1
Introduction
Reactive
oxygen, nitrogen, and sulfur species, referred to as ROS,
RNS, and RSS, respectively, are produced during normal cell function
and in response to various stimuli. An imbalance in the metabolism
of these reactive intermediates results in the phenomenon known as
oxidative stress. If left unchecked, oxidative molecules can inflict
damage on all classes of biological macromolecules and eventually
lead to cell death. Indeed, sustained elevated levels of reactive
species have been implicated in the etiology (e.g., atherosclerosis,
hypertension, diabetes) or the progression (e.g., stroke, cancer,
and neurodegenerative disorders) of a number of human diseases.
1
Over the past several decades, however, a new
paradigm has emerged in which the aforementioned species have also
been shown to function as targeted, intracellular second messengers
with regulatory roles in an array of physiological processes.
2
Against this backdrop, it is not surprising that
considerable ongoing efforts are aimed at elucidating the role that
these reactive intermediates play in health and disease.
Site-specific,
covalent modification of proteins represents a prominent
molecular mechanism for transforming an oxidant signal into a biological
response. Amino acids that are candidates for reversible modification
include cysteines whose thiol (i.e., sulfhydryl) side chain is deprotonated
at physiological pH, which is an important attribute for enhancing
reactivity. While reactive species can modify other amino acids (e.g.,
histidine, methionine, tryptophan, and tyrosine), this Review will
focus exclusively on cysteine, whose identity as cellular target or
“sensor” of reactive intermediates is most prevalent
and established.
3
Oxidation of thiols results
in a range of sulfur-containing products, not just disulfide bridges,
as typically presented in biochemistry textbooks. An overview of the
most relevant forms of oxidized sulfur species found in vivo is presented
in Chart 1.
Chart 1
Biologically Relevant Cysteine Chemotypesa
a
Red, irreversible modifications.
Green, unique enzyme intermediates. Note: Additional modifications
can form as enzyme intermediates including thiyl radicals, disulfides,
and persulfides.
Sulfur occupies a unique position
in biology because of its ability
to adopt a wide range of oxidation states (−2 to +6) and chemically
unique forms or “chemotypes”
3a
each with distinct pathways of formation, physical and reactivity
properties. Redox reactions of cysteine residues can lead to an array
of post-translational modifications that are an important mechanism
for the regulation of proteins from all major functional categories
(e.g., enzymes, contractile, structural, storage, and transport proteins).
Among these modifications are reversible, regulatory disulfides, thiosulfinates, S-glutathionylation,
sulfenic acids, sulfenamides, sulfinamides, S-nitrosylation, and persulfides in conjunction
with largely
irreversible species, such as sulfinic acids, sulfonic acids, and
sulfonamides that are often viewed as hallmarks of oxidative stress
and disease.
4
In regards to terminology,
we note that the “-yl-“ particle in the terms above
has gained widespread use in recent years
5
as an analogy to other post-translational modifications, such as
phosphorylation or acetylation, and is not intended to indicate a
specific mechanism of S-group attachment or a radical-associated
process.
The reversibility of many oxidative post-translational
modifications
(oxPTMs) of cysteine thiols highlights their ability to function as
a binary “switch”, regulating protein function, interactions
and localization, akin to phosphorylation. Given this analogy, and
the discovery of biological RO/N/S-generating systems, it not surprising
that investigation of cell signaling pathways involving oxidation
of cysteine residues has emerged as an extremely active area of research.
However, elucidating the functional role of cysteine oxPTMs in normal
physiology and disease has been hampered, in part, because of the
difficultly in detecting these modifications in complex biological
systems with chemical specificity. After a brief introduction reprising
major RO/N/S species produced by cells and mechanisms of thiol oxidation,
we focus this review on different oxPTMs of protein cysteine thiols,
with particular emphasis on those chemical properties that differentiate
one modification from another. In keeping with this general theme,
we review recent progress in using chemical approaches to develop
probes that enable selective and direct detection of individual modifications
within their native cellular environment. Along the way, we complement
this discussion with examples from the literature that highlight ways
in which cysteine oxidation can be used to control protein function
and cell signaling pathways.
2
Cysteine Reactivity and Oxidant
Sensitivity
Ionization constants (pK
a) for the
low-molecular weight thiols, cysteine (Cys), and glutathione (GSH),
are 8.3 and 8.8, respectively. However, pK
a values for cysteine residues in proteins can be strongly influenced
by the local environment. For example, the two active-site cysteines
in the DsbA disulfide oxidoreductase have pK
a values of 3.5 and 10.
6
Low pK
a protein thiols, particularly those ionized
at physiological pH, are often referred to as “reactive cysteines”.
7
Features of the protein environment that can
facilitate thiol ionization include proximity to positively charged
amino acids,
8
hydrogen bonding,
9
and location at the N-terminal end of an α-helix
(Ncap).
10
For example, Ncap effects on cysteine reactivity have recently been noted
in the thiol peroxidase, peroxiredoxin 1 (Prx1),
11
and the epidermal growth factor receptor (EGFR) kinase.
11b,12
Although the molecular basis remains incompletely understood,
empirical
observations indicate that not all cysteine residues in an individual
protein are equally sensitive to oxidation. Since thiolates are much
stronger nucleophiles than thiol groups, one key factor in oxidization
susceptibility is low pK
a. This fact is
highlighted by the observation that many biological oxidants, such
as hydrogen peroxide (H2O2), react exclusively
with the thiolate anion.
13
On the other
hand, as noted by Winterbourn and Hampton, low pK
a is not the only determinant of oxidant reactivity.
14
To illustrate this point, one need only to consider
the 1 000 000-fold difference in reaction rate constants
of H2O2 with the active site cysteine of peroxiredoxin
2 (pK
a ≈ 5–6; 2 × 107 M–1 s–1)
15
and protein tyrosine phosphatases (PTPs), such as PTP1B
(pK
a ≈ 5.4; 20 M–1 s–1).
16
Structural
and functional studies suggest that the superior reactivity of Prx2
is due to a protein environment that is preorganized to activate both
the peroxidatic cysteine and the peroxide substrate, as well as to
stabilize the transition state for the SN2 substitution
reaction.
11a,15
In short, low pK
a protein thiols are prime candidates for oxidation, but
it is also important to recognize that “reactive cysteine”
and “oxidant-sensitive cysteine” are not always synonymous
with one another. A more extensive discussion of this topic has been
presented by Winterbourn and colleagues.
2d,14
2.1
Methods to Identify Low-pK
a Cysteine Residues
From first principles, we
know that cysteine reactivity depends on features of the local protein
microenvironment; however, there is still much to learn about sequence
and structural motifs that are associated with lowering cysteine thiol
pK
a.
9b
One
approach to understand these features is to generate a comprehensive
list of proteins that harbor low pK
a cysteines
and collate this information with sequence and three-dimensional (3D)
structural data. To this end, a number of methods have been developed
to identify low pK
a cysteine residues
in proteins.
Computational methods to identify reactive cysteines
in the proteome are often based on the conservation of redox-active
cysteine residues, particularly those required for catalysis.
17
Chemical methods typically employ reagents such
as N-ethylmaleimide (NEM, 1) or iodoacetamide
(IAM, 2), which form covalent adducts with sulfhydryl
groups by Michael addition or nucleophilic substitution (SN2), respectively (Chart
2). The reaction of
NEM with thiols is faster than IAM and less dependent on pH.
18
However, IAM is more specific for thiols than
NEM, which can modify side chain amines, such as histidine and lysine,
when used in large excess or at basic pH.
19
Since the thiol primarily reacts with IAM as the unprotonated thiolate
anion, this reagent is most frequently used to identify low pK
a cysteines, also referred to as the “reactive
thiol proteome”.
18,20
Both NEM and IAM can
be conjugated to biotin or fluorophores to facilitate enrichment of
labeled proteins, followed by one or two-dimensional (1 or 2D) gel
electrophoresis with subsequent identification by liquid chromatography-tandem
mass spectrometry (LC-MS/MS). In one recent example, N-(biotinoyl)-N′-(iodoacetyl)ethylenediamine,
commonly referred to as biotinylated iodoacetamide (BIAM), was used
to identify surface-exposed reactive cysteine residues in Saccharomyces cerevisiae.
21
In
yet earlier examples, BIAM and 5-iodoacetamido-fluorescein were used
at low micromolar concentrations and mildly acidic pH to label reactive
thiols.
22
The majority of methods for profiling
reactive cysteine residues use the alkylating reagent at a single
concentration; however, a recent study by Weerapana et al. employed
a range of IAM concentrations and differential isotopic labeling to
identify reactive cysteines.
23
Identifying
low pK
a cysteine thiols affords a list
of proteins that are candidates for redox-mediated modification, but
additional studies are required to evaluate oxidant sensitivity.
Chart 2
Protein Thiols React with N-Ethylmaleimide (NEM, 1, Equation 1) and Iodoacetamide
(IAM, 2, Equation
2) by Michael Addition or SN2 Displacement, Respectively
3
Reactive
Oxygen Species (ROS) in Biological
Systems
Among biologically relevant and abundant ROS (Chart 3), superoxide (O2
•–) and
H2O2 appear most important in receptor-mediated
signaling. Although rates of cellular O2
•– production can be high, in most mammalian cells the steady-state
concentration is estimated to be in the low picomolar range (note
that cellular concentrations and half-lives for ROS are approximate
and can vary considerably depending on the cell type, nutritional
and environmental conditions, as well as the stage of the cell-cycle).
24
This is due to the rapid rate constant for spontaneous
dismutation of O2
•– to H2O2 and molecular oxygen (∼105 M–1 s–1) or as catalyzed by the superoxide
dismutase (SOD) enzyme family, which is 104 times as fast
(∼109 M–1 s–1).
25
In turn, antioxidant enzymes, such
as peroxiredoxin (Prx), catalase (CAT), and glutathione peroxidase
(GPx), maintain steady-state intracellular H2O2 levels in the nanomolar to low micromolar
range.
24b,26
Compared to other ROS in Chart 3, H2O2 is a mild oxidant and has the longest cellular
half-life
(∼1 ms).
2a,24b,26,27
Owing to its relative stability and selective
reactivity, H2O2 appears well suited for a second
messenger role.
Chart 3
Formation and Transformation of Biologically Relevant
Reactive Oxygen
Species (ROS)a
a
Superoxide (O2
•–), formed predominantly from the mitochondrial
electron transport chain and NADPH oxidase enzyme complexes (not shown),
is dismutated to hydrogen peroxide (H2O2) and
oxygen by superoxide dismutases (SOD). H2O2 is
in turn metabolized by catalases, peroxiredoxins, and glutathione
peroxidases. Additionally, H2O2, alone or in
concert with O2
•—, can react with
trace metal ions (Fe2+ or Cu+) to generate hydroxyl
radical (•OH) via Fenton or Haber–Weiss chemistry,
respectively. In phagosomes, H2O2 serves as
a substrate for myeloid peroxidase to produce hypochlorous acid (HOCl)
and water. Color intensity correlates to relative ROS reactivity.
The relative stability and uncharged nature of
H2O2 may permit its diffusion through membranes,
though this diffusion
would be less rapid than that of gases, such as nitric oxide (•NO) and hydrogen sulfide
(H2S). Recent studies
indicate that aquaporins, a family of small (24–30 kDa) pore-forming
integral membrane proteins, can also mediate H2O2 transport.
28
Underscoring its diffusible
nature and relative stability, H2O2 is known
to function as a mobile paracrine signal to regulate plant cell differentiation
29
as well as recruitment of immune cells for wound
healing in eukaryotes.
30
By contrast, the
negatively charged O2
•— does not
freely diffuse across membranes (though evidence for its translocation
via anion channels has been reported
31
).
The protonated form of O2
•– (HO2
• pK
a ≈
4.9) is membrane permeable but is only present in low amounts at physiological
pH (<0.2% at pH 7.4). Nonetheless, HO2
• may be relevant in phagocytes where O2
•– may reach a steady-state concentration of ∼25 μM.
32
H2O2 alone, or in
concert with O2
•–, can also react
with trace metal ions
(Fe2+ or Cu+) to generate the hydroxyl radical
(•OH) via Fenton or Haber–Weiss chemistry,
respectively (Chart 3).
33
Unlike O2
•– and H2O2, whose production and metabolism are regulated
processes, there are no known enzyme antioxidants for •OH neutralization. The •OH
is a strong oxidant
and reacts indiscriminately at diffusion-limited rates with protein,
DNA, and lipid biomolecules,
24b,34
which contributes to
its short cellular half-life (∼1 ns).
24b
In healthy cells, •OH formation is low since H2O2 metabolism and metal ion concentrations
are
both tightly regulated to avoid toxicity. Conversely, pathologies
that are associated with aberrant H2O2 metabolism
or the presence of adventitious uncomplexed metal ions are often associated
with increased •OH production and oxidative damage.
For instance, mutations in Cu,Zn-SOD linked to familial amyotrophic
lateral sclerosis (FALS) enhance •OH formation by
Fenton and Haber–Weiss reactions and contribute to motor neuron
degeneration.
35
3.1
ROS Production
and Metabolism
The
subsections below outline important biological sources of ROS, which
are formed as byproducts of respiration or by the action of enzymes.
Although our discussion is focused primarily on the initial species
generated by reduction of oxygen (O2
•– and H2O2) important secondary products, such
as hypohalous acids (HOX) are also briefly covered. The interested
reader is also directed to these sources for more information about
the regulation of ROS metabolism
26,36
and methods
for ROS detection.
37
3.1.1
Mitochondrial
Sources of ROS
The
mitochondrial electron transport chain (ETC) funnels electrons from
reduced metabolic components (NADH and FADH2) in the mitochondrial
matrix through four protein complexes (I–IV) in which molecular
oxygen serves as the terminal electron acceptor and is reduced to
water (Figure 1a). The energy released during
electron transfer is used to establish a proton gradient across the
inner mitochondrial membrane that is harnessed to drive the production
of the primary cellular energy source, adenosine-5′-triphosphate
(ATP) via ATP synthase (complex V). This is an imperfect system, however,
and electrons can leak prematurely from the ETC at complexes I and
III resulting in the univalent reduction of molecular oxygen to O2
•– in either the matrix (complex
I and III) or the intermembrane space (complex III) (Figure 1a).
26,38
It is estimated that 0.15–2%
of molecular oxygen consumed is converted to O2
•– by the mammalian ETC.
38b,39
While this figure may
seem low, mammals consume a large amount of oxygen resulting in the
constitutive production of a significant amount of O2
•– (and H
2
O2 through O2
•– dismutation). For
example, mutant mice lacking mitochondrial manganese-SOD (Mn-SOD)
exhibit neonatal lethality resulting from neurodegeneration and cardiomyopathy,
which may be rescued by small-molecule scavengers of O2
•–.
40
Deletion
of individual SOD genes is also detrimental to bacteria
41
and yeast
42
survival
further highlighting the impact of O2
•– production in the ETC. Clearly, mitochondria are significant contributors
to cellular H2O2 generation by dismutation of
O2
•– from the ETC.
Figure 1
Biological sources of
reactive oxygen species (ROS). (a) The mitochondrial
electron transport chain (ETC). Four protein complexes (I–IV)
funnel electrons (black arrows) from NADH and succinate in the matrix
to ultimately reduce molecular oxygen to water and establish a proton
gradient (gray arrows) that is harnessed by complex V to generate
ATP. Electrons can leak prematurely from the ETC at complexes I and
III (red arrows) to generate superoxide (O2
•–) in either the matrix or intermembrane space. (b) p66 (Shc) facilitates
pro-apoptotic O2
•– or H2O2 production in the mitochondria. In response to UV irradiation
or growth factor deprivation, p66 (Shc) localizes to the mitochondria
where it interacts with complex III to divert electrons from cytochrome
c directly to molecular oxygen to generate O2
•– or H2O2. This H2O2 can
translocate to the cytoplasm (not shown) where it can influence signaling,
and can regulate opening of the mitochondrial permeability transition
pore (mPTP), which initiates mitochondrial swelling and apoptosis.
(c) NOX enzyme complexes assemble at distinct regions of the plasma
membrane or intracellular membranes to regulate localized ROS production
in response to diverse signals. Receptor stimulation initiates the
recruitment of specific coactivating proteins or calcium to one of
seven NOX catalytic cores. Once activated, NOX enzymes funnel electrons
from NADPH in the cytoplasm through FAD and heme cofactors across
the membrane to generate O2
•– (NOX1-2)
or H2O2 (Duox1-2) on the extracellular/lumenal
face. O2
•– is dismutated to H2O2 and oxygen either spontaneously or as enhanced
by SOD, which can translocate across the membrane by diffusion or,
more likely, through aquaporin channels to regulate protein activity
and signaling in the cytoplasm.
The amount of mitochondrial-derived O2
•– is variable
43
and regulated by a number
of factors, such as oxygen concentration, proton motive force,
44
ETC efficiency,
45
and the availability of electron donors. Pathologies that include
neurodegenerative disorders, cancer, and diabetes are associated with
mitochondrial dysfunction and enhanced ROS production.
46
Mitochondrial stress and ROS-dependent AMP kinase
activation have also been implicated in maternally inherited hearing
loss.
47
Recent studies in mice and yeast
have revealed an evolutionarily conserved mechanism that cells use
to control mitochondrial O2
•– production.
48
This is accomplished by adjusting the flux through
metabolic pathways that regulate the flow of electrons into the ETC.
Interestingly, these studies show that ROS-dependent inactivation
of pyruvate kinase or a switch in isoform expression can redirect
metabolic flow through the pentose phosphate pathway, which makes
the reduced nicotinamide adenine dinucleotide phosphate (NADPH) required
to maintain cellular redox homeostasis.
Extrinsic and intrinsic
signals can also regulate mitochondrial
O2
•– production. This process
is strictly dependent on the adaptor protein p66(Shc), which regulates
the level of ROS, apoptosis induction, and lifespan in mammals.
49
Cell signals including growth factor deprivation,
oxidative stress, or UV irradiation induce translocation of p66(Shc)
into the mitochondria where it promotes electron transfer from Complex
III directly to oxygen, enhancing O2
•— production (Figure 1b).
50
After conversion to H2O2 through
dismutation, this ROS diffuses into the cytoplasm where it decreases
the activity of FoxO3, a transcription factor that regulates the expression
of mitochondrial antioxidant enzymes, including Mn-SOD and catalase.
51
The reduction in antioxidant capacity further
increases mitochondrial oxidative stress and enhances the pro-apoptotic
function of p66(Shc).
52
Of note, mutant
Mn-SOD heterozygous knockout mice exhibit marked sensitization of
the mitochondrial permeability transition pore (mPTP) and premature
induction of apoptosis.
53
Mice lacking
p66(Shc) live ∼30% longer and show increased resistance to
oxidative stress and age-related pathologies, marking it as a potential
therapeutic target for diseases that are associated with oxidative
damage.
26,49,50,54
Several studies suggest an additional role for mitochondrial
ROS in immune system function.
55
For instance,
a recent report demonstrated recruitment of mitochondria to phagosomes
in infected activated murine macrophages and that mitochondrial-derived
ROS was required for microbial killing.
56
Mice lacking p66(Shc) also exhibit decreased O2
•– production in macrophages, highlighting another potential role for
p66(Shc)-regulated mitochondrial ROS production.
57
Although a thorough review of the plant literature
in this area
is beyond the scope of this review, we would be remiss if we did not
note that in plant cells O2
•– is
also produced in the mitochondria by the ETC, as well as other subcellular
compartments, such as chloroplasts and peroxisomes through photorespiration.
58
The amount of ROS generated via photorespiration
can increase in response to environmental constraints, including biotic
and abiotic stresses. The interested reader is referred to the following
extensive reviews for additional information on this topic.
59
3.1.2
Enzymatic Generation
of ROS
In
addition to mitochondrial sources of O2
•–, this reactive intermediate can be generated as a byproduct during
the catalytic cycle of numerous enzymes, such as “nonspecific”
peroxidases (i.e., haem-containing peroxidases capable of using H2O2 to oxidize a
range of substrates), as well as
xanthine and aldehyde oxidases.
3a,60
Electron leakage from
NADPH cytochrome P450 reductases present in the endoplasmic reticulum
(ER) can also generate O2
•– during
hormone and drug metabolism.
61
The autoxidation
of glyceraldehydes, reduced flavin mononucleotide (FMNH2), and reduced flavin adenine
dinucleotide (FADH2) can
also produce O2
•–, albeit with
slow reaction kinetics.
24b,24c
As noted above, the
dismutation of O2
•– provides a
major source of H2O2 in cells. In addition,
there are numerous enzymes that produce H2O2 without the intermediacy of O2
•–, including xanthine, glucose, lysyl, monoamine, and d-amino
acid oxidases, as well as the peroxisomal pathway for beta-oxidation
of fatty acids.
62
The contribution of these
sources of O2
•– and H2O2 to redox signaling remains to be determined.
In activated phagocytes of the immune system, myeloperoxidase- and
eosinophil peroxidase-catalyzed oxidation of halide (Cl–, Br–, I–) and pseudohalide
(SCN–) ions converts H2O2 to the corresponding
hypohalous acid (HOX), such as hypochlorous acid (HOCl) (Chart 3).
2d,32e,63
HOXs react preferentially with thiols and methionine residues and
these potent oxidants are generally believed to be responsible for
much of the bactericidal activity of neutrophils. The reaction of
HOCl with O2
•– is also known to
generate •OH and is proposed to serve as the primary
source of •OH in neutrophils.
64
The interested reader is referred to the following sources
for additional information on this unique class of oxidants.
2d,32e
A variety of extracellular signals including peptide growth
factors,
cytokines, and G-protein-coupled receptor (GPCR) agonists and, more
recently, mechanical distortion in cardiomyocytes
65
trigger deliberate production of ROS through activation
of NADPH oxidase (NOX) complexes.
66
NOX-derived
ROS is required for propagation of many pathways
12,65,67
and the maintenance of essential stem cell
populations in the brain.
68
NOX complexes
produce ROS with one of seven enzymatic cores (NOX1-5, Duox1, and
Duox2) that exhibit differential cell- and tissue-specific expression
patterns. As illustrated in Figure 1c, activation
of NOX requires association of a flavin adenine dinucleotide (FAD)
cofactor, distinct membrane and cytoplasmic coactivator proteins (Nox1-4,
Duox1, and Duox2) or binding of calcium to the intracellular domain
(Nox5, Duox1 and Duox2).
36a,36b,36d
As follows, NOX activation can be tightly controlled by signal-mediated
recruitment of these coactivating proteins
69
or cofactors,
69c,70
which are likely to be pathway-
and isoform-specific.
The activated NOX transports an electron
from cytoplasmic NADPH
through FAD and heme cofactors across plasma and intracellular membranes
to produce O2
•– on the extracellular/lumenal
face (Figure 1c).
36a,36b,36d,71
O2
•– is then dismutated to H2O2 and molecular oxygen, either spontaneously or
via extracellular SOD,
72
though some NOX
isoforms (Duox1 and Duox2) are equipped with an extracellular peroxidase
domain that is believed to directly mediate two-electron reduction
of molecular oxygen to H2O2.
73
Translocation of electrons from the cytoplasm across biological
membranes with the concomitant release of protons from NADPH results
in local acidification proportional to oxidant production. In neutrophil
phagosomes, where NOX2 is estimated to produce O2
•– at steady-state levels of 25 μM,
32d
sustained NOX2 activity is coupled to voltage-gated proton channels
to mitigate local acidification.
74
A similar
dependence on a voltage-gated proton channel has been demonstrated
for prolonged NOX activation in active B cells.
75
The efflux of electrons also results in net positive charge
accumulation on the ROS-producing face, which may promote electron
transfer through NOX. Recently, a nonselective cation (Ca2+, Na+, K+) channel called,
TRPM2, was shown
to be activated by NOX-derived ROS.
76
TRPM2
activation depolarized the plasma membrane, which dampened NOX-mediated
ROS production in phagosomes. This finding presents a novel mechanism
by which cells can regulate the amplitude and duration of NOX activity.
Within a given signaling pathway, identifying which NOX isoform
is acting as the primary ROS source is usually accomplished by determining
the relative expression level of each isoform using isoform-specific
antibodies
12
or by overexpressing the isoform
of interest.
77
However, inherent differences
in antibody affinity and specificity issues can complicate these determinations,
and protein overexpression does not reflect native conditions. Many
cell types express multiple NOX isoforms, making it difficult to discern
isoform-specific roles in a given signaling pathway, as knockout or
siRNA knockdown studies are not always feasible. The participation
of NOX in a given signaling pathway is commonly assessed using a number
of small molecule inhibitors, including apocynin or the flavin analog,
diphenyleneiodonium (DPI). These results should be interpreted with
caution, as both compounds have been shown to have off-target effects
in some cell types.
78
Isoform-specific
NOX inhibitors would greatly assist in dissecting the role of individual
NOX family members in signaling pathways.
79
For example, a peptide inhibitor that is highly specific for NOX2
has been used to study its role in vascular O2
•– production in mice
80
and during mechanical
distortion in cardiomyocytes.
65
High-throughput
screens have also identified small-molecule inhibitors of NOX1
81
and NOX2.
82
H2O2 that results from NOX activation can
enter the cytoplasm through diffusion, or as recently shown, by transport
through aquaporin channels where it can mediate distinct physiological
responses, such as proliferation, differentiation, and apoptosis.
26,83
Since H2O2 that is produced extracellularly
or in the luminal space must enter the cytoplasm to modulate intracellular
signaling pathways, one key question is how can its effects be localized?
Much remains to be understood about this important aspect of redox
signaling, however, one possible answer is that aquaporins are directed
to lipid raft membrane microdomains
84
that
are also enriched for NOX. Indeed, NOX isoforms are both temporally
and spatially localized to distinct membrane regions via lipid rafts,
36b
activated receptors,
12,70
and focal adhesions.
85
Depending on the
stimuli and cell type, NOX family members also localize to distinct
subcellular compartments, such as the ER
86
and nucleus.
87
As will be discussed in
more detail below, the localized activities of NOX, as well as antioxidant
enzymes that metabolize ROS may also help restrict H2O2 to regions where signaling
proteins are similarly localized.
3.1.3
ROS-Metabolizing Enzymes
As stated
above, dismutation of O2
•– by
SOD produces H2O2. The peroxiredoxin (Prx) and
glutathione peroxidase (GPx) families are primarily responsible for
the metabolism of H2O2 in cells. These enzymes
decompose H2O2 to form water and molecular oxygen
in a mechanism involving the oxidation of an active site cysteine
(or selenocysteine in GPxs from higher eukaryotes).
88
The enzymes are recycled back to their active, reduced
form by thioredoxin/thioredoxin reductase (Trx/TrxR) or glutathione/glutathione
reductase (GSH/GR) systems using reducing equivalents from NADPH.
Another H2O2-metabolizing enzyme, known as catalase,
is present mainly in peroxisomes. Plants synthesize high concentrations
of ascorbate,
59b
which is used as a substrate
by ascorbate peroxidases to regulate H2O2 bioavailability
in these systems.
89
Ascorbate peroxidases
are subsequently reduced by a complex metabolic pathway, known as
the glutathione-ascorbate cycle.
90
A growing
list of antioxidant enzymes, including Prxs, are themselves subject
to redox regulation, which could permit localized accumulation of
H2O2 for signaling while simultaneously limiting
the range of H2O2 diffusion.
11b,91
3.2
Modification of Protein Cysteine Thiols by
ROS
The reaction of ROS with protein thiols provides a mechanism
by which cells can “sense” changes in the redox balance.
Though H2O2 is most often associated with a
second messenger role, there is also evidence to suggest that O2
•– functions in this capacity. For
instance, a recent study demonstrated that disparate gradients of
O2
•– and H2O2 differentially regulated plant root proliferation and differentiation,
respectively implicating distinct activities for these ROS.
92
O2
•– is
a relatively unreactive radical and its primary cellular targets appear
to be other radical species, such as nitric oxide (•NO) or metals. In proteins, O2
•– can react with iron–sulfur clusters and heme centers leading
to release and/or oxidation of iron.
13
Numerous
iron–sulfur cluster- and heme-containing proteins are sensitive
to O2
•–, including aconitase,
93
the bacterial transcription factor SoxR,
94
guanylate cyclase,
95
and myeloperoxidase.
96
Reactivity at
protein metal centers is not unique to O2
•–, however, as metal-dependent peroxide sensors like Bacillus
subtilis PerR have also been reported.
2a,83c,97
In contrast to redox switches based on peroxide-sensitive
cysteine residues, PerR senses H2O2 by metal-catalyzed
oxidation of histidine residues involved in coordinating Fe2+ (note that the mechanism
involves reduction of H2O2 by Fe2+ to generate •OH, which
then reacts rapidly with histidine). H2O2 may
also modify tryptophan and tyrosine residues through a radical-based
mechanism, but such reactions are much less favored and may not be
physiologically relevant.
98
H2O2 can directly oxidize the thioether group of
methionine to yield two diastereomeric methionine sulfoxide products;
99
however, a large body of evidence identifies
cysteine as the most sensitive amino acid residue to H2O2-mediated oxidation. The
two-electron oxidation of a
thiolate by H2O2 yields sulfenic acid, which
is increasingly implicated in a number of important biochemical transformations.
Second-order rate constants for this reaction can vary dramatically
in proteins (e.g., 20–107 M–1 s–1).
14
Once formed, the sulfenic
acid is subject to several alternative fates (Figure 2). Depending on the microenvironment,
the sulfenic acid modification
can be stabilized as observed in human serum albumin (HSA)
100
and more than 40 protein crystal structures.
9b,101
In this regard, there are several factors that appear to stabilize
protein sulfenic acids, including the absence of thiols proximal to
the site of formation or inaccessibility to low-molecular-weight thiols,
such as GSH (γ-l-Glu-l-Cys-Gly).
3b
Reaction of sulfenic acid with a protein thiol
or GSH yields an inter/intramolecular disulfide bridge or protein-S-GSH disulfide,
respectively. Alternatively, in some proteins
lacking a neighboring cysteine, a nitrogen atom of a backbone amide
can react with sulfenic acid, forming a cyclic sulfenamide.
102
The formation of disulfide and sulfenamide
states protects against irreversible overoxidation, as S–S
and S–N bonds can be reduced through the activity of Trx/TrxR
or GSH/glutaredoxin (Grx)/GR systems.
103
Sulfenic acid can also be reduced directly by the Trx system, through
hydride transfer (H–) from FADH2 in a
reaction catalyzed by NADH oxidase and NADH peroxidase enzymes from Streptococcus
faecalis,
104
or
through the DsbD/DsbG system in the bacterial periplasm.
105
In the presence of excess H2O2, sulfenic acid can be further oxidized to sulfinic
(RSO2H) and sulfonic (RSO3H) oxyacids, though the observed
rate constants for such reactions are generally slower (0.1–100
M–1 s–1) than the initial thiolate
oxidation event (Figure 2).
15,104b,106
Figure 2
Oxidative modification of cysteine residues
by hydrogen peroxide
(H2O2). The initial reaction product of a low
pK
a protein thiolate with H2O2 yields a sulfenic acid, whose stability is determined,
in part, by its accessibility to additional thiols. Reaction with
a second cysteine in the same or neighboring protein yields a disulfide.
Alternatively, reaction with the low molecular weight thiol, glutathione
(GSH) affords a specialized mixed disulfide called a glutathione disulfide.
In some proteins in which a neighboring cysteine is not present, nucleophilic
attack of a backbone amide on the sulfenic acid yields a cyclic sulfenyl
amide. Each of these oxoforms can be reduced by the GSH/glutaredoxin
or thioredoxin/thioredoxin reductase systems to regenerate the reduced
thiolate (not shown). In the presence of excess H2O2, such as under conditions of
oxidative stress, the sulfenic
acid can be hyperoxidized to the largely irreversible sulfinic and
sulfonic acid forms (red box).
HOXs, such as HOCl, also mediate two-electron oxidation of
cysteine.
These reactions proceed through X+ transfer to give an
unstable sulfenyl halide, which rapidly hydrolyzes to sulfenic acid
(>107 M–1 s–1 for
HOBr
and HOCl).
107
HOXs are aggressive oxidants
and halogenating agents, which react with a wide range of cellular
targets, including methionine, histidine, tryptophan, lysine, tyrosine,
the protein backbone, nucleic acids and fatty acids. On the whole,
the modifications of biomolecules that are mediated by HOX are numerous
and highly damaging, which makes these oxidants highly effective toxic
defense molecules that can be exploited by the human immune system
to fight off microbial infection. As a final comment in this section,
we note that the oxidation of cysteine thiols can also occur by one-electron
redox pathways to give thiyl radicals, which undergo distinct sets
of reactions. These transformations are briefly discussed in section 5 below (Reactive
Sulfur Species (RSS) in Biological
Systems) and we also refer the interested reader to the following
sources for additional information.
3a,14,108
3.3
Methods for Detecting ROS-Modified
Cysteines
The reversible nature of cysteine sulfenic acid,
disulfide and S-glutathionylation makes them well
suited to control protein
function during cell signaling. With the discovery of Sulfiredoxin
(Srx) proteins,
109
which can convert the
sulfinic acid modification back to the thiol form, cysteine sulfinic
acids have also emerged as a potential regulatory mechanism. Consequently,
there has been considerable effort to develop methods to study changes
in protein cysteine oxPTM. These techniques include indirect and direct
methods for detection. The majority of indirect methods to detect
cysteine oxidation rely upon the loss of reactivity with thiol-modifying
reagents (Figure 3a) or restoration of labeling
by reducing agents such as dithiothreitol (DTT) (Figure 3b). The latter method requires
a complete blocking of free
thiols with alkylating agents prior to the reduction step and is therefore
limited to studies in cell lysates or with purified proteins.
Figure 3
General overview
of indirect and direct chemical methods to study
protein oxidation. (a) Loss of labeling of oxidized thiols by an alkylating
agent indirectly monitors protein oxidation. In response to oxidant
treatment, susceptible cysteines are oxidized (purple) and thus are
less reactive with alkylating agents such as NEM or IAM. Use of a
biotinylated or fluorophore-conjugated alkylating agent permits detection
by avidin blot or in-gel fluorescence, in which oxidized proteins
exhibit a loss of signal. (b) Differential alkylation of reduced and
oxidized thiols indirectly monitors protein oxidation. Free thiols
(blue) are blocked with an alkylating agent such as NEM or IAM, reversibly
oxidized thiols (purple) are reduced with a reducing agent such as
DTT or TCEP, and nascent thiols are labeled with a second alkylating
agent conjugated to biotin or a fluorophore. Oxidized proteins exhibit
enhanced signal by avidin blot or in-gel fluorescence. (c) Direct
chemical method to detect specific cysteine oxoforms. Samples are
treated with a biotin or fluorophore-conjugated probe that selectively
reacts with a distinct cysteine chemotype (purple) in which signal
by avidin blot or in-gel fluorescence increases with increased protein
oxidation.
More recently, chemical
biology approaches have facilitated the
development of small molecule- and protein-based methods for direct
detection of distinct oxidative cysteine modifications (Figure 3c). In the event that
these small molecules are
cell permeable, specific cysteine modifications can be detected directly
in their native environment without cell disruption (i.e., lysis).
This is an attractive approach since it preserves labile cysteine
modifications and maintains the integrity of subcellular organelles.
The latter is especially important as organelles like the nucleus,
mitochondria, and cytoplasm have more reduced redox potentials whereas
the secretory system and the extracellular space are more oxidizing
environments.
110
Not surprisingly, cell
lysis disrupts these individual redox environments and can result
in substantial protein oxidation artifacts. The net result is to increase
the challenges related to detecting low abundance modifications and
in deciphering their biological significance. Likewise, cell disruption
can hamper the detection of labile or transient cysteine modifications.
Methods to decrease oxidation artifacts in lysates have been reported,
but these are often dependent upon the addition of trichloroacetic
acid (which denatures proteins and can lead to acid-catalyzed overoxidation
of labile modifications such as sulfenic acid) or on the addition
of ROS-metabolizing enzymes to the lysis buffer.
111
Even with these considerations, lysis buffers can never
accurately mimic the intracellular redox potential, thereby exposing
redox-sensitive proteins to oxygen and a different redox environment.
Direct detection methods may also be associated with their own limitations
as the addition of a small-molecule probe to cells could alter the
biological function under investigation. This issue can be addressed,
at least in part, by adding the probe to cells after signal pathway
activation and/or by monitoring the effect of probe addition on relevant
downstream biological markers.
12
Another
important consideration with direct detection methods is the rate
at which probes react with the modified cysteine residue. If the reaction
is slow, transient cysteine oxidation events may be missed. Conversely,
if the reaction is too fast it could diminish the chemical selectivity
of the probe or disrupt the biological process under study. In this
way, moderately reactive probes for detecting individual oxidative
cysteine modifications may be viewed as “spectators”,
which sample the redox-signaling environment with minimal biological
impact. Increasing the concentration of probe can also compensate
for modest rates of reaction, but appropriate controls must be performed
to ensure that the underlying biology is not disturbed.
Collectively,
indirect and direct methods to monitor cysteine oxidation
have enabled the discovery of many proteins that can undergo redox
modification in a wide range of organisms and different cell types.
To highlight the progress made over the past few years in the redox
biology field, the following subsections will independently address
the chemical properties of ROS-mediated cysteine modifications and
methods for their detection. We also discuss selected examples from
the recent literature that highlight the ways in which distinct cysteine
modifications can mediate critical biological events.
3.3.1
Indirect Approaches for Detecting ROS-Sensitive
Cysteines
Several methods have been developed to monitor
global changes in cysteine oxidation, but do not reveal the chemical
nature of the modification. One of the most commonly used reagents
for this purpose is the BIAM alkylating reagent. In these experiments,
the diminished nucleophilicity of the oxidized cysteine residue results
in lower reactivity with BIAM and correlates with a loss of protein
labeling (Figure 4a). An adaptation of this
methodology that permits simultaneous identification and quantification
of oxidant-sensitive cysteine thiols employs an acid-cleavable BIAM-based
isotope-coded affinity tag (ICAT).
112
In
this method, free thiols are differentially alkylated with isotopic
versions of the ICAT reagent and the extent of cysteine oxidation
is determined by the ratio of light (12C) and heavy (13C) ICAT label by LC-MS/MS (Figure
4b).
Figure 4
Indirect chemical methods to study general cysteine oxidation.
(a) Loss of labeling of oxidized thiols by biotinylated-iodoacetamide
(BIAM) indirectly monitors protein oxidation. Oxidized cysteines (purple)
exhibit decreased reactivity with BIAM than reduced thiols, and are
observed as a loss of signal by avidin blot. (b) Isotope-coded affinity
tag (ICAT) reagents determine the ratio of oxidized thiols. Samples
are untreated or subjected to oxidant. Free thiols are subsequently
labeled with a light (12C) ICAT reagent in the untreated
sample and with a heavy (13C) ICAT reagent in the oxidant-treated
sample. As in panel a, reactive thiols (purple) exhibit decreased
labeling upon oxidation. The samples are mixed, trypsinized, and enriched
via the biotin affinity tag on the ICAT reagent. Eluted peptides are
analyzed by LC-MS and heavy and light ICAT-labeled peptides are chemically
identical, but differ in mass by 9 Da. The fraction of a thiol oxidized
in the sample is determined by the ratio of heavy (13C)
to light (12C) signal intensity, whereby thiols that are
susceptible to oxidation (purple) will exhibit decreased signal intensity
with the heavy ICAT reagent.
A subsequent alternative approach incorporates treatment
with a
reducing agent into the workflow (Figure 3b).
Such protocols require free thiol alkylation, a reduction step with
DTT or tris(2-carboxyethyl)phosphine (TCEP), and labeling of nascent
thiols with a tagged alkylating agent, such as BIAM. In this approach,
changes in cysteine oxidation are detected as differences in sample
BIAM alkylation as assessed by avidin blot and oxidized proteins can
be identified by enrichment and LC-MS/MS analysis (Figure 3b). In addition to BIAM,
alternative biotinylated
or fluorophore-modified alkylating reagents can be used to differentially
alkylate thiols and these methodologies have been used to monitor
protein oxidation in response to exogenous oxidants (e.g., H2O2 or diamide)
21,113
or to ROS-promoting
stimuli (e.g., peptide growth factors).
114
A similar workflow has also been used to identify substrates of
the Trx/TrxR and GSH/Grx/GR systems.
113
Alternatively, protein substrates of the aforementioned reducing
systems can be identified through their inclusion in the reduction
step.
21
For instance, BIAM-alkylated nascent
thiols will represent oxidized proteins that were selectively reduced
by the Trx/TrxR or GSH/Grx/GR systems. Together with the ICAT technology,
this method has been used to identify protein disulfide targets of
the Trx/TrxR system in plant extracts.
115
In addition to studying the oxidized proteome, changes in total
thiol content in protein and low molecular weight thiols, including
GSH and homocysteine, can be indicative of fluctuations in biological
redox balance and, in some cases, serves as a diagnostic function
for disease. In this vein, an active area of research is the development
of sensitive probes to monitor fluctuations in total thiol content.
116
3.3.2
Direct and Selective
Approaches for Detecting
ROS-Sensitive Cysteines
3.3.2.1
Disulfides
Disulfide bond formation
in proteins is a widely recognized cysteine modification that has
important roles in protein folding and stability. Under normal cellular
conditions, disulfide bond formation occurs largely in the extracellular
space or the endoplasmic reticulum (ER). In this organelle, a class
of enzymes called protein disulfide isomerases (PDI) inserts disulfides
into nascent proteins that are destined for export to the extracellular
milieu.
117
By comparison, disulfide bonds
are rare and generally transiently formed in the cytoplasm, mitochondria,
or nucleus where thiol-dependent reductases maintain a reducing environment.
Exceptions exist, however, as the sulfhydryl oxidase Erv1 and oxidoreductase
Mia40 form a relay system that introduces disulfide bonds in substrate
proteins in the mitochondrial inner membrane.
118
Under oxidative stress conditions the intracellular redox
balance can shift to support disulfide bond formation in reducing
compartments until redox homeostasis is restored.
A major route
of disulfide formation is by thiol condensation with sulfenic acid
(Figure 2). These processes can occur either
intra- or intermolecularly, and the rate of disulfide bond formation
is dependent, in part, upon the distance between the two cysteine
residues. Estimated rate constants for intra- and intermolecular disulfide
bond formation are 10 s–1 and 105 M–1 s–1, respectively.
119
Once formed, disulfides are relatively stable
to most physiological nucleophiles and are generally cleaved by other
thiols as in thiol-disulfide exchange (nucleophilic substitution)
reactions (Figure 5).
120
The thiol in a disulfide with the lower pK
a will be the better leaving group and often dictates which
cysteine is released in thiol-disulfide exchange. Indeed, this strategy
is employed by the thiol-disulfide exchange catalysts in the cell,
such as protein disulfide isomerases (PDI).
121
Disulfides can also be oxidized to generate a thiosulfinate, which
can subsequently react with a thiol to give disulfide and sulfenic
acid products (Figure 5). The prevalence or
biological significance of the thiosulfinate is unknown, however,
it is interesting to note that this species forms as an intermediate
during Srx-catalyzed sulfinic acid reduction of Prxs.
122
Although the intermediate thiosulfinate is
formed via a mechanism distinct from disulfide oxidation, its formation
implies that the thiosulfinate may be a physiologically relevant,
yet understudied modification. Further oxidation of a disulfide yields
a thiosulfonate (Figure 5), which releases
a disulfide and sulfinic acid subsequent to reaction with a thiol.
Thiosulfonates have not been detected in cells, but could possibly
be formed as an enzyme intermediate in sulfonic acid reduction akin
to sulfinic acid reduction via sulfiredoxin, though an enzyme capable
of catalyzing such a reaction is currently unknown.
3a
Figure 5
Possible fates of protein disulfides. Once formed, a protein disulfide
(inter- or intramolecular) can undergo thiol-disulfide exchange with
a third cysteine within the same or neighboring protein (eq 1). Herein,
pK
a of the disulfide thiols and thiol
accessibility influence which cysteine is expelled. In the presence
of high concentrations of H2O2, disulfides can
additionally be oxidized to the thiosulfinate and thiosulfonate forms,
though these reactions are very slow. Because of the potential for
resonance stabilization or decreased pK
a, subsequent reaction of these intermediates with a third cysteine
affords a disulfide and a sulfenic acid (eq 2) or sulfinic acid (eq
3). The biological relevance of the thiosulfinate and thiosulfonate
modifications is unknown due to a lack of means to study these oxoforms,
however, a thiosulfinate forms as an intermediate during the sulfiredoxin
catalytic cycle.
Global studies to identify
proteins that undergo disulfide bond
formation implicate this modification in the regulation of, among
others, redox homeostasis, chaperone activity, metabolism, transcriptional
regulation, and protein translation.
111b,113
Once formed,
disulfides can impact enzyme activity, subcellular localization, as
well as protein–protein interactions.
71
For example, the activity of certain PTPs is inhibited by disulfide
bond formation involving the active site cysteine and the so-called
backdoor cysteine.
106b,123
This regulatory mechanism is
also observed in certain members of the caspase family of cysteine
proteases.
124
Numerous studies have demonstrated
an increase in protein phosphorylation in response to receptor activation
that is dependent upon endogenous H2O2 production.
12,65,67,68
Owing to this observation and their conserved catalytic cysteine
residue, PTPs were initially proposed as the major cellular targets
of signaling-derived H2O2.
125
Kinases are now also believed to be redox regulated, though
in many cases the molecular details are much less well characterized.
Nonetheless, it has been established that serine/threonine kinases
PKG1α
126
and ATM
127
are activated by intermolecular disulfide formation between
homodimers that, in the case of PKG1α, enhances its affinity
for target proteins. By contrast, intermolecular disulfide formation
between Src tyrosine kinase monomers appears to inhibit kinase activity,
128
though Src has also been shown to be activated
by H2O2.
129
Differential
regulation by H2O2 may be explained, in part,
by modification of multiple cysteine residues. For example, oxidative
inhibition of Src involves Cys277, which is not conserved in all Src
family kinases.
128
The Src-family kinase
Lyn, which encodes a glutamine at the site corresponding to Cys277,
is activated by ROS in neutrophils suggesting that oxidative activation
of this enzyme involves a different cysteine residue.
30e
Additional proteins whose activity have recently
been shown to be modulated by disulfide bond formation include the
bacterial chaperone Hsp33,
130
the nonspecific
cation channel TRPA1,
131
and the glycolytic
enzyme pyruvate kinase M2 (PKM2).
48a
Disulfide bond formation can also influence the subcellular localization
of a protein and/or protein–protein interactions. For example,
intramolecular disulfide formation in the Saccharomyces cerevisiae transcription factor
Yap1 induces a conformational change that masks
the nuclear export signal (NES) and precludes interaction with the
nuclear export receptor, Crm1. This results in nuclear accumulation
of Yap1 and active transcription of genes involved in the oxidative
stress response.
132
Intramolecular disulfide
formation in the small molecular chaperone, DnaJb5 and the class II
histone deacetylase, HDAC4 results in sequential dissociation of the
DnaJb5-HDAC4 complex, unmasking of the HDCA4 NES to mediate its cytoplasmic
localization and derepression of target genes involved in hypertrophy
(Figure 6a).
71,133
A recent
study by Shacter and colleagues indicates that oxidative stress-induced
formation of two intramolecular disulfides in the actin-regulatory
protein, cofilin leads to dissociation of the actin-cofilin complex.
Additionally, oxidation of cofilin enables its mitochondrial accumulation
(by an unresolved mechanism) where it can interact with the mPTP to
promote mitochondrial swelling, cytochrome c release, and ultimately
induction of apoptosis (Figure 6b).
134
Figure 6
Disulfide-mediated redox regulation of subcellular localization
and protein–protein interactions. (a) Model for redox regulation
of cardiac hypertrophy by HDAC4. The class II histone deacetylase
HDAC4 normally deacetylates histones to suppress expression of genes
involved in cardiac hypertrophy. Nuclear localization of HDAC4 is
mediated by its association with importin α (Imp) via a multiprotein
complex including the small molecular chaperone DnaJb5, the thioredoxin
binding protein TBP-2, and thioredoxin (Trx1). In response to oxidant,
HDAC4 and DnaJb5 undergo intramolecular disulfide bond formation,
which causes dissociation and nuclear export of the complex permitting
derepression of genes involved in hypertrophy. Upon removal of H2O2, Trx1 is believed
to reduce the disulfides in
HDAC4 and DnaJb5 to restore assembly and nuclear localization of the
complex (not shown). (b) Model for redox regulation of apoptosis by
cofilin. Cofilin associates with actin in the cytoplasm to disassemble
actin filaments for cytoskeletal reorganization. In the presence of
H2O2, two intramolecular disulfides form in
cofilin permitting its relocation to the mitochondria by an unresolved
mechanism. In the mitochondria, cofilin interacts with the mPTP to
stimulate pore opening, mitochondrial swelling, cytochrome c release,
and ultimately induction of apoptosis.
Methods to detect protein disulfide formation often use reducing
and nonreducing SDS-PAGE gel electrophoresis (Figure 7a). Intermolecular disulfides
are detected as reducing agent-sensitive
protein complexes that migrate at a molecular mass equal to the that
of the two oxidized proteins, as seen for PKG1α,
126
Src,
128
and ATM
127
dimers (Figure 7a, right).
Intramolecular disulfide bond formation can also lead to altered migration
on gels, as observed for S. cerevisiae thiol peroxidase
Gpx3,
66c,135
PKM2,
48a
or PTEN
(Figure 7a, left).
123b
Cysteine residues involved in disulfide bond formation can also
be identified by the differential alkylation-type approach mentioned
above. In this method, thiols are alkylated prior to sample separation
by nonreducing SDS-PAGE; the protein band corresponding to the oxidized
proteins of interest is then reduced in-gel with DTT or TCEP, and
nascent thiols are labeled with a second alkylating agent. The protein
is then digested in-gel and the differentially alkylated cysteine
residues are identified by LC-MS/MS analysis.
127,134
Figure 7
Methods
for detection and identification of protein disulfides.
(a) Differential migration of proteins containing intra- and intermolecular
disulfide bonds. Samples are resolved under nonreducing SDS-PAGE conditions.
Intramolecular disulfides can facilitate enhanced protein migration
in some proteins as compared to the reduced species (left). Intermolecular
disulfide complexes migrate at the combined molecular weight of the
individual proteins (right). (b) Redox 2D-PAGE. Protein samples are
first separated by nonreducing gel electrophoresis to separate disulfide-bonded
complexes by size (top). The proteins are subsequently reduced in-gel
with DTT, alkylated with NEM or IAM, and separated in the second dimension
under reducing conditions (down). Proteins that are not involved in
intermolecular disulfide complexes run at the diagonal. Proteins involved
in disulfide complexes migrate off the diagonal and can be identified
by in-gel digestion and LC-MS/MS (not shown). (c) OxICAT method combines
the ICAT technology with differential alkylation of reduced and oxidized
thiols to permit quantification of oxidized residues. Cell lysates
are generated in the presence of trichloroacetic acid and detergents
to facilitate exposure of all protein cysteines while inhibiting thiol/disulfide
exchange. Reduced thiols (blue) are subsequently blocked with the
light (12C) ICAT reagent (blue), oxidized proteins (purple)
are reduced with TCEP, and nascent thiols are alkylated with the heavy
(13C) ICAT reagent (purple). Samples are trypsinized and
labeled peptides are avidin enriched. Eluted peptides are analyzed
by LC-MS and heavy and light ICAT-labeled peptides are chemically
identical, but differ in mass by 9 Da. The percentage of a particular
thiol that is oxidized in a sample is determined by the ratio of heavy
(13C) to light (12C) signal intensity from the
corresponding peptide. While TCEP can reduce all reversible oxoforms
(e.g., disulfides, sulfenic acid, S-nitrosothiols),
sulfenic acids and S-nitrosothiols are often acid-labile
and likely lost during sample preparation. As such, OxICAT is likely
most suitable to detect cysteines involved in disulfide bonds.
The differential migration of
disulfide-containing proteins by
nonreducing and reducing gel electrophoresis have also been exploited
to develop the only direct and high-throughput method to identify
oxidant induced, disulfide-bonded protein complexes. This approach,
termed diagonal SDS-PAGE
136
or redox 2D-PAGE
137
involves sequential nonreducing/reducing two-dimensional
SDS-PAGE (Figure 7b). The protein mixture is
first resolved by nonreducing gel electrophoresis to separate complexes
by size, followed by excision of a narrow gel strip in the sample
lane over the entire molecular weight range. The proteins are then
reduced and alkylated in-gel to prevent disulfide bond reformation,
the gel strip laid at a 90° angle across a second gel, and the
proteins are subsequently resolved under reducing conditions. Proteins
that are not involved in disulfide bond formation will lie in a diagonal
line on the 2D gel, whereas proteins that form disulfide bonds will
appear as distinct spots above or below the diagonal line. Protein
identity is subsequently determined by LC-MS/MS analysis. A major
limitation of this method, as with all 2D SDS-PAGE based methods,
is that it cannot reliably visualize or produce analytical quantities
of low abundance proteins that are present in less than 1000 copies
per cell.
138
Nonetheless, this procedure
has been used to detect disulfide-linked proteins in whole cell lysates
derived from oxidant-treated rodent nerve cell cultures
139
and cardiac myocytes.
140
As outlined above, redox 2D-PAGE identifies proteins that form disulfides
but does not provide information as to which proteins form which complexes.
An alternative approach is to first isolate the protein of interest
using a protein-specific antibody or affinity tag. This procedure
permits identification of proteins that form disulfides with a protein
of interest, and was recently used to identify of a novel reducing
system in the bacterial periplasm.
105
One limitation of the redox SDS-PAGE approach is that it does not
provide quantitative information about the extent or fraction of cysteine
oxidized under a given condition. To enable identification and quantification
of reversibly oxidized protein cysteine residues, including disulfides,
the Jakob group has reported an extension of the ICAT technology,
known as OxICAT (Figure 7c).
111b
Lysates are first generated in the presence of TCA to precipitate
proteins and prevent thiol/disulfide exchange. Free thiols are then
alkylated with a light (12C) ICAT reagent, followed by
reduction of with TCEP, which serves to reduce reversible modifications
(Chart 1). Nascent thiols are subsequently
labeled with a heavy (13C) ICAT reagent, protein samples
are digested and ICAT-modified peptides are isolated by avidin affinity
chromatography. The eluted peptides are then analyzed by LC-MS/MS
and the extent of oxidation for a particular cysteine is determined
by the ratio of the heavy to light MS signals. While this procedure
is not specific for disulfide-bonded cysteines per se, sulfenic acids
and S-nitrosothiols are exquisitely sensitive to
changes in pH and may be lost during sample preparation.
104a,141
Consequently, the OxICAT method seems best suited for disulfide
detection, including both protein and low molecular weight (e.g.,
S-glutathionylation) disulfides.
3.3.2.2
S-Glutathionylation
The thiol-containing tripeptide,
GSH is maintained at millimolar
concentrations inside cells. Under normal conditions, 98% or more
of GSH is maintained in its reduced state, however, in oxidative stress-associated
disorders like cancer and neurodegenerative diseases, an appreciable
amount of the GSH pool exists in the oxidized state, GSSG.
142
The GSH/Grx/GR system maintains protein thiols
in their reduced state through thiol-disulfide exchange and redox
reactions. Additionally, GSH undergoes nucleophilic addition and displacement
reactions to purge the cell of toxic electrophilic and oxidizing reagents
as catalyzed by glutathione S-transferase (GST),
glyoxalase, GR, and Grx.
143
Protein
S-glutathionylation can occur during reduction of disulfides by the
GSH/Grx/GR system and is readily reversible. When the GSH/GSSG redox
balance shifts toward a more oxidizing state, protein S-glutathionylation can function
as a regulatory mechanism or protect
against irreversible oxidation.
120
If the
GSH/Grx/GR system is compromised during oxidative stress, the accumulation
of S-glutathionylated proteins can occur and has been associated with
aging.
144
Within the context of redox signaling,
protein S-glutathionylation can take place through two possible mechanisms:
(i) thiol–disulfide exchange of GSSG with a thiolate or (ii)
condensation of GSH with a sulfenic acid (Figure 8) or an S-nitrosothiol. In a study
of sulfenic
acid-modified HSA, S-glutathionylation was estimated to occur with
a rate constant of 2–100 M–1 s–1.
106c
Thiol–disulfide exchange
between GSSG and a protein thiolate is very slow,
145
but may be catalyzed by Grx, which appears to promote S-glutathionylation
of the ETC complex I.
146
In this case,
Grx-mediated S-glutathionylation may occur through free radical formation.
147
Specificity in S-glutathionylation may depend
upon the steric properties, surrounding environment, and oxidation
sensitivity of the cysteine. Like disulfides, S-glutathione protein
adducts are stable to nonthiol nucleophiles. Deglutathionylation is
catalyzed by members of the Grx family,
148
but Srx,
149
Trx,
150
and PDI
150a
may also perform
this function, albeit with decreased efficiency.
151
Figure 8
Mechanisms for glutathionylation. Protein glutathionylation products
can be formed by (a) thiol/disulfide exchange of a protein thiolate
with oxidized glutathione (GSSG) or (b) condensation of GSH with a
protein sulfenic acid.
Enzymes such as trypsin,
152
collagenase,
153
and fructose-1,6-bisphosphatase
154
are activated by S-glutathionylation, whereas
glyceraldehyde 3-phosphate dehydrogenase (GAPDH),
155
26S proteasome,
156
cysteine
protease caspase-1,
157
and ETC complex
I
158
are inactivated by this modification.
As previously mentioned, many PTPs are regulated by intramolecular
disulfide bond formation at their catalytic cysteine.
159
However, some PTPs do not contain a second
cysteine proximal to their active site. In some of these cases, for
example in PTP1B, the phosphatase undergoes S-glutathionylation
to guard against hyperoxidation (defined as oxidation to irreversible
sulfinic and sulfonic acid states).
160
In
addition to regulating enzyme activity, S-glutathionylation can also
influence protein–DNA and protein–protein interactions.
For instance, S-glutathionylation of cysteines in the DNA binding
domain of transcriptional regulator, p53 weakens its association with
DNA.
161
Similarly, S-glutathionylation
of the transcriptional regulator, interferon regulatory factor 3 (IRF3)
inhibits its interaction with CBP/p300 coactivators and prevents activation
of target genes involved in induction of an antiviral response.
162
To date, several methods have been developed
to detect protein
S-glutathionylation based on immunological, metabolic labeling, and
differential alkylation approaches.
138
A
common method to detect S-glutathionylation in proteins employs an
antibody specific for the protein-S-GSH adduct.
162,163
This antibody is amenable to immunoprecipitation, Western blot on
nonreducing gels, and immunofluorescence analysis. The anti-GSH antibody
has also been used in conjunction with 2D SDS-PAGE, where samples
are separated by isoelectric focusing in the first dimension and by
molecular weight in the second dimension, with ensuing MALDI-TOF MS
to identify S-glutathionylated proteins in HeLa cells.
163b
Given the differences in the surrounding environment
of the modified cysteine, a limitation of the antibody is that not
all protein-S-GSH adducts are detected with the same
affinity.
164
An alternative immunological
approach, called GST overlay, exploits the specificity and affinity
of GST for GSH. In this method, Western blots from nonreducing SDS-PAGE
gels are exposed to biotinylated-GST, which recognizes and binds selectively
to protein-S-GSH disulfides; biotin-GST is subsequently
detected by avidin blot (Figure 9a).
165
Protein S-glutathionylation can also be monitored
indirectly by differential alkylation. In this workflow, free thiols
are alkylated, protein-S-GSH adducts are selectively
reduced by Grx, and nascent thiols are tagged by a biotinylated or
fluorescent alkylating reagent (Figure 9b).
166
In theory, this approach could also be coupled
to the OxICAT method to measure the extent of protein-S-GSH disulfides.
Figure 9
Methods to detect protein S-glutathionylation. (a) GST
overlay.
Samples are separated by nonreducing SDS-PAGE to preserve protein-GSH
disulfides. Blots are subsequently treated with biotinylated glutathione
S-transferase (GST), which binds selectively to GSH and permits detection
of S-glutathionylated proteins by avidin blot. (b) Indirect differential
alkylation of S-glutathionylated proteins. Free thiols are blocked
with NEM or IAM, protein-GSH disulfides are selectively reduced with
glutaredoxin (Grx), and nascent thiols are labeled with an alkylating
agent conjugated to biotin or a fluorophore. S-Glutathionylated proteins
are detected by avidin blot or in-gel fluorescence. (c) Biotinylated
glutathione ethyl ester (BioGEE, 3) enables in situ detection
of glutathionylated proteins. N,N-Biotinyl glutathione disulfide (4) permits detection
of proteins that become S-glutathionylated by thiol/disulfide exchange
with GSSG.
Approaches have been
developed to facilitate detection of S-glutathionylated
proteins in cells. One such method involves inhibiting protein synthesis
with cycloheximide, which does not affect GSH synthesis, with subsequent
metabolic labeling of the GSH pool through
35
S-cysteine incorporation.
167
Cells are
subsequently lysed in the presence of a thiol alkylating agent to
minimize thiol-disulfide exchange, samples are separated under nonreducing
conditions, and analyzed by radiography. This technique has been used
to identify proteins, such as enolase and 6-phosphogluconolactonase,
that undergo S-glutathionylation in human T lymphocytes
exposed to exogenous oxidants (e.g., H2O2 and
diamide).
5b
Alternatively, Finkel, Eaton
and colleagues have used biotinylated-GSH ethyl ester (BioGEE, 3)
155,167
and N,N-biotinyl
glutathione disulfide (4)
164
(Figure 9c) to monitor protein S-glutathionylation in lysates, isolated cells, and
tissues. While
the biotin tag facilitates enrichment and identification of proteins
that undergo S-glutathionylation, limitations of
these methods include steric occlusion of biotinylated GSH analogues
and poor cellular trafficking of biotinylated probes.
168
3.3.2.3
Sulfenic Acids
Because of their
reactive nature, sulfenic acids are often deemed unstable intermediates
en route to additional cysteine modifications (Figure 2 and Chart 4). The formal oxidation
state of the sulfur atom in a sulfenic acid is 0, enabling it to function
as both a weak nucleophile and a soft electrophile (Chart 4 and 5, eq 1).
3b
The dual nature of its reactivity is clearly
illustrated by the condensation of two sulfenic acids to generate
a thiosulfinate (Chart 4). Thiosulfinate formation
via sulfenic acid condensation may be most facile when sulfenate and
sulfenic acid states are equally present.
119b
As previously discussed, the prevalence of thiosulfinates in cells
is currently unknown; however, given the abundance of cellular thiols,
interfacing of two sulfenic acids is likely to be a rare event.
3a
Chart 4
Sulfenic Acids Exhibit Both Nucleophilic
and Electrophilic Character,
As Illustrated by Condensation of Two Sulfenic Acids to Afford a Thiosulfinate
(Black Box)a
a
As a nucleophile (purple boxes),
sulfenic acids can undergo SN2 displacement with halogenated
compounds, such as 4-chloro-7-nitrobenzo-2-oxa-1,3-diazole (NBD-Cl, 5), reaction with
alkynes (6) and alkenes (7) to form the corresponding sulfoxides, and reaction with
two equivalents of triphenylphosphines (8) to afford
the free thiol and oxidized phosphine (not shown). Sulfenic acids
can also function as an electrophile (green boxes) to react with thiols
to yield a disulfide, and with 1,3-cyclohexadiones including dimedone
(9), to yield a thioether adduct. As an electrophile,
sulfenic acids can also react with hydrazines (10) to
yield the thiol and azo compound (not shown), or with amines (11) to yield sulfenyl
amides.
Chart 5
Reaction
Schemes of Condensation of Two Sulfenic Acids to Yield a
Thiosulfinate (Equation 1) and Electrophilic Reaction of Sulfenic
Acid with Dimedone (9, Equation 2) and Hydrazines (10, Equation 3)
Analogous to the reactivity of sulfur in a cysteine thiol,
the
nucleophilic character of a sulfenic acid is likely to be influenced,
in part, by pK
a. Studies of sulfenic acids
in small molecules have shown that electron-withdrawing substituents
reduce the pK
a to favor sulfenate formation
and enhance the stability of this species.
169
The pK
a of sulfenic acids in proteins
could be similarly modulated to regulate their stability and reduce
its reactivity toward a thiol. Stabilization of the sulfenate anion
through decreased pK
a could also enhance
the nucleophilic character of the sulfur atom, marking potential sites
of cysteine hyperoxidation.
The pK
a of sulfenic acids in small
molecules has been estimated to be in the range of 4.5–12.5.
104a,170
The pK
a of protein sulfenic acids has
not been as extensively studied, but two measurements have been made,
both with bacterial Prxs. There are three classes of Prxs: typical
2-Cys, atypical 2-Cys, and 1-Cys Prxs. Both typical and atypical 2-Cys
Prxs form sulfenic acid at their active site cysteine after reaction
with H2O2, which then condenses with a second
cysteine in the same (atypical) or neighboring (typical) Prx to generate
a disulfide that is reduced by Trx/TrxR to complete the catalytic
cycle.
171
1-Cys Prxs do not contain a resolving
cysteine and the sulfenic acid intermediate may be reduced by GSH
or ascorbate.
172
The first pK
a measurement reported for the sulfenyl group of a protein
sulfenic acid was obtained using a mutant form of 2-Cys Prx from Salmonella typhimurium,
AhpC in which the resolving cysteine
was changed to serine. Key to the success of these experiments, the
sulfenic acid and sulfenate forms exhibit distinct spectral shifts
in AhpC, allowing a pK
a determination
of 6.1.
173
Consistent with this measurement,
a tryptophan fluorescence study revealed a pK
a of 6.6 for the sulfenic acid in a 1-Cys Prx from Mycobacterium tuberculosis.
106a
Analogous to cysteine thiolate reactivity with H2O2, the propensity for sulfenic acid
to undergo further
oxidation
to sulfinic acid can be strongly influenced by the local protein environment.
Relative to their prokaryotic counterparts, 2-Cys Prxs from eukaryotic
organisms appear uniquely sensitive to hyperoxidation and may be related,
at least in part, to sulfenic acid pK
a.
91,174
For example, oxidation of bacterial peroxiredoxin
AhpE sulfenic acid by H2O2 occurs at 40 M–1 s–1, whereas HSA sulfenic acid
reacts at 0.4 M–1 s–1.
106a,106c
While the pK
a of the protein sulfenic
acids were not reported in these studies, it is interesting to note
that initial formation of sulfenic acid was also significantly slower
in HSA (2.7 M–1 s–1)
106c
compared to AhpE (8.2 × 104 M–1 s–1).
106a
To better understand how some protein environments facilitate
sulfenic acid oxidation, additional physical organic and computational
studies of both small-molecule and protein model systems will be required.
Sulfenic acids have been identified in the catalytic cycle of multiple
enzymes, including Prx, NADH peroxidase, and methionine sulfoxide-
and formylglycine-generating enzymes.
66c,71,106a
Formation of sulfenic acid has also been linked to
oxidative stress-induced transcriptional changes in bacteria due to
altered DNA binding of OxyR and OhrR and changes in the activity of
the yeast Prx and Yap1 protein.
66c,175
Less is known
about the mechanisms that underlie sulfenic acid-mediated regulation
of mammalian protein function and signaling pathways; however, cysteines
from several transcription factors (i.e., NF-κB, Fos, and Jun),
or proteins involved in cell signaling or metabolism (e.g., GAPDH,
GR, PTPs, kinases, and proteases) can be converted to sulfenic acid
in vitro. Sulfenic acid formation has also been implicated in the
regulation of apoptosis, immune cell activation and proliferation,
and growth factor (GF) signaling pathways.
12,123c,176
Although sulfenic acids
are often transient, an advantage to studying
this modification is that it represents the initial product of two-electron
oxidants with the thiolate anion and can therefore serve as a marker
for oxidant-sensitive cysteine residues. A variety of indirect and
direct chemical methods have been developed to detect protein sulfenic
acid modifications (also termed sulfenylation
5a,12
). An early indirect chemical method that was reported involves thiol
alkylation, reduction of sulfenic acids by arsenite, and labeling
of nascent thiols with biotinylated NEM (Figure 10a).
177
This methodology was subsequently
used to profile sulfenic acid formation in rat kidney cell extracts;
178
however, as with other indirect differential
alkylation methods, a significant limitation is the debatable selectivity
of the arsenite-mediated reduction step.
179
Figure 10
Methods to detect protein sulfenic acids. (a) Indirect differential
alkylation of protein sulfenic acids. Free thiols (blue) are blocked
with NEM or IAM, protein sulfenic acids (purple) are reduced with
arsenite, and nascent thiols are labeled with biotinylated NEM (NEM-Biotin).
Sulfenylated proteins are detected by avidin blot where increased
protein oxidation is observed as an increase in signal intensity.
(b) Direct in situ labeling of protein sulfenylation. Cells are treated
with or without stimulant (e.g., oxidant, growth factor) and subsequently
incubated with azido or alkyne dimedone analogues, such as 18 to chemically modify
sulfenylated proteins. Afterward, excess probe
is removed, cell lysates are generated, and probe-modified proteins
are conjugated to biotin or a fluorophore by a coupling reaction (e.g.,
Staudinger ligation or Huisgen [3 + 2] cycloaddition). The samples
can then be avidin enriched and subjected to proteomics analysis or
analyzed by avidin blot or in-gel fluorescence where increased protein
sulfenylation correlates to enhanced signal intensity. (c and d) High-throughput
immunological detection of dimedone (9)-modified proteins
using arrays. (c) Proteins immobilized on a microarray that are susceptible
to sulfenylation are irreversibly modified by 9. The
protein–dimedone adduct forms an epitope for selective detection
by the antibody. (d) Cells are treated with or without stimulant (e.g.,
oxidant, growth factor) and are subsequently incubated with 9 to irreversibly modify
sulfenylated proteins. Subsequent
to cell lysis, proteins within a given signaling pathway are immobilized
on an antibody array and dimedone-modified proteins are detected by
addition of the antibody. (e) Isotope-coded dimedone 2-iododimedone
(ICDID) permits quantification of protein sulfenylation. Sulfenic
acids are labeled by d
6-dimedone (21, purple), then excess reagent is removed and free thiols
are labeled by d
0-2-iododimedone (22, blue) generating chemically identical adducts that differ
by 6 Da. The samples are trypsinized and analyzed by LC-MS where the
extent of sulfenic acid occupancy is determined by the ratio of d
6-dimedone to d
0-dimedone peak intensities. (f) Quantification and site-identification
of protein sulfenic acids with d
6-DAz-2
or d
6-DYn-2 and an acid-cleavable linker
(ACL) coupling reagent. Sulfenic acids are labeled with d
0-DAz-2 (16) in the untreated sample and
with d
6-DAz-2 (19) in the
oxidant-treated sample. Excess probe is removed and the samples are
combined and biotinylated by coupling with the alkyne-ACL (23) to generate chemically
identical adducts that differ by 6 Da. The
sample is then trypsinized, avidin enriched, and trifluoroacetic acid-eluted
peptides are analyzed by LC-MS/MS where the increase in sulfenic acid
modification in response to oxidant is determined by the ratio of
d
6
to d
0
peak
intensities. Biotin can complicate spectra and decreases peptide recovery,
and the removal of biotin with the ACL permits increased sample elution
and direct identification of modified peptides in the MS.
Direct methods for sulfenic acid detection have
been developed
that take advantage of the chemical reactivity of this oxyacid. Nucleophilic
substitution of halonitroarenes, such as 4-chloro-7-nitrobenzo-2-oxa-1,3-diazole
(NBD-Cl, 5), and nucleophilic addition to electron-deficient
alkynes (6), alkenes (7), and triphenyl
phosphines (8) are reported to trap sulfenic acids (Chart 4).
169
Of these, the most
commonly used in the detection of protein sulfenic acids is NBD-Cl.
This reagent reacts with thiols, sulfenic acids, and at higher pHs,
amine-containing residues, but the resulting products are distinguished
on the basis of their spectral properties and molecular weight.
180
As NBD-Cl can react with a variety of protein
functional groups, this reagent appears best suited for use with recombinant
proteins, especially those with a single cysteine residue.
16
Consequently, NBD-Cl does not have utility in
global detection of protein sulfenic acids in complex protein mixtures,
necessitating the development of methods for selective detection that
exploit the electrophilic properties of the sulfur atom in sulfenic
acid.
As first reported by Benitez and Allison in 1974, protein
sulfenic
acids react with cyclic 1,3-diketone carbon nucleophiles, like 5,5-dimethyl-1,3-cyclohexadione
(dimedone, 9) and with hydrazines (10) or
amines (11) (Chart 4 and 5)
181
Dimedone has proven
useful in revealing the requirement for protein sulfenic acid modifications
in the S. cerevisiae Yap1-Gpx3 H2O2-sensing pathway,
66c
T cell activation,
123c
and EGFR signaling.
12
Unlike sulfur, nitrogen, or phosphorus-based nucleophiles, under
aqueous conditions cyclic 1,3-diketones do not cross react with cysteine
thiols, sulfinic acid, or other functional groups commonly found in
biomolecules, making this reaction an extremely attractive avenue
for developing chemically selective detection methods. All chemoselective
methods for detecting protein sulfenic acids reported to date depend
upon this chemistry.
138
Two recent reports
expand the scope of reactive templates to 1,3-cyclopentadione
182
and linear β-ketoester
183
analogues (though caution should be exercised with linear
derivatives since they have been reported to cross react with amines,
such as lysine
184
). The lack of an enrichment
or visualization “handle” for protein-S-dimedone adducts subsequently motivated the
development of biotinylated
(12,13)
185
and
fluorophore-conjugated (14) analogues
185b,186
(Chart 6). These probes have been used in
a proteomic study with isolated rat hearts
185a
and to identify AKT2 as a target of PDGF-induced H2O2.
187
Depending upon the application,
one potential drawback for direct conjugation of any probe to biotin
or a fluorophore is that the bulky chemical tags can reduce cell-permeability.
168b
Naturally, not all conjugated probes are entirely
impermeant (e.g., DCFH diacetate, DCP-Bio1) however, comparative studies
show time and again that tagged derivatives often suffer from diminished
cell uptake and trafficking properties.
168,188
Alternative mechanisms of uptake are possible (e.g., active transport
of BioGEE), but may limit probe distribution to specific cellular
compartments. A further consideration when functionalizing probes
with large chemical tags is that increased steric bulk can lead to
a significant bias in protein target labeling.
188a,189
The poor permeability of many biotin- and fluorophore-tagged probes
typically necessitates labeling of proteins in lysates and is, therefore,
subject to the aforementioned limitations. In this context, it is
also important to bear in mind that labile or transient sulfenic acid
modifications may be further oxidized or insufficiently trapped during
the lysis procedure.
Chart 6
Biotin, Fluorophore, and Chemical Handle Derivatives
of Dimedone
A subsequent alternative
approach that has emerged is the development
of azido- and alkyne-functionalized dimedone analogous (Chart 6), termed DAz-1 (15),
168b,180c
DAz-2 (16),
190
DYn-1 (17), and DYn-2 (18),
12
which enable the trapping and tagging of protein sulfenic acid modifications
directly in living cells. In later steps, proteins covalently modified
by DAz or DYn probes can be coupled to biotin or fluorophores by Staudinger
ligation
191
or Huisgen [3 + 2] cycloaddition
reactions (Chart 7, eqs 1 and 2, and Figure 10b).
192
Application of
DAz-2 to identify proteins that undergo sulfenic acid modifications
in HeLa cells identified upward of 200 candidates, including the majority
of known sulfenic acid-modified proteins.
190
Cross-comparison of these data with those from disulfide and S-glutathionylation
proteomes revealed modest overlap between these “redoxomes”,
suggesting that a significant portion of sulfenic acid modifications
may not be intermediates en route to S-thiolated forms and, instead,
can be stabilized by the protein local environment.
138
Alternatively, or in addition, it is also possible that
(i) lysate-based approaches employed in the S-thiolation proteomic
studies resulted in fewer identifications and, therefore, lower overlap
with the “sulfenome”, and (ii) the modest rate constant
for the reaction of many dimedone analogues with sulfenic acid (103 M–1 min–1)
111a
may not be sufficient to trap especially transient
modifications. Azido dimedone analogues have also been used to show
that sulfenic acid modification of the thiol peroxidase, Gpx3 is essential
for yeast to sense oxidative stress
66c
and
to identify a unique reducing system in the bacterial periplasm that
protects single cysteine residues from oxidation.
105
Chart 7
Bioorthogonal Coupling Reactions Staudinger Ligation
and Huisgen
[3 + 2] Cycloadditiona
a
Staudinger ligation
(equation
1) functionalizes an azide-modified molecule, while Huisgen [3 + 2]
cycloaddition (equation 2) couples azide-modified or alkyne-modified
proteins to detection tags. Alkyne acid cleavable linker (Yn-ACL, 23) reagent for
Huisgen [3 + 2] cycloaddition with azide-modified
proteins. Blue, acid cleavable moiety.
More
recently, DYn-2 was used in global profiling studies to reveal
dynamic protein sulfenylation during EGF signaling in human epidermoid
A431 cells and to identify the EGFR kinase as a prominent target of
endogenous signaling H2O2 (Figure 11).
12
Three PTPs involved
in the regulation of EGFR signaling, PTP1B, PTEN, and SHP2 were also
shown to undergo sulfenic acid modification in response to EGF stimulation
of cells. Interestingly, PTPs and EGFR displayed differential sensitivity
to oxidation by EGF-induced endogenous H2O2 that
correlated with the relative proximity of each enzyme to the oxidant
source itself, NOX2 (Figure 11). This study
was the first of its kind to provide evidence for sulfenic acid modification
of PTP in cells during growth factor signaling. Prior studies performed
in lysates had led to speculation as to the likelihood of PTP oxidation
due to their modest reactivity with H2O2 and
their low abundance in comparison to the abundant and reactive Prxs.
2d,123c,193
Interestingly, while this study
found ER-localized PTP1B to be only moderately sensitive to H2O2 derived from plasma
membrane-bound NOX2, a study
by Keaney and colleagues indicates that this oxidation reaction becomes
relevant during ER-localized NOX4 activation.
194
As it is unlikely that the intrinsic reactivity of the
active site cysteine in PTP1B differs in these two systems, these
data suggest that proximity of PTP1B (and other proteins) to the NOX
oxidant source may be an important determinant of target selectivity.
Hence, the apparent sensitivity and physiological relevance of PTP1B
oxidation, and protein thiols in general, is likely to be a signaling
pathway and cell type-specific phenomena.
Figure 11
Redox regulation of
epidermal growth factor (EGF) signaling by
protein sulfenylation. Binding of EGF to the EGF receptor (EGFR) facilitates
receptor dimerization, activation (not shown), and promotion of NOX2
complex assembly. NOX2-derived H2O2 translocates
into the cytoplasm likely through channels such as aquaporins where
it has been shown to regulate the activity of proteins involved in
the EGFR signaling cascade. EGFR and the phosphatases SHP2, PTEN,
and PTP1B were all found to be sulfenylated in response to EGF stimulation,
albeit with differential sensitivities (ranked 1–4 in order
of decreasing susceptibility). The sensitivity of each protein to
oxidation correlates to their relative proximity to the oxidant source.
In this way, EGFR, which forms a complex with NOX2, exhibited the
highest sensitivity. Moreover, the EGFR-associated phosphatase SHP2
exhibited increased susceptibility to sulfenylation as compared to
the cytoplasmic phosphatase PTEN, which regulates the levels of PIP3, and PTP1B, which
is localized to the cytoplasmic face of
the endoplasmic reticulum (ER). Co-localization of antioxidant enzymes
such as peroxiredoxins (Prx) to the signaling regions is thought to
limit the range of H2O2 diffusion (green area).
Interestingly, NOX-derived reactive oxygen species (ROS) have also
been shown to inactivate Prxs by hyperoxidation (PrxII) or phosphorylation
(PrxI).
11b
These regulatory mechanisms
have been proposed to permit localized accumulation of ROS for redox
regulation of proteins located near the oxidant source (pink area).
In 2009, our group reported the
first immunological method for
detecting protein sulfenic acid modifications. Antibodies were elicited
by a synthetic hapten mimicking dimedone-modified cysteine conjugated
to KLH (Figure 10c) and are highly specific
and sensitive for detecting protein-S-dimedone adducts by Western
blot and immunofluorescence.
195
Application
of this immunochemical approach to protein arrays and breast cancer
cell lines revealed considerable differences in the level of protein
sulfenic acid modifications among tumor subtypes (Figure 10c). This method has also
been used to demonstrate
the cysteine sulfenylation and colocalization of oxidized proteins
with NOX2 during EGF signaling.
12
Subsequently,
in 2011, Eaton and colleagues reported a similar antibody and used
this reagent to study sulfenic acid modification of GAPDH in cardiac
myocytes exposed to exogenous H2O2.
196
A future application of these antibodies will
be to combine their use with antibody arrays to facilitate unbiased
investigation of protein sulfenic acid modifications in signaling
pathways (Figure 10d).
Beyond detection,
one approach to determine which protein sulfenic
acid modifications are relevant to signaling in normal cells as well
as in pathological processes is to quantify the extent of oxidation.
To this end, our laboratory has recently developed two methods to
facilitate relative quantification of sulfenic acid modifcations:
(1) isotope-coded dimedone and 2-iododimedone (ICDID) (Figure 10e),
197
and (2) isotopically
light and heavy derivatives of DAz-2 (19) and DYn-2 (20) (Chart 6).
198
The ICDID workflow uses deuterium-labeled dimedone (d
6-dimedone, 21) to trap sulfenic
acids, followed by alkylation of free thiols with 2-iododimedone (22). Importantly,
the covalent adducts afforded by these two
reagents are structurally/chemically identical, and have identical
efficiencies of ionization. Nevertheless, the thiol and sulfenic acid-tagged
species are differentiated from each other by 6 Da and the extent
of sulfenic acid modification at a cysteine is determined from the
ratio of heavy to light isotope-labeled peak intensities (Figure 10e). Alternatively,
isotopically light and heavy
forms of DAz-2 can be used to monitor relative changes in sulfenic
acid modification. This strategy has been combined with an acid cleavable
linker (ACL) that is suitable for Huisgen [3 + 2] cycloaddition coupling
(Chart 7).
198
With
this method, samples are labeled by heavy or light DAz-2, combined
and conjugated to the alkyne-biotin ACL reagent (Yn-ACL, 23) digested with trypsin,
enriched on avidin cartridges, and tagged
peptides are eluted by trifluoroacetic acid (TFA)-mediated cleavage.
Peptides are then mapped by LC-MS/MS analysis to identify the sulfenic
acid-modified protein and map the site of modification. The relative
change in protein sulfenic acid modfication between two samples is
determined by the ratio of heavy to light isotope-labeled peak intensities
(Figure 10f).
Analogous to irreversible
electrophilc inhibitors that modify semiconserved
cysteines residues in protein tyrosine kinases (PTKs) currently in
phase II and III cancer clinical trials,
199
we envision the development of nucleophile-functionalized small
molecules that target a sulfenic acid-modified cysteine in a specific
protein. Our design strategy is to conjugate the nucleophile “warhead”
to a high affinity ligand that binds proximal to the target cysteine
sulfenic acid. As proof of principle, we have developed small molecules
that target PTPs, termed redox-based probes (RBPs, Figure 12, 24–26), comprised
of three parts: (i) a cyclohexanedione nucleophile, (ii) a chemical
scaffold that binds to the conserved PTP active site, and (iii) an
azide (or alkyne) chemical reporter to facilitate downstream detection
and isolation of labeled PTPs (Figure 12a).
189
The RBPs exhibited enhanced binding and sensitivity
for detecting sulfenic acid modification of the catalytic cysteine
in the YopH and PTP1B phosphatases, compared to the parent compound,
DAz-1 (15), which lacks the additional binding element.
The RBP approach should facilitate cellular investigations of PTP
redox regulation. Methods to study PTP redox modulation are often
thwarted by issues of low abundance and studies of this nature would
greatly benefit from a targeted approach, as exemplified by RBPs.
Neel and colleagues have also reported an indirect immunochemical
method for global proteomic assessment of the PTP “redoxome”
that relies on performic acid hyperoxidation of cysteine oxyacids
(Figure 12b).
200
Figure 12
Targeted approaches to monitor cysteine oxPTM with in specific
or family of proteins. (a) Redox-based probes (RBPs, 24–26) for protein tyrosine phosphatases
(PTPs)
are comprised of three parts: a warhead that permits chemoselective
reaction with sulfenic acid, a PTP-directed scaffold that exhibits
enhanced affinity for target binding, and an azide chemical reporter
to facilitate downstream detection and isolation of labeled protein.
(b) Indirect two-stage immunochemical approach to monitor PTP oxidation.
In stage 1, free thiols in one aliquot of sample are alkylated with
NEM, oxidized cysteines are reduced with DTT, and nascent thiols are
hyperoxidized to sulfonic acid with pervanadate. The proteins are
subsequently trypsinized, and sulfonic acid-modified peptides are
isolated with a monoclonal antibody that recognizes hyperoxidized
PTPs. In stage 2, a second aliquot of sample is reduced with DTT and
all thiols are oxidized to sulfonic acid with pervanadate and processed
as in stage 1. The enriched peptides are analyzed by LC-MS/MS and
the extent of PTP oxidation is determined by the ratio of stage 1
to stage 2 signal intensities. (c) Conformation-sensing single-chain
variable antibodies that selectively recognize the unique conformation
of the sulfenyl amide oxoform of PTP1B.
In addition to pan-PTP recognition, RBPs can be refined to
target
a single member of the PTP family. Such a reagent would not only be
useful to study redox-regulation of a specific PTP, but might possibly
serve as lead compounds for the development of a new class of therapeutics
to ameliorate diseases associated with aberrant PTP activity, as in
diabetes.
201
In support of this approach,
Tonks and colleagues recently reported the development of antibodies
as single-chain variable fragments that selectively recognize the
unique conformation that PTP1B adopts when its activate site cysteine
exists in the sulfenamide form (Figure 12c).
202
These conformation-sensing antibodies were
able to trap PTP1B in the inactive conformation permitting sustained
insulin signaling in human embryonic kidney (HEK) cells. Lastly, the
RBP or “nucleophilic inhibitor” approach can be extended
to other classes of proteins that contain a redox-sensitive cysteine,
such as EGFR.
12
3.3.2.4
Sulfinic
Acids
In the presence
of excess oxidant, sulfenic acid can be oxidized to sulfinic acid
(Figure 2). The formal oxidation number of
the sulfur atom in sulfinic acid is +2. On this basis, this oxyacid
might be expected to have enhanced electrophilicity compared to sulfenic
acid. However, sulfinic acid does not undergo self-condensation or
nucleophilic attack by thiols. This can be explained by the increase
in partial positive charge on the sulfur in sulfinic acid, which converts
the sulfur atom into a harder electrophile making it less prone to
reaction with soft nucleophiles, such as thiols. With a pK
a of 2, sulfinic acid is deprotonated
at physiologic pH and can exist in two resonance forms (Chart 8).
203
Sulfinic acids
function as nucleophiles (Chart 9), reacting
largely from sulfur to undergo alkylation (27), as well
as nucleophilic addition to activated alkenes (28), aldehydes
(29), lactones (30), α,β-unsaturated
compounds (31), and diazonium salts (32)
to give the corresponding sulfones.
3b,204
The preceding
reactions are established for sulfinic acids under synthetic organic
conditions, but it is not established whether all of these reactions
would take place with protein sulfinic acids. The reactions in Chart 9 exhibit a wide
range of rates and some go to completion
on the hour time scale (27,
205
28,
206
30,
207
31
208
), while others, such as 29 and 32,
209
undergo rapid equilibrium-based transformations.
Of note, the reaction of sulfinic acid with aldehydes serves as the
basis for the Schiff’s test. As an ambidentate nucleophile,
sulfinic acid can also react at oxygen as illustrated by nucleophilic
attack of the sulfinate oxygen on the γ-phosphate of ATP (33) to form the sulfinic
acid phosphoryl ester intermediate
in the Srx catalytic cycle (Chart 9).
122a,210
Chart 8
Resonance Structures of Sulfinic Acids
Chart 9
Sulfinic Acids Function as Soft Nucleophiles Reacting Primarily
from
the Sulfur to Undergo Alkylation (27) or Nucleophilic
Addition to Activated Alkenes (28), Aldehydes (29), Lactones (30), α,β-Unsaturated
Compounds (31), and Diazonium Salts (32)a
a
Sulfinic acids can also function
as nucleophiles involving reactivity from the oxygen as exemplified
by the sulfiredoxin (Srx)-catalyzed reaction with ATP (33) to yield a transient sulfinic
acid phosphoryl ester. Though not
shown, reactions 27 and 28 undergo acid-catalyzed
SN1 reactions and thus require the protonated sulfinic acid species.
Cysteine sulfinylation can also modify protein
metal binding properties.
Oxidized sulfur ligands are weaker donors and can increase the Lewis
acidity of the liganded metal center, which influences affinity and
coordination. In matrix metalloproteases (MMPs), sulfinic acid oxidation
of a zinc-coordinated active site cysteine thiolate activates protease
function, in part by reducing the ability to coordinate the zinc cation.
211
In contrast, nonheme FeIII coordination
in nitrile hydratases (NHases) is accomplished by a unique CXXCXC
binding sequence in which two cysteines are present as the sulfinic
and sulfenic acid states.
154,212
Cysteine oxidation
is necessary for hydratase activity, and the increased Lewis acidity
of the FeIII afforded by cysteine oxidation is believed
to regulate the affinity of a catalytic water molecule for the metal
center.
3b,212a,213
A similar
coordination motif has also been identified in the unique noncorrin
cobalt center in a NHase from Pseudonacardia thermophila(214) and in thiocyanate hydrolase.
215
It has also been suggested that cysteine oxidation
alters the metal coordination from zinc (thiol) to iron or cobalt
(sulfenic/sulfinic acid). This preference may not be as strictly defined
as once thought, however, as a peptide mimetic inhibitor of neurotoxin
F from Clostridium botulinum was recently shown to
coordinate to an essential zinc by a cysteine sulfinate ligand.
216
To date, the cysteine sulfinic acid modification
has been most
extensively characterized in Prxs and the Parkinson’s disease
protein, DJ-1.
217
Eukaryotic Prxs appear
most susceptible to sulfinic acid modification,
91,174
a feature that evolutionarily coincides with Srx expression, the
only known sulfinic acid reductase.
109a,218
Srx was recently
identified in cyanobacteria,
219
which also
appear to have 2-Cys peroxiredoxins that are susceptible to hyperoxidation.
220
Recent work shows that Srx-mediated reduction
of Prx proceeds by a sulfinic acid phosphoryl ester that undergoes
nucleophilic attack by Srx Cys84 to form a thiosulfinate intermediate
that is subsequently resolved by Srx Cys48 to release Prx sulfenic
acid and oxidized Srx, which are both recycled by the Trx/TrxR system.
122a,210,221
The reaction of Srx with sulfinic
acid is slow (k
cat ≈ 0.2 min–1), and it is currently unknown whether any accessory
proteins enhance this reaction in vivo.
109b
The biological reversibility of sulfinic acid (at least in
some
proteins) hints at a regulatory function, analogous to a disulfide
or sulfenic acid. In this vein, it has been proposed that reversible
inactivation of Prx by sulfinic acid modification facilitates the
accumulation of endogenous H2O2 to regulate
signaling events in the so-called “floodgate hypothesis”.
91
While Prx II appears to be particularly sensitive
to hyperoxidation, it has recently been shown that phosphorylation
inactivates Prx I,
11b
with both mechanisms
of Prx inactivation serving to facilitate localized accumulation of
H2O2 for signaling purposes. Reversible Prx
oxidation has also been proposed to regulate eukaryotic circadian
rhythms, though the molecular details remain largely unknown.
222
Evidence of a regulatory role for reversible
sulfinic acid Prx
inactivation also stems from the observation that many signaling pathways,
including neuronal N-methyl-d-aspartate
(NDMA) receptor activity
223
and macrophage
activation by lipopolysaccharides
224
induce
Srx expression. In both cases, induction of Srx was dependent upon
redox-regulated transcription factors, AP-1 and Nrf2.
225
Srx can also translocate to the mitochondria,
to reduce hyperoxidized Prx III and protect against oxidative damage
and apoptosis.
226
The molecular details
are not entirely clear, however, it is possible that Srx-mediated
reactivation of Prx III maintains low mitochondrial ROS levels to
prevent opening of the mPTP. Srx overexpression also stabilizes PTEN
and PTP1B,
109c
which is reminiscent of
Prx I-mediated protection of PTEN tumor suppressor activity.
227
The aforementioned studies suggest an important
biological role for the reversibility of Prx hyperoxidation. Nonetheless,
further studies are required, including the development of an Srx
knockout mouse model to assess the physiological relevance of Prx
reactivation. It should also be noted that Srx may also carry out
other biological functions independent of sulfinic acid reduction.
109c,224
Although sulfinic acid has gained recognition as a regulatory
modification,
the full scope of its biological formation remains poorly understood,
due in part, to the lack of methods that are suited to general detection.
Methods to detect protein sulfinic acids include the a molecular mass
increase of 32 Da,
228
acidic electrophoretic
gel shifts,
335,336,228
and antibodies that recognize a sulfinic/sulfonic acid peptide from
a specific protein.
200,229
Such approaches facilitate study
of sulfinic acids in individual proteins, but have limited utility
in global analysis. As thiols are good nucleophiles, a challenge to
developing chemical methods for sulfinic acid detection lies in is
its behavior as a weak nucleophile. An alternative approach is to
design a reaction in which the product of the reaction with sulfinic
acid is uniquely stabile. Along these lines, our lab has recently
investigated the reaction sulfinic acids with aryl-nitroso compounds
(Chart 10, eq 1). The initial sulfinic acid-derived N-sulfonyl hydroxylamine product
is reversible, but can
be trapped by ester-functionalized aryl-nitroso 34 to
give an irreversible N-sulfonylbenzisoxazolone adduct
(Chart 10, eq 1).
203a
The reaction of 34 with a thiol yields a sulfenamide
species that can be cleaved with nucleophiles (Chart 10, eq 2) and, importantly, 34
does not cross react
with other sulfur and nonsulfur containing biological functional groups.
Chart 10
Chemoselective Approach to Detect Sulfinic Acidsa
a
Reaction of a sulfinic acid
with ester-protected aryl-nitroso compound 34 and subsequent
intramolecular nucleophilic attack of the nitroso anion intermediate
on the ester yields a stable N-sulfonylbenzisoxazolone
adduct (equation 1). In contrast, analogous reaction with a thiol
yields a sulfenyl amide adduct that is susceptible to subsequent reaction
with a second thiol or reducing agent (equation 2).
3.3.2.5
Sulfonic Acids
While the sulfinic
acid modification is relatively stable, it can undergo further oxidization
to give sulfonic acid, the most highly oxidized thiol species (Figure 2). Like sulfinic
acid, the sulfur atom in sulfonic
acid (formal oxidation number of +4) functions as a hard electrophile
and does not undergo self-condensation or nucleophilic attack by thiols.
With a pK
a <2, the
sulfonic acid is a both strong acid and a weak base, which makes it
a good leaving group in SN1, SN2, E1, and E2
reactions.
3b
Moreover, organic sulfonic
acids can undergo nucleophilic attack on alkenes (35),
alkynes (36), and allenes (37) to generate
the corresponding sulfonic acid esters (Chart 11), where the reaction initiates exclusively
at the oxygen.
230
However, it is currently unknown whether any
of the reactions presented in Chart 11 are
amenable to protein studies.
Chart 11
Reactions of Sulfonic Acidsa
a
Sulfonic acids function as
soft nucleophiles and react exclusively from the oxygen atom to undergo
acid-catalyzed reaction with alkenes (35), alkynes (36), and allenes (37) to generate
the corresponding
sulfonic acid esters. These reactions occur only in organic solvent
as the resulting products are largely unstable in water.
The sulfonic acid modification has been characterized in
a small
group of proteins, including mammalian Cu,Zn-SOD, where it has been
speculated that damage resulting from hyperoxidation plays an important
role in diseases like familial amyotrophic lateral sclerosis.
229b
Sulfonic acid is also present in mammalian
cells as the naturally occurring low molecular weight compound, taurine.
This biomolecule plays functions as a general osmolyte and modulator
of neuronal activity.
231
As with sulfinic
acid, elucidation of the biological and pathological role of sulfonic
acid modification has been hindered by a lack of means to selectively
detect this oxyacid. Recently, a method has been developed that permits
selective enrichment of sulfonic acid-modified peptides using poly
arginine (PA)-coated nanodiamonds as high affinity probes.
232
BSA, used as a model system in this study,
was oxidized with performic acid, digested, and sulfonic acid-containing
peptides were enriched and eluted from PA-coated nanodiamonds with
phosphoric acid, with subsequent identification of oxidized peptides
by MALDI-MS analysis (Figure 13a). This methodology
might have an application in the characterization of protein sulfonic
acids in cell lysates by first alkylating reduced and reversibly oxidized
thiols with IAM or NEM. Sulfinic acid and sulfonic acid-modified peptides
might also identified through a scheme involving performic acid oxidation
(Figure 13b). A limitation of this method,
however, is that sulfonic acid-modified peptides are in competition
with phosphorylated peptides for binding to the PA-coated nanodiamonds,
233
though this could potentially be reduced by
phosphatase treatment of lysates.
Figure 13
Method for selective enrichment of sulfonic
acid-modified peptides.
(a) All cysteine residues are oxidized to sulfonic acid with performic
acid. Proteins are then trypsinized and sulfonic acid-modified peptides
are enriched using polyarginine (PA)-coated nanodiamonds (ND). Eluted
peptides are analyzed by LC-MS/MS. (b) A plausible extension of the
PA-ND enrichment technology to identify sulfinylated and sulfonylated
cysteines. Samples are first treated with a reducing agent to reduce
all reversibly oxidized cysteines (purple), and alkylated with NEM
or IAM. Irreversibly oxidized cysteines (green) are subsequently oxidized
to sulfonic acid with performic acid. The sample is then trypsinized,
sulfonic acid-modified peptides are enriched with PA-ND, and eluted
peptides are analyzed by LC-MS/MS to identify sites of hyperoxidation.
4
Reactive
Nitrogen Species (RNS) in Biological
Systems
The prototypical RNS produced in biological systems
is nitric oxide
(•NO). In cells, the estimated steady state concentration
and half-life of this species is 100 pM–5 nM and ∼0.1–2
s, respectively.
234
Although •NO is more stable than H2O2 in cells, protein
and small molecule NO-donors are believed to be a relevant source
of •NO in biological systems. In general, •NO is a modestly reactive radical and does
not inflict indiscriminate
damage on biomolecules. Due to its gaseous and neutral nature, •NO is four times more
soluble in membranes than in
aqueous solution,
235
which permits its
diffusion across membranes and, in this context, •NO can function as an autocrine
and a paracrine signal within a 100–200
μm radius of the production site.
236
For example, •NO was recently shown to function
as a paracrine signal to regulate active T cell expansion in lymph
nodes.
237
Initially deemed toxic, •NO was later identified as the first gas known to act
as a biological second messenger in mammals where it regulates vasodilation/relaxation
of underlying smooth muscle cells.
238
Since
these seminal discoveries, roles for •NO have been
established in a range of biological processes including proliferation,
apoptosis, angiogenesis, and host defense.
239
•NO appears to be metabolized by autoxidation
to nitrite (NO2
–) and nitrate (NO3
–), which occurs about 30-fold faster within
the interior of lipid bilayers than in aqueous solution (Chart 12).
240
•NO can also react rapidly (1.1 × 109 M–1 s–1) with nitrogen dioxide (•NO2) to generate
additional nitrosating compounds such
as dinitrogen trioxide (N2O3) (Chart 12).
241
The production
of N2O3 is a trimolecular reaction with oxygen
and two molecules of •NO and is, therefore, not
favorable at low •NO concentrations (Chart 12). In turn, these •NO oxidation
products play important roles in physiological and pathological processes.
241,242
In addition to autoxidation, •NO reacts rapidly
(1010 M–1 s–1) with
O2
•
— to generate peroxynitrite
(ONOO–), which is reactive and damaging to biomolecules,
analogous to •OH.
242c,243
Though it
will not be discussed further here, ONOO– is an
important RNS in many biological settings; the interested reader is
referred to the following source for additional information.
242c
Chart 12
Formation and Transformation of Biologically
Relevant Reactive Nitrogen
Species (RNS)a
a
Metabolism of nitric oxide
(•NO) ultimately involves oxidation to nitrite (NO2
–) and nitrate (NO3
–) and there is evidence for this process being reversible (dashed
arrow). Though a generally unreactive radical, •NO can react with molecular oxygen
to generate nitrogen dioxide radical
(•NO2). Radical-radical coupling of •NO2 with a second molecule of •NO affords dinitrogen
trioxide (N2O3) via an
ultimately trimolecular reaction. The trimolecular nature of this
reaction makes N2O3 generation poorly favored
at low •NO concentrations. In the presence of O2
•–, •NO can react
to yield peroxynitrite (ONOO–). Color intensity
correlates to relative RNS reactivity.
4.1
•NO Production and Metabolism
4.1.1
•NO Synthases (NOS)
Enzymatic •NO production is predominantly mediated
by the heme- and flavin-containing •NO synthases
(NOSs), which catalyze the formation of •NO from
NADPH, molecular oxygen, and l-arginine (Figure 14a).
244
The linear arrangement
of NOSs reveal three domains: the N-terminal oxygenase domain, C-terminal
reductase domain, and the connecting calmodulin (CaM)-binding site.
The oxygenase domain contains the heme and (6R)-5,6,7,8-tetrahydrobiopterin
(BH4) cofactors, and the l-arginine binding site,
while the reductase domain has a binding site for NADPH and houses
the FAD and FMN flavin cofactors (Figure 14b).
245
•NO is produced
by the flow of electrons derived from NADPH through the flavins in
the reductase domain to the heme in the oxygenase domain, where oxygen
and l-arginine are bound. NOS functions as a dimer in which
the large (3000 Å) dimer interface in the oxygenase domain includes
the BH4 binding site and is stabilized by a zinc ion that
is coordinated by two cysteine residues in a conserved CXXXXC motif
per monomer.
246
Dimerization helps to structure
the active-site pocket containing the heme cofactor and the l-arginine binding site,
and there is evidence for electron flow occurring
between monomers (Figure 14b).
247
Figure 14
Nitric oxide (•NO) production
by nitric oxide
synthases (NOS). (a) Reaction catalyzed by NOS. (b) Linear arrangement
of NOS. NOS contain three distinct domains: the N-terminal oxygenase
domain (gray), the C-terminal reductase domain (light blue), and the
connecting calmodulin (CaM) binding site (purple). All three NOS isoforms
encode a C-terminal regulatory tail and endothelial NOS (eNOS) and
neuronal NOS (nNOS) additionally contain an autoinhibitory region
in the reductase domain. Activation of eNOS and nNOS requires binding
of Ca2+/CaM whereas inducible NOS (iNOS) is expressed with
Ca2+/CaM tightly bound. The production of •NO by NOS involves translocation of electrons
from NADPH bound in
the reductase domain through the FAD and FMN cofactors. The electrons
are then transferred to the heme prosthetic group in the oxygenase
domain where l-arginine and molecular oxygen bind. The tetrahydrobiopterin
(BH4) cofactor in the oxygenase domain appears to regulate
the nature of reactive intermediates generated by NOS (e.g., •NO versus O2
•–).
The CaM binding site, autoinhibitory region, and regulatory tail are
believed to regulate enzyme activity by influencing the efficiency
of electron transfer between the reductase and oxygenase domains.
All NOS isoforms function as homodimers that are stabilized by a zinc
ion coordinated by two cysteine residues from each monomer and there
is evidence suggesting that electrons are transferred between monomers
(as depicted). NOS enzymes additionally contain sequences such as
PDZ domains (deep blue) that facilitate protein–protein interactions,
which are involved in subcellular targeting of NOS and for mediating
direct interactions with protein targets of •NO.
There are three known NOS isoforms
that exhibit 51–57% sequence
homology among the human enzymes: inducible NOS (iNOS), endothelial
NOS (eNOS), and neuronal NOS (nNOS). iNOS is expressed in a wide range
of cell types and tissues including phagocytic cells where it produces •NO for cytotoxic
purposes. eNOS is expressed primarily
in vascular endothelial cells where •NO functions
in a paracrine manner to regulate vasodilation. Lastly, nNOS is expressed
primarily in the brain where •NO is involved in
neurotransmission.
245d,245e,248
The NOS isoforms can be classified as those that exhibit constitutive
(eNOS, nNOS) and inducible (iNOS) expression as well as those that
are activated in a Ca2+-dependent (eNOS, nNOS) and independent
(iNOS) manner. All NOS isoforms have a C-terminal tail that appears
to regulate enzyme activity.
249
Moreover,
eNOS and nNOS additionally contain an autoinhibitory loop in the flavin-binding
domain that is believed to hinder efficient electron transfer between
the FAD and FMN cofactors.
250
In response
to receptor activation, a number of growth factors, cytokines, and
G-protein coupled receptor (GPCR) agonists have been shown to induce
an increase in intracellular calcium, which binds tightly to CaM.
251
The Ca2+/CaM complex then binds
to the CaM binding site in NOS and is believed to induce a conformational
change involving the C-terminal tail and the autoinhibitory loop to
optimally orient the reductase and oxygenase domains for efficient
electron transfer, which is the rate limiting step in •NO production.
249,250,252
In contrast to eNOS and nNOS, iNOS, whose expression is controlled
by cytokines and interleukins, is expressed with tightly bound Ca2+/CaM and thus functions
independent of the intracellular
calcium concentration. Rate constants for •NO production
range from 200 min–1 for iNOS, which produces high
concentrations of •NO over the course of hours in
immune cells, to 100 min–1 for nNOS and 20 min–1 for eNOS.
249,253
The diverse •NO production rates suggest structural and regulatory differences
between isoforms that influence inherent electron flux rates. Moreover,
NOS isoforms appear to be regulated, in part, by the rate of product
(•NO) release.
254
In addition to intrinsic factors, extrinsic factors such as phosphorylation
and protein–protein interactions also regulate NOS activity.
245e,248
The serine/threonine kinase AKT has been shown to phosphorylate
eNOS in the reductase domain and the C-terminal regulatory tail.
255
Accessibility of these phosphorylation sites
appears to be regulated by Ca2+/CaM binding, and both AKT-mediated
eNOS phosphorylation and sustained eNOS activation were recently found
to be necessary for the tumorigenic properties of oncogenic Ras.
256
Additional extrinsic factors that regulate
NOS activity are protein–protein
interactions, as illustrated by Ca2+/CaM-mediated eNOS
and nNOS activation. Interestingly, eNOS and nNOS have also been shown
to interact with the ATP-dependent molecular chaperone, Hsp90, which
may facilitate Ca2+/CaM-induced conformational changes.
257
Protein–protein interactions have also
been shown to mediate membrane localization of the cytoplasmic NOS
enzymes for cell signaling. In contrast to certain NOX complexes,
which have been shown to assemble at activated membrane receptors,
12,70
NOS appears to preassociate with receptors and distinct membrane
microdomains prior to ligand stimulation. For example, nNOS has a
unique N-terminal PDZ domain, which mediates protein–protein
interactions and directs intracellular proteins into multiprotein
complexes.
258
In neuronal cells, nNOS is
targeted to postsynaptic sites through binding of its PDZ domain to
corresponding domains of proteins including PSD-95 and PSD-93.
259
PSD-95 binds to the NMDA receptor (NMDAR) thereby
mediating a link between the receptor and nNOS, and this complex forms
in the absence of NMDA.
260
By an independent
mechanism, eNOS is localized to the membrane through direct interaction
with the bradykinin 2 receptor (B2R).
261
All three NOS isoforms contain the conserved sequence FXXFXXXXW,
which is a putative caveolin binding site, in their oxygenase and
reductase domains.
246b,262
In endothelial cells and cardiac
myocytes, eNOS is localized to caveolae, a specialized form of lipid
raft, by direct interaction with caveolin-1 and caveolin-3.
263
Interestingly, eNOS is held in an inactive
conformation by its interaction with caveolin and B2R, which is released
upon Ca2+/CaM-binding or receptor activation, respectively.
261,264
Membrane localization of eNOS regulates •NO production
in endothelial cells by mediating l-arginine availability.
In endothelial cells, eNOS forms a complex with the cationic amino
acid transporter CAT-1 and arginosuccinate lyase (ASL).
265
CAT-1 is responsible for arginine transport
266
and ASL works in concert with arginosuccinate
synthase (ASS1) to recycle l-citrulline, the amino acid product
formed by NOS, back to l-arginine. Additionally, ASL funnels l-arginine imported by
CAT-1 to eNOS.
265
In this way, eNOS complex formation with CAT-1 and ASL regulates •NO production by
modulating local substrate availability,
somewhat like the regulation of flavin availability through NOX complex
formation with riboflavin kinase.
70
Under certain circumstances, NOS can form O2
•– instead of •NO.
267
Such
conditions include the absence of the BH4 cofactor
268
and uncoupling of electron transfer within
NOS via conformational changes that permit direct oxygen interaction
with the flavins in the reductase domain
264b
or S-glutathionylation of Cys689 and Cys908 in
the reductase domain of eNOS.
163a
The more
recent finding that S-glutathionylation influences
reactive intermediate production by NOS is interesting given that
coproduction of ROS and RNS can result in generation of the aggressive
oxidant ONOO– and might be a mechanism to deter
ONOO– production.
163a
Although further studies are required to determine whether ROS mediate
NOS glutathionylation, the cross-regulation proposal is also supported
by the observation that •NO can inhibit NOX-mediated
oxidant production in plant immune cells.
269
•NO has been shown to regulate a range of
processes
including proliferation, apoptosis, angiogenesis, host defense, and
regulation of vasodilation.
239
To elucidate
the role of unique NOS isoforms in regulating these diverse biological
processes, mouse models of NOS deficiency have been generated.
270
Additionally, much effort has been aimed at
developing selective small molecule inhibitors for individual NOS
isoforms.
245d
Selective inhibitors have
been developed for iNOS that act in competition with l-arginine
in which selectivity is achieved through interactions with the novel
substrate-binding site in this isoform, as compared to nNOS and eNOS.
271
Two iNOS inhibitors have been used to probe
the roles of this isoform in several animal models of diseases in
which iNOS has been implicated. More recently, a therapeutic role
for iNOS selective inhibitors has been shown for lung regeneration
in a mouse model of full-established emphysema.
272
Selective iNOS inhibitors have also been used in clinical
studies for medical conditions involving lung damage including chronic
obstructive pulmonary disease (COPD) and asthma.
273
The continued development of NOS inhibitors will further
our understanding of distinct roles for each of these isoforms in
diverse biological processes and will certainly continue to uncover
additional avenues for therapeutic intervention for diseases where
NOS are implicated.
In plants, •NO has been
shown to be involved
in seed germination, root growth, respiration, stromal closure, and
adaptive responses to biotic and abiotic stresses.
274
Although the existence of bona fide NOS isoforms in plants
remains controversial, the cytoplasmic enzyme nitrate reductase is
a recognized source of •NO in these organisms, as
reviewed elsewhere.
275
Alternative pathways
for NOS-independent •NO production have also been
identified in different plant cell compartments, such as peroxisomes,
mitochondria, and the apoplasm.
275
The discovery that RNS are produced as second messengers to regulate
a number of biological processes has spurred the development of methods
to specifically detect these species in cells. Historically, •NO production has been
detected indirectly by monitoring
its oxidation products, namely N2O3, NO2
–, and NO3
– by colorimetric, spectroscopic and fluorescent means.
276
The field has more recently seen the development
of direct methods to specifically detect not only •NO, but also ONOO– and nitroxyl
(HNO) by exploiting
the unique reactivity of each of these species. These methods include
nanotube-,
277
cell-,
278
protein-,
279
small molecule-,
280
and electrochemical-based
281
assays. To date, no RNS probes are available that permit
species detection in specific subcellular compartments or organelles.
Improvements to the current technology including reversibility are
required for regio- and spatiotemporal resolution of RNS production
and the interested reader is referred to the following review for
additional information.
282
4.1.2
•NO-Metabolizing Enzymes
Unlike O2
•– and H2O2, for which ROS metabolizing enzymes exist to regulate
their levels, far less is known about enzymatic regulation of •NO availability. As
previously mentioned, •NO autooxidizes to NO2
– and NO3
–, however, it was recently shown that •NO oxidation to NO2
– can also be catalyzed
by the abundant plasma multicopper oxidase, ceruloplasmin.
283
NO2
– and NO3
– have traditionally been thought of as
inert byproducts of •NO; however, there is increasing
evidence for enzymatic reduction of NO2
— to regenerate •NO by xanthine oxidase,
284
by nitrate reductase in plants,
275
or through reaction with deoxyhemoglobin in
the vasculature.
285
NO2
– reduction could also facilitate •NO release at sites distant from NOS. Along these
lines, fatty acids
and proteins modified by •NO can similarly be reduced
to release •NO or act to transfer •NO to sites distal from NOS. Through protein–protein
interactions,
NOS has been found to localize to the plasma membrane, endoplasmic
reticulum, sarcoplasmic reticulum, and sarcolemmal caveolae where
NOS regulates a distinct set of proteins in each location.
286
This has spurred the hypothesis that NOS is
placed where it is needed for local action of •NO,
akin to NOX.
287
However, it is possible
that the aforementioned alternative mechanisms of •NO release and transport may extend
•NO signaling
to subcellular regions that are inaccessible by NOS, such as the nucleus,
288
or may enhance the paracrine activity of •NO.
289
4.2
Modification of Protein Cysteine Thiols by
RNS
Similar to O2
•–, •NO is a relatively unreactive radical and its primary
targets in cells include other radical species such as O2
•– and metals. Indeed, the propensity for •NO to coordinate to metals has been exploited
in the
development of •NO-specific small molecule fluorescent
detectors.
280a−280i
The first identified cellular target of •NO was
soluble guanylyl cyclase (sGC) in which •NO activates
sGC through binding reversibly to the prosthetic heme.
290
In endothelial cells, the •NO produced migrates through the vasculature to activate
sGC in the
underlying vascular smooth muscle cells to promote vasodilation.
291
•NO-mediated sGC activation
also stimulates mitochondrial biogenesis in brown adipose tissue.
292
In addition to sGC, •NO can
regulate other heme-containing proteins including ETC Complex IV,
where •NO binding inhibits cellular respiration
and ROS production under hypoxic conditions.
293
•NO can also control protein function through
iron–sulfur clusters, as documented for bacterial transcriptional
regulators, such as NsrR, SoxR, and FNR.
294
This form of regulation is thought to occur via •NO-mediated iron–sulfur cluster nitrosylation
and degradation.
295
It was recognized early on in the field
that, in addition to regulating protein function by coordination to
metal-based prosthetic groups, •NO could covalently
modify protein cysteines, a modification subsequently termed S-nitrosylation.
5c
Analogous to other oxPTMs, specificity in modification
appears to be imparted by cysteine reactivity, local protein environment,
and proximity to the oxidant source.
3b,12,194
In contrast to NOX signaling, (or, more likely, as
is less well established for NOX signaling) the proximity of protein
targets of •NO to the RNS source is frequently imparted
by direct interaction with NOS. As discussed above, NOS enzymes contain
structural features that facilitate protein–protein interactions,
and a number of NOS-interacting proteins including caspase-3,
296
cyclooxygenase-2,
297
and the postsynaptic scaffolding protein PSD-95,
260
have been shown to be S-nitrosylated after NOS activation.
Though still an active area of research, three prominent mechanisms
have been proposed to account for de novo S-nitrosothiol
formation, none of which involve direct reaction of •NO with thiols (Figure 15a–c).
As mentioned
above, •NO can be converted to the nitrosating compound
N2O3 (Chart 12). The
initial reaction of •NO with molecular oxygen to
generate •NO2 and subsequent radical–radical
combination of •NO with •NO2 permits N2O3 production with a rate
constant of 109 M–1 s–1.
241
N2O3 can subsequently
react with a protein or low molecular weight thiolate to yield an S-nitrosothiol (Figure
15a). Given
the requirement for two molecules of •NO in this
reaction, the route is not favorable at low concentrations of this
species. Alternatively, •NO2 or other
radical species such as O2•–
298
can promote the one-electron oxidation of a
protein or low molecular weight thiolate to generate a thiyl radical
that can undergo radical–radical combination with •NO to yield the S-nitrosothiol
(Figure 15b). While evidence exists to support both of these
mechanisms,
299
a third route has been postulated
to account for S-nitrosylation of some proteins.
This mechanism, which has been demonstrated for both hemoglobin
300
and nitrophorin,
301
relies on the propensity of •NO to bind to heme
prosthetic groups in which the heme-bound NO undergoes reductive nitrosylation
of the heme prosthetic group and autotransfer to a thiol within the
same protein to generate an S-nitrosothiol (Figure 15c). Though it will not be further
discussed here, •NO and •NO-derived species can also
modify other amino acids, including tyrosine.
Figure 15
Formation and subsequent
reactions of S-nitrosothiols.
Three prominent mechanisms for S-nitrosothiol formation
include (a) reaction of a protein or low molecular weight thiolate
with N2O3, (b) formation of a thiyl radical
upon initial reaction of a thiolate with •NO2 and other radical species and subsequent
radical–radical
combination with •NO, and (c) autotransfer of heme-bound +NO to a nearby cysteine
thiolate as has been demonstrated
for hemoglobin and nitrophorin. (d) Once formed, an S-nitrosothiol can react with
a neighboring cysteine residue either
within the same or an adjacent protein, or with GSH (not shown) to
undergo transnitrosylation (eq 1) or disulfide bond formation (eq
2). Alternatively, an S-nitrosothiol can be hydrolyzed
to release the free thiolate and nitrite (HNO2) (eq 3)
or a sulfenic acid and HNO (eq 4). In each case, the pK
a of the sulfur in the S-nitrosothiol,
in part, influences which product is formed. In most cases, transnitrosylation
and release of a free thiolate are favored upon reaction with a second
cysteine or water, respectively due to the high pK
a of the HNO leaving group.
Like sulfenic acid, the formal oxidation number of the sulfur
atom
in S-nitrosothiol is 0; despite this apparent similarity,
there are many important differences between these modifications.
The S-nitrosothiol group is not ionizable,
302
can undergo hydrolysis to give sulfenic acid,
303
or react with a thiol (Figure 15d). Interestingly, reaction of an S-nitrosothiol
with a protein thiol or GSH does not always yield the mixed disulfide,
but can instead (and perhaps more frequently) facilitate a process
known as transnitrosylation (Figure 15d). The
ability to undergo transnitrosylation is due to the different chemical
properties of +NO compared to the hydroxyl in sulfenic
acid, and will be discussed further in the following subsection. Transnitrosylation
is increasingly viewed as another physiologically relevant mechanism
for S-nitrosothiol formation
5c,287,304
and studies of protein S-nitrosylation often use +NO donors such as
GSNO, S-nitrosocysteine (SNOC), and S-nitroso-N-acetyl-d,l-penicillamine (SNAP).
260,297,305
In vitro rate constants for
de novo thiol S-nitrosylation in human and bovine serum albumin are
on the order of 103 to 104 M–1 s–1.
306
In contrast,
in vitro S-nitrosylation rate constants for glutathione and other
low molecular weight thiols are on the order of 105 to
107 M–1 s–1.
306,307
Since de novo S-nitrosothiol formation depends
on the combined reactivity of two •NO, molecular
oxygen, and a thiol (Chart 12, Figure 15a and b), GSH and abundant proteins such as
Trx,
albumin, and hemoglobin could be primary targets of S-nitrosylation.
Indeed, as previously mentioned, the aforementioned protein and low
molecular weight thiols can function as +NO donors, and
are proposed to extend •NO signaling to proteins
distal to its site of production both within cells and as a paracrine
signal.
288,289
The tendency for a particular cysteine
residue to undergo transnitrosylation
appears to be regulated, in part, through steric (e.g., accessibility to +NO donors)
and electrostatic factors.
287,305h,308
Computational studies to identify
a consensus sequence for S-nitrosylation have uncovered an acid–base
motif, located distal to the modified cysteine in the protein tertiary
structure among some S-nitrosothiols.
304,309
This charged
nature of the acid–base motif has been proposed to engage in
protein–protein and protein-GSH interactions to facilitate
transnitrosylation. A recent structural study, has revealed to two
additional sequence motifs proximal to the S-nitrosothiol
that facilitate reduction by Trx, though whether these particular
cysteines also participate in transnitrosylation from nitrosylated
Trx has not been established.
304
Beyond
these putative protein–protein interaction motifs, manual inspection
of S-nitrosothiol sites incidate that S-nitrosylated
cysteines may be directly flanked by an acid–base motif that
enhance reactivity or decrease pK
a.
310
An additional feature of the environment surrounding
S-nitrosylation sites is hydrophobicity.
311
Hydrophobic protein surfaces could potentially concentrate nonpolar •NO and molecular
oxygen, permitting the formation of
N2O3 directly at the site of S-nitrosylation.
Neither the local acid–base motif nor the hydrophobic environment
are uniformly conserved, which is consistent with a similar lack of
sequence bias for sulfenylation
9b
and may
be reflective of the numerous mechanisms for de novo and transnitrosylation.
304,312
Given the propensity for S-nitrosothiols
to participate
in transnitrosylation reactions, reducing systems are in place to
reverse S-nitrosylated thiols and the interested
reader is referred to the following reviews for a more thorough coverage
of the topic.
287,313
GSH can reduce a S-nitrosothiol to give the free thiol and GSNO. In turn, GSNO is
reduced
to regenerate GSH and release HNO by GSNO reductases (GSNOR).
314
GSNOR acts exclusively on GSNO and deficiency
of this enzyme increases the steady-state level of protein S-nitrosylation (which
can be further enhanced by iNOS activation
and may support a physiological role for GSNO as an +NO
donor).
314a,315
Protein S-nitrosothiols
can also be reduced by the Trx/TrxR system.
305e
Enzymes with primary functions unrelated to protein S-nitrosylation
may also act as denitrosylases, including PDI, xanthine oxidase, and
SOD,
313
though the physiological relevance
of these activities remains unclear.
Since its discovery, S-nitrosylation
has been implicated in the
regulation of proteins involved in cellular trafficking,
316
muscle contractility,
317
apoptosis,
305a,305e
circulation,
318
neural transmission,
260,319
and host
defense.
320
However, it is important to
keep in mind that most S-nitrosylated proteins that have been identified
to date are derived from studies with exogenous +NO donors
employed at unphysiological concentrations (though protein S-nitrosylation
from endogenous •NO production has been observed
in neurons and immune cells, the latter of which produce high concentrations
of •NO for bactericidal purposes).
245d,248
In plants, attempted microbial invasion triggers the hypersensitive
response, a programmed execution of plant cells at the sites of infection.
This process involves the generation of NOS-derived •NO with subsequent production
of NOX-derived O2
•– and the chemical messenger, salicylic acid.
320
Interestingly, S-nitrosylation of NPR1, a master regulator
of salicylic acid-mediated defense genes, promotes its oligomerization
and cytoplasmic retention. S-nitrosylation is reversed by Trx in a
salicylic acid-stimulated manner to facilitate NPR1 monomerization
and nuclear translocation.
305f
Whether
microbial invasion in plants induces cell death appears to be regulated,
in part, by the extent of •NO and O2
•– production, which together produce the more
reactive ONOO–. Interestingly, both NOS
321
and NOX
269
have
been found to be inhibited by S-nitrosylation, shedding
light on a potential regulatory mechanism to control ROS and RNS coproduction
in immune responses, which could be conserved across species. More
recently, it was shown in a mouse model of Clostridium difficile infection that host-derived
•NO S-nitrosylates
and inhibits clostridial small molecule-activated glucosylating toxins,
thereby preventing toxin cleavage and cell entry.
322
This represents a unique mechanism for •NO-mediated pathogen detoxification.
In addition to being involved
in the immune response, NOS may also
play a role in synaptic plasticity (the strength of connection between
two neurons), which is relevant to processes, such as spatial learning.
323
nNOS is recruited to the membrane prior to
synaptic signaling through its interaction with PSD-95, which physically
links nNOS to NMDAR (Figure 16).
259a,324
Stimulation of NMDAR triggers calcium entry and activates •NO production via the proximal
nNOS. PSD-95 is localized to the membrane
through a dynamic reversible cycling of S-palmitoylation, a posttranslational
lipid modification, of two N-terminal cysteine residues.
325
It was recently shown that nNOS activation
mediates S-nitrosylation of these same cysteine residues in PSD-95
thereby preventing S-palmitoylation and reducing PSD-95 and hence
nNOS membrane localization subsequent to neuron activation (Figure 16).
260
This study highlights
the intriguing possibility that differential modification of cysteines
may represent a general paradigm in cell signaling and, in this context,
S-nitrosylation of PSD-95 may function to regulate the duration of
NMDA signaling. NMDAR activation also regulates the recruitment of
AMPA receptors (AMPAR) to the synapse to propagate signaling. PSD-95
regulates AMPAR through its interaction with stargazin
326
and was recently shown to be S-nitrosylated
in response to NMDA signaling, thereby enhancing its binding to AMPAR
(Figure 16).
319a
Lastly, nNOS-derived •NO can also regulate neural
cells at the level of gene transcription. For example, S-nitrosylation
of histone deacetylase 2 was found to induce its release from chromatin,
permitting increased acetylation of histones surrounding genes involved
in neural development and promoting transcription.
305d
Figure 16
Regulation of neuronal signaling by S-nitrosylation. PSD-95
is
a scaffolding protein that localizes to postsynaptic densities by
reversible S-palmitoylation of two cysteine residues in the N-terminal
region. In the absence of ligand, nNOS is physically linked to the
NMDA receptor (NMDAR) at the neuronal plasma membrane via a mutual interaction with
PSD-95. PSD-95 similarly localizes stargazin
near NMDAR. NMDA binding to NMDAR facilitates calcium entry, which
activates nNOS mediating •NO production and subsequent
S-nitrosylation of PSD-95 on the same cysteine residues that undergo
S-palmitoylation. S-nitrosylation thereby prevents PSD-95 lipidation
and decreases membrane association of PSD-95 and, hence, nNOS. Stargazin
is similarly S-nitrosylated, which enhances its interaction with the
AMPA receptor facilitating its recruitment to the postsynaptic densities.
Expression of NOS isoforms is
regulated by Ca2+/CaM
binding. S-Nitrosylation of calcium transporters has been increasingly
demonstrated, revealing a potential positive feedback loop. In one
instance, S-nitrosylation of ryanodine receptors in skeletal muscle
327
and neurons
328
releases
intracellular calcium stores to potentiate signaling that, in the
latter case, are required for neural synaptic plasticity and can also
contribute to neuronal cell death. S-Nitrosylation can also regulate
entry of extracellular calcium as S-nitrosylation of transient receptor
potential (TRP) cation channels mediates a conformational change in
endothelial cells that opens the pore to permit calcium entry,
329
which may similarly function as a positive
feedback loop to potentiate NOS activity.
In addition to promoting
cell signaling responses, dysregulation
of S-nitrosylation has been implicated in disease,
including neurodegenerative disorders.
256,287,305a,305b,330
The E3 ubiquitin ligase parkin, which regulates the degradation
of proteins important to survival of dopamine neurons, is S-nitrosylated
in a mouse model of Parkinson’s disease (PD) and in brains
of patients with PD.
305b
Parkin S-nitrosylation
inhibits its ubiquitin ligase activity, which impairs ubiquitination
of its substrate proteins and may contribute to the degenerative process.
It has also been shown that PDI, an ER-resident enzyme that facilitates
proper protein folding and protects neuronal cells against ER dysfunction,
is S-nitrosylated in brain samples manifesting sporadic PD or Alzheimer’s
disease.
305c
PDI S-nitrosylation inhibits
its activity, resulting in activation of ER stress pathways (including
the unfolded protein response) and abrogates PDI-mediated attenuation
of neuronal cell death triggered by ER stress, which could contribute
to neurodegenerative disorders. More recently, amyloid-β, a
key mediator in Alzheimer’s disease was found to induce •NO production, which triggered
mitochondrial fission,
synaptic loss, and neuronal damage.
305a
This effect was attributed, in part, to S-nitrosylation of dynamin-related
protein 1 (Drp1), a protein involved in regulation of mitochondrial
fission. S-nitrosylated Drp1 is increased in brains of human Alzheimer’s
disease patients where it is postulated to contribute to disease pathogenesis.
4.3
Methods for Detecting RNS-Modified Cysteines
The discovery of protein S-nitrosylation has spurred the development
of methods for its detection.
138,331
Initial indirect chemiluminescent,
colorimetry and electrochemical approaches relied upon detection of
NO liberated from S-nitrosothiols by mercury.
332
However, these methods are artifact prone because
of interference from other metabolites in the sample, such as NO2
–. Moreover, indirect spectroscopic methods
report only on the total amount of S-nitrosothiols
and do not permit identification of the target proteins. While NO
is released with metal treatment, the protein thiol remains coordinated
to mercury and strategies to identify metal-coordinated proteins have
been reported.
304,333
A limitation to this method
is that other metal-interacting modifications, including protein-S-GSH disulfides
and, perhaps cysteine sulfenic acid and
persulfide, can similarly be detected and complicate selective analysis.
While alternative methods for S-nitrosothiol detection
have since been developed, these indirect spectroscopic methods are
still used to quantify the total amount of S-nitrosylated protein
in purified samples.
305g
S-Nitrosylated
proteins can also be identified using an anti-S-nitrosocysteine
antibody or by MS, however, these methods are not well suited to identify S-nitrosothiols
in complex protein mixtures and do not facilitate
enrichment of oxidized proteins.
Chemical methods for direct
and selective detection of protein S-nitrosylation have also been
reported. In contrast to sulfenic acids, which have one electrophilic
center, S-nitrosothiols contain two, allowing nucleophiles
to attack the sulfur or nitrogen (the reaction site is influenced
by the relative stability (i.e., pK
a)
of the leaving group). As previously discussed, the pK
a of a free thiol group is ∼8.5 but can be significantly
modulated in the protein environment.
334
The pK
a of HNO, the alternative leaving
group, is approximated at 11.4.
242a,335
On the basis
of these relative pK
a values, in the majority
of cases, the thiol is predicted as the preferred leaving group. This
leaving group preference provides a chemical rationale for transnitrosylation;
however, S-nitrosothiols can also form en route to
disulfide bonds (Figure 15d). In these cases,
it is possible that features of the S-nitrosothiol
environment favor disulfide bond formation by increasing the electrophilicity
of the sulfur through pK
a modulation (e.g.,
increasing the thiol pK
a) or by promoting
protonation of HNO in the S-nitrosothiol. The latter
would be analogous to protonation of a hydroxyl group prior to nucleophilic
attack to facilitate expulsion as water. Importantly, the disparate
electrophilic centers in sulfenic acids and S-nitrosothiols
can be exploited to permit chemical discrimination between these forms.
To date, the most popular procedure for S-nitrosothiol
detection, is known as the biotin switch technique (BST).
336
As shown in Figure 17a, the BST is an indirect method that involves blocking free
thiols
by S-methylthiolation with methylmethane thiosulfonate (MMTS, 38), selective reduction
of S-nitrosothiols
with ascorbate, and labeling nascent thiols with N-[6-(biotinamido)-hexyl]-3′-(2′-pyridyldithio)-propionamide
(biotin-HPDP). The reaction of thiols with biotin-HPDP yields a mixed
disulfide adduct that can be detected by avidin blot. Additionally,
the biotin handle permits enrichment of labeled proteins for proteomics
analysis. As with the two previously described indirect chemical methods,
the success of the BST is dependent upon complete blocking of free
thiols and the selectivity and efficiency of the reducing agent. Though
the mechanism is not entirely clear, S-nitrosothiol
reduction may involve nucleophilic attack of ascorbate (39) at the electrophilic nitrogen
center to release the thiol (Chart 13). In accordance with the leaving group bias
of
transnitrosylation versus disulfide bond formation, some S-nitrosothiols cannot be
reduced efficiently by ascorbate,
331a,337
which might be due to competing reaction at the electrophilic sulfur
center. Recently, the use of ascorbate as a selective reductant for S-nitrosothiols
has been questioned, because of the observation
that ascorbate can reduce some disulfides
179,338
and sulfenic acid, as recently shown for some 1-Cys Prxs.
172
In one instance, sinapinic acid was used in
place of ascorbate as it does not appear to react with disulfides.
339
Despite the limitations of the BST, this technique
is routinely used in diverse protein systems and led to important
advances in S-nitrosylation research.
226,260,269,288,329
Chart 13
Predicted Mechanism for the Reaction of Ascorbate with S-Nitrosothiol
Figure 17
Indirect and direct chemical methods for S-nitrosothiol
detection. (a) The biotin switch technique (BST) is an indirect differential
alkylation method that involves blocking free thiols (blue) with methylmethane
thiosulfonate (MMTS, 38), reducing S-nitrosothiols (purple) with ascorbate, and labeling
nascent thiols
with Biotin-HPDP. The samples are then analyzed by nonreducing avidin
blot in which S-nitrosylation correlates to increased
signal intensity. (b) Quantification of protein S-nitrosylation with
d-Switch. The d-Switch technique combines the BST with isotopically
labeled NEM in which free thiols (blue) are blocked with d
0-NEM, S-nitrosothiols (purple) are reduced
with ascorbate and labeled with d
5-NEM.
The samples are subsequently separated by SDS-PAGE, digested in-gel
with trypsin, and the resulting peptides are analyzed by LC-MS/MS.
The extent of protein S-nitrosylation is determined by the ratio of d
5-NEM to d
0-NEM
signal intensity. (c–e) Triarylphosphine-based methods to directly
modify S-nitrosothiols. (c) Triarylphosphine reagent 40 reacts with S-nitrosothiols
to yield a
disulfide-bonded biotin adduct. (d) Compound 41 is oxidized
upon reaction with S-nitrosothiols to yield a fluorescent
compound that comments on the presence of S-nitrosothiols,
but does not covalently modify oxidized proteins. (e) Water-soluble
triarylphosphine 42 appears to form a stable S-alkylphosphonium adduct as monitored
by 31P
NMR and mass spectrometry.
Improvements to the BST have been made that involve biotin
enrichment
of trypsin-digested peptides,
310,340
resin-assisted capture,
341
fluorescence labeling,
342
and a microarray-based assay.
305h
The latter case shows a small percentage of false positives and
does not cover the entire proteome; nonetheless, it permits rapid
identification of candidate S-nitrosylated proteins
and allows for the direct comparison and assessment of chemically
distinct +NO donors. More recently, Thatcher and colleagues
developed a quantitative approach termed d-Switch that combines the
BST with isotopically labeled NEM (d
5-NEM)
(Figure 17b).
343
Future adaptations of the d-Switch technique could incorporate a
biotin affinity handle to permit sample enrichment analogous to ICAT.
Methods for direct chemical modification of S-nitrosothiols
have also been reported. For example, triarylphosphines have shown
promise as chemical probes for S-nitrosylation
344
and the interested reader can find additional information
about this chemistry from the following review.
331b
In the first demonstration of this approach, a small-molecule S-nitrosothiol model
underwent reductive ligation with a
triarylphosphine ester.
344
A variation
on this theme involves reductive ligation of an S-nitrosothiol with a biotinylated
triarylphosphine thioester (40) in a THF-PBS system to generate a disulfide linkage
with
biotin (Figure 17c).
345
The triarylphosphine reduction reaction has also been adapted to
generate a new fluorescent probe (41) to monitor S-nitrosothiol content in recombinant
proteins, but it is
not currently amenable to identification of S-nitrosylated proteins
in complex mixtures (Figure 17d).
346
King and colleagues have reported the water-soluble
triarylphosphine (42) that reacts with S-nitrosothiols to give a stable S-alkylphosphonium
adduct detectable by 31P NMR and MS (Figure 17e).
347
A future modification of
this reagent could incorporate an affinity handle for protein enrichment
(though the anionic nature of 42 likely precludes membrane
permeability for cellular studies). Interestingly, while the S–P
bond is usually labile, steric hindrance imparted by the substituted
aryl ligands and aromatic stabilization of the phospho cation is believed
to stabilize the S-alkylphosphonium adduct. Future
work with triarylphosphine reagents will need to address cross reactivity
with disulfides and sulfenic acids. From the perspective of selectivity,
only the strategy presented in Figure 17c would
be able to rigorously discriminate between S-nitrosothiols
and disulfides or sulfenic acids, as biotin disulfide formation would
be unique to this species (Chart 14).
Chart 14
Reaction
Mechanism of Triarylphosphine 40 with a Protein S-Nitrosothiol to Yield a Disulfide-Bonded
Biotin Adduct
(Equation 1) and Potential Reaction of Sulfenic Acids or Disulfides
with 40 (Dashed Arrows) Should Not Yield the Same Adduct
(Equation 2)
5
Reactive
Sulfur Species (RSS) in Biological
Systems
As we have seen, oxidation of protein and low molecular
weight
thiols generates a wide range of sulfur-containing products including
disulfides, thiosulfinates, sulfenic acids, and S-nitrosothiols. Each modification
is capable of propagating redox
transformations that involve oxidation of other thiols analogous to
ROS and RNS (Figures 2, 5, and 15d). As follows, these chemically reactive
forms of cysteine can be classified as reactive sulfur species (RSS).
213,348
Several “nonspecific” peroxidases, such as horseradish
peroxidase, can also oxidize thiol substrates by one-electron oxidation
to form thiyl radicals, which also represent an important class of
RSS.
349
Once formed, this radical species
can participate in a variety of chemical reactions. Of particular
note, thiyl radicals can react with a thiolate affording a disulfide
radical anion intermediate, which culminates later in disulfide and
O2
•– formation.
In addition
to reactive cysteine species in proteins, inorganic
sulfur-containing species are also classified as RSS. The prototypical
inorganic RSS is hydrogen sulfide (H2S), which is the most
stable reactive intermediate considered in this review with a half-life
on the minute time-scale.
350
Along with •NO and carbon monoxide, H2S is produced
in biological systems where it functions as a gasotransmitter to regulate
diverse biological processes as an autocrine, paracrine, and endocrine
signal.
351
H2S is a weak acid
with a pK
a1 and pK
a2 of 6.9 and >12 and, therefore, exists primarily in the
dissociated
thiolate form (HS—) at physiological pH (though
H2S is commonly used to refer to all species: H2S, HS—, S2-).
352
Similar to other reactive intermediates, H2S
was first recognized as a toxic species when it was found to emanate
from sewers, and is produced as a toxic byproduct of industrial processes.
Research over the past two decades, however, has implicated H2S in a number of physiological
and pathological systems. Roles
for H2S in biology were initially suggested in vasodilation/relaxation,
subsequently as a synaptic modulator and neuroprotectant, and as a
regulator of inflammation.
350,351,352b,353
The latter has motivated the