Redox Signaling by Reactive Electrophiles and Oxidants

Abstract

The concept of cell signaling in the context of nonenzyme-assisted protein modifications by reactive electrophilic and oxidative species, broadly known as redox signaling, is a uniquely complex topic that has been approached from numerous different and multidisciplinary angles. Our Review reflects on five aspects critical for understanding how nature harnesses these noncanonical post-translational modifications to coordinate distinct cellular activities: (1) specific players and their generation, (2) physicochemical properties, (3) mechanisms of action, (4) methods of interrogation, and (5) functional roles in health and disease. Emphasis is primarily placed on the latest progress in the field, but several aspects of classical work likely forgotten/lost are also recollected. For researchers with interests in getting into the field, our Review is anticipated to function as a primer. For the expert, we aim to stimulate thought and discussion about fundamentals of redox signaling mechanisms and nuances of specificity/selectivity and timing in this sophisticated yet fascinating arena at the crossroads of chemistry and biology.

Figure 1

The compartmentalized/subcellular generation of various chemotypes of ROS and RES. Some of the most important sources of ROS in biological systems include NOX enzymes localized on plasma membrane and membranes of cellular organelles, mitochondrial ETC, and metabolic enzymes in the ER and peroxisomes. Enzymatic and non-enzymatic oxidation of fatty acids are the primary sources of RES in mammalian cells.

Figure 2

Sites of ROS generation in eukaryotic cells (A) NOX enzymes generate O₂ ^•−by a single electron reduction of ³O₂.(144,145) NOX enzymes (NOX2 shown here), consist of multi-subunit complexes comprising transmembrane subunits (gp91^phox/NOX2 and p22^phox) and various cytosolic regulatory domains. Stimulation with cytokines or microorganisms results in phosphorylation and subsequent recruitment of the cytosolic components p67^phox, p47^phox, and p40^phox and a Rac1 GTP to the transmembrane subunits. The p67^phox initiates electron transfer by accepting two electrons from NADPH. The electrons are relayed through FAD and then sequentially to two heme cofactors (depicted as orange circles) housed in the transmembrane gp91^phox domain. O₂ acts as the terminal electron acceptor generating two molecules of O₂ ^•−per molecule of NADPH used. GDI: guanosine nucleotide dissociation inhibitor. (B) Complex I and Complex III are the two important sites of O₂ ^•−generation in the mitochondrial electron transport chain (mETC). At Complex I, O₂ ^•−is generated by the reaction of ³O₂ with reduced FMN. O₂ ^•−generation at Complex I is favored under conditions of high proton motive force and when CoQ is reduced, resulting in a reverse electron transfer from reduced CoQ to FMN site at Complex I. ³O₂ reduction at Complex III is mediated by ubisemiquinone housed in this subunit. Complex I releases O₂ ^•−primarily in the mitochondrial matrix whereas Complex III generated O₂ ^•−is released in both the mitochondrial matrix and the intermembrane space. (C) The catalytic cycle of microsomal monooxygenase (MMOs) CYP450. CYP450 is a heme-iron-containing protein. In the resting state, iron is in a hexa-coordinated ferric form (Fe³⁺)–equatorial sites are taken up by the heme cofactor (denoted by rhombus); a cysteine thiol from the protein, and a water molecule occupy the apical sites (i). Substrate binding displaces the water molecule, and a subsequent one-electron reduction by NAD(P)H via FAD/FMN-containing CYP450 reductase (CPR) generates the penta-coordinated ferrous complex (Fe²⁺) (iii). Addition of O₂ to (iii) generates a hexa-coordinated Fe³⁺⁻O₂ oxycomplex (iv). MMOs in the eukaryotic ER generate ROS because of inefficient utilization of activated O₂ for substrate oxidation. Two uncoupling reactions that result in the generation of ROS are shown. The first route generates O₂ ^•−due to the decay of one-electron reduced oxycomplex (iv). A possible route of H₂O₂ generation includes protonation and subsequent uncoupling of the peroxycomplex (v). Adapted with permission from ref. (146). Copyright 2004 Elsevier.

Figure 2

Sites of ROS generation in eukaryotic cells (A) NOX enzymes generate O₂ ^•−by a single electron reduction of ³O₂.(144,145) NOX enzymes (NOX2 shown here), consist of multi-subunit complexes comprising transmembrane subunits (gp91^phox/NOX2 and p22^phox) and various cytosolic regulatory domains. Stimulation with cytokines or microorganisms results in phosphorylation and subsequent recruitment of the cytosolic components p67^phox, p47^phox, and p40^phox and a Rac1 GTP to the transmembrane subunits. The p67^phox initiates electron transfer by accepting two electrons from NADPH. The electrons are relayed through FAD and then sequentially to two heme cofactors (depicted as orange circles) housed in the transmembrane gp91^phox domain. O₂ acts as the terminal electron acceptor generating two molecules of O₂ ^•−per molecule of NADPH used. GDI: guanosine nucleotide dissociation inhibitor. (B) Complex I and Complex III are the two important sites of O₂ ^•−generation in the mitochondrial electron transport chain (mETC). At Complex I, O₂ ^•−is generated by the reaction of ³O₂ with reduced FMN. O₂ ^•−generation at Complex I is favored under conditions of high proton motive force and when CoQ is reduced, resulting in a reverse electron transfer from reduced CoQ to FMN site at Complex I. ³O₂ reduction at Complex III is mediated by ubisemiquinone housed in this subunit. Complex I releases O₂ ^•−primarily in the mitochondrial matrix whereas Complex III generated O₂ ^•−is released in both the mitochondrial matrix and the intermembrane space. (C) The catalytic cycle of microsomal monooxygenase (MMOs) CYP450. CYP450 is a heme-iron-containing protein. In the resting state, iron is in a hexa-coordinated ferric form (Fe³⁺)–equatorial sites are taken up by the heme cofactor (denoted by rhombus); a cysteine thiol from the protein, and a water molecule occupy the apical sites (i). Substrate binding displaces the water molecule, and a subsequent one-electron reduction by NAD(P)H via FAD/FMN-containing CYP450 reductase (CPR) generates the penta-coordinated ferrous complex (Fe²⁺) (iii). Addition of O₂ to (iii) generates a hexa-coordinated Fe³⁺⁻O₂ oxycomplex (iv). MMOs in the eukaryotic ER generate ROS because of inefficient utilization of activated O₂ for substrate oxidation. Two uncoupling reactions that result in the generation of ROS are shown. The first route generates O₂ ^•−due to the decay of one-electron reduced oxycomplex (iv). A possible route of H₂O₂ generation includes protonation and subsequent uncoupling of the peroxycomplex (v). Adapted with permission from ref. (146). Copyright 2004 Elsevier.

Figure 3

ROS interconversion in eukaryotic cells. ROS generated in cells are highly dynamic and rapidly interconvert. O₂^{• −} generated by enzymatic pathways such as NOX enzymes and mETC by reduction of ground-state ³O₂, rapidly dismutates to yield H₂O_2. SODs also catalyze this conversion. O₂^{• −} also reacts with ^•NO yielding ONOO⁻, which can further decompose to generate ^•OH. H₂O₂ generates ^•OH in presence of trace metals such as reduced iron. Reactive radicals such as ^•OH initiate lipid peroxidation by abstracting a bis-allylic hydrogen from PUFAs (see Figure 5a). Myeloperoxidase enzyme (MPO) in neutrophils and macrophages(203) utilize H₂O₂ to generate HOCl that can react with ⁻OOH to generate singlet oxygen (¹O₂) in phagocytic cells. ¹O₂ is also directly generated by the excitation of ³O₂ by UV radiation.

Figure 4

A selection of endogenously-generated carbonyl-containing reactive electrophilic species (RES) in biological systems.

Figure 5

Enzymatic generation of lipid-derived RES.(250) COX-1/COX-2-catalyzes the formation of PGH₂ using arachidonic acid (AA) as the substrate. PGH₂ acts as an intermediate for the generation of PGD₂ and PGE₂ by the action of PGD synthase and PGE synthase, respectively. PGH₂ is also an intermediate for the generation of a number of other bioactive non-electrophilic molecules (not shown). PGD₂ and PGE₂ undergo spontaneous dehydration at C₍₉₎ and C₍₁₁₎ to yield PGJ₂ and PGA₂, respectively. PGJ₂ can then isomerize at C₍₁₂₎–C₍₁₃₎–C₍₁₄₎ to generate Δ¹²-PGJ₂ and further dehydrate at C₍₁₅₎ to 15d-PGJ₂.

Figure 6

Non-enzymatic generation of reactive lipid-derived electrophiles (LDEs): The generation of HNE (xiii) and A₂-(xvi) and J₂-isoprostanes (xvii) from ω-6-FAs such as AA (i) is shown here. Reactive radicals such as ^•OH initiate lipid peroxidation by abstracting a bis-allylic hydrogen from AA (i). O₂ addition at either C₍₁₅₎ or C₍₁₁₎ generates the corresponding peroxy radical (ii and viii), which abstracts an H radical (possibly from another AA molecule thus propagating the radical chain reaction) to generate the hydroperoxy intermediates 15-HpETE (iii) and 11-HpETE (ix), respectively. The abstraction of hydrogen at C₍₁₀₎ of 15-HpETE forms a radical intermediate which migrates to C₍₁₂₎, reacts with O₂, yielding the dihydroperoxide intermediate (iv). This intermediate then undergoes Hock cleavage to yield HNE as one possible product. 11-HpETE is shown to first undergo a Hock cleavage to generate the nonenal (x), which undergoes oxygenation to yield the hydroperoxide (xii) and its further reduction to generate HNE (xiii). AA peroxidation also generates isoprostanes. The peroxy radical intermediate (viii) at C₍₁₁₎ can undergo cyclization, further oxygenation, and rearrangement to yield E₂-(xiv) and D₂-isoprostanes (xv). Dehydration of the E₂- and D₂-isoprostanes result in J₂ (xvi) and A₂-isoprostanes (xvii), respectively. Analogous peroxidation pathway from ω-3-FA such as DHA generates HHE.

Figure 7

Generation of NO₂-FA from the alkyl portion of unsaturated FAs. Under low ³O₂, ^•NO₂ undergoes radical adduction with unsaturated FAs (i) (such as linoleic acid) to generate a nitroalkyl radical intermediate (ii). This intermediate can either abstract a hydrogen to generate nitroalkane (iii) or further react with ^•NO₂ to generate a nitro/nitrite intermediate (iv). Subsequent loss of nitrous acids (HONO) yields a nitroalkene (v) whereas hydrolysis of the intermediate generates a hydroxy-nitro FA (vi). Under high ³O₂, a lipid hydroperoxide (vii) is formed.

Figure 8

Nucleophilic amino acid and amino-acid derived molecules (in order of decreasing nucleophilicity)

Figure 9

Reversible and irreversible RES conjugates: pK_a of the C-H proton(s) α-to the carbonyl or nitro group affects the reversibility of the thiol conjugates. In the case of the reduced HNE–thiol adduct, the source of the reductant is currently unknown, but this modification has been reported by independent laboratories for the RES-sensor Keap1 following global treatment of cells with HNE.(259,511) This modification was also detected following T-REX-mediated HNE-delivery to another RES-sensor, Akt3.(512) See also Figure 13.

Figure 10

Properties of a privileged redox sensor. (A) Although lowering of pK_a is widely credited with increasing nucleophilicity of the sensing cysteine residues, other factors must be responsible for privileged redox sensing ability. Indeed, thiolate nucleophiles formed from thiols with higher pK_a are more nucleophilic compared to those formed from low pK_a thiols. Thiolate formed from higher pK_a thiols have a higher HOMO resulting in a better overlap with the LUMO of the electrophile making them more nucleophilic. Other factors such as the extent of solvent exposure and the microenvironment of a sensor cysteine also affect privileged redox sensing (B). In Akt3, a privileged RES sensor, C119 located in the flexible linker region of the enzyme and surrounded by charged amino acids is a privileged HNE sensing residue. PH: Pleckstrin homology domain. Inset: Logo demonstrates the high sequence conservation of C119 and the surrounding charged residue reflecting their functional importance. Sequence alignment of 35 vertebrate Akt3 sequences was performed using Mega7.0. Logo was created using WebLogo (Berkeley).

Figure 11

Oxidation of cysteine by various oxidants generates sulfenic acid. Sulfenic acid can also act as an intermediate to other cysteine modifications including glutathionylation, disulfide bond formation, and sulfenamide species.

Figure 12

(A) Schematic of sulfenylamide formation in enzyme catalytic pocket. Sulfenylamide formation at the catalytic sites of enzymes such as PTP1B prevents them from irreversible hyperoxidation. (B) A sulfenamide intermediate is also formed by the drug omeprazole.

Figure 13

Protein alkylation by (A) monofunctional RES, 9-nitroleic acid, versus (B) modifications by the multifunctional RES, HNE. See also Figure 9.

Figure 14

(A) A selection of fluorescent probes for measuring ROS in cells. (B) Mechanism of Hydroxyethidine (HE) oxidation. Oxidation by superoxide generates 2-OH-E⁺ with distinct spectral properties compared to Ethidium (E⁺) generated by a two-electron oxidation step by hydride acceptors. (C) Oxidation of Amplex Red (an H₂O₂ sensor) or reduction of resazurin (a cell viability reagent) leads to the fluorescent product resorufin.

Figure 15

(A) A selection of boron “:ate”-based fluorescent-probes for measuring H₂O₂ in cells. (B) Mechanism of boron “ate” probe oxidation by H₂O_2.

Figure 16

(A) Examples of genetically-encoded H₂O₂ sensors. The reversibility of genetically-encoded sensors makes them an excellent tool to measure intracellular flux of H₂O₂. Top panel: Orp1-roGFP2 consists of a Orp1, a member of the yeast glutathione peroxidase, fused the redox-sensitive roGFP2. The peroxidatic cysteine on Orp (C36) forms a sulfenic acid in presence of H₂O₂, which is rapidly condensed by the resolving C82 of Orp1. This redox relay is transferred to the conjugated roGFP2 through a thiol-disulfide exchange. Disulfide bond formation of roGFP2 results in conformational change and a gain in fluorescence. Oxidized Orp1 is reversible by Trx. Oxidized roGFP2 can be reversed by Grx. Bottom panel: HyPer uses a circularly permutated YFP protein fused in between the regulatory domain of OxyR, a bacterial H₂O₂ -sensing transcription factor. The peroxidatic cysteine of OxyR (C199) is oxidized by H₂O₂ and resolved by C208 forming a disulfide bond. The consequent change in conformation of the conjugated cpYFP results in increase in fluorescence. Trx: Thioredoxin; Grx: Glutaredoxin. (B) A selection of EPR probes used for detecting radical RES and ROS generation in cells. Inset: Mechanism of radical trapping by cyclic nitrone spin traps to generate a stable nitroxide radical (C) Examples of fluorescent probes for the detection of free RES in cells. Inset: Top: Mechanism of 2-aza-cope reaction based fluorescent probes for detecting formaldehyde. Bottom: Mechanism of Mbo, a ‘turn-on’ fluorescent probe for methylglyoxal. Other hydrazine-based probes for detecting malondialdehyde and formaldehyde work on a similar principle.

Figure 16

(A) Examples of genetically-encoded H₂O₂ sensors. The reversibility of genetically-encoded sensors makes them an excellent tool to measure intracellular flux of H₂O₂. Top panel: Orp1-roGFP2 consists of a Orp1, a member of the yeast glutathione peroxidase, fused the redox-sensitive roGFP2. The peroxidatic cysteine on Orp (C36) forms a sulfenic acid in presence of H₂O₂, which is rapidly condensed by the resolving C82 of Orp1. This redox relay is transferred to the conjugated roGFP2 through a thiol-disulfide exchange. Disulfide bond formation of roGFP2 results in conformational change and a gain in fluorescence. Oxidized Orp1 is reversible by Trx. Oxidized roGFP2 can be reversed by Grx. Bottom panel: HyPer uses a circularly permutated YFP protein fused in between the regulatory domain of OxyR, a bacterial H₂O₂ -sensing transcription factor. The peroxidatic cysteine of OxyR (C199) is oxidized by H₂O₂ and resolved by C208 forming a disulfide bond. The consequent change in conformation of the conjugated cpYFP results in increase in fluorescence. Trx: Thioredoxin; Grx: Glutaredoxin. (B) A selection of EPR probes used for detecting radical RES and ROS generation in cells. Inset: Mechanism of radical trapping by cyclic nitrone spin traps to generate a stable nitroxide radical (C) Examples of fluorescent probes for the detection of free RES in cells. Inset: Top: Mechanism of 2-aza-cope reaction based fluorescent probes for detecting formaldehyde. Bottom: Mechanism of Mbo, a ‘turn-on’ fluorescent probe for methylglyoxal. Other hydrazine-based probes for detecting malondialdehyde and formaldehyde work on a similar principle.

Figure 17

Indirect methods to detect oxidative and electrophilic modifications of proteins (A) Oxidative and electrophilic modification of proteins renders them unreactive to a biotin/fluorophore conjugated alkylating agent such as N-ethyl maleimide (NEM)-Fluorescein. Modified proteins are detected from the loss in signal intensity using SDS-PAGE analysis (B) Similar to (A) except unmodified proteins are first capped using a broadly-reactive alkylating agent such as iodoacetamide (IAM, denoted by black cap). The oxidative modification (X) on proteins is then reduced using a strong reducing agent such as DTT. A fluorophore/biotin-conjugated probe is used to label the reduced proteins. An increase in signal intensity is detected using SDS-PAGE/western blot corresponding to the increase in modification of protein with increasing oxidant concentration. (C) Isotope-labeling methods coupled with mass spectrometric approaches allow quantitative determination of protein modification. In methods such as competitive-isoTOP ABPP, the experimental sample is treated with the desired RES (red/orange circle) and the control sample is treated with the vehicle. The unreacted cysteines in the protein lysate are then capped with an alkylating agent with an alkyne handle (IA alkyne; IAA which caps most but not necessarily all remaining cysteines) which enables click-coupling with isotope-labeled and TEV-protease-cleavable-biotin azide. The samples are then mixed 1:1, enriched using streptavidin beads, the bound proteins are eluted using TEV protease, and a MS analysis performed subsequent to trypsin digestion. Proteins labeled with RES won’t react with alkyne-tagged alkylating agents resulting in a loss in signal intensity in MS spectrum.

Figure 18

(A) A selection of dimedone-based nucleophilic probes and (B) electrophilic probes for detecting sulfenic acid formation. (C) Aryl nitroso compounds enable detection of sulfinic acid although their reactivity with reduced thiol may limit their ability to detect sulfinic acid in biological systems. Capping reduced thiols with alkylating agents prior to probe treatment may mitigate this problem. A biotinylated aryl nitroso probe has recently been used for detection of sulfinic acid modified proteins in biological samples.

Figure 19

(A) A selection of nucleophilic probes used for aldehyde capture of proteins-modified with endogenous electrophilic signals. Biotinylated and fluorescently-labeled hydrazide can be used for aldehyde capture for detection by WB/mass spectrometry and in-gel fluorescence, respectively. (B) Biotinylated and fluorescently-labeled LDEs can also be used to directly enrich or detect protein targets. Alternatively, alkyne and azide functional groups provide largely non-intrusive handles to assess protein targets of electrophiles. (C) Schematic of direct methods for profiling LDE sensitive cysteines in the proteome. Cells/lysates are treated with an excess of alkyne-functionalized RES (red diamond). Modified proteins are either observed using in-gel fluorescence after click-coupling with fluorophore-azide or enriched using streptavidin beads post click coupling with biotin-azide. Subsequent tandem-MS reveals the modified peptides, and depending on the experimental context, the site(s) of modification.

Figure 20

Workflow of G/T-REX. (A) G-REX employs expression of HaloTag in live cells to bind a non-invasive photocaged precursor to HNE (Ht-PreHNE; see inset) and release maximally 5 μM HNE in situ upon photoactivation. This method can be coupled to proteomics to globally profile privileged RES sensors. (B) T-REX is built on a proximity-targeting concept of a freely-diffusible RES (demonstrated thus far for various native bioactive LDEs such as HNE) to a specific sensor protein-of-interest (POI). As such, T-REX enables low-occupancy on-target RES-modifications at a user-defined time and space in living systems which can be directly linked to functional redox responses (thus far proven compatible with live cells, fish, and worms). Under these native electrophile-limited settings, competition is set up between innate diffusivity of RES and reactivity (kinetic privilege) of the sensor protein/cysteine. T-REX involves incubating the live cells/animals expressing functional HaloTagged-POI with sub-micromolar bioinert bifunctional photocaged precursors to an LDE such as HNE (see Inset) non-invasively, followed by rinsing out the excess probe unbound to Halo, and subsequent light exposure. The latter rapidly liberates the LDE in the amount at maximum stoichiometric to in vivo concentration of Halo-POI within the coordination shell of the Halo-POI. Built-in controls ensure/account for no off-target labeling/responses.(252,259,389,512,609,957,958) Signaling consequences resulting from the on-target LDE modification of a privileged LDE-sensor POI have been shown to be evaluated using various downstream readouts. Inset: One terminus of the bifunctional photocaged probe (exemplified for HNE) consists of a hexyl chloride linker (brown) that can covalently conjugate with high specificity and affinity to a HaloTag (gray) genetically fused to a POI (blue). The other terminus consists of photocaged-LDE which upon photouncaging using a low-power UV lamp generates LDE such as HNE.

Figure 21

Small molecules bearing electrophilic pharmacophores approved by the FDA (year approved in parenthesis) or currently in clinical trials in the treatment of indicated disease. HER2⁺: human EGFR-2-positive. MS: Multiple sclerosis. NSCLC: Non-small cell lung carcinoma.

Figure 22

Elevated ROS and RES levels are markers of various diseases including diabetes, multiple sclerosis, and neurodegenerative disorders. Activation of ROS/RES detoxification pathways such as the Nrf2/AR has shown promise in ameliorating these diseases. However, pathways other than Nrf2/AR may play important role in mediating the beneficial effects of broad-specificity electrophilic drugs/drug candidates such as Tecfidera.

Figure 23

Nrf2/AR signaling is at the crossroads of Wnt and Akt/FOXO signaling pathways. Nrf2, the master transcriptional regulator of the AR, is controlled by multiple redox-sensitive proteins. Keap1 negatively regulates AR signaling by targeting Nrf2 for degradation under non-stress conditions. Nrf2 is also regulated by the Akt signaling axis: activation of the Akt pathway triggers GSK3β-mediated phosphorylation of Nrf2, resulting in the degradation of Nrf2 mediated by β-TrCP. Recent studies have identified the role of β-catenin, another substrate of β-TrCP, in regulating Nrf2/AR. Nrf2 inhibits β-catenin-mediated Wnt signaling, and β-catenin stimulates Nrf2-dependent AR.