysed in a buffer containing 0.26 M sucrose, 20 mM Tris-

HCl, pH 7.6, 40 mM KCl, 20 mM MgCl2, 0.8 mM EDTA, 0.05 mM spermine, 0.05

mM spermidine, 6 mM β-mercaptoethanol, and 1.6 % Triton X-100 using a Dounce

homogenizer.

To collect the crude ribosomes, whole cell and mitochondrial lysates (2 mL)

were layered onto a 34 % sucrose cushion (4 mL) in buffer (50 mM Tris-HCl, pH 7.6,

60 mM KCl, 20 mM MgCl2, and 6 mM β-mercaptoethanol) and centrifuged in a Type

40 rotor (Beckman Coulter) at 40,000 rpm for 16 h. The post-ribosomal supernatant

was fractionated into six separate layers (designated L1 to L6) for analysis and the

pellet was collected as a crude ribosomal fraction (Fig. 4.4). The crude ribosome

preparations including mitochondrial and cytoplasmic ribosomes for whole cell lysate

and only mitochondrial ribosomes for the mitochondrial lysate were resuspended in

50 µL of Base Buffer III (50 mM Tris-HCl, pH 7.6, 60 mM KCl, 20 mM MgCl2, 1

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mM DTT, and protease inhibitor cocktail (Sigma-Aldrich). Ribosome suspensions

were stored at -80oC for further analyses.

4.3.4 RNase A Treatment of Mitochondrial Ribosomes

In order to confirm the direct or indirect interaction of new MRPs with rRNA

of mitochondrial ribosome, approximately, ~5 A260 units of a crude preparation of

ribosomes obtained from bovine liver were incubated in the absence or presence of

RNase A and loaded onto separate 10-30 % linear sucrose gradients in buffer

containing 40 mM KCl, 20 mM MgCl2, 50 mM Tris-HCl pH 7.6 and 1 mM DTT

(28). After centrifugation, equal volumes (25 μL) of gradient fractions were separated

on 12 % SDS-polyacrylamide gel and probed with corresponding antibodies after

transferring to PVDF membranes (Fig. 4.6).

4.3.5 Immunoblotting Analysis

Ribosome samples collected from HeLa and bovine mitochondria including

sucrose gradient fractions, purified 55S ribosomes and 28S and 39S subunits were

separated on 12 % SDS-polyacrylamide gels. Proteins were transferred to PVDF

membranes which were probed with rabbit polyclonal anti-CHCHD1 antibody at a

1:1000 dilution (Abcam), rabbit anti-AURKAIP1 antibody at a 1:1000 dilution

(Sigma-Aldrich), goat anti-CRIF1 antibody at a 1:500 dilution (Santa Cruz

Biotechnology), mouse monoclonal anti-MRPS29 and anti-HSP60 antibodies at

1:5000 dilutions (BD Transduction Laboratories), and mouse anti-OXPHOS

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(MITOPROFILE ® Total OXPHOS Antibody Cocktail, Mitosciences Inc.) at 1:5000

dilution (Abcam Inc.), or rabbit polyclonal human anti-MRPL47 and mouse

polyclonal human anti-MRPS18-2 antibodies at 1:5000 dilutions (Covance Inc.) for

16 h at 4ºC. The secondary antibodies, donkey anti-goat IgG (Santa Cruz

Biotechnology) for CRIF1, goat anti-mouse IgG (Pierce Biochemicals Inc.) for

MRPS29, HSP60 and OXPHOS, and goat anti-rabbit IgG for CHCHD1, AURKAIP1,

and MRPL47, were all used at 1:5000 dilutions. The membranes were developed

using SuperSignal West Pico Chemiluminescent (Pierce Biochemicals Inc.) according

to the protocol provided by the manufacturer.

4.3.6 [35S]-Methionine Labeling of Mitochondrial Translation Products In Vivo

Human embryonic kidney 293T (HEK293T) cell lines were cultured in

DMEM (Cellgro, Mediatech Inc.) supplemented with 10 % bovine calf serum

(Hyclone), 100 IU/ml penicillin and 100 μg/ml streptomycin, at 37oC and 5 % CO2 in

a humidified atmosphere. Cells were transfected with control siRNA (sc-44235) and

siRNA against CHCHD1 (sc-90488), AURKAIP1 (sc-72472), and CRIF1 (sc-97804)

from Santa Cruz Biotechnology using Lipofectamine™ 2000 (Invitrogen) according

to the protocol provided by the manufacturer. The transfected cells were grown in

transfection medium for 2 days prior to labeling with [35S]-methionine. Labeling

experiments were performed in the presence of dialyzed serum (25 mM Tris-HCl, pH

7.4, 137 mM NaCl, and 10 mM KCl) and minimum essential DMEM medium which

does not contain methionine, glutamine, and cysteine (10,29,30). Cells were incubated

with emetine containing medium for 5 min to arrest cytoplasmic protein synthesis and

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0.2 mCi/ml of [35S]-methionine (Perkin Elmer) containing medium was added to label

the mitochondrially-encoded proteins. After a 2 h incubation, cells were lysed in

buffer containing 50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1 mM EDTA, 1 mM

EGTA, 0.1 % SDS and 0.5 % NP-40 supplemented with 1 mM PMSF and protease

inhibitor cocktail (Sigma-Aldrich). Whole cell lysates (40 μg) were electrophoresed

through 12 % SDS-polyacrylamide gel. The gels were dried on 3MM chromatography

paper (Whatman) and then total intensities of the signals were quantified by

phosphorimaging analyses (Fig. 4.8) (31). Results from four independent experiments

were analyzed using the ANOVA test. Values of P<0.05 were considered statistically

significant. siRNA mediated knockdown efficiency of corresponding mitochondrial

ribosomal protein was confirmed with immunoblotting analysis of whole cell lysate or

crude ribosomal fraction prepared as stated above (Fig. 4.7A).

4.3.7 Reverse Transcription Polymerase Chain Reaction (RT-PCR)

Total RNA was isolated from siRNA transfected HEK293T cells by using

RNeasy Mini Kit from Qiagen and the cDNA was synthesized using the

ThermoScriptTM RT-PCR system (Invitrogen). Primers used: CHCHD1 forward 5'-

ACCTCTCATTCTAGCTAACCGCGT -3', reverse 5'-

AGACTCTCCCAGGGTTTCCTGTAT -3'; AURKAIP1 forward 5'-

TCCACCGCAATCCTACCAGTGT -3', reverse 5'-

CGAACTTGATCTGCTTGCGTCTCA -3'; CRIF1 forward 5'-

GATGATTGTGAACTGGCAGCAGCA -3', reverse 5'-

CGCCTCCTTCTTCCGTTTCTGTTT -3'; ND6 forward 5'-

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GAGTGTGGGTTTAGTAATGGGGTTTGTGGGG -3', reverse 5'-

CCTATTCCCCCGAGCAATCTC -3'; COI forward 5'-

ATTTAGCTGACTCGCCACACTCCA -3', reverse 5'-

TAGGCCGAGAAAGTGTTGTGGGAA -3'; COII forward 5'-

ATGGCACATGCAGCGCAAGTA -3', reverse 5'-

CTATAGGGTAAATACGGGCCC-3'; ATP6 forward 5'-

TAATACGACTCACTATAGATGAACGAAAATATGT -3', reverse 5'-

TTTTTTTTTTTTTTTTTTTTTTCATTGTTGGGTGGTGATTAG -3'; 12s rRNA

forward 5'- AATAGGTTTGGTCCTAGCCTAGCC -3’, reverse 5'-

GTTCGTCCAAGTGCACTTTCCAG -3'; MRPS29 forward 5'-

ATGGACCGACACGGGTATTGTACC -3', reverse 5'-

AAGGCCATGGGGAAATACAGTC -3'; MRPL47 forward 5'-

AAACGGGGTACCGAGATGGCTGCGGCCGGTTTGGCCC -3', reverse 5'-

CCGCTCGAGTTAATGGTGATGGTGATGATGGACAAGACTTGACTTTTGGG

C -3'; glyceraldehyde 3-phosphate dehydrogenase (GAPDH) forward 5'-

GTCTTCACCACCATGGAGAAGG -3', reverse 5'-

ATGAGGTCCACCACCCTGTTGC -3';. Reactions were performed according to the

instructions provided by the manufacturer. Samples were visualized using ChemiDoc

XRS system employing Quantity One® 1D analysis software (Bio-Rad). Quantitative

graph was constructed by relative signal intensities to GAPDH sample.

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4.4 Results and Discussion

4.4.1 Identification of 55S Ribosomal Proteins by Tandem Mass Spectrometry

The majority of the protein components of mammalian mitochondrial

ribosomes were identified by our group as well as several other groups using

proteomic strategies a decade ago (5-9,32,33). However, due to the limited

availability of bovine or rat protein and DNA sequence information, some of the

ribosomal and/or ribosome-associated proteins were not detected and identified by

matching the tandem MS data to the publicly available ESTs or protein databases

available at that time. An analysis of the protein composition of a large

macromolecular complex requires the preparation of the complex under conditions

that are strong enough to remove contaminants but are gentle enough to prevent the

loss of protein components that are more loosely bound. In order to reevaluate the

protein composition and eliminate transiently associated proteins, mammalian

mitochondrial ribosomes were purified under two different salt and detergent

conditions from bovine liver (Fig. 4.1) (23). The 10-30 % linear sucrose gradient

separation of mitochondrial ribosomes was carried out at 20 mM Mg2+ to collect 55S

particles (Fig. 4.1).

The protein compositions of the 55S preparations prepared under the two

different salt and detergent conditions were compared on SDS-polyacrylamide gels

(Fig. 4.2A). In general, the gel pattern was similar in both ribosomal preparations;

however, the sample prepared at high salt/detergent concentrations contained

relatively lower amounts of high molecular weight proteins. The proteins in these

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bands may be subunits of the large enzyme complexes co-sedimenting with the

mitochondrial ribosome fractions (Fig. 4.2 and Fig. 4.3). The gels were sliced into

thirty fractions and in-gel tryptic digestion of each gel piece was analyzed by capLC-

MS/MS. Their peptide/protein contents were determined by matching MS/MS spectra

to an in-house protein sequence database generated by the combination of ~ 30,000

bovine proteins found in the Swiss-Prot and UniProtKB databases. The complete list

of the previously unidentified proteins was created by excluding MRPs identified

earlier, mitochondrial metabolic pathway proteins, and oxidative phosphorylation

proteins. In these analyses, CHCHD1, AURKAIP1, PTCD3, ICT1, CRIF1, and

C7orf30 were repeatedly found as additional ribosome-associated proteins with high

confidence in both low and high salt and detergent preparations (Table 4.1). With the

exception of large complexes of metabolic enzymes, specifically 2-oxoglutarate and

pyruvate dehydrogenases and ATP synthase F1 subunits sedimenting with the

ribosomes, proteins consistently found in both low and high salt and detergent

preparations of 55S ribosomes were considered as possible components of the

mitochondrial translational machinery and/or ribosomal proteins (Table 4.1).

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Figure 4.1: An experimental scheme for the purification of mitochondrial ribosomes and 28S and 39S subunits.

In order to obtain purified 55 ribosomes and 28S and 39S subunits using sequential linear sucrose gradients, bovine

mitochondrial lysates were prepared at different ionic and detergent conditions to remove mitochondrial metabolic enzymes co-

sedimenting with the mitochondrial ribosome. Experimental details are included in Materials and Methods.

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Figure 4.2: SDS-PAGE analyses of purified bovine 55S ribosomes and 28S and 39S subunits.

A. Approximately 0.5 A260 units of mitochondrial ribosomes (55S) purified at

two different salt and detergent concentrations (Low and High, see Materials and

Methods for details) were separated by 14 % SDS-polyacrylamide gels and stained

with Coomassie Blue. B. Purified mitochondrial (55S) ribosomes (~10 A260) prepared

at high salt and detergent conditions were dissociated into the small (28S) and large

(39S) subunits by sedimentation through another 10-30 % linear sucrose gradient in

the presence of 2 mM Mg2+. The same amounts (0.5 A260) of subunits and 55S

ribosomes was separated by SDS-polyacrylamide gel which was excised into 30

pieces for in-gel digestion of protein bands with trypsin for LC-MS/MS analysis to

identify their protein content. These samples were prepared for analysis by B.

Kumcuoglu and N. Abu. Gel fractions containing the peptides detected from newly

identified MRPs are marked on the image.

A B

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Figure 4.3: Mitochondrial ribosomes prepared at different salt and detergent

concentrations.

Approximately 50-60 A260 units of low (A) and high (B) salt and detergent

preparations of crude ribosomes were separated on 10-30 % linear sucrose gradients to

sediment purified 55S ribosomes (fractions containing purified 55S ribosomes are

shown by arrows). Different subsets of metabolic enzyme complexes were removed by

sedimentations performed at low and high salt and detergent conditions; however, 55S

fractions shown by arrows mainly contained the components of the mitochondrial

translation machinery. Each fraction corresponding to dissociated small (28S) and

large (39S) subunits and intact (55S) ribosomes was separated by 12 % SDS-

polyacrylamide gels. The 55S fractions shown by arrows were collected for the

preparation of purified 28S and 39S subunits. Dr. G. Akpinar contributed to this

experiment.

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4.4.2 Subunit Assignments of Newly Identified Ribosome-associated Proteins

In order to categorize newly identified proteins as either MRPs or proteins

involved in translation, subunit assignments of these proteins was essential. Bovine

55S ribosomes prepared at high salt and detergent conditions (Fig. 4.2A) were

sedimented on another 10-30 % linear sucrose gradient containing 2 mM Mg2+ to

promote dissociation of the small (28S) and large (39S) subunits (Fig. 4.3). Purified

28S, 39S, and 55S samples were separated on SDS-polyacrylamide gel and stained

with Coomassie Blue (Fig. 4.2B). Although the amount of high molecular weight

proteins found in the 55S ribosome preparation decreased, a few of them still

remained associated with the 28S and 39S subunits in the second sucrose gradient

(Fig. 4.2B). The Coomassie Blue stained gel clearly showed the differential

protein/MRP distribution in the 28S and 39S fractions (Fig. 4.2B). Each lane was cut

into thirty equal pieces; in-gel tryptic digestions were carried out and the resulting

peptides analyzed by capLC-MS/MS and database searches were performed as

described for the analysis of 55S ribosomal proteins (Fig. 4.2B).

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Table 4.1: Peptide sequences of new mitochondrial ribosomal proteins identified

from capLC-MS/MS analyses of in-gel tryptic digestions of 28S, 39S, and 55S

proteins.

Name Sequence Score m/z Mr(expt) CHCHD1 SIQEDLGELGSLPPR 112 805.7 1609.4 KPILKPNKPLILANHVGER 97 713.2 2136.7 SIQEDLGELGSLPPRK 85 870.4 1738.7 AURKAIP1 EAPPGWQTPK 83 556.7 1109.6 AGLKEAPPGWQTPK 74 740.7 14.79.4 PTCD3 ADVHTFNSLIEATALVVNAK 150 1057.8 2113.7 TWDKVAVLQALASTVHR 145 948.4 1894.9 AHTQALSMYTELLNNR 143 931.9 1861.7 DLELAYQVHGLLNTGDNR 124 1014.9 2027.9 GSSLIIYDIMDEITGK 118 878.3 1754.6 DEGADIAGTEEVVIPK 113 821.7 1641.5 QMVAQNVKPNLQTFNTILK 110 1094.5 2186.9 DPDDDMFFQSAMR 102 788.2 1574.4 LTADFTLSQEQK 95 691 1379.9 SDLKEEILMLMAR 91 775.4 1548.9 EALGDLTALTSDSESDSDSDTSKDK 90 863.8 2588.4 VAVLQALASTVHR 88 683 1364.1 NELLNEFMDSAK 80 706.3 1410.5 ASSSPAQAVEVVK 79 636.7 1271.4 TFSPKDPDDDMFFQSAMR 72 723.6 2167.9 AGHQLGVTWR 63 563.4 1124.7 LEMIPQIWK 55 579.4 1156.8 WNNILDLLK 54 564.9 1127.9 EEILMLMAR 51 553.6 1105.2 ICT1 DMIAEASQPATEPSKEDAALQK 124 1165.9 2329.9 FHLASADWIAEPVR 119 806.8 1611.5 SAYSLDKLYPESR 95 766.3 1530.7 QGNDDIPVDR 90 565.3 1128.5

AGELILTSEYSR 79 670.4 1338.8 VPGDAKQGNDDIPVDR 69 565.6 1693.9 GADTAWRVPGDAK 60 672.6 1343.2

CRIF1 AAAMAAAAAQDPADSETPDS 116 930.9 1859.8 HGAASGVDPGSLWPSR 115 797.5 1593.1 EQLLELEAEER 77 680.3 1358.6 FQELLQDLEK 62 632.3 1262.7 MPQMIENWR 54 602.9 1203.8 C7orf30 YTDYFVIGSGTSTR 100 784.0 1566.0 FDIDMLVSLLR 84 670.3 1338.6 Table is generated by Dr. E. Koc and Dr. H. Koc.

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In general, the same set of proteins, which was consistently found in capLC-

MS/MS analyses of 55S ribosomes prepared at different ionic and detergent

conditions, was also identified in 28S and 39S subunits. The proteins transiently

associated with the 55S ribosome including mitochondrial EF-Tu and ribosome

release factor (RRF) were released when the 55S ribosome preparation was

dissociated into 28S and 39S ribosome in the second sucrose gradient at 2 mM Mg2+.

Six new proteins (CHCHD1, AURKAIP1, PTCD3, ICT1, CRIF1, and C7orf30) were

consistently observed in 28S or 39S subunits and 55S ribosomes making it crucial to

determine whether these proteins are bona fide ribosomal proteins or ribosome-

associated proteins. As marked in Fig. 4.2B, the gel slices where the peptides detected

from these proteins by capLC-MS/MS analyses were in good agreement with their

expected molecular masses after removal of mitochondrial import signals (34). In

addition, the experimental emPAI scores described by Ishihama et al. (27) were

calculated to demonstrate the relative protein abundance in each subunit using the

ratio of peptides detected by capLC-MS/MS analyses to the number of observable

peptides obtained from in silico digestion of a protein (Table 4.2). The GenBank™

and Swiss-Prot access numbers of the newly identified MRPs used in the emPAI

determination are listed in Table 4.3. The agreement between the emPAI scores for

the 55S ribosome and either 28S or 39S subunit values for each protein (except

PTCD3 and ICT1) clearly shows that these proteins were associated with the

ribosome and its subunits at conditions used in our experiments (Table 4.2). Slight

variations in emPAI scores could be due to the variations in excision of gel slices,

extraction of peptides and/or data dependent acquisition of peptides by the capLC-

MS/MS system. Some of the characteristics of these new ribosomal proteins are

described below.

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Table 4.2: Relative distribution of new mitochondrial ribosomal proteins using

emPAI (Exponentially Modified Protein Abundance Index) values.

Subunit CHCHD1 AURKAIP1 PTCD3 ICT1 CRIF1 C7orf30

28S 3.30 0.27 169.61 0.97 0.49 0.22

39S ND 0.19 0.64 1.09 2.02 0.48

55S 3.14 0.27 22.77 0.86 2.11 0.30

Peptides (ion score cut off of ≥ 45) were used in emPAI calculations.

ND; not detected.

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Table 4.3: GenBank™ and Swiss-Prot accession numbers of new mitochondrial

ribosomal proteins found in various species.

MRP Protein Human Bovine Mouse

MRPS37 CHCHD1 Q96BP2 Q2HJE8 XP_852408

MRPS38 AURKAIP1 Q9NWT8 Q0VCJ1 XM_843641

MRPS39 PTCD3 Q96EY7 Q2KI62 XP_532975

MRPL59 ICT1 Q14197 Q3T116 XP_533118

MRPL60 CRIF1 Q8TAE8 A1A4P4 XP_533898

MRPL61 C7orf 30 Q96EH3 Q0P562 Q9CWV0

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CHCHD1: Although a signal peptide cleavage site was not predicted by MitoProt II,

the probability of a mitochondrial localization of bovine CHCHD1 is 81% (34). The

full length bovine CHCHD1 is about 13.6 kDa. Mass spectrometry analysis of tryptic

peptides extracted from the protein bands excised from 28S, 39S, and 55S lanes

resulted in identification of CHCHD1 peptides in 28S and 55S samples but not in 39S

subunits (Table 4.2). The emPAI values calculated using detected CHCHD1 peptides

clearly suggest that this protein is mainly associated with the small subunit of

mitochondrial ribosomes (Table 4.2). Although the mammalian mitochondrial

CHCHD1 has a homolog in yeast mitochondrial ribosome, MRP10, which has about

20% sequence identity (35,36), here we provide the first experimental evidence for

CHCHD1 being a component of the animal mitochondrial small subunit. This is

confirmed with immunoblotting analysis of purified mitochondrial ribosomal subunits

in Fig. 4.5.

AURKAIP1: AURKAIP1, also known as Aurora-A-interacting protein (AIP), was

first described as a regulator of Aurora-A kinase, which is a Ser/Thr kinase involved

in cell cycle progression and tumorigenesis (37-39). The unknown domain found in

AURKAIP1 homologs is described as DUF1713 and this unknown domain has high

homology with the C-terminal domain of yeast COX24. The calculated molecular

mass of the full length AURKAIP1 is 22.4 kDa; however, the mature protein migrates

at about 16 kDa (Fig. 4.2B). It is possible that AURKAIP1 has a longer signal peptide

than the predicted signal peptide which provides a 99% possibility for the

translocation of this protein into mitochondria. Translocation of AURKAIP1 into the

mitochondria has also been experimentally validated by the Human Protein Atlas

(HPA) project (40). AURKAIP1 was repeatedly detected in the capLC-MS/MS

analyses of low and high salt preparation of bovine mitochondrial ribosomes and

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subunits. The calculated emPAI values for peptides obtained from purified 28S and

39S preparations suggested association of AURKAIP1 with the small subunit since its

emPAI value for 28S subunit is 30 % higher than that of the 39S subunit (Table 4.2).

This observation was also supported by immunoblotting analysis performed for

AURKAIP1 distribution in these samples in Fig. 4.5.

PTCD3: We first reported PTCD3 as an mRNA-binding protein associated with the

28S subunit as a PET309 homolog due to the presence of the pentatricopeptide

repeats (41). Association of PTCD3 with the small subunit has also been reported in

addition to its essential role in mitochondrial translation (42). We have discovered

that it is one of the 28S subunit proteins interacting with mitochondrial initiation

factor 3 (43). The full length bovine PTCD3 is 77.8 kDa with a pI of 6.0, possibly the

largest protein and one of the most acidic components of the 55S ribosome. It is

highly conserved among its animal mitochondrial homologs (Fig. 4.2B). Due to the

size and lower pI of the PTCD3 compared to the basic proteins of the mitochondrial

ribosome, nineteen unique peptides were detected in capLC-MS/MS analyses

(Table 4.1). In agreement with the earlier observations, PTCD3 peptides were mainly

detected in protein bands excised from 28S and 55S lanes (Table 4.2). Its subunit

localization is also supported by the emPAI values shown in Table 4.2; specifically,

the high experimental emPAI value for peptides identified in the 28S subunit sample

as well as in the 55S sample indicates that PTCD3 is a ribosomal component

associated with both 28S subunit and 55S ribosomes. The presence of a trace amount

of PTCD3 in 39S preparations could be due to the presence of a small amount of the

28S subunits in the large subunit fractions analyzed. This cross-fractionation is not

uncommon even for MRPs with clear bacterial homologs (6). High emPAI scores of

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the 28S subunit and 55S ribosomes enabled us to conclude that PTCD3 is a

mitochondrial-specific MRP mainly associated with the 28S subunit.

ICT1: This protein belongs to the polypeptide release family of proteins and is the

first example of a peptidyl-tRNA hydrolase activity provided by an integral

component of the large subunit (44). ICT1 is essential for the hydrolysis of peptidyl-

tRNAs on prematurely terminated mRNAs lacking a stop codon (44,45). Tryptic

peptides detected by capLC-MS/MS analyses of protein bands are listed in Table 4.1.

ICT1 peptides were detected in the 28S, 39S, and 55S samples, making the calculated

emPAI values of the ICT1 in 28S and 39S subunits very close to each other. The

experimental emPAI scores were not different enough to assign ICT1 to a particular

subunit (Table 4.2). The human ICT1 was previously shown to be associated with the

large subunit (44); however, our MS data does not clearly support association of ICT1

with the large subunit. Here, it is plausible to suggest that ICT1 is located near the

interface region of the large subunit and a fraction of ICT1 also sediments with the

small subunit.

CRIF1: The calculated molecular mass of the full length bovine CRIF1 is 25.7 kDa.

The mitochondrial localization signal peptide is predicted to be in the first 28 amino

acid residues with a 90 % possibility which was calculated by MitoProt II (34). The

calculated molecular weight for the mature CRIF1 is 22.9 kDa and this value is in

agreement with the migration of the protein in SDS-polyacrylamide gel detected by

capLC-MS/MS (Fig. 4.2B). The emPAI values calculated from five unique CRIF1

peptides detected in the 39S and 55S samples strongly suggest that the mitochondrial

CRIF1 is a large subunit protein (Table 4.2). Association of CRIF1 with the large

subunit is also demonstrated with immunoblotting assay of sucrose gradient fractions

and of the purified subunits from these fractions (Fig. 4.5).

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C7orf30: Bovine C7orf30 is a 234 amino acid residue long protein with a predicted

signal peptide cleavage site within the first 45 residues. C7orf30 has a homolog in

bacteria, ybeB. C7orf30 was recently discovered to be essential for the assembly or

stability of mitochondrial ribosomes by interacting with the large subunit of the

ribosome analogous to bacterial ybeB (46,47). Its role in translation is confirmed by

siRNA knock-down studies in human cell lines causing an overall reduction in

expression of mitochondrially expressed proteins (46,47). In agreement with these

recent studies, a fraction of C7orf30 remained associated with the 55S ribosome and

39S subunits in the LC-MS/MS analyses of the ribosome and its subunits prepared

under different salt conditions (Table 4.2).

4.4.3 Localization of CHCHD1, AURKAIP1, and CRIF1 into the Mitochondria

and Their Roles in Mitochondrial Translation

In the capLC-MS/MS analyses of purified bovine mitochondrial ribosomal

proteins presented above, we clearly demonstrated that CHCHD1 and AURKAIP1 are

small subunit proteins and CRIF1 is a highly conserved large subunit protein of the

55S ribosome. However, various ectopically expressed forms of CRIF1 in human cell

lines have been shown to localize to the nucleus and to be involved in transcriptional

activation (48). In order to evaluate the mitochondrial ribosome association of

CHCHD1, AURKAIP1, and CRIF1, HeLa whole cell and mitochondrial lysates

prepared under non-denaturing conditions were centrifuged through a 34 % sucrose

cushion, a step analogous to that used in the preparation of bovine liver mitochondrial

ribosomes (Fig. 4.4). This process enriched the multimeric complexes of human cell

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and mitochondrial lysates resistant to high salt and non-ionic detergent treatment at

the bottom layer and pellet, mainly as ribosomes. In immunoblotting analyses of

postribosomal supernatant layers and crude ribosomal pellets obtained from the whole

cell and mitochondrial lysates, new MRPs were found mainly in the pellet along with

two mitochondrial ribosomal proteins; MRPS29 and MRPL47, confirming their co-

localization with the mitochondrial ribosome and/or the other large complexes in

mitochondria (Fig. 4.4).

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Figure 4.4: Sedimentation of new MRPs with large complexes in human cell lines and

mitochondria.

A. Whole cell lysate (WCL) and B. mitochondrial lysate (MITO) obtained from

human cell lines were layered on sucrose cushion preparations. After 16 h centrifugation,

post-ribosomal supernatant layers (Layers 1 to 6) and crude ribosome pellet (P) were

collected and analyzed by immunoblotting probed with CHCHD1, AURKAIP1, CRIF1,

MRPS29, MRPL47, and HSP60 antibodies.

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Following the enrichment of multimeric complexes of mitochondria, the crude

bovine mitochondrial ribosomal pellet was layered and sedimented through a 10-30 %

linear sucrose gradient to separate free subunits and 55S ribosomes (Fig. 4.5A). Again

the immunoblotting analysis of sucrose gradient fractions with antibodies against

MRPS29, and MRPL47 confirmed the association of new MRPs only with the

mitochondrial ribosome (Fig. 4.5A). Free-pools of new MRPs in early fractions of the

sucrose gradient are absent which implies that each of these proteins is an integral part

of the ribosome rather than a ribosome-associated factor which dissociates from the

ribosomes as observed for EF-Tu, RRF, or C7orf30. In order to confirm the

association of CHCHD1 and AURKAIP1 with the small subunit and CRIF1 with the

large subunit, fractions corresponding to the 55S ribosome were combined and

dissociated into 28S and 39S subunits by lowering the Mg2+ concentration prior to

their separation on another linear sucrose gradient (Fig. 4.5B). Immunoblotting

analyses of purified subunits and 55S ribosome displays overlapping signals of

CHCHD1 and AURKAIP1 with MRPS29 antibody in 28S and 55S confirming their

association with the small subunit, and of CRIF1 with MRPL47 antibody in the 39S

and 55S fractions indicating its association with the large subunit (Fig. 4.5B). Further

evidence for RNA-dependent association of these new MRPs with the mitochondrial

ribosome was provided by the RNase A-treatment of the 55S ribosome prior to the

loading on the sucrose gradient. As shown by the immunoblotting analysis, signals

from the new MRPs completely disappeared in the sucrose gradient fractions of 55S

ribosomes treated with RNase A similar to the other mitochondrial ribosomal proteins

while the HSP60 signal remained in these fractions (Fig. 4.6). The data obtained by

capLC-MS/MS and immunoblotting analyses together suggest that CHCHD1 and

181

AURKAIP1 are newly described components of the small subunit while CRIF1 is a

bona fide component of the large subunit of mitochondrial ribosomes.

182

Figure 4.5: Distribution of new MRPs in 28S and 39S subunits.

A. Crude bovine mitochondrial ribosomes prepared at high salt conditions

(~10 A260) were layered on a 10-30 % sucrose gradient. Sedimentation of intact

ribosomes and ribosomal subunits into the sucrose gradient were analyzed by

immunoblotting using MRPS18-2, MRPS29, and MRPL47 antibodies to mark the

small (28S) and large subunit (39S) locations. CHCHD1 and AURKAIP1 co-

sediments with the 28S subunit and 55S ribosomes, while CRIF1 co-sediments

mainly with the 39S subunit and 55S ribosomes. B. Purified 28S and 39S subunits

(~0.2 A260) obtained from dissociation of 55S fractions shown in (A) at 2 mM Mg2+

were separated on a 12 % SDS-polyacrylamide gel and analyzed by immunoblotting

using CHCHD1, AURKAIP1, CRIF1, MRPS29, and MRPL47 antibodies to confirm

subunit distribution of the new MRPs.

183

Figure 4.6: RNA-dependent association of new MRPs with the 55S ribosome.

Same amounts of crude bovine mitochondrial ribosomes (~5 A260), Control

(A) and RNAse A-treated (B), were sedimented on 10-30 % sucrose gradients.

Sedimentation of the new MRPs with ribosomal subunits and 55S ribosomes in the

absence and presence of RNAse A was analyzed by immunoblotting using CHCHD1,

AURKAIP1, CRIF1, MRPS29, MRPL47, and HSP60 antibodies. Equal volumes of

sucrose gradient fractions were also analyzed by 12 % SDS-polyacrylamide gel.

184

Mitochondrial ribosomes synthesize thirteen essential subunits of the

OXPHOS complexes, including subunits of Complex I (ND1, ND2, ND3, ND4,

ND4L, ND5 and ND6), Complex III (Cytb), Complex IV (COI, COII, COIII), and

Complex V (ATP6 and ATP8). Given the association of the new MRPs with the

mitochondrial ribosome described above, we next examined the effect of CHCHD1,

AURKAIP1, and CRIF1 knock-down on the de novo synthesis of mitochondrially-

encoded proteins by [35S]-methionine labeling in the presence of the cytoplasmic

protein synthesis inhibitor, emetine. For this purpose, HEK293T cells were

transfected with control siRNA and corresponding specific siRNAs to CHCHD1,

AURKAIP1, and CRIF1. The specific knock-down of CHCHD1, AURKAIP1, and

CRIF1 in HEK293T cell lines transfected with siRNA was confirmed by

immunoblotting analysis of whole cell lysate for AURKAIP1, MRPS29, MRPL47,

and HSP60 and crude ribosomal fraction for CHCHD1 and CRIF1 using

corresponding antibodies (Fig. 4.7A) in addition to the RT-PCR assay (Fig. 4.7B).

Next, the expression of [35S]-methionine labeled mitochondrial proteins were

analyzed by SDS-PAGE (Fig. 4.8). In cells transfected with siRNAs to new MRPs,

mitochondrial protein synthesis was decreased by at least 30 % compared to the cells

transfected with control siRNA while the total protein content of the cell lysates were

comparable (Fig. 4.8). Given that a considerable amount of new MRPs remained in

siRNA knock-down cells, the 30% reduction in overall mitochondrial translation is

significant (Fig. 4.8). In addition, when we analyzed the steady state levels of

oxidative phosphorylation subunits in these samples, the reduction in Complex IV

subunit, COII, which is a mitochondrial encoded protein, but not in the subunits of

Complex V, Complex III, and Complex II, which are nuclear encoded, implies the

essentiality of these proteins in mitochondrial protein synthesis as a member of

185

ribosomes (Fig. 4.9). Clearly, the inhibition of protein synthesis in knock-down cells

was due to a reduction in expression of new MRPs since the reprobing of the same

membrane with MRPS29, MRPL47, and HSP60 antibodies showed no difference in

total ribosome or mitochondrial contents (Fig. 4.7A). In order to confirm that the

observations in knock-down cells were not due to changes in the transcription levels

of mitochondrial proteins or ribosomal RNA, transcription levels of ND6, COI, COII,

ATP6, 12S rRNA, MRPS29, MRPL47, and GAPDH were assayed by RT-PCR. Their

levels in knock-down cells were not altered when compared to the cells transfected

with control siRNA (Fig. 4.7B). Altogether, these observations suggest that

CHCHD1, AURKAIP1, and CRIF1 all have essential roles in mitochondrial protein

synthesis as a component of the small and large subunit proteins in the 55S ribosome.

186

Figure 4.7A: siRNA mediated knock-down of new MRPs.

Expression of new MRPs in control HEK293T and siRNA transfected cells

was analyzed by immunoblotting analysis using antibodies against CHCHD1,

AURKAIP1, CRIF1, HSP60, MRPS29, and MRPL47. HSP60 antibody was used as a

loading control for total mitochondrial and MRPS29 and MRPL47 antibodies are for

ribosome contents. Signal intensities were graphed relative to siCon.

* : crude ribosome preparations were used instead of cell lysate as in others.

187

Figure 4.7B: RT-PCR analysis of mitochondrial- and nuclear-encoded transcripts.

Transcript levels of CHCHD1, AURKAIP1, and CRIF1 were shown as an

indication of knock-down efficiency in cell lines transfected with control siRNA and

siRNAs for corresponding new MRP. Mitochondrial encoded ND6, COI COII, ATP6,

and 12S rRNA, and nuclear encoded MRPS29 and MRPL47 were examined by RT-

PCR reactions in the same samples as well. RT-PCR reactions with GAPDH were

performed as a positive control. Signal intensities were quantified relative to siCon and

given as a graph.

188

Figure 4.8: Effect of new MRPs knock-down on mitochondrial protein synthesis.

A. De novo synthesis of mitochondrial proteins was evaluated in control and

siRNA transfected HEK293T cells (CON: control and knock-down of corresponding

MRP) by pulse labeling of proteins in the presence of [35S]-methionine and a

cytosolic translation inhibitor, emetine. A. representative electrophoretic pattern of the

de novo synthesized translational products is presented. ND1, -2, -3, -4, -4L, -5, and -

6 are subunits of Complex I; Cytb is a subunit of Complex III; COI, -II, and -III are

subunits of Complex IV; ATP6 and ATP8 are subunits of Complex V. Coomassie

Blue staining of the same gel was performed to ensure equal protein loading in the

gel. B. The combined intensities of 13 mitochondrially-encoded proteins from each

lane were used as an overall quantitation of mitochondrial protein synthesis.

Shown is the mean ± SD of four independent experiments. Results were analyzed by

one way ANOVA test; *, P < 0.05.

* * *

189

Figure 4.9: Effect of new MRPs knock-down on OXPHOS subunits.

Immunoblotting analysis of whole cell lysates prepared from control siRNA

and new MRP specific siRNA transfected HEK293T cells to examine the steady state

levels of nuclear encoded subunits of Complex V, III, and II and mitochondrial

encoded subunit of Complex IV, COII. The same immunoblot was used to probe with

MRPS29, MRPL47, and HSP60 to ensure equal loading.

190

4.4.4 Nomenclature

Two-dimensional gel analysis of the mitochondrial ribosomal proteins in

bovine liver suggested the presence of as many as 33 ribosomal proteins in the 28S

subunit and 52 ribosomal proteins in the 39S subunit (5,6). These proteins were

designated S1 through S33 for the 28S subunit and L1 through L52 for the 39S

subunit in order of decreasing molecular mass. However, this nomenclature system

did not provide consistency for designation of same proteins from different

organisms. Hence, we adopted a system of nomenclature, in which proteins with

prokaryotic homologues are given the same number as the corresponding ribosomal

protein in E. coli. Proteins without bacterial homologues are given the next available

number. Bacterial ribosomes have proteins designated S1 through S21 in the 28S

subunit and L1 through L36 in the large subunit. Additionally, previously identified

mammalian ribosomal proteins without bacterial homologues were designated S22

through S36 for the 28S subunit and L37 through L57 for the 39S subunit. As a result,

newly identified small subunit proteins; CHCHD1, AURKAIP1, and PTCD3, will be

named as MRPS37, MRPS38, and MRPS39 and ICT1 and CRIF1, as newly identified

large subunit proteins MRPL58 and MRPL59, respectively (Table 4.4).

191

28S and 39S: small and large subunit of mitochondrial ribosome 30S and 50S: small and large subunit of bacterial ribosome New Class: mitochondria-specific ribosomal proteins

Table 4.4: List of mammalian mitochondrial ribosomal proteins with their bacterialhomologs

28S 30S New Class 39S 50S New Class Missing S1 MRPS22 MRPL1 L1 MRPL37 MRPS2 S2 MRPS23 MRPL2 L2 MRPL38 Missing S3 MRPS24 MRPL3 L3 MRPL39 Missing S4 MRPS25 MRPL4 L4 MRPL40 MRPS5 S5 MRPS26 Missing L5 MRPL41 MRPS6 S6 MRPS27 Missing L6 MRPL42 MRPS7 S7 MRPS28 MRPL12 L7/L12 MRPL43 Missing S8 MRPS29 MRPL9 L9 MRPL44 MRPS9 S9 MRPS30 MRPL10 L10 MRPL45 MRPS10 S10 MRPS31 MRPL11 L11 MRPL46 MRPS11 S11 MRPS32 MRPL13 L13 MRPL48 MRPS12 S12 MRPS33 MRPL14 L14 MRPL49 Missing S13 MRPS34 MRPL15 L15 MRPL50 MRPS14 S14 MRPS35 MRPL16 L16 MRPL51 MRPS15 S15 MRPS36 MRPL17 L17 MRPL52 MRPS16 S16 MRPS37 (CHCHD1) MRPL18 L18 MRPL53 MRPS17 S17 MRPS38 (AURKAIP1) MRPL19 L19 MRPL54 MRPS18-1 S18 MRPS39 (PTCD3) MRPL20 L20 MRPL55 MRPS18-2 S18 MRPL21 L21 MRPL56 MRPS18-3 S18 MRPL22 L22 MRPL57 (RP_63) Missing S19 MRPL23 L23 MRPL58 (ICT1) Missing S20 MRPL24 L24 MRPL59 (CRIF1) MRPS21 S21 Missing L25 MRPL60 (C7orf30) MRPL27 L27 MRPL28 L28 MRPL47 L29 MRPL30 L30 Missing L31 MRPL32 L32 MRPL33 L33 MRPL34 L34 MRPL35 L35 MRPL36 L36

4.5 Conclusions and Future Directions

Ribosomal proteins have roles in ribosome assembly, substrate binding, and/or

different stages of translation (initiation, elongation, and termination). The MRPs with

bacterial homologs are expected to have conserved functions in ribosome structure and

translation. For the mammalian mitochondrial-specific proteins without known homologs

in other ribosomes, it is highly challenging to determine whether a ribosome-associated

protein is a genuine ribosomal protein or a factor transiently associated with the

ribosome. The criteria used in this study, however, allowed us to assign the newly

identified proteins as mitochondrial-specific ribosomal proteins due to their distribution

similar to that of the MRPs with bacterial homologues in highly purified subunit and

ribosome preparations (Table 4.1). With the newly identified MRPs, the number of

mammalian mitochondrial ribosomal proteins is brought to 31 for the small subunit and is

increased to 51 for the large subunit (Table 4.4) (5,6,11,33,49). It is not feasible to assign

specific roles for the majority of mitochondrial-specific proteins without structural

information and their relative locations in the ribosome; however, many of them have

been shown to be essential for mitochondrial protein synthesis and/or function. For

instance, the yeast homologue of MRPS37 (CHCHD1), MRP10, was found to be

indispensible for mitochondrial translation (50,51). A respiratory defect caused by a null

mutant of MRP10 was recovered by the reintroduction of the MRP10 gene in a wild-type

mitochondrial DNA background (35). Similarly, MRPS38 (AURKAIP1) is a possible

homolog of yeast COX24p involved in processing and translation of COX I mRNA (52).

193

Unlike the mitochondrially encoded genes in yeast, mammalian mitochondrial genes do

not contain introns to be processed or require minimal processing. MRPS37 and MRPS38

are ribosomal small subunit proteins involved in translation since their knock-down in

cells impairs the mitochondrial translation of thirteen essential proteins of oxidative

phosphorylation. However, their specific role in mammalian mitochondrial translation

remains to be discovered.

siRNA knock-down of MRPS39 (PTCD3) decreased Complex III and Complex

IV activities possibly by directly affecting mRNA binding to the mitochondrial ribosome

(42). In fact, PTCD3 was identified as one of the IF3mt interacting proteins in the small

subunit (43). Although the majority of the MRPs forming the platform of the mRNA-

binding path in the 28S subunit are bacterial homologs such as MRPS7, MRPS11,

MRPS18, and MRPS21, the shoulder region and the mRNA-gate of the 28S subunit are

mainly formed by mitochondrial-specific MRPs (28). Therefore, it is possible that

MRPS39 is one of the proteins forming the mRNA-binding path interacting with the 5’-

ends of mitochondrial mRNAs.

MRPL58 was initially identified as immature colon carcinoma transcript 1 (ICT1)

since it was one of the transcripts differentially expressed in colorectal tumors that

deviate from the normal maturation pathway in colon epithelium (53,54). Later, it was

discovered to be an unusual member of a release factor family involved in termination of

mitochondrial translation with a codon-independent peptidyl-tRNA hydrolase activity

associated with the mitochondrial ribosome (44,55). It was also recently reported that

MRPL58 is essential for cell viability since its knock-down resulted in reduced

194

cytochrome c oxidase activity, mitochondrial membrane potential, and eventually

apoptotic cell death (45).

The other new large subunit protein MRPL59 (CRIF1) has been identified as a

transcription co-factor which controls the G1/S phase of the cell cycle (56). It also

negatively regulates the stability of a transcription factor NRF2 (nuclear respiratory

factor 2), which promotes expression of many mitochondrial proteins and proteins

involved in oxidative damage, by ubiquitination (48). Here, the majority of these studies

were performed using the ectopically expressed form of CRIF1 with a tag at its amino

terminus; therefore, it is possible that the majority of tagged CRIF1 may not get

incorporated into the 55S ribosome due to masking of mitochondrial targeting signal and

is subsequently translocated into the nucleus. Another explanation could be that a very

small fraction of CRIF1 is located into the nucleus to manifest its function as a

transcription co-factor. However, when the tag is placed at protein’s carboxy terminus,

the protein is targeted to mitochondria (57). Recently, it has been reported to be

interacting with ATAD3 as a component of mitochondrial nucleoprotein complexes

confirming CRIF1 to be localized in mitochondria (57). Endogenous levels of CRIF1 has

been shown to be dramatically reduced in epithelial cell cancers in thyroid and breast

which is in agreement with possible mitochondrial dysfunction reported in many different

cancer types (56). In this study, we provided very strong evidence for CRIF1 being a

bona fide ribosomal protein and an essential component of the mitochondrial translation

machinery in vivo.

195

Given that the mitochondrial translational machinery and its components are

essential for the expression of OXPHOS subunits, studies related to the identification of

new components of the translational machinery and their specific roles in translation have

the utmost importance in understanding the energy production by OXPHOS. A

completed picture of the mitochondrial translation machinery will also allow us to assess

mitochondrial dysfunction manifested as neurodegenerative diseases, aging, diabetes, and

cancer.

4.6 Acknowledgment

This work was supported in part by the National Institutes of Health grants

[R01GM32734 to L.L.S., R01GM071034 to E.C.K.].

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57. He, J., Cooper, H. M., Reyes, A., Di Re, M., Sembongi, H., Litwin, T. R., Gao, J., Neuman, K. C., Fearnley, I. M., Spinazzola, A., Walker, J. E., and Holt, I. J. (2012) Nucleic Acids Res

Chapter 5

Concluding Remarks and Future Directions

The mitochondria provide most of the cellular energy in the form of ATP through

respiratory chain complexes in eukaryotic cells. During the transfer of electrons through

these complexes, mitochondria generate reactive oxygen species (ROS) which is

implicated in cellular death and aging. Given that mitochondrial ribosomes are

responsible for translating the essential subunits of the respiratory chain complexes

crucial for oxidative phosphorylation, post-translational modifications of ribosomal

proteins may be one mechanism of regulating ATP production in order to maintain

energy homeostasis in the eukaryotic cell. This is plausible since acetyl-CoA, NAD+, and

ATP are the mitochondrial metabolites which are involved in acetylation, deacetylation,

and phosphorylation of target proteins, respectively. Our group previously reported the

implication of phosphorylated ribosomal proteins in regulation of mitochondrial

translation (1). In addition, recent studies have demonstrated that reversible acetylation is

involved in the regulation of mitochondrial enzymes, akin to their phosphorylation (2-7).

Overall, our findings in chapter 2 suggest that MRPL10 is the major acetylated

ribosomal protein in mitochondria, which is a bona fide substrate for NAD+-dependent

deacetylase, SIRT3. Similar to the bacterial ribosomes, MRPL10 serves the structural

surface for MRPL12 binding to the L7/L12 stalk region on the ribosome. MRPL12 plays

a significant role in translation by recruiting initiation, elongation, and release factors to

the ribosome (8-10). In bacterial ribosomes, acetylation of RPL12 was found to stabilize

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the interaction between stalk proteins due to increased hydrophobicity by acetylation

(11). The location of acetylated lysine residues on MRPL10, which were mapped using

mass spectrometry-based proteomic analyses, prompted us to reveal the role of

acetylation in modulation of protein-protein interactions in the L7/L12 stalk and

mitochondrial protein synthesis (Fig. 5.1). In SIRT3 knock-out mice, increased

acetylation status of mitochondrial ribosome enhanced MRPL12 binding to the ribosome

and recruitment of elongation factors to increase translation. On the other hand, SIRT3

over-expression reduced MRPL12 binding to ribosomes by deacetylation of MRPL10. In

addition, deacetylation of the mitochondrial ribosome impairs the mitochondrial protein

synthesis and suppresses the activities of respiratory chain Complexes I and IV resulting

in reduced ATP production in cells overexpressing SIRT3. We therefore propose a model

where the SIRT3-dependent reversible acetylation of MRPL10 may cause changes in

MRPL12 binding to the ribosome by neutralizing the positively charged lysine residues

to regulate mitochondrial protein synthesis (Fig. 5.1).

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Figure 5.1: Model for the role of MRPL10 acetylation on MRPL12 binding to

ribosome.

The mitochondrial L7/L12 stalk is formed by MRPL10 (green), MRPL11 (yellow),

and MRPL12 (blue), which are highly conserved between mammalian mitochondria and

bacteria. SIRT3 removes acetyl groups in an NAD+-dependent reaction producing

nicotinamide (NAM) and O-acetyl-ADP-ribose (O-Acetyl-ADPR). Deacetylated lysine

residues located at the N-terminal domain of MRPL10 and lysine residues in MRPL12

were shaded in red. Crystal structure model of the T. thermophilus 50S subunit (PDB:

2WRL) and solution structure of the E. coli L7/L12 dimers (PDB: 1RQU) were used to

model location of multiple copies of MRPL12 on the ribosome. The 50S ribosomal 23S

and 5S rRNA, and ribosomal proteins were colored in cyan and pink, respectively.

Structural models were generated by PyMol software (DeLano Scientific LLC) (12).

Courtesy of Dr. E. Koc.

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Overall data suggest that modulation of MRPL12 binding to the ribosome might

be one of the crucial mechanisms that regulate the synthesis of thirteen essential

components of OXPHOS complexes and ATP production in response to SIRT3 activity

in mitochondria.

In order to reveal the direct correlation between MRPL10 acetylation and

MRPL12 binding to ribosome and mitochondrial translation, site-directed mutagenesis

studies were performed on selected three acetylated residues, Lys124, Lys169, and

Lys196, to generate Arg and Gln mutants mimicking hypoacetylation and

hyperacetylation, respectively. However, due to the technical difficulties in our

preliminary studies, the in vitro and in vivo translation assays performed with MRPL10

mutants were not conclusive (Chapter 2). It is also possible that MRPL10 might have

additional acetylated lysine residues, which are involved in the modulation of MRPL10

interaction with MRPL12 on the stalk. One approach might be followed in the future

experiments is to knock-out RPL10 in bacteria with recombination techniques and to

supplement the bacteria with various RPL10 and MRPL10 plasmids including their

lysine mutants to construct a hybrid ribosomal system (13). In this system, stability of

MRPL10 expressed in bacteria would be enhanced due to its incorporation into the

bacterial ribosomes. Since the bacteria lack RPL10, the resulting ribosomes would be

mostly the hybrid ribosomes containing MRPL10. This process would also maintain the

activity of MRPL10. In our petDUET® overexpression system (Invitrogen Inc.),

recombinant MRPL10 and its mutants over-produced in bacteria was mostly insoluble

possibly because of its hydrophobic nature even though MRPL12 was also included in

the expression system to promote and stabilize MRPL10-MRPL12 complex formation.

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Even the soluble fraction of the partially purified MRPL10-MRPL12 complex did not

restore the translation activity of bacterial stripped ribosomes possibly due to the co-

purification of heat-shock proteins in my preliminary studies as opposed to our earlier

published studies (8).

The source of protein acetylation in mitochondria was not known and believed to

be a non-enzymatic reaction due to the abundance of acetyl-CoA in mammalian

mitochondria until recently (14). An alternative view was that mitochondrial proteins

could be acetylated prior to their import into mitochondria. However, a recent report on

the acetylation of ATP synthase F0 subunit 8, which is encoded and synthesized in

mitochondria, strongly suggested the presence of an acetyltransferase in mitochondria

(7). In a recent report, a component of the mitochondrial acetyltransferase system,

GCN5L1, was demonstrated to catalyze the acetylation of respiratory chain targets and

inhibit their enzymatic activities (15). This enzyme reverses the effects of sirtuins in

mitochondria. We will perform experiments with GCN5L1 to determine whether it is

involved in the acetylation of MRPL10 and, therefore, in the modulation of L7/L12 stalk

composition. The identification of the other components of the mitochondrial

acetyltransferase which are responsible for the acetylation of mitochondrial ribosomal

proteins is also plausible by using mass spectrometry-based proteomics and biochemical

approaches in mammalian mitochondria.

Succinate dehydrogenase, which is involved in both the electron transport chain

and TCA cycle, was identified as a novel substrate of SIRT3 in our two-dimensional gel

electrophoresis separation followed by mass spectrometry and immunoblotting analyses

of SIRT3 knock-out mice liver mitochondria. SIRT3 dependent deacetylation of SdhA

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subunit of the enzyme increases its activity. Acetylated lysine residues on the enzyme are

located on the surface close to the substrate binding site. In another study, additional

acetylated lysine residues on succinate dehydrogenase were reported to be involved in the

regulation of its enzymatic activity (16). It is possible that there are still additional

acetylated lysine residues to be discovered or acetylation of lysines occurs in a sequence-

dependent manner to modify the surface of SdhA. Since succinate dehydrogenase

participates in both respiratory chain and TCA cycle, it is of great interest to study how

acetylated lysine residues are coordinated and how it contributes to the regulation of its

function during aging or under different conditions, such as exposure to hypoxia and

reactive oxygen species, and even in cells lacking mitochondrial DNA.

Identification of additional proteins and factors involved in translation is

important for deep understanding of the regulation of mitochondrial protein synthesis

machinery in mammals. Therefore, the discovery of three additional mitochondrial

ribosomal proteins described in Chapter 4 provides a more complete picture of

mitochondrial ribosomes and protein synthesis. To reveal their specific roles in

mitochondrial translation, further structural information of mitochondrial ribosomes is

required. Generation of highly specific antibodies against mitochondria-specific

ribosomal proteins and cryo-EM analysis with improved resolution may provide possible

roles of the newly identified ribosomal proteins. On the other hand, there are still

questions that need to be addressed in terms of the translation process. For instance, it is

still unclear how mitochondrial mRNAs are positioned on the ribosomes during

translation initiation and the factors that are involved in this process remain to be

discovered. Furthermore, the coordination of mitochondrial translation with other

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processes including replication and transcription of mitochondrial genome is still under

investigation to identify fundamental relevance of these processes (17,18).

Mitochondrial ribosomes, which are in the process of translating a subset of

respiratory chain subunits, were found to be associated with inner membrane through

Oxa1L (19). It is currently unknown where exactly the protein synthesis occurs in

mitochondria but translating ribosomes are believed to be enriched in intracristal regions

of inner membrane (20). Visualization of individual translating mitochondrial ribosomes

in the organelle is not possible due to limitations in optical resolution of fluorescence

microscopy. However, recent improvement in stimulated emission depletion microscopy

(STED), which was employed to visualize VDAC isoforms in mitochondria, might be

sufficient to detect and analyze ribosomes in mitochondria (21). With the availability of

highly specific antibodies against mitochondrial ribosomal proteins, this technique might

provide the visualization of ribosomes during translation. Heterogeneous population of

mitochondria in cells challenges the imaging techniques when heteroplasmy, which is the

co-existence of normal and mutant mtDNAs, needs to be studied in single mitochondrion.

In addition, heterogeneous population of mitochondria or even mitochondrial ribosomes

might be generated by genetic and biochemical modifications. For instance,

overexpression of a mutant mitochondrial ribosomal protein might provide a subset of

ribosomes with that mutant protein incorporated (or even a subset of mitochondria where

mutant protein is imported). Its effect on mitochondrial functions may not be significant

since abundant normal ribosomes could compensate this deficiency. Analysis of single

mitochondrion containing the target mutant protein and even further the analysis of

mitochondrial ribosomes having the mutant protein incorporated may provide the

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significant insight about the role of that particular mutation in mitochondrial protein

synthesis and mitochondrial dysfunction. Flow cytometry (FC), laser capture

microdissection (LCM), and optical tweezers (OT) are the main techniques employed to

separate single mitochondrion (22). FC is a quantitative tool employed in the analysis of

single mitochondrion under different conditions, such as mtDNA composition, membrane

potential, and ROS production (23-25). LCM is optically controlled removal of a single

mitochondrion from specific cells or tissue. The analysis of mtDNA from individual cells

or cancer tissue is automated with this technique (26,27). OT uses a highly focused

infrared laser beam to trap single mitochondrion from a single cell. This technique

provided the detection of heteroplasmy at mitochondrion level (28). Overall, these

techniques to study single mitochondrion are expected to be advanced enough to analyze

ribosomes from single mitochondrion.

In conclusion, acetylation of protein synthesis machinery and respiratory chain

complexes in mammalian mitochondria broadens our knowledge of the role of post-

translational modifications in mitochondrial respiration. Post-translational modifications

of mitochondrial proteins and their interrelationships might enable us to devise novel

strategies to understand and modulate mitochondrial function/dysfunction in disease

states in our future studies.

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4. Ahn, B. H., Kim, H. S., Song, S., Lee, I. H., Liu, J., Vassilopoulos, A., Deng, C. X., and Finkel, T. (2008) Proc Natl Acad Sci U S A 105(38), 14447-14452

5. Schwer, B., Bunkenborg, J., Verdin, R. O., Andersen, J. S., and Verdin, E. (2006) Proc Natl Acad Sci U S A 103(27), 10224-10229

6. Choudhary, C., Kumar, C., Gnad, F., Nielsen, M. L., Rehman, M., Walther, T. C., Olsen, J. V., and Mann, M. (2009) Science 325(5942), 834-840

7. 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(4), 607-618

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20(5), 723-731 11. Gordiyenko, Y., Deroo, S., Zhou, M., Videler, H., and Robinson, C. V. (2008) J

Mol Biol 380(2), 404-414 12. Brunger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P., Grosse-

Kunstleve, R. W., Jiang, J. S., Kuszewski, J., Nilges, M., Pannu, N. S., Read, R. J., Rice, L. M., Simonson, T., and Warren, G. L. (1998) Acta Crystallogr D Biol Crystallogr 54(Pt 5), 905-921

13. Gaur, R., Grasso, D., Datta, P. P., Krishna, P. D., Das, G., Spencer, A., Agrawal, R. K., Spremulli, L., and Varshney, U. (2008) Mol Cell 29(2), 180-190

14. Guarente, L. (2011) Cold Spring Harb Symp Quant Biol 76, 81-90 15. Scott, I., Webster, B. R., Li, J. H., and Sack, M. N. (2012) Biochem J 443(3), 655-

661 16. Finley, L. W., Haas, W., Desquiret-Dumas, V., Wallace, D. C., Procaccio, V.,

Gygi, S. P., and Haigis, M. C. (2011) PLoS One 6(8), e23295 17. Surovtseva, Y. V., Shutt, T. E., Cotney, J., Cimen, H., Chen, S. Y., Koc, E. C.,

and Shadel, G. S. (2011) Proc Natl Acad Sci U S A 108(44), 17921-17926 18. He, J., Cooper, H. M., Reyes, A., Di Re, M., Sembongi, H., Litwin, T. R., Gao, J.,

Neuman, K. C., Fearnley, I. M., Spinazzola, A., Walker, J. E., and Holt, I. J. (2012) Nucleic Acids Res

19. Haque, M. E., Elmore, K. B., Tripathy, A., Koc, H., Koc, E. C., and Spremulli, L. L. (2010) J Biol Chem 285(36), 28353-28362

20. Christian, B. E., and Spremulli, L. L. (2011) Biochim Biophys Acta http://dx.doi.org/10.1016/j.bbagrm.2011.11.009

21. Neumann, D., Buckers, J., Kastrup, L., Hell, S. W., and Jakobs, S. (2010) PMC Biophys 3(1), 4

22. Pflugradt, R., Schmidt, U., Landenberger, B., Sanger, T., and Lutz-Bonengel, S. (2011) Mitochondrion 11(2), 308-314

23. Fuller, K. M., and Arriaga, E. A. (2003) Curr Opin Biotechnol 14(1), 35-41

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25. Cavelier, L., Johannisson, A., and Gyllensten, U. (2000) Exp Cell Res 259(1), 79-85

26. Aldridge, B. A., Lim, S. D., Baumann, A. K., Hosseini, S., Buck, W., Almekinder, T. L., Sun, C. Q., and Petros, J. A. (2003) Biotechniques 35(3), 606-607, 609-610, 612

27. Kraytsberg, Y., Bodyak, N., Myerow, S., Nicholas, A., Ebralidze, K., and Khrapko, K. (2009) Methods Mol Biol 554, 315-327

28. Reiner, J. E., Kishore, R. B., Levin, B. C., Albanetti, T., Boire, N., Knipe, A., Helmerson, K., and Deckman, K. H. (2010) PLoS One 5(12), e14359

Appendix A

The Effect of GCN5L1 Knockdown in Hep3B and HIB1B Cells.

GCN5L1, (general control of amino acid synthesis 5)-like 1, was recently

identified as an essential molecular component of the mitochondrial acetyltransferase

complex in HepG2 cells, human liver hepatocellular carcinoma cell line (1). It was

demonstrated to reverse the effects of SIRT3 on mitochondrial protein acetylation (1).

To examine the possible effect of GCN5L1 on mitochondrial protein synthesis,

we performed siRNA-mediated knockdown of GCN5L1 in Hep3B cells. We employed

Hep3B cells, human hepatoma cell lines, because of the high acetylation state of proteins,

which was expected to display significant reduction in protein acetylation with GCN5L1

knockdown. Whole cell lysate (WCL) and mitochondrial lysate (ML) prepared from

Hep3B cell lines transfected with control siRNA (Con) and GCN5L1 siRNA (siG) were

analyzed with immunoblotting assays to examine protein acetylation with anti-acetyl Lys

antibody, GCN5L1 knockdown with anti-GCN5L1 antibody, two forms of MRPL12 with

anti-MRPL12 antibody, and steady-state levels of OXPHOS subunits with MitoProfile®

total OXPHOS antibody cocktail (Fig. A1.1). Protein acetylation levels did not present

much difference between control and GCN5L1 knockdown samples. This might be due

to the growth condition of the cells since cells grown in rich media and under starvation

conditions might demonstrate different levels of protein acetylation. One of the

observations in GCN5L1 knockdown cells was their slower growth rate compared to the

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control cells; however, this needs to be confirmed by employing cell proliferation assays

with WST-1. On the other hand, when these experiments were repeated with anti-acetyl

Lys antibody, the results were inconclusive due to the inconsistent performance of the

antibody. As a result of similar protein acetylation levels in Hep3B samples,

mitochondrial protein synthesis did not display difference in steady state levels of

OXPHOS subunits, especially mitochondrially-encoded Complex IV subunit (Fig. A1.1).

Anti-GCN5L1 antibody did not recognize the acetyltransferase subunit in whole cell

lysate but in mitochondrial lysate it demonstrated a reduction in GCN5L1. It is possible

that the amount of GCN5L1 in knockdown cells was enough to maintain the acetylation

state of the proteins. In addition, immunoblotting analysis of the mitochondrial lysate

from GCN5L1 knockdown cells revealed the decrease in the shorter form of MRPL12

(Fig. A1.1). This observation needs to be confirmed by immunoblotting analysis of

enriched ribosomal fractions prepared from the GCN5L1 knockdown cells with anti-

MRPL12, anti-MRPL10, and anti-acetyl Lys antibodies.

One of the limitations in these experiments was the amount of sample required for

analyses. Mitochondrial lysates or enriched ribosomal fractions need to be prepared by

using cells from two 6-well plates for each sample set (Con vs siG) since the sample yield

for downstream analysis would not be enough for more than a couple of immunoblotting

assays.

Pulse labeling of mitochondrial translation products in Hep3B with GCN5L1

knockdown did not reveal a conclusive result due to the inefficient labeling of translation

products with [35S]-methionine (Fig. A1.2). On the other hand, pulse labeling assay with

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HIB1B cells (CON) and HIB1B cells overexpressing SIRT3 (SIRT3) with GCN5L1

knockdown did not demonstrate a significant effect on the mitochondrial translation

when compared GCN5L1 knockdown sample to control sample (Con) (Fig. A1.2).

However, there was a significant difference between control HIB1B transfected with

control siRNA and SIRT3 overexpressing HIB1B transfected with GCN5L1 siRNA. This

is probably due to the concerted activity of overexpressed SIRT3 and GCN5L1

knockdown, reducing the synthesis of mitochondrially-encoded proteins (Fig. A1.2).

These experiments need to be repeated to confirm the effect of GCN5L1 on

mitochondrial protein synthesis.

Overall, the efficient knockdown of GCN5L1 in cell lines and its effect on

mitochondrial translation activity might provide a more complete picture of the

regulation of ribosomal activity in mammalian mitochondria. The modulation MRPL12

stalk composition on mitochondrial ribosomes in respond to changes in expression and

activity of GCN5L1 might confirm the importance of MRPL10 acetylation and its role in

mitochondrial protein synthesis.

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Figure A1.1: The effect of GCN5L1 knockdown on acetylation and OXPHOS

subunits in Hep3B cells.

Immunoblotting analyses of whole cell lysates (WCL) and mitochondrial lysates

(ML) prepared from control siRNA (Con) and GCN5L1 specific siRNA (siG) transfected

Hep3B cells was performed to examine the protein acetylation, steady-state levels of

nuclear encoded subunits of Complex V, III, and II and mitochondrial encoded subunit of

Complex IV, COII, and MRPL12. GCN5L1 was not detected in WCL, while it was

reduced in ML. The same immunoblot was used to probe with MRPS29, MRPL47, and

HSP60 to ensure equal loading.

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Figure A1.2: The effect of GCN5L1 knockdown on mitochondrial protein synthesis

in Hep3B cells.

De novo synthesis of mitochondrial proteins was evaluated in control siRNA

(Con) vs GCN5L1 siRNA (siG) transfected Hep3B and HIB1B control (CON) and

SIRT3 overexpressing (SIRT3) cells by pulse labeling of proteins in the presence of

[35S]-methionine and a cytosolic translation inhibitor, emetine. Coomassie Blue staining

of the same gel was performed to ensure equal protein loading in the gel. The combined

intensities of 13 mitochondrially-encoded proteins from each lane were used as an overall

quantitation of mitochondrial protein synthesis.

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References:

1. Scott, I., Webster, B. R., Li, J. H., and Sack, M. N. (2012) Biochem J 443(3), 655-661

Appendix B

Strong-Cation Exchange (SCX) Chromatography Purification of Mitochondrial

MRPL10-MRPL12 Stalk Complex

The importance of Lys124, Lys162, and Lys196 in the interaction between

MRPL10 and MRPL12 was examined by employing site-directed mutagenesis by

generating LysAla in addition to LysArg and LysGln to mimic hypoacetylation

and hyperacetylation, respectively, as described in Materials and Methods of Chapter 2.

In our purification approach with nickel-nitrilotriacetic acid (Ni-NTA) affinity

chromatography followed by strong-cation exchange (SCX) chromatography (Fig. 2.24),

the impurities and improperly folded MRPL10-MRPL12 complexes were possibly the

factors affecting the activity of reconstituted ribosomes. We attempted to improve the

purification of MRPL10-MRPL12 complex to examine the effects of the acetylated

lysine residues on MRPL12 binding and ribosomal activity (Fig. A2.1). Ni-NTA affinity

column purification of MRPL10-MRPL12 was performed by including 5 mM of ATP

and 2 mM MgCl2 in the buffers used to prepare bacterial cell lysate. This reduced the

association of contaminating proteins, such as groEL, HSP70, and EF-P with MRPL10-

MRPL12 complex (1,2). In addition, the buffer component of the solutions used in Ni-

NTA affinity chromatography purification was replaced with Tris-HCl (pH 7.6) (see Ini

in Fig. A2.1). The Ni-NTA column purified samples were then applied to the SCX gravity

column and eluted by using the conditions given in Materials and Methods section in

216

Chapter 2 (Step gradient salt of KCl; a linear gradient 50 mM –150 mM for 30 min, 400

mM – 500 mM for 10 min, and 500 mM – 1 M for 10 min). Fractions were monitored by

UV detection at 260 nm and then analyzed by SDS-polyacrylamide gel electrophoresis.

The fractions containing the MRPL10-MRPL12 complex (labeled with * in Fig. A2.1)

were collected, combined, and dialyzed for further analysis in activity assays (Fig. A2.2).

SDS-polyacrylamide gel analysis of dialyzed samples displayed highly purified

MRPL10-MRPL12 complex to be used in poly(U)-directed poly(Phe) synthesis assays to

examine the effect of acetylated Lys residues and their mutants in the activity of hybrid

ribosomes (Fig. A2.2). The experiments involving the activity assays with the purified

MRPL10-MRPL12 complex are currently under progress.

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Figure A2.1: Strong-cation exchange (SCX) chromatography purification of

mitochondrial MRPL10-MRPL12 stalk complex.

Ni-NTA chromatography-purified wild type and mutant MRPL10-MRPL12

complexes (Ini) were applied on SCX column to remove contaminants (FT: flow-

through, 1: first fraction) where fractions were monitored with a UV detector at 260 nm

to trace the proteins during application of step gradient of salt, KCl (dashed line). In the

elution profile, stalk complexes containing MRPL10 wild type (WT) were labeled with

blue solid line and lysine mutants, Ala (A), Gln (Q), Arg (R) were marked in black, red,

and green, respectively. Fractions were analyzed on 14 % SDS-polyacrylamide gels to

locate MRPL10-MRPL12 complexes. *: Fractions were combined and dialyzed to be

used in activity assays.

218

References:

1. Thain, A., Gaston, K., Jenkins, O., and Clarke, A. R. (1996) Trends Genet 12(6), 209-210

2. Han, M. J., Cimen, H., Miller-Lee, J. L., Koc, H., and Koc, E. C. (2011) Protein Expr Purif 78(1), 48-54

Figure A2.2: Dialyzed mitochondrial MRPL10-MRPL12 stalk complex.

The fractions containing MRPL10-MRPL12 stalk complex from SCX column

were combined and dialyzed against a dialysis buffer containing 50 mM Tris-HCl, pH

7.6, 200 mM KCl, 1 mM DTT, and 10 % glycerol for a total of 4 h at 4oC to reduce the

KCl amount. These samples will be employed to determine the effect of acetylated Lys

residues and their mutants in the activity of hybrid ribosomes in poly(U)-directed

poly(Phe) synthesis assays.

VITA: HUSEYIN CIMEN

EDUCATION PhD Pennsylvania State University, Biochemistry and Molecular Biology. August 2012

Advisor: Assist. Prof. Emine C. Koc, Co-advisor: Assist. Prof. Hasan Koc BS Middle East Technical University, Molecular Biology and Genetics, May 2006

Advisor: Prof. Dr. Feride Severcan PUBLICATIONS 2012 _Koc EC*, Cimen H*, Kumcuoglu B, Abu N, Akpinar G, Haque ME, Spremulli LL, Koc H.

Identification and Characterization of CHCHD1, AURKAIP1, and CRIF1 as New Members of the Mammalian Mitochondrial Ribosome. Nucleic Acids Res. (Manuscript Submitted). (*: contributed equally).

_ Zhu JH, Gusdon AM, Cimen H, Van Houten B, Koc E, Chu CT. Impaired Mitochondrial Biogenesis Contributes to Depletion of Functional Mitochondria in Chronic MPP+ Toxicity: Dual Roles for ERK1/2. Cell Death Dis. 2012 May 24;3:e312. doi: 10.1038/cddis.2012.46.

2011 _Haque ME, Koc H, Cimen H, Koc EC, Spremulli LL. Contacts between mammalian mitochondrial translational initiation factor 3 and ribosomal proteins in the small subunit. Biochim Biophys Acta. 1814(12), 1779-84

_Surovtseva YV, Shutt TE, Cotney J, Cimen H, Chen SY, Koc EC, Shadel GS. Mitochondrial ribosomal protein L12 selectively associates with human mitochondrial RNA polymerase to activate transcription. Proc Natl Acad Sci USA. 108(44),17921-6

_Han MJ, Cimen H, Miller-Lee JL, Koc H, Koc EC. Purification of human mitochondrial ribosomal L7/L12 stalk proteins and reconstitution of functional hybrid ribosomes in Escherichia coli. Protein Expr Purif. 78(1), 48-54.

2010 _Yang Y*, Cimen H*, Han MJ*, Shi T, Deng JH, Koc H, Palacios OM, Montier L, Bai Y, Tong Q, Koc EC. NAD+-dependent deacetylase SIRT3 regulates mitochondrial protein synthesis by deacetylation of the ribosomal protein MRPL10. J Biol Chem. 285(10), 7417-29. (*: contributed equally)

_Cimen H, Han MJ, Yang Y, Tong Q, Koc H, Koc EC. Regulation of succinate dehydrogenase activity by SIRT3 in mammalian mitochondria. Biochemistry. 49(2), 304-11.

2009 _Miller JL, Cimen H, Koc H, Koc EC. Phosphorylated proteins of the mammalian mitochondrial ribosome: implications in protein synthesis. J Proteome Res. 8(10), 4789-98.

ABSTRACTS AND PRESENTATIONS2012 _Cimen H, Koc H, Koc EC. A newly identified protein regulates translation in mammalian

mitochondria. Experimental Biology 2012 meeting, San Diego, CA 2011 _Cimen H, Han MJ, Tong Q, Koc H, Koc EC. SIRT3 regulates mitochondrial translation by

modulating MRPL12 binding to ribosome. Experimental Biology 2011 meeting, Washington, DC 2009 _Cimen H, Yang Y, Shi T, Han MJ, Deng JH, Koc H, Palacios OM, Montier L, Bai Y, Tong Q, Koc

EC. NAD+-dependent deacetylase SIRT3, regulates mitochondrial protein synthesis by deacetylation of the ribosomal protein, MRPL10. Mitochondrial Medicine 2009, Vienna, VA (Selected for an oral presentation and received Third-Place Runner Cash Award, presented by Dr. Emine C. Koc)

2008 _Cimen H, Yang Y, Shi T, Han MJ, Deng JH, Koc H, Palacios OM, Montier L, Bai Y, Tong Q, Koc EC. Regulation of mitochondrial protein synthesis by a mitochondrial NAD+-dependent deacetylase SIRT3. Translational Control Meeting, Cold Spring Harbor, NY

FELLOWSHIPS, SCHOLARSHIPS, AND AWARDSApril 2012 2012 ASBMB Annual Meeting Grad/ Postdoc Travel Award Summer 2011 Braucher Pela Fay Scholarship, Dept. of Biochemistry and Molecular Biology,

Pennsylvania State University April 2011 2011 ASBMB Annual Meeting Grad/ Postdoc Travel Award 2007-2008 Homer F. Braddock Fellowship, Dept. of Biochemistry and Molecular Biology,

Pennsylvania State University