Protein Carbonylation E-Book

• Herausgeber: John Wiley & Sons
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 777

Veröffentlichungsjahr: 2017

Beschreibung

Protein carbonylation has attracted the interest of a great number of laboratories since the pioneering studies at the Earl Stadtman's lab at NIH started in early 1980s. Since then, detecting protein carbonyls in oxidative stress situations became a highly efficient tool to uncover biomarkers of oxidative damage in normal and altered cell physiology. In this book, research groups from several areas of interest have contributed to update the knowledge regarding detection, analyses and identification of carbonylated proteins and the sites where these modifications occur. The scientific community will benefit from these reviews since they deal with specific, detailed technical approaches to study formation and detection of protein carbonyls. Moreover, the biological impact of such modifications in metabolic, physiologic and structural functions and, how these alterations can help understanding the downstream effects on cell function are discussed. * Oxidative stress occurs in all living organisms and affects proteins and other macromolecules: Protein carbonylation is a measure of oxidative stress in biological systems * Mass spectrometry, fluorescent labelling, antibody based detection, biotinylated protein selection and other methods for detecting protein carbonyls and modification sites in proteins are described * Aging, neurodegenerative diseases, obstructive pulmonary diseases, malaria, cigarette smoke, adipose tissue and its relationship with protein carbonylation * Direct oxidation, glycoxidation and modifications by lipid peroxidation products as protein carbonylation pathways * Emerging methods for characterizing carbonylated protein networks and affected metabolic pathways

Leseprobe

Table of Contents

Cover

Title Page

List of Contributors

Preface

1 Reactive Oxygen Species Signaling from the Perspective of the Stem Cell

1.1 Introduction

1.2 ROS Regulation

1.3 ROS Signaling

1.4 ROS and Stem Cells

1.5 ROS, Metabolism, and Epigenetic Influence

1.6 Stem Cells and Mitochondria

1.7 ROS and Stem Cell Aging

1.8 Concluding Remarks

References

2 Analysis of Protein Carbonylation

2.1 Introduction

2.2

In Vivo

Carbonylation Reactions

2.3 Analytical Derivatization of Carbonylated Groups

2.4 Selective Purification and/or Detection of Carbonylated Proteins and Peptides

2.5 Oxidative Stress‐Based PTMS Not Involving Carbonylation

2.6 Conclusion

References

3 Diversity of Protein Carbonylation Pathways

3.1 Introduction

3.2 Pathways of Protein Carbonylation

3.3 Analytical Methods for Detection of Total and Specific Protein Carbonylation

3.4 Protein Susceptibility to Different Carbonylation Pathways and Modifications Cross‐Talk

3.5 Conclusion

Acknowledgments

References

4 Protein Carbonylation by Reactive Lipids

4.1 Introduction

4.2 Chemistry of Protein Carbonylation by Reactive Lipid Aldehydes

4.3 Antigenicity of Protein Carbonyls

4.4 Thiolation of Protein Carbonyls

4.5 Reductive Amination‐Based Fluorescent Labeling of Protein Carbonyls

4.6 Conclusion

References

5 Mechanism and Functions of Protein Decarbonylation

5.1 Protein Carbonylation

5.2 Primary Protein Carbonylation in Cell Signaling

5.3 Discovery and Mechanisms of Protein Decarbonylation

5.4 Proposed Functions of Protein Decarbonylation in Oxidative Stress and Redox Signaling

Acknowledgments

References

6 Carbonylated Proteins and Their Metabolic Regulation

6.1 Metabolic Regulation and Reactive Oxygen Species

6.2 ROS and Protein Carbonylation

6.3 Metabolic Control and Characteristics of Carbonylated Proteins

6.4 Protein Targets of Carbonylation and Implications in Human Health

6.5 Technologies and Methods for Characterizing Protein Carbonylation

6.6 Emerging Multifunctional Reagents for Protein Carbonylation Analysis via MS

6.7 Emerging Methods for Characterizing Carbonylated Protein Networks and Affected Pathways

6.8 Conclusion

References

7 Oxidative Stress and Protein Carbonylation in Malaria

7.1 Introduction

7.2 Oxidative Stress during Malaria Infection

7.3 Protein Carbonylation in

Plasmodium

and Oxidative Targeting of Antimalarials

7.4 Oxidative Dysfunction in Host Tissues

7.5 Host Tolerance to Malaria by Modulation of Oxidative Stress Responses

7.6 Perspectives

References

8 Protein Carbonylation in Brains of Subjects with Selected Neurodegenerative Disorders

8.1 Introduction to Protein Carbonylation

8.2 Relationship between ROS and Oxidative Stress

8.3 An Overview of Some Neurodegenerative Diseases

8.4 Role of Protein Carbonylation in Brains of Subjects with AD

8.5 An Introduction to Tauopathies

8.6 An Introduction to Amyotrophic Lateral Sclerosis

8.7 Discussion

References

9 Cigarette Smoke‐Induced Protein Carbonylation

9.1 Introduction

9.2 Protein Carbonylation in Human Smokers

9.3 Protein Carbonylation in Cultured Human Cell Models of Exposure to CS

9.4 Limitations and Congruence of

In Vivo

and

In Vitro

Human Studies

9.5 Conclusion and Future Perspectives

Acknowledgments

References

10 Chronic Obstructive Pulmonary Disease and Oxidative Damage

10.1 Introduction

10.2 Protein Oxidation in Tissues

10.3 Antioxidants in Skeletal Muscle Fibers

10.4 Implications of Protein Carbonylation in COPD Skeletal Muscle Dysfunction

10.5 Muscle Protein Carbonylation and Exercise in COPD Patients

10.6 Protein Carbonylation in Muscles Exposed to Chronic Cigarette Smoke

10.7 Protein Carbonylation in Cancer Cachexia Models

10.8 Protein Carbonylation as a Predisposing Mechanism of Lung Cancer in COPD

10.9 Conclusion and Future Perspectives

Acknowledgments

References

11 Protein Carbonylation in Aging and Senescence

11.1 Introduction

11.2 Protein Oxidation during Aging

11.3 Chemistry of Protein Carbonylation and Fate of Carbonylated Proteins

11.4 Protein Carbonyls in Cellular Aging Models

11.5 Protein Carbonylation in Aging Organisms

11.6 Concluding Remarks

References

12 Adipose Carbonylation and Mitochondrial Dysfunction

12.1 Introduction

12.2 Reactive Oxygen Species (ROS)

12.3 Oxidative Stress and Obesity

12.4 Detection of Protein Carbonylation

12.5 Outcomes of Protein Carbonylation

12.6 Concluding Remarks

Acknowledgments

References

13 Protein Carbonylation in Plants

13.1 Introduction

13.2 Turnover of Reactive Oxygen Species in Plants

13.3 Methods Used in Plants for Quantifying and Identifying Carbonylation Sites

13.4 Protein Carbonylation in Plants

13.5 Protein Carbonylation in Plant Mitochondria

13.6 Protein Carbonylation in Seeds

13.7 Perspectives

Acknowledgments

References

14 Specificity of Protein Carbonylation and Its Relevance in Aging

14.1 Introduction

14.2 Specificity of Protein Oxidative Damage

14.3 Protein Carbonylation in Aging

14.4 Concluding Remarks

Acknowledgments

References

Index

Series Editors

End User License Agreement

List of Tables

Chapter 03

Table 3.1 Overview of main analytical methods used for detection of carbonylated proteins.

Chapter 06

Table 6.1 Examples of proteins affected by carbonylation and their disease association.

Table 6.2 Some “multifunctional” reagents for protein carbonylation detection.

Chapter 07

Table 7.1 Carbonylated

Plasmodium falciparum

proteins at erythrocytic stages.

Table 7.2 Differentially carbonylated human erythrocyte proteins.

Chapter 08

Table 8.1 Brain proteins identified to be carbonylated during the progression of Alzheimer disease.

Chapter 09

Table 9.1 PCO in human smokers detected by different methods.

Table 9.2 PCO in human cell models of exposure to CS detected by different methods.

Chapter 12

Table 12.1 Proteomic studies of carbonylated targets in adipose tissue.

Chapter 13

Table 13.1 Comparison of mitochondrial proteins with HNE modification, carbonylation, tryptophan oxidation, and oxidative stress‐induced breakdown.

Chapter 14

Table 14.1 Nucleotide‐binding proteins represent a high percentage among carbonylated targets identified in aging, diseases, and other oxidative stress situations.

List of Illustrations

Chapter 01

Figure 1.1 Redox sensors critical for stem cell fate: ROS regulation of signaling molecules and transcription factors and their effect on ROS regulation.

Chapter 02

Figure 2.1 Forms of methionine and cysteine involved in regulation. The presence of enzymes in a biological system that reduce oxidized forms of cysteine and methionine is an important component of regulation.

Figure 2.2 Modes of acquisition and production of reactive oxygen species in biological systems.

Figure 2.3 Free radical mediated cleavage of the polypeptide backbone. Panel (a) illustrates the two major pathways by which proteins are cleaved following α‐carbon initiation of oxidation. Pathway (a) occurs via an α‐amidation route, while that in (b) transpires with diamide formation. Panel (b) shows that the α‐carbon initiated cleavage mechanism can also be directed by the structure of the amino acid side chain in the case of proline. Carbonyl group formation in these reactions is highlighted with circles. Panel (c) illustrates backbone cleavage at prolyl residues. An important lesson in this figure is that multiple reaction products are formed that lack distinguishable features, such as a carbonyl group.

Figure 2.4 Amino acid side chain oxidation. Lysine, arginine, and proline oxidation in Panels (a), (b), and (d), respectively, is initiated adjacent to nitrogen in the side chain of these amino acids. Note that glutamine semialdehyde is the end product of both arginine and proline oxidation, making product origin impossible to identify directly. Carbonyl groups formed during oxidation are highlighted by circles. Threonine oxidation also occurs in the side chain but without free radical initiation (Panel (c)). Proline oxidation in Panel (d) fails to produce a carbonyl group.

Figure 2.5 Formation and Michael addition of 4‐hydroxy‐2‐nonenal (4‐HNE) to amino acid residues in proteins. 4‐HNE is potentially formed from PUFAs through multiple routes; one being illustrated in Panel (a). Panel (b) illustrates the structure of products formed in the addition of 4‐HNE to amino acid residues in proteins. Note that the carbonyl group of 4‐HNE (circle) is retained in the addition products.

Figure 2.6 Protein glycation with an example of subsequent oxidation and protein cross‐linking.

Figure 2.7 Carbonyl derivatization through Schiff base formation and reduction.

Figure 2.8 Derivatizing agents.

Figure 2.9 Reversed phase liquid chromatography of anthranilamide labeled peptides derived from

in vitro

oxidized hemoglobin. Subsequent to

in vitro

hydrogen peroxide oxidation of hemoglobin, the sample was labeled with anthranilamide, reduced with NaCNBH

4

, and trypsin digested before chromatographic analysis of the digest.

Figure 2.10 Amino acids modifications encountered in proteins that are not selected via derivatization.

Chapter 03

Figure 3.1 Overview of three main protein carbonylation pathways. Metal‐catalyzed direct oxidation of Lys, Arg, Pro, Thr, and Trp residues (a); lipid peroxidation product (LPP)‐mediated Michael addition on Lys, Cys, and His residues (b); and reactions of sugar oxidation products with Lys, Arg, Cys, and His residues (c).

Figure 3.2 Schematic representation of ELISA assay for quantification of total protein content. Carbonylated proteins are immobilized on ELISA plate and derivatized with DNPH. After subsequent washing steps to remove unbound DNPH, derivatized proteins probed with anti‐DNP antibodies followed by incubation with secondary antibodies labeled with horseradish peroxidase (POD). Addition of specific substrate initiate POD‐catalyzed reaction and signal proportional to the protein carbonyl content can be recorded.

Figure 3.3 Schematic representation of mass spectrometry‐based identification of carbonylated proteins including modification sites and types. (a) Complex biological samples containing carbonylated proteins separated by liquid chromatography or SDS‐PAGE and protein fractions digested with trypsin. Carbonyl‐containing peptides derivatized with carbonyl‐specific tags and used for enrichment. (b) For mass spectrometry‐based identification, mixture of enriched carbonylated peptides is ionized (e.g., ESI) and analyzed by the first mass analyzer. Selected precursor ions can be fragmented using collision‐induced dissociation and resultant fragments detected by the second mass analyzer. Assignment of recorded signals allows identification of peptide sequence, modification type, and site.

Figure 3.4 Possible cross‐talks between protein carbonylation and other regulatory PTMs. “Carbonylatable” amino acid residues such as Cys, Lys, and Thr are substrates for several other PTMs including redox signaling via reversible Cys oxidation, acetylation (Ac), methylation (Me), and ubiquitination (Ub) of Lys as well as Thr phosphorylation‐mediated signaling.

Chapter 04

Figure 4.1 Lipid peroxidation‐derived aldehydes. (a) Key structural moieties of lipid peroxidation‐derived aldehydes. R: alkyl chains. (b) Structures of representative lipid peroxidation‐derived aldehydes.

Figure 4.2 The structures (

1–6

) of adducts with carbonyl functionality.

Figure 4.3 Thiolation of FDP‐lysine.

Figure 4.4 Reductive amination‐based fluorescent labeling of HNE–cysteine adducts. (a) A procedure for pyridylamination of HNE–cysteine adducts. (b) HPLC analysis of the pyridylaminated

R

‐HNE‐ and

S

‐HNE‐cysteine adducts.

Chapter 05

Figure 5.1 Endothelin‐1 (ET‐1) promotes protein carbonylation. Cultured bovine pulmonary artery smooth muscle cells were treated with ET‐1 (30 nM) for 0, 5, or 10 min. Cell lysates were prepared and derivatized with DNPH. Carbonylated protein levels were monitored by Western blot with the DNP antibody. (a) Representative Western blot is shown. (b) The bar graph shows means ± SEM (

n

 = 6) of percent of total carbonyl content relative to untreated control as determined by densitometry. The symbol a denotes the value that is significantly different from untreated control at 0 min at

p

 < 0.05. Reprinted by permission from Wong et al. [25].

Figure 5.2 H

2

O

2

promotes protein carbonylation. Cultured bovine pulmonary artery smooth muscle cells were treated with H

2

O

2

(0.5 μM) for 0, 5, or 10 min. Cell lysates were prepared and derivatized with DNPH. Carbonylated protein levels were monitored by Western blot with the DNP antibody. Reprinted by permission from Wong et al. [25].

Figure 5.3 Endothelin‐1 (ET‐1) promotes carbonylation and subsequent degradation of annexin A1. (a) Cultured bovine pulmonary artery smooth muscle cells were treated with ET‐1 (30 nmol/L) for 10 min. Cell lysates were prepared, derivatized with DNPH, immunoprecipitated with the DNP antibody, and subjected to Western blotting with the annexin A1 antibody. (b) Cells were pretreated with MG132 (30 µmol/L) for 30 min before ET‐1 treatment for 30 min. Annexin A1 protein levels were determined by Western blotting. Reprinted by permission from Wong et al. [25].

Figure 5.4 Proposed model of the role of annexin A1 carbonylation in the regulation of cell growth and survival. Growth factors promote the production of H

2

O

2

, leading to metal‐catalyzed oxidation and protein carbonylation of annexin A1. Carbonylated annexin A1 is degraded by the proteasome. Since the function of annexin A1 is to decrease cell growth and promote cell death, the degradation of this protein results in cell growth and cell survival.

Figure 5.5 Endothelin‐1 (ET‐1) promotes protein carbonylation and subsequent decarbonylation. Cultured bovine pulmonary artery smooth muscle cells were treated with ET‐1 (30 nM) for 0–30 min. Cell lysates were prepared and derivatized with DNPH. Carbonylated protein levels were monitored by Western blot with the DNP antibody. (a) Representative Western blot is shown. (b) The bar graph shows means ± SEM (

n

 = 6) of percent of total carbonyl content relative to untreated control as determined by densitometry. Symbols a and b denote values that are significantly different from untreated control and the value from cells treated with endothelin‐1 for 10 min, respectively, at

p

 < 0.05. Reprinted by permission from Wong et al. [25]. Carbonylation and decarbonylation phases are indicated by arrows.

Figure 5.6 Effects of siRNA knockdown of Trx and Grx1 on PDGF‐induced carbonylation and decarbonylation in cultured cells. Cultured human pulmonary artery smooth muscle cells were transfected with (a) Trx or (b) Grx1 siRNA for 2 days, followed by PDGF stimulation for 10 or 30 min. Cell lysates were prepared, derivatized with DNPH, immunoprecipitated with the DNP antibody, and then immunoblotted with the peroxiredoxin‐6 (Prx6) antibody. Bar graphs represent means ± SEM of the ratio of carbonylated Prx6 to Prx6 protein level expressed in arbitrary unit (a.u.) (

n

 = 6 for (a);

n

 = 4 for (b)). The symbol * denotes that the values are significantly different from each other at

p

 < 0.05. The symbol ns denotes that the two values are not significantly different. Reprinted by permission from Wong et al. [29].

Figure 5.7 Scheme depicting the role of Grx1 in protein decarbonylation. A protein molecule depicted by a circle is carbonylated (=O) and reversibly decarbonylated. Grx1 appears to be involved in the decarbonylation reaction.

Figure 5.8 A model in which carbonylated peroxiredoxin (Prx) plays a role in cell signaling. Ligand–receptor interactions activate Nox, which in turn produces ROS including H

2

O

2

, leading to metal‐catalyzed carbonylation of Prx (in particular Prx2 and Prx6). This carbonylation inhibits the activity of Prx to eliminate H

2

O

2

, leading to enhanced H

2

O

2

levels.

Figure 5.9 Iron‐catalyzed oxidations of arginine and proline residues, both of which result in the formation of glutamyl semialdehyde (with a carbonyl group) as proposed by Amici et al. [15].

Figure 5.10 Proposed arginine–proline conversion in the protein structure that may occur in the biological system.

Figure 5.11 Arginine–glutamic acid conversion and proline–glutamic acid conversion in the protein structure.

Chapter 06

Figure 6.1 Lipid peroxidation of α,β‐unsaturated fatty acids resulting in products such as acrolein, malondialdehyde, 2‐hexenal, 4‐hydroxy‐trans‐2,3‐nonenal (4‐HNE), and 4‐oxo‐trans‐2,3‐nonenal (4‐ONE).

Figure 6.2 Metal‐catalyzed oxidation (MCO) using ions such as Fe

3+

as a mechanism for protein carbonylation.

Figure 6.3 Metabolic control of reactive carbonyl‐containing compounds.

Figure 6.4 Structure of aminoxyTMT reagent.

Figure 6.5 Immunoblot detection (anti‐TMT) of liver mitochondria treated with HNE and labeled with aminoxyTMT.

Figure 6.6 MS/MS spectra of a BSA peptide modified with HNE at Cys and labeled with aminoxyTMT.

Figure 6.7 Extracting quantitative information on a peptide of interest from DIA MS data. DIA collects complex spectra of fragments from multiple peptides. In the example, data from fragments (labeled 1, 2, and 3) from a peptide of interest (called “peptide A”) are extracted. An ion assay library is used that contains the expected fragment ion information for any given peptide of interest. The overlapping chromatographic peaks from the extracted fragment ions (1, 2, and 3 in this example) are reconstructed, and area under the curve (AUC) is measured for quantitative measurements between different samples or conditions of interest.

Chapter 07

Figure 7.1 Life cycle of

Plasmodium

species infecting humans. Malaria infection starts by the

Plasmodium

sporozoites injected by a feeding mosquito (an

Anopheles

spp. female). Sporozoites migrate to liver hepatocytes where, asymptomatically, they divide and mature into merozoites. The symptoms start with the blood‐stage sub‐cycle upon erythrocytes invasion by the liver‐released merozoites. Merozoites grow up and develop in erythrocytes through ring, trophozoite, and schizont stages to finally multiply into new merozoites. Daughter merozoites released invade other erythrocytes to continue the intraerythrocytic cycle. During infection, this asexual blood‐stage sub‐cycle is responsible for the clinical pathologies of malaria. Some intraerythrocytic parasites develop into gametocytes, which can be ingested by a mosquito during a new feeding. Sexual reproduction of the parasites in the mosquito gut leads to the generation of new sporozoites ready to infect a new human host. (Taken from Ref. [14]).

Figure 7.2 Antioxidant defense system in

Plasmodium

during the intraerythrocytic cycle. The major source of reactive oxygen species (ROS) is the breakdown of host hemoglobin in the parasite food vacuole. Although most of the released heme (FP IX) is biocrystallized to hemozoin, some of it reaches the cytoplasm, generating superoxide anions, which are detoxified mainly by cytosolic Fe superoxide dismutase to yield H

2

O

2

. Due to the lack of catalase in

Plasmodium

, reduction of H

2

O

2

to H

2

O is achieved by the thioredoxin or GSH redox cycles. The dual enzyme glucose‐6‐phosphate dehydrogenase/6‐phosphogluconolactonase supplies NADPH to both redox cycles.

De novo

GSH is readily provided by γGlu‐Cys synthase. In

Plasmodium

, the thioredoxin and GSH redox cycles are linked via plasmoredoxin, which is also used as substrate thioredoxin, providing flexibility to the system in enhanced oxidative stress. Reduced GSH takes part in detoxifying heme, a product of hemoglobin digestion. As for other eukaryotic organisms, the parasite is capable of transporting GSSG, GSH conjugates, and GSH in the presence of drugs via multidrug resistance transporters (MRP).

Figure 7.3 The redox proteome of the malaria parasite and its host. Upper panel: redox proteome patterns across the intraerythrocytic stages of

Plasmodium falciparum

modified from an original figure in Ref. [84]. Carbonyl immunoblots for synchronized cultures of rings, trophozoites, and schizonts show a relative constant number of spots across the stages (between 122 and 141). Spot intensities diminished from the ring and trophozoite stages to the schizont stage. A large proportion of the carbonylated spots are matched across the cycle, with more than 50% of the spots detected for a given stage also seen in extracts corresponding to other stages. Lower panel: 2D‐PAGE and corresponding oxyblots of membrane proteins from

P. falciparum

‐infected G6PD B (normal genotype) and G6PD A

−

(deficient phenotype) erythrocytes modified from an original figure in [89]. Upon malaria infection, 22 and 49 carbonylated spots are respectively detected in G6PD normal (G6PD B) and G6PD‐deficient (G6PD A

−

) erythrocytes. Carbonylated protein spots from the membrane fraction of the infected G6PD A

−

erythrocytes correspond to 15 different proteins that are only observed in this condition. Among these, five are free hemichromes and three hemichromes bound as complexes to other oxidative stress response enzymes. Labeled with a square are the new hemichromes generated upon the increased oxidative stress in the infected erythrocytes from deficient G6PD A

−

subjects. Six other proteins associated to stress response were also exclusively found carbonylated in infected G6PD A

−

erythrocytes.

Chapter 08

Figure 8.1 Beta scission from peptide backbone.

Figure 8.2 Amino acid side chain oxidation.

Figure 8.3 Chemical reaction of α‐ketoglutarate and 2,4‐DNPH to form 2,4‐DNP.

Figure 8.4 (a) Disproportionation (dismutation) of superoxide; (b) formation of superoxide anion; (c) oxidation by xanthine oxidase; (d) an example of Fenton chemistry using ferrous iron; and (e) formation of hydroxyl radical from hydrogen peroxide and superoxide anion.

Figure 8.5 Pyruvate kinase enzymatic reaction.

Figure 8.6 Catalytic conversion of 2‐phosphoglycerate to phosphoenolpyruvate by enolase.

Figure 8.7 Enzymatic reaction of GAPDH.

Figure 8.8 Catalytic cleavage of 1,6‐fructose bisphosphate to produce DHAP and G3P.

Figure 8.9 Enzymatic reaction of phosphoglycerate mutase.

Figure 8.10 Triose phosphate isomerase enzymatic reaction.

Figure 8.11 Structures of methionine and its oxidized forms.

Chapter 09

Figure 9.1 The health consequences causally linked to cigarette smoking and exposure to secondhand smoke.

Figure 9.2 Whole cigarette smoke (WCS) exposure apparatus. (a) WCS exposure chamber (British American Tobacco). (b) Schematic of WCS exposure apparatus used in many cultured cell studies. Reprinted with permission from Wang et al. [99].

Figure 9.3 Immunofluorescence analysis of CSE‐induced formation of protein carbonyls in ECV‐304 cells. Protein carbonylation was assessed by an immunocytochemical DNPH assay as described in Section 9.2 in untreated (a) and CSE‐treated ECV‐304 cells (b–d). Immunoreactivity was evident in cells exposed to 2.5% (b), 5% (c), and 10% (d) CSE. Representative microphotographs of three independent experiments are shown. Original magnification: 63×.

Chapter 10

Figure 10.1 Oxidative stress results from the imbalance between the production of oxidants and the effects of antioxidants in favor of the former.

Figure 10.2 ROS that are not scavenged by cellular antioxidants oxidize key cellular structures such as membrane lipids, nuclear DNA, and proteins. Oxidative damage of proteins exerts different effects such as alteration of enzyme activity and DNA binding of transcription factors and may also render the proteins more susceptible to be degraded.

Figure 10.3 Schematic representation of the formation of reactive oxygen species (ROS) derived from molecular oxygen. Nitric oxide is synthesized by nitric oxide synthases. Peroxynitrite develops from the near‐diffusion‐limited reaction between nitric oxide and superoxide anion. Nitrogen dioxide develops from the reaction between nitric oxide and molecular oxygen.

Figure 10.4 The different molecular sources contributing to ROS production in skeletal muscle fibers are listed in the scheme.

Figure 10.5 Schematic representation of the molecular targets of reactive oxygen and nitrogen species (ROS and RNS, respectively) in the skeletal muscle fibers. Nitric oxide synthases (NOS), especially the inducible isoform (iNOS), is an important contributor to muscle dysfunction, through the formation of high levels of nitric oxide (NO·). NO· reacts with superoxide anion (O

2−

) to form the highly reactive species peroxynitrite (ONOO

−

). Peroxynitrite may directly oxidize proteins (via oxidative stress), or it may also modify aromatic amino acids such as tyrosine, thus leading to the formation of 3‐nitrotyrosine and nitrosative stress. High levels of ROS and RNS selectively target different structures within the myofibers such as the ryanodine receptors; sarcoplasmic reticulum, especially Ca

2+

‐ATPase; plasma membrane Ca

2+

‐ATPase; creatine kinase; myosin content; myosin‐ATPase; microtubules; and Ca

2+

sensitivity of actin.

Chapter 11

Figure 11.1 Cellular sites of ROS/RNS production. Any enzymatic reaction where a transfer of electrons occurs is a potential source for the production of ROS as a by‐product. The transfer of electrons from electron donor to electron acceptor might be incomplete. Thus electrons are leaked and react immediately with molecular oxygen that results in the production of ROS. The electron transport chain during the oxidative phosphorylation in the mitochondria is a significant producer of ROS. Also during the peroxisomal β‐oxidation of fatty acids, ROS are formed. Other potential sources are the detoxification reactions mediated by the cytochrome P450 monooxygenase system, which is located in the endoplasmic reticulum. Additionally, the lumen of the endoplasmic reticulum provides an oxidative environment for the formation of disulfide bonds (oxidative protein folding). During this formation the enzymatic‐driven electron transfer from thiol groups to molecular oxygen produces ROS. Furthermore the catalytic reactions of membrane‐bound as well as cytoplasmic proteins such as aldehyde oxidase, xanthine oxidase, and nitric oxide synthase can generate ROS or RNS.

Figure 11.2 Superoxide generation during oxidative phosphorylation. The mitochondrial respiratory chain consists of five protein complexes that are located at the inner mitochondrial membrane. During the transfer of electrons from complex to complex, energy is released. This energy is used for the generation of a proton gradient. During this process protons (H

+

) are transferred from the mitochondrial matrix to the intermembrane space. These protons are used by ATP synthase (complex V) for ATP synthesis. Complexes I and III are the main sites of mitochondrial superoxide (O

2

·−

) formation. CoQ, coenzyme Q10; CytC, cytochrome C.

Figure 11.3 Possible fates of carbonylated proteins. Carbonylated proteins are substrates for the degradation by the mitochondrial Lon protease and the 20S proteasome. If the degradation systems are overwhelmed, carbonylated proteins accumulate and generate cross‐linked protein aggregates such as lipofuscin. These aggregates are not chemically inert; they rather increase cellular ROS formation. Furthermore, they are able to inhibit the 20S proteasome. Therefore, the uptake of protein aggregates into lysosomes acts as a protective mechanism. Previously, protein carbonyls were considered as irreversible damaged protein structures. However, there is some evidence that the direct oxidation of proteins (primary protein carbonyls) is reversible by a decarbonylation mechanism.

Figure 11.4 Summary of the fate and effects of carbonylated proteins. ROS/RNS leads to oxidative stress that is able to damage all cellular macromolecules (proteins, lipids, sugars, nucleic acids). Protein carbonyls arise either by the direct oxidative attack of proteins or by the reaction with oxidized lipids and oxidized or reducing sugars (reactive carbonyl species; RCS). The rapid elimination of protein carbonyls takes place by degradation via the 20S proteasome or the Lon protease. Without degradation protein carbonyls tend to cross‐link leading to the formation of protein aggregates. Depending on the extension of protein aggregate accumulation, apoptosis or necrosis of cells can occur. In most cases protein carbonylation is known to affect the function of proteins. Otherwise also cellular signalling pathways can be modulated. Thus it was shown that HNE‐mediated modification of Kelch‐like ECH‐associated protein 1 (Keap1) activates the nuclear transcription of genes that encode antioxidative enzymes

via

Nrf2 release. Keap1, Kelch‐like ECH‐associated protein 1; Nrf2, nuclear factor (erythroid‐derived 2)‐like 2.

Chapter 12

Figure 12.1 Metabolism of ROS and outcomes of oxidative stress. Superoxide is metabolized via SOD to yield hydrogen peroxide that is subsequently detoxified by PRDX, GPX, or catalase. Alternatively, hydrogen peroxide can undergo Fenton chemistry forming hydroxyl radicals, leading to the production of reactive lipid aldehydes. Upon formation, reactive aldehydes can either undergo detoxification by phase I and phase II enzymes or covalently modify protein side chains (Lys, His, and Cys), leading to protein carbonylation.

Figure 12.2 Mechanism of formation of reactive α,β‐unsaturated aldehydes from PUFA. Formation mechanism of (a) 4‐HNE from arachidonic acid (AA) and (b) 4‐HHE from docosahexaenoic acid (DHA).

Figure 12.3 Oxidative stress and antioxidant expression in obese adipose tissue. Expression of major antioxidant enzymes is downregulated in the transition from the lean to obese state. As a result, ROS is channeled toward the formation of reactive lipid aldehydes and protein carbonylation.

Figure 12.4 Methods for detection of carbonylated proteins.

Figure 12.5 KEGG analysis of mitochondrial protein carbonylation in adipocytes. Schematic representation of key mitochondrial pathways that contain carbonylated proteins. The number represents the fraction of pathway proteins, based on KEGG analysis, that are known to be carbonylated. Data from Curtis et al. [11].

Chapter 13

Figure 13.1 Proposed sequence of events leading from the formation of superoxide in the electron transport chain in the inner mitochondrial membrane, via damage to aconitase and release of Fe ions, to oxidative modification of a protein in the matrix space. The individual events are marked with numbers: (1) formation of superoxide at Complexes I and III, (2) interaction of superoxide with aconitase and release of Fe ions from the FeS center, (3) unspecific binding of Fe ions to other proteins, (4) hydrogen peroxide formation by dismutation of superoxide, (5) generation of hydroxyl radical by Fenton reaction at the bound Fe and (6) local oxidation of biomolecules, for example, carbonylation of a protein, by the hydroxyl radical.

Chapter 14

Figure 14.1 Subcellular location of carbonylated proteins. Proteins from pathways or functions with fewer than five members were grouped as “Others.” Each group includes proteins with two or more possible locations.

Figure 14.2 Physiological function of carbonylated proteins. Proteins from physiological functions with fewer than five members were grouped as “Miscellaneous.”

Guide

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WILEY SERIES ON MASS SPECTROMETRY

Series Editors

Dominic M. DesiderioDepartments of Neurology and BiochemistryUniversity of Tennessee Health Science Center

Joseph A. LooDepartment of Chemistry and BiochemistryUCLA

Founding Editor

Nico M. M. Nibbering (1938–2014)Dominic Desiderio

A complete list of the titles in this series appears at the end of this volume.

Protein Carbonylation

Principles, Analysis, and Biological Implications

Edited by Joaquim Ros

University of Lleida

Lleida, Spain

This edition first published 2017© 2017 John Wiley & Sons, Inc.

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Library of Congress Cataloging‐in‐Publication Data

Names: Ros, Joaquim, 1955– editor.Title: Protein carbonylation : principles, analysis, and biological implications / edited by Joaquim Ros.Description: 1st edition. | Hoboken, NJ : John Wiley & Sons, Inc., 2017. | Includes bibliographical references and index.Identifiers: LCCN 2017004060 (print) | LCCN 2017005655 (ebook) | ISBN 9781119074915 (hardback) | ISBN 9781119374961 (pdf) | ISBN 9781119374954 (epub)Subjects: | MESH: Protein CarbonylationClassification: LCC QP517.P76 (print) | LCC QP517.P76 (ebook) | NLM QZ 160 | DDC 572/.633–dc23LC record available at https://lccn.loc.gov/2017004060

Cover Image: courtesy of Joaquim RosCover design: Wiley

List of Contributors

Somaieh Afiuni‐ZadehDepartment of Biochemistry, Molecular Biology and BiophysicsUniversity of MinnesotaMinneapolis, MNUSA

Esther BarreiroPulmonology Department and Muscle and Lung Cancer Research GroupInstitut Hospital del Mar d’Investigacions Mèdiques (IMIM)‐Hospital del Mar, Health and Experimental Sciences (CEXS), Universitat Pompeu Fabra (UPF), Barcelona Biomedical Research Park (PRBB)BarcelonaSpainandCentro de Investigación en Red de Enfermedades Respiratorias (CIBERES), Instituto de Salud Carlos III (ISCIII)BarcelonaSpain

José M. BautistaHospital 12 de Octubre Research Institute, Avda. de Cordoba s/nMadridSpainandDepartment of Biochemistry and Molecular BiologyUniversidad Complutense de Madrid, Ciudad UniversitariaMadrid, Spain

David A. BernlohrDepartment of Biochemistry, Molecular Biology and BiophysicsUniversity of Minnesota‐Twin CitiesMinneapolis, MN, USA

Joel S. BurrillDepartment of Biochemistry, Molecular Biology and BiophysicsUniversity of Minnesota‐Twin CitiesMinneapolis, MN, USA

D. Allan ButterfieldDepartment of Chemistry and Sanders‐Brown Center on AgingUniversity of KentuckyLexington, KYUSA

Elisa CabiscolDepartament de Ciències Mèdiques Bàsiques, IRB LleidaUniversitat de LleidaLleida, CataloniaSpain

Graziano ColomboDepartment of BiosciencesUniversity of MilanMilanItaly

Isabella Dalle‐DonneDepartment of BiosciencesUniversity of MilanMilanItaly

Amalia DiezHospital 12 de Octubre Research Institute, Avda. de Cordoba s/nMadridSpainandDepartment of Biochemistry and Molecular BiologyUniversidad Complutense de Madrid, Ciudad UniversitariaMadridSpain

Maria FedorovaInstitute of Bioanalytical ChemistryFaculty of Chemistry and MineralogyLeipzigGermanyandCenter for Biotechnology and BiomedicineUniversität LeipzigLeipzigGermany

Maria Lisa GaravagliaDepartment of BiosciencesUniversity of MilanMilanItaly

Saghi GhaffariDepartment of Developmental & Regenerative BiologyDivision of Hematology, Oncology, Department of MedicineBlack Family Stem Cell InstituteTisch Cancer InstituteIcahn School of Medicine at Mount SinaiNew York, NYUSA

Timothy J. GriffinDepartment of Biochemistry, Molecular Biology and BiophysicsUniversity of MinnesotaMinneapolis, MN, USA

Tilman GruneDepartment of Molecular ToxicologyGerman Institute of Human Nutrition Potsdam‐Rehbruecke (DIfE)NuthetalGermany

Amy K. HauckDepartment of Biochemistry, Molecular Biology and BiophysicsUniversity of Minnesota‐Twin CitiesMinneapolis, MNUSA

Jesper F. HavelundDepartment of Molecular Biology and GeneticsAarhus UniversitySlagelseDenmarkandDepartment of Biochemistry and Molecular BiologyUniversity of Southern DenmarkOdense MDenmarkandInstitute of Molecular MedicineUniversity of Southern DenmarkOdense CDenmark

Tobias JungDepartment of Molecular ToxicologyGerman Institute of Human Nutrition Potsdam‐Rehbruecke (DIfE)NuthetalGermany

Jeannette KönigDepartment of Molecular ToxicologyGerman Institute of Human Nutrition Potsdam‐Rehbruecke (DIfE)NuthetalGermanyRaymond LiangDepartment of Developmental & Regenerative BiologyIcahn School of Medicine at Mount SinaiNew York, NYUSA

María LinaresHospital 12 de Octubre Research Institute, Avda. de Cordoba s/nMadridSpain

Ashraf G. MadianBiotechnology and Aseptic Sciences GroupGlobal Technology Services, Hospira a Pfizer CompanyLake Forest, ILUSA

Aldo MilzaniDepartment of BiosciencesUniversity of MilanMilanItaly

Ian Max MøllerDepartment of Molecular Biology and GeneticsAarhus UniversitySlagelseDenmark

Dalay H. OlsonDepartment of Biochemistry, Molecular Biology and BiophysicsUniversity of Minnesota‐Twin CitiesMinneapolis, MNUSA

Antonio PuyetHospital 12 de Octubre Research Institute, Avda. de Cordoba s/nMadridSpainandDepartment of Biochemistry and Molecular BiologyUniversidad Complutense de Madrid, Ciudad UniversitariaMadridSpain

Tanea T. ReedDepartment of ChemistryEastern Kentucky UniversityRichmond, KYUSA

Fred E. RegnierChemistry DepartmentPurdue UniversityWest Lafayette, INUSA

Adelina Rogowska‐WrzesinskaDepartment of Biochemistry and Molecular BiologyUniversity of Southern DenmarkOdense MDenmark

Joaquim RosDepartament de Ciències Mèdiques Bàsiques, IRB LleidaUniversitat de LleidaLleida, CataloniaSpain

Yuichiro J. SuzukiDepartment of Pharmacology and PhysiologyGeorgetown University Medical CenterWashington, DCUSA

Jordi TamaritDepartament de Ciències Mèdiques Bàsiques, IRB LleidaUniversitat de LleidaLleida, CataloniaSpain

Koji UchidaGraduate School of Bioagricultural SciencesNagoya UniversityNagoyaJapan

Ao ZengNovilytic LLCWest Lafayette, INUSA

Preface

Protein carbonylation has attracted the interest of a great number of laboratories since its pioneering studies at the Earl Stadtman’s lab at NIH started in the early 1980s. Since then, detecting protein carbonyls in situations of oxidative stress has become a highly efficient tool to uncover biomarkers of oxidative damage in normal and altered cell physiology. Carbonylated proteins suffer from structural alterations that can impair function or, in certain cases, can have a regulatory role. For these reasons, identification of carbonylated proteins and the site of carbonylation are essential pieces in elucidating the mechanism of altered cellular function occurring under endogenous or exogenous oxidative stresses.

In this book, research groups from several areas of interest have contributed to update the knowledge on the detection, analyses, and identification of carbonylated proteins and the sites where these modifications occur.

I am sure that the scientific community will benefit from these reviews since they deal with specific, detailed technical approaches to study the formation and detection of protein carbonyls. Moreover, the biological impact of such modifications in metabolic, physiologic, and structural functions and how these alterations can help us understand the downstream effects on cell function are discussed.

Finally, I want to express my gratitude to Rodney Levine for his help in designing the book and convincing the authors to contribute a chapter. Without him this book would not have been as successful as the final result shows.

1Reactive Oxygen Species Signaling from the Perspective of the Stem Cell

Saghi Ghaffari1,2,3,4and Raymond Liang1

1Department of Developmental & Regenerative Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA

2Division of Hematology, Oncology, Department of Medicine, Icahn School of Medicine at Mount Sinai, New York, NY, USA

3Black Family Stem Cell Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA

4Tisch Cancer Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA

CHAPTER MENU

1.1 Introduction

1.2 ROS Regulation

1.3 ROS Signaling

1.4 ROS and Stem Cells

1.4.1 Adult Stem Cells

1.4.2 Embryonic Stem Cells

1.5 ROS, Metabolism, and Epigenetic Influence

1.6 Stem Cells and Mitochondria

1.7 ROS and Stem Cell Aging

1.8 Concluding Remarks

References

1.1 Introduction

Stem cells maintain tissue integrity and homeostasis by regenerating damaged or lost cells throughout life. Impaired stem cell function may promote defective response to stress, aging, and cancer. Work in the past decade has uncovered the critical role that redox signaling plays in the biology of stem cells. A major part of this work has taken place in blood‐forming (hematopoietic) stem cells (HSCs) that are broadly used as a model system for adult stem cells. This chapter overviews the investigations of redox regulation of stem cells in the past decade.

1.2 ROS Regulation

ROS are generated from the reduction of molecular oxygen by one electron. ROS species are composed mainly of superoxide anions (O2−), hydrogen peroxide (H2O2), and hydroxyl radicals (OH−). The superoxide anion is highly reactive and is rapidly reduced to H2O2 by the antioxidant enzyme superoxide dismutase (SOD) [1]. H2O2 can be further reduced to H2O and O2 by cellular antioxidants. ROS react adversely with and damage DNA, lipids, and proteins, the cumulative effects of which may cause cellular alterations or death. Overall ROS‐mediated damage to macromolecules is thought to contribute to the physiological effects of aging [2]. ROS are also considered to be essential components in multiple biological processes as second messengers intimately implicated in the physiological regulation of signaling pathways [3]. Alterations of ROS generation versus scavenging, that is creating the redox milieu, may lead to disease as a result of either too much direct ROS damage (e.g., DNA mutations) or perhaps by impaired function of physiologically relevant ROS‐dependent signaling pathways (e.g., myeloproliferative disorder; see succeeding text).

The main source of ROS in the cell is mitochondrial respiration. The generation of proton motive force by the electron transport chain—which leads to ATP production through ATP synthase in a process known as oxidative phosphorylation—is responsible for mitochondrial respiration. However, a small fraction, approximately 0.1–0.2% of O2, consumed by mitochondria form ROS through the premature electron flow to O2 mainly through complexes I and III [4]. The cell type, the environment, and ultimately the activity of mitochondria can influence greatly the precise proportion of ROS generated from mitochondrial respiration [5]. Thus, modulations of mitochondrial activity as well as metabolism in general regulate ROS levels; for instance, reduced ROS levels are achieved by decreasing the rate of mitochondrial respiration via minimizing oxidative phosphorylation. Furthermore, processes that regenerate oxidized glutathione, such as the pentose phosphate pathway, repress ROS levels. Another major source of ROS, in addition to mitochondria, is the membrane‐bound protein NADPH oxidase (NOX), which consumes NADPH to generate O2 and subsequently H2O2. NOX generation of ROS has antimicrobial effects in host defense. In addition, NOX are also important for producing ROS in non‐phagocytic cells to influence cellular signaling including growth factor (GF) signaling [6]. This includes increased NOX4‐mediated ROS production in stem cells [7]. Notably differentiation of mesenchymal stem cells (MSCs) toward adipocytes or neuron‐like cells has also been shown to employ NOX4‐mediated H2O2 signaling as well as mitochondrial ROS [8, 9]. Elevated ROS in MSCs on the other hand reduces their engraftment potential and induces apoptosis after transplantation [7, 10].

Under normal physiological conditions, the generation of ROS is tightly regulated by the ROS scavenging system. ROS scavengers are antioxidant enzymes that can neutralize ROS by directly reacting with and accepting electrons from ROS. When ROS production outpaces ROS scavenging, an excessive accumulation of ROS occurs, leading to oxidative stress and adverse effects on multiple cellular components including proteins, lipids, and nucleotides. To counteract this, the cell contains multiple types of antioxidants specific to different species of ROS, which helps to prevent pathological levels of ROS and to repair oxidative damage to cellular components. These include SOD, catalase, peroxiredoxins (PRX), thioredoxin (TRX), glutathione peroxidase (GPX), and glutathione reductase (GR). Glutathione, a tripeptide, is one of the most abundant antioxidants synthesized by the cell. Oxidized proteins and H2O2 are reduced by glutathione through the glutaredoxin and TRX system. Other key antioxidants include SOD and catalase, which reduce O2− and H2O2, respectively. The subcellular localization of antioxidants at areas of high ROS generation, such as within the mitochondria, may further enhance the efficiency of ROS scavenging.

1.3 ROS Signaling

Despite their deleterious properties, cumulating evidence in the past three decades has established ROS as pivotal signals in cell fate regulation [11, 12]. There is little doubt that oxygen radicals serve as signaling messengers that variably influence cellular behavior [13, 14]. ROS reaction with proteins such as transcription factors, kinases, and phosphatases alters processes that regulate cell cycle, apoptosis, quiescence, or differentiation [15–17]. GF and oncogenic signaling [18–23] are some examples of ROS signaling. ROS also influence transcriptional activity and likely epigenetics [24–26]. The main ROS species involved in intracellular signaling are Hydrogen peroxide (H2O2) mostly due to their relatively longer half‐life and ability to easily diffuse through membranes relative to other types of ROS [27]. H2O2 is also among ROS species with substrate specificity that generates reversible oxidation that is likely to trigger signaling cascade in in vivo physiological settings [12].

ROS signal via direct modification of proteins by amino acid oxidation, the most common of which is oxidation of cysteine residues [28]. ROS signaling to amino acids can cause functional changes in a range of proteins. Proteins directly modified by ROS—known as redox sensors—undergo a conformational change as a result of oxidative modification that influences their function, stability, subcellular localization, interactions with other proteins, and other critical processes. A major example is provided by ROS modulation of protein tyrosine phosphatases (PTP) [1]. It has been shown recently that ROS‐mediated inhibition of PTP1B (encoded by PTPN1) in oncogenic‐induced senescent cells results in the upregulation of cell cycle inhibitor p21CIP, cell cycle arrest, and senescence by a mechanism involving miRNAs. These studies showed that argonaute that regulates miRNA loading is a target of PTP1B whose repression results in tyrosine phosphorylation of argonaute and reduced loading of miRNAs targeting p21CIP leading to cell cycle arrest and senescence [29]. These studies illustrate the extent of ROS signaling impact and further reiterate the function of ROS as rheostat in cell signaling [30]; in addition by establishing a link between ROS, inhibition of phosphatases, and regulation of miRNAs, these studies expand the scope of ROS‐mediated modulations of signaling pathways.

ROS regulation of protein function is complicated by many feedback loops. While ROS can modify protein function, a growing network of proteins modulates ROS levels. These include PTEN and sirtuins (SIRTs) (specifically SIRT1 and SIRT3), ataxia telangiectasia mutated (ATM), p38 mitogen‐activated protein kinase (MAPK), mammalian target of rapamycin (mTOR), and protein kinase B (AKT) protein kinases as well as the multifunctional apurinic/apyrimidinic (AP) endonuclease1/redox factor‐1 (APE/Ref‐1) protein. Transcription factors such as nuclear factor kappa B (NFκB) mediate ROS transactivation of the hypoxia‐inducible factor 1 alpha (HIF‐1α) [31]; Forkhead box O (FOXO) family; nuclear factor (erythroid‐derived 2)‐like 2, also known as NFE2L2 or NRF2; PR domain containing 16 (PRDM16); and p53 tumor suppressor [32–37]. Among these, many proteins considered as redox sensors that also modulate ROS levels have key functions in the regulation of stem cell fate (reviewed in [13, 38]) (Figure 1.1). For instance, changes of ROS and p53 activity by thioredoxin‐interacting protein (TXNIP) may be implicated in hematopoietic stem cell (HSC) function specifically with age [39]. The polycomb group member BMI1 also regulates stem cell function, modulates ROS levels, and is implicated in regulating mitochondrial function [40–42]. Some of these have also been implicated in the regulation of mitochondrial biogenesis or metabolism.

Figure 1.1 Redox sensors critical for stem cell fate: ROS regulation of signaling molecules and transcription factors and their effect on ROS regulation.

1.4 ROS and Stem Cells

Current findings raise the possibility that ROS modulations influence signaling pathways that ultimately impinge on key transcription factors. In turn these factors readjust ROS levels by regulating the expression of antioxidant, metabolic, and mitochondrial genes. Transcription factors that are essential for stem cell machinery and critical for cellular redox state include HIF, FOXO, PRDM16, NRF2, and p53. This model postulates that ROS function as rheostat especially in cells that are highly sensitive to levels of ROS [30] such as stem cells that maintain low ROS levels.

1.4.1 Adult Stem Cells

Adult stem cells including stem cells of the hematopoietic system, skin, muscle, brain, and intestine share two key properties: (i) they are capable of self‐renewing divisions to generate other stem cells and (ii) are multipotent, able to give rise to all cells within their tissue of origin. Adult stem cells replace differentiated cells and replenish damaged and lost tissue during fetal life and throughout life after birth. Adult stem cells with very few exceptions are mainly quiescent under homeostatic conditions as has been definitively shown for stem cells of the skin and hematopoietic system [43–46] (reviewed in Ref. [30]). Quiescence of stem cells is critical for their self‐renewal property. In response to damage or loss and in contrast to homeostasis, stem cells proliferate extensively to regenerate their tissue of origin. To adapt to either quiescence or the highly proliferative state, stem cells have adopted metabolic plasticity. While the precise nature of the stem cell metabolic program remains elusive, levels of ROS appear to both reflect the stem cell metabolic state and have profound effects on stem cell behavior [13]. This is of major importance since perturbations in stem cell properties are associated with degenerative diseases and aging.

Multipotent hematopoietic progenitors in Drosophila exhibit higher ROS levels relative to their downstream progenies [47]. This property is shared with mammalian myeloid blood progenitors relative to their upstream HSC. In this in vivo drosophila model, burst of endogenous ROS in hematopoietic progenitors primes the larval lymph gland for differentiation [47]. In agreement with an in vivo ROS function in mediating hematopoietic cell fate, accumulated ROS in primary hematopoietic progenitors in the context of loss of transcription factor FOXO3 leads to myeloproliferation [48]. In mammals, stem cells of the hematopoietic system contain low ROS levels [49]. Among major known HSC regulators of ROS are transcription factors FOXO (FOXO3) and ATM protein kinase. FOXOs are evolutionarily conserved regulators of redox state that inhibit oxidative stress in quiescent cells by direct transcription of antioxidant genes including SOD and catalase [50–56]. FOXO’s control of the redox homeostasis is also via the pentose phosphate pathway [57]. The redox control contributes to FOXO regulation of aging and longevity [53–55]. In the hematopoietic system, in addition to stem cells, FOXO3 regulates redox state in primary erythroblasts and myeloid progenitors [48, 58].

Increased ROS in HSC is associated with HSC differentiation and increased production of their immediate progenitors [49]. Notably, HSC are highly enriched in glutathione S‐transferase enzymes that mediate detoxification of xenobiotics and defense against environmental stress and cellular damage [59]. Dormant HSCs are acutely sensitive to oxidative stress, a cellular state instigated by an imbalance between the generation and the detoxification of ROS [33, 36, 37, 60–62]. In many cases unbalanced ROS accumulation is associated with impaired HSC function in vivo [60, 63, 64]. Some of the main examples are provided by ATM kinase (Atm)−/− HSC, loss of Foxo1/3/4 (Forkhead box O 1/3/4) transcription factors, or just Foxo3 deletion [33, 36, 37, 60]. In many cases such as in Atm−/− HSC, increased ROS levels mediate defects of stem cell activity [60]. However, in contrast to ATM−/− HSC, elevated ROS do not mediate the defective long‐term repopulation activity—that is, the ultimate measurement of in vivo blood stem cell activity—of Foxo3−/− HSC [60, 65]. ATM and FOXO3 are in a cross talk in which ATM enzymatic activity and expression are regulated by FOXO3 [48, 66]; FOXO3 is required for HSC mitochondrial metabolism [65], while the role of ATM in mitochondrial regulation of HSC is less clear. Control of redox balance and metabolic gene transcription by FOXO3 is also implicated in the maintenance of neural stem cells (NSCs) [57, 67]. However NSCs require high ROS to maintain their self‐renewal and the regulation neurogenesis properties [68]. Although FOXOs are also critical for embryonic stem cell (ESC) pluripotency, this function does not seem to be through regulation of oxidative stress in ESCs [69].

NRF2 is a ubiquitously expressed transcription factor and a master regulator of antioxidant response and mitochondrial biogenesis. Loss of NRF2 results in relative expansion of HSCs and increased generation of their progenitors without any impact on HSC self‐renewal. This has been attributed to cell intrinsic hyper‐proliferation and is associated with modulations of cell migration and homing [70]. Unexpectedly the defective HSC function in these mice is associated with normal ROS levels; on the other hand ROS levels are increased upon restoration of NRF2−/− HSC function [71]. In addition, enhanced NRF2 signaling increases hematopoietic stem and progenitor cell function [70, 71] and mitigates irradiation‐induced myelosuppression and mortality [71]. These studies suggest that despite the association that is commonly observed between ROS levels and HSC function [63, 72–77], elevated ROS do not always result in HSC defective function; these conclusions are analogous to that derived from Foxo3−/− HSC studies [65, 70, 78]. Current findings point to unhealthy mitochondria rather than ROS as potential mediators of stem cell defects [65, 79] in the case of Foxo3−/− HSC. Given the importance of both NRF2 and FOXO3 for mitochondrial function [65, 72–75, 80], it is conceivable that lack of association between ROS elevation and defective HSC function phenotype might indicate active involvement of mitochondria in NRF2−/− HSC as has been proposed for Foxo3−/− HSC [65]. Similar NRF2 functions are described in lung stem cells. In mouse and human airway basal stem cells (ABSCs), intracellular flux from low to moderate ROS levels is required for stem cell self‐renewal and proliferation. The stem cell self‐renewal involves modulations of ROS levels that activate NRF2 and Notch pathways [81]. NRF2 bears interesting functions in cancer stem cells that involve its interactions with the cell cycle inhibitor p21 (Cdkn1a) that competes with Keap1 for NRF2 binding [82, 83] and stabilizes NRF2 in TGF‐beta‐responsive squamous cell carcinoma stem cells [84]. This binding increases glutathione metabolism and NRF2 antioxidant response that render cells drug resistant. Decreasing NRF2 increases drug‐induced apoptosis in these cancer stem cells without significantly modifying their low cycling profile [84]. In resting drosophila intestinal stem cells, NRF2 (CncC) is constitutively active in maintaining low ROS levels [85]. Increased degradation of NRF2 by Keap1 enhances intestinal stem cell proliferation. Loss of NRF2 increases ROS levels and accelerates age‐related degeneration of the intestinal epithelium.

These studies raise the possibility that HSC defects are not directly mediated by ROS elevation when mitochondrial function is defective [70, 71, 78, 86–92]. In these settings as observed in Foxo3−/− and Nrf2−/− HSC, ROS elevation might only be secondary to changes in mitochondrial function, a signal that might be indicating the unhealthy state of mitochondria and mediating only some (e.g., DNA damage) of stem cell defects [65, 70, 71, 93]. ROS elevation in hematopoietic progenitors induces myeloproliferation in vivo [48]. Importantly, scavenging ROS in vivo improves myeloproliferation in the context of human leukemias [94, 95].

1.4.2 Embryonic Stem Cells

ESCs originate from the inner cell mass of the mammalian blastocyst and possess the ability to differentiate all three germ layers of the embryo under defined in vitro conditions [96]. ESCs are highly resistant to oxidative stress [97] but, undergo apoptosis when exposed continuously to high ROS levels. Their genomic integrity and clonal recovery is maintained when cultured under physiological oxygen levels (2%) [98], whereas prolonged hypoxic environment leads to increased ROS and apoptosis [99].

ESCs have a shortened G1 cell cycle phase which enable them to self‐renew rapidly. ESC self‐renewal relies mainly on glycolysis and the pentose phosphate pathway, with oxidative phosphorylation clearly suppressed [100–104]. The rapid generation of ATP and the precursors for nucleotide biosynthesis by glycolysis and the pentose phosphate pathway, respectively, enable the rapid DNA replication and ESC growth [105]. Undifferentiated pluripotent ESC in contrast to their lineage‐committed progenies relies on enhanced lactate production and an uncoupling of electron transport chain flux from ATP production, suggesting their dependence on glycolysis. This is associated with an immature mitochondrial morphology and a more reduced redox environment, further supporting the notion that ESC avoids dependence on mitochondrial metabolism [104, 106]. Forced activation of oxidative phosphorylation by knockdown of uncoupling protein 2 (UCP2), that limits pyruvate entry into the mitochondrial oxidative phosphorylation pathway, as well as by metabolites that activate this pathway leads to loss of stem cell properties and increased differentiation or apoptosis [104]. Enhancing glycolysis or inhibition of oxidative phosphorylation may also be achieved through hypoxia‐induced HIF activation that results in improved proliferation and maintenance of ESCs while repressing differentiation similar to experiments described earlier [107, 108]. In all cases, improved stem cell maintenance is associated with decreased ROS levels. The high sensitivity of mouse ESC to endogenous ROS is in part mediated by deacetylase sirtuin 1 (SIRT1) coordination of p53 activity toward (inhibition) antioxidants with its regulation of pluripotency factor Nanog expression [109]. These functions might also be related to SIRT1 regulation of ESC mitochondria [110]. These findings support the idea that ESC fate may be directly modified by ROS modulation of metabolism. They also suggest that in ESC as in cancer cells glycolysis supports the biosynthetic demands of highly proliferative cells [105].