Site-specific and redox-controlled S-nitrosation of
thioredoxin
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Barglow, K. T. et al. “Site-specific and Redox-controlled Snitrosation of Thioredoxin.” Proceedings of the National Academy
of Sciences 108.35 (2011): E600–E606. Web.
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http://dx.doi.org/10.1073/pnas.1110736108
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PNAS PLUS
Site-specific and redox-controlled
S-nitrosation of thioredoxin
Katherine T. Barglowa,1, Charles G. Knutsonb,1, John S. Wishnokb, Steven R. Tannenbaumb,c, and Michael A. Marlettaa,d,2
a
Department of Chemistry and QB3 Institute, University of California, Berkeley, CA 94720; dDepartment of Molecular and Cell Biology, University of
California, Berkeley, CA 94720; bDepartment of Biological Engineering, Massachusetts Institute of Technology, Cambridge, MA 02139; and
c
Department of Chemistry, Massachusetts Institute of Technology, Cambridge, MA 02139
Protein S-nitrosation on cysteine residues has emerged as an
important posttranslational modification in mammalian cells. Previous studies have suggested a primary role for thioredoxin (Trx)
in controlling protein S-nitrosation reactions. Human Trx contains
five conserved Cys, including two redox-active catalytic Cys (Cys32
and Cys35) and three non-active-site Cys (Cys62, Cys69, and Cys73),
all of which have been reported as targets of S-nitrosation. Prior
reports have studied thermodynamic end points of nitrosation reactions; however, the kinetics of Trx nitrosation has not previously
been investigated. Using the transnitrosation agent, S-nitrosoglutathione, a kinetic analysis of the selectivity and redox dependence
of Trx nitrosation at physiologically relevant concentrations and
times was performed, utilizing a mass spectrometry-based method
for the direct analysis of the nitrosated Trx. Reduced Trx (rTrx) was
nitrosated 2.7-times faster than oxidized Trx (oTrx), and rTrx was
nitrosated selectively on Cys62, whereas oTrx was nitrosated only
on Cys73. These sites of nitrosation were confirmed at the peptide
level using a novel modification of the biotin-switch technique
called the reductive switch. These results suggest separate signaling pathways for Trx-SNO under different cellular redox states.
cysteine ∣ nitric oxide ∣ S-nitrosoglutathione ∣ transnitrosation
P
rotein S-nitrosation, the modification of thiols to nitrosothiols
(SNOs), has emerged as an important posttranslational modification (1). S-nitrosation of a thiol that is essential for structure
or function (e.g., catalytic Cys, metal-binding Cys, and Cys involved in key structural disulfide bonds) can lead to dramatic
functional and cellular outcomes. A prime example is the S-nitrosation of the catalytic cysteine in caspase-3 (Casp3) leading to
inhibition of apoptosis (5). S-Nitrosation of Casp3 involves a
cascade initiated by thioredoxin (Trx) (2–8), a ubiquitous, 12-kDa
cytosolic protein that contains five Cys (Cys32, Cys35, Cys62,
Cys69, and Cys73). We previously reported that human Trx1 is
selectively nitrosated on Cys73 both in vitro and in Jurkat cells
and that nitrosated Trx can form a complex with proapoptotic
Casp3 and selectively transfer the nitroso moiety to the active-site
Cys of Casp3 (4). This transnitrosation event blocks Casp3 catalytic activity and thus inhibits apoptosis as mentioned above (5).
These data represented an example of highly specific transnitrosation signal transduction mediated by a protein–protein interaction occurring in cells. Because Trx interacts with numerous
proteins and also maintains cellular redox homeostasis (9), Trx
may have a central role in transnitrosation reactions in cells.
Biological nitrosation begins with the formation of nitric oxide
(NO). NO reacts at diffusion-controlled limits in a radical–radical
recombination reaction with thiyl radicals of glutathione (γ-GluCys•-Gly, GS•), yielding S-nitrosoglutathione (GSNO) (10).
Additionally, aerobic solution decomposition of NO involves the
formation of N2 O3 , an efficient nitrosating agent that reacts
avidly with nucleophiles such as water (to form nitrite) (11), and
with glutathione (GSH) to form GSNO (12, 13). GSH is present
at millimolar concentrations in all mammalian cells and is important for maintaining a reduced intracellular environment (14).
GSNO has been detected both intra- and extracellularly in mamwww.pnas.org/cgi/doi/10.1073/pnas.1110736108
mals and is the most abundant SNO in cells, with concentrations
estimated at 1–5 μM (15–17). Once formed, SNOs, like GSNO,
can undergo a transnitrosation reaction with another thiol leading to the transfer of NO. Although GSNO reacts promiscuously
with protein thiols in vitro (18), the process is more selective
in vivo, with nitrosation often limited to a specific Cys within the
protein and modulation of GSNO levels controlling protein nitrosation (19). Formation of a specific protein–protein or protein–
ligand complex may align the SNO and thiol for intermolecular
transfer, suggesting a possible mechanism for transnitrosation
signaling specificity. Cascading transnitrosation reactions between
different, successive interacting partners is an intriguing paradigm
for SNO-mediated signal transduction in cells.
The two redox-active, catalytic Cys (Cys32 and Cys35) of Trx
reduce Cys disulfides on substrate proteins, producing a Cys32Cys35 disulfide on the now oxidized Trx (oTrx) (9). This disulfide
can then be reduced in vivo by the NADPH-dependent Trx reductase (TrxR) to regenerate reduced Trx (rTrx). The relative levels
of rTrx and oTrx are likely controlled by the redox state of the cell
and are related to both NADPH and GSH concentrations. The
three non-active-site cysteines (Cys62, Cys69, and Cys73) are
highly conserved in mammals but without a specific function.
Cys73 is the site for dimerization between Trx molecules, but this
appears to be an in vitro artifact.
There are inconsistent results pertaining to the specificity
of GSNO-dependent nitrosation of Trx: Nitrosation has been reported on all three of the non-active-site Cys. A crystal structure
of Trx was reported with a nitrosated Cys62 (20, 21), whereas
other studies indicate that Cys69 (3, 22, 23) or Cys73 (3–5, 7, 24)
are primary sites of nitrosation. Several groups reported that the
redox state of the active-site influences nitrosation of nonactive-site Cys. However, the reported redox-dependent, Cys specificity on Trx differs considerably under different experimental
conditions (3, 7). Wu et al. reported no detectable nitrosation on
rTrx and abundant nitrosation of oTrx on Cys73 (7), whereas
Hashemy and Holmgren reported that rTrx is nitrosated at
Cys69 and Cys73, with oTrx nitrosating only on Cys73 (3). Trx also
catalyzes the removal of nitroso groups (denitrosation) from
other protein thiols via the catalytic Cys (Cys 32 and Cys 35)
(2, 6, 8, 25). Given the reducing activity of Trx/TrxR/NADPH
(26), it is not surprising that Trx can act to denitrosate at high
concentrations. Despite these reports, the role of Trx in controlling cellular nitrosation/denitrosation, and the mechanistic contribution of each of the Trx Cys, remains unclear.
Author contributions: K.T.B., C.G.K., J.S.W., S.R.T., and M.A.M. designed research; K.T.B.
and C.G.K. performed research; K.T.B., C.G.K., J.S.W., S.R.T., and M.A.M. analyzed data;
and K.T.B., C.G.K., J.S.W., S.R.T., and M.A.M. wrote the paper.
The authors declare no conflict of interest.
This is a Contributed submission.
1
K.T.B. and C.G.K. contributed equally to this work.
2
To whom correspondence should be addressed. E-mail: marletta@berkeley.edu.
This article contains supporting information online at www.pnas.org/lookup/suppl/
doi:10.1073/pnas.1110736108/-/DCSupplemental.
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Contributed by Michael A. Marletta, July 2, 2011 (sent for review June 3, 2011)
Detection of protein SNOs is difficult. The commonly used
methods suffer from low sensitivity and high background. The
biotin-switch method and variations on the original procedure
(19, 25, 27–34) involve denaturation of the protein, alkylation
of free Cys, and precipitation of the protein. SNOs are then
reduced with ascorbate followed by the reaction of the newly
liberated Cys residues with a reactive group containing a tag for
later identification. The biotin-switch method is limited by high
background, low sensitivity, and nonspecific loss of protein during
sample workup (35). Interpretation of results is complicated by
differential ascorbate reduction and labeling rates of different
Cys-SNOs and Cys, respectively (35, 36). Furthermore, the sites
of nitrosation may be scrambled or lost upon denaturation. The
biotin switch and alternate methods (e.g., MS or UV detection)
(3, 7), are typically performed at supraphysiological concentrations of protein (10–100 μM) and GSNO (100 μM–1 mM) to aid
detection. Additionally, long reaction times (>10 min) are used,
which likely drives the reaction to a thermodynamic equilibrium;
thus, kinetic information on nitrosation is lost.
To gain a better understanding of the Trx-GSNO reaction
under biologically relevant conditions, we developed an in vitro,
MS-based methodology for the rapid and direct detection of
protein-S-nitrosation, allowing kinetic analysis of the transnitrosation reaction between GSNO and Trx while simultaneously
monitoring Trx redox status. We studied the selectivity and redox
dependence of this reaction under relevant cellular concentrations of Trx and GSNO. rTrx and oTrx not only show different
rates of nitrosation with GSNO, but also strikingly different
reaction selectivity, with nitrosation of oTrx on Cys73 and rTrx on
C62. We confirmed these findings with a peptide-level analysis,
using a modification of the biotin-switch technique that we call
the reductive switch, in which labeling is performed on native
protein to prevent loss or scrambling of nitrosation upon denaturation. Together, these techniques allowed a thorough analysis
of both the kinetics and the selectivity of Trx nitrosation by
GSNO. These results may have implications regarding selective
signaling under oxidative stress/hypoxic conditions.
Results
Direct Detection of Trx Nitrosation and Redox State with TOF-MS. The
redox state of the Trx active-site Cys may affect nitrosation (3, 7).
To explore the effect of the Trx redox state on the GSNO-dependent rate of nitrosation, conditions were first developed to fully
oxidize and reduce the Trx active-site Cys prior to reaction with
GSNO. The oxidation state of recombinant human Trx1 was
determined by TOF-MS analysis by deconvolution of the protein
envelope to determine protein mass. Treatment of humanTrx
(recombinant his10 -tagged protein) with Tris(2-carboxyethyl)
phosphine (TCEP) produced rTrx with a mass of 14,126.6
(Fig. 1 A and D). Oxidized Trx is often prepared by treatment
with H2 O2 (3, 7); however, incubation of Trx with 1 mM H2 O2
resulted in apparent thiol oxidation beyond the active-site Cys
(m∕z of 14,124.1) as well as in intermolecular disulfide bonds,
yielding a dimeric oTrx (Fig. 1 B and E). Therefore, more moderately oxidizing conditions were investigated. Treatment of Trx
with 1.2 equivalents of insulin (a natural Trx substrate) yielded
oTrx (m∕z of 14,124.7) that remained monomeric (Fig. 1 C
and F). Insulin treatment is a physiologically relevant way of generating oTrx in vitro. This 2-Da shift between rTrx and oTrx (loss
of two protons upon formation of the disulfide) was readily observed by TOF-MS analysis. When uniformly [15 N]-labeled oTrx
[molecular weight (MW) of 14,297.4)] was mixed in a 1∶1 ratio
with [14 N]-rTrx (MW of 14,126.6), both [15 N]-oTrx and [14 N]-rTrx
underwent 1-Da mass shifts to 14,298.3 and 14,125.6, respec-
Fig. 1. Direct detection of rTrx, oTrx, and Trx-SNO. (A) rTrx (TCEP treated) protein envelope. (B) oTrx (H2 O2 treated) showing evidence for dimeric protein in the
protein envelope (dimer exhibits double the number of peaks within a given interval due to higher charge states). (C) oTrx (insulin treated) protein envelope.
(D) Deconvoluted spectrum of rTrx. (E) Deconvoluted spectrum of oTrx (H2 O2 treated). (F) Deconvoluted spectrum of oTrx (insulin treated). (G) Deconvoluted
spectrum of [14 N]-rTrx treated 1∶1 with [15 N]-oTrx, showing the shift of both [14 N]-rTrx (Left) and [15 N]-oTrx (Right) to a 50% reduced state (MW of 14,125.6 and
14,298.3, respectively). Black lines represent 50% oxidized, dashed gray lines represent fully oxidized, and solid gray lines represent fully reduced. (H) Deconvoluted spectra for a representative time course of GSNO (10 μM) incubation with rTrx (1 μM) showing the singly nitrosated (+29) peak and lack of double
nitrosation (+58). The peak at 14,167 (+41) did not increase over time upon GSNO treatment; it is likely the Trx:acetonitrile adduct. (I) Deconvoluted spectrum of
polynitrosated Trx (treatment of 100 μM Trx with 1 mM GSNO for 15 min).
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Treatment of oTrx and rTrx with GSNO. Having determined that
Trx nitrosation could be directly detected without derivatization,
separate kinetic analyses on the rate of nitrosation for rTrx and
oTrx by GSNO were carried out. Both rTrx and oTrx (1 μM) were
readily nitrosated upon treatment with 10 μM GSNO. The rate
of nitrosation for rTrx was greater than that for oTrx, with pseudofirst-order rate constants of 3.53 ± 0.15 and 1.33 0.05 s−1 ,
respectively (Fig. 2A). No evidence of polynitrosation was observed. oTrx-SNO was 2 Da lower in mass from rTrx-SNO
(Fig. 2C), indicating that the initial redox state of Trx was reflected in the nitrosated species, without interconversion between
rTrx to oTrx (as would be expected if rTrx first oxidized to oTrx
and was then nitrosated). Rather, oTrx-SNO and rTrx-SNO are
two distinct species that form at different rates.
Over the course of the reaction, rTrx partially oxidized, forming low amounts of oTrx (<10% of total unmodified Trx, Fig. 2B).
Oxidation of the rTrx active site following reaction with GSNO
has been reported previously (3). On examination of the rTrx
active-site oxidation with GSNO, GSSG, and GSH, it appears
that GSSG, a contaminant in commercially available GSNO, is
the relevant oxidant in vitro (Fig. 3 and Figs. S3 and S4).
Nitrosation Specificity of rTrx Versus oTrx. Given the striking differences in nitrosation rate between rTrx and oTrx, the redox-dependent specificity of Cys nitrosation was determined. A library of
Trx Cys mutants was designed to remove possible sites of nitrosation. The mutants made included C62S, C69S, C73S, C69S/C73S,
Barglow et al.
PNAS PLUS
Fig. 2. Nitrosation of rTrx and oTrx. (A) Extent of nitrosation of rTrx and
oTrx reactions of 1 μM Trx with 10 μM GSNO as detected by TOF-MS. Data
shown are the average (with standard deviation) of the percent nitrosation
determined by the ratio of the +29 (singly nitrosated) peak to total Trx.
Data were collected as two independent runs with n ¼ 5 for each time point.
(B) Oxidation of rTrx at the active site (Cys32-Cys35) over the course of
the reaction with GSNO. Percent oxidation for each time point was calculated
as the mass shift from fully reduced (with a 2-Da shift for fully oxidized).
(C) Average masses (4-min time point, n ¼ 5 with SD) of rTrx, oTrx, rTrxSNO, and oTrx-SNO.
and C32S/C35S and a series of kinetic experiments with GSNO
were performed. Pseudo-first-order rate constants (Fig. 3C) were
calculated based on the percent of nitrosated Trx. As previously
described, mutant Trx (1 μM) was reacted with GSNO (10 μM)
for 1, 2, 3, and 4 min, desalted using the RapidFire DS Module™,
and analyzed by TOF-MS. For all analyzed proteins, only a single
NO addition was observed under these conditions.
The rTrx mutants C73S (3.26∕s) and C69S (3.59∕s), both nitrosated at rates similar to wild type (3.53∕s), with the double mutant
C73S/C69S (2.92∕s) slightly slower (Fig. 3A, black bars; Fig. 3C
contains rate constants with error for each). Strikingly, rTrx-C62S
(0.56∕s) nitrosated significantly more slowly (>6-fold), suggesting
that the primary site of nitrosation for rTrx is Cys62. The oTrx
mutant C73S demonstrated the slowest nitrosation rate (0.73∕s),
with C69S (1.13∕s) also nitrosating slightly slower than wild type
(1.33∕s) (Fig. 3A, gray bars). A C69S/C73S double mutant
showed no detectable nitrosation, and C62S nitrosated slightly
faster than wild type (1.92∕s). Based on these observations, Cys73
is the primary site of nitrosation for oTrx, with a secondary site
at Cys69. Cys62, the presumed site of nitrosation for rTrx, does
not appear to be nitrosated on oTrx. The specificity of oTrx
nitrosation is markedly different from rTrx. The active-site mutant C32S/C35S was used to ensure that the use of insulin and
TCEP did not influence the rate of nitrosation; indeed, there
was no significant difference between rates of the insulin treated
(1.53∕s) and TCEP treated (1.64∕s) C32S/C35S mutant (Fig. 3B)
Peptide-Level Confirmation of Cys Selectivity for Nitrosation of
oTrx and rTrx. To confirm the nitrosation selectivity suggested
by the TOF-MS analysis of intact Trx and Trx mutants, a modified
version of the biotin-switch assay, dubbed the reductive switch,
was employed to identify the individual sites of nitrosation
(Fig. S5). Briefly, the reaction was initiated by incubating Trx
(1 μM) with GSNO (10 μM), and quenched over time (3, 4,
5 min) with deuterium5 -labeled N-ethyl maleimide (d5 -NEM,
heavy label). Nonheavy labeled Cys were subsequently reduced
with DTTand counterlabeled with nonisotopic NEM (light label).
Samples were then trypsinized and analyzed by liquid chromatography (LC)-TOF-MS to determine ratios of heavy to light labeling for each Cys-containing peptide. Using DTT, which both
quenched the d5 -NEM and reduced SNOs and other oxidized
Cys, in place of ascorbate, increased the reproducibility of the
assay by eliminating precipitation or buffer exchange steps that
can lower protein recovery. Because Trx was labeled in the native
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tively, after 1 min of reaction (Fig. 1G). This result demonstrates
that the average mass of the deconvoluted Trx peak represents
the relative percentage of rTrx and oTrx in a mixture. Furthermore, oTrx is a substrate for rTrx, and in solution these species
are in rapid equilibrium.
Having determined that oTrx and rTrx could be quantitatively
differentiated, the transnitrosation of Trx by GSNO and the redox
state of Trx over the course of reaction were simultaneously
measured. The RapidFire DS Module™ (Agilent Technologies),
a dual-valve, desalting system (Fig. S1) was used for rapid removal of buffer salts, GSH, GSH disulfide (GSSG), and GSNO
from the reaction mixtures, followed by direct injection for
analysis by TOF-MS. The rapid and in-line desalting allowed the
monitoring of the reaction at the anticipated physiological concentrations of both Trx (1 μM) and GSNO (10 μM) at short time
points (1–4 min). The resolution and sensitivity of the TOF-MS
system were sufficient to detect nitrosated proteins above 2%
of total Trx. During the reaction time course, the deconvoluted
base peak for rTrx (14,126.6) decreased in intensity, while a +29
peak emerged (MW of 14,155.4, indicating the gain of NO and
loss of hydrogen) (Fig. 1H). The peak at 14,167 (+41) did not
increase over time upon GSNO treatment; it is likely the acetonitrile adduct of Trx. Under these conditions, there was no
evidence for glutathionylation [+306, which has been detected by
others using aged GSNO stocks that contain decomposition products (37)] or for doubly (+58) or triply (+87) nitrosated Trx,
which is consistent with the specificity implied by selective GSNO
binding and transfer.
Reactions carried out at higher concentrations of GSNO
(1 mM) and rTrx (100 μM) for longer periods of time (15 min)
produced increased amounts of polynitrosation (Fig. 1I) and led
to complete disulfide formation at the active site (rTrx to oTrx).
The dominant Trx species was doubly nitrosated, and significant
proportions of protein with three, four, and five NO additions
were also observed. Because Trx contains only five Cys, even the
active-site Cys (thought to be involved in denitrosation) became
nitrosated under these conditions, possibly due to the simultaneous nitrosation of both Cys32 and Cys35 before the disulfide
could form. At shorter times (4 min, Fig. S2), significant amounts
of single, double, and triple nitrosation were observed.
were comparable with the direct analysis of the intact, nitrosated
protein.
Using the reductive-switch method, independent measurements of the rates of nitrosation on the Cys73- and on Cys62/
Cys69-containing peptides (these two Cys were on a single tryptic
peptide) (Fig. 4B) could be made. For the wild-type protein, rTrx
showed the highest rate of nitrosation on the Cys62/Cys69-containing peptide (2.77∕s, Fig. 4A, white bars), whereas oTrx had
the fastest rate on the Cys73-containing peptide (1.47∕s, Fig. 4A,
gray bars), consistent with the intact protein results. The Cys62/
Cys69-containing peptide showed little nitrosation in any of the
oxidized proteins tested (wild type, C62S, or C73S all had rates
<0.9∕s), whereas the reduced proteins (wild type, C73S) nitrosated readily on the Cys62/Cys69-containing peptide (2.77∕s
and 3.58∕s respectively, Fig. 4, white bars). Considerable attenuation of the nitrosation rate on the Cys62/Cys69-containing peptide of rTrx-C62S mutant (0.73∕s) supports the finding that
Cys62, and not Cys69, is the target of nitrosation on rTrx.
Unexpectedly, the rate of nitrosation on the Cys73-containing
peptide of rTrx (2.23∕s) was faster overall than the nitrosation of
the Cys73-containing peptide of oTrx (1.47∕s) (Fig. 4B). For the
C62S mutant, this selectivity was reversed, and nitrosation on the
Cys73-containing peptide was faster for oTrx-C62S (2.8∕s) than
Fig. 3. Nitrosation rates of Cys mutants of Trx. (A) Plot of calculated rates of
nitrosation for rTrx (black) and oTrx (gray). Rates were calculated assuming
pseudo-first-order kinetics for Trx with an excess of GSNO, as the negative of
the linear slopes of the plot of ln (Trx consumption) versus time. Trx consumption was calculated as the average (with standard deviation) of 100-% nitrosation (determined by the ratio of the +29 (singly nitrosated) peak to total
Trx). Rates were calculated from a single experiment with n ¼ 5 at each data
point (C69S, C73S/C69S, C32S, C32S/C35S), or as the average from injections
performed over multiple days (wild-type, C62S, C73S). (B) Plot of calculated
nitrosation rates for the active-site mutant C32S/C35S, which served as a
control for the reduction/oxidation conditions used in A. (C) Table of the
rates shown in A and B.
(nondenatured) state, local protein structure improved SNO
stability (as it is thought to in vivo) during the initial alkylation
(heavy label) step; even in the folded state, nearly complete d5 NEM labeling was seen in control experiments with rTrx (>90%).
A potential limitation of the analysis is that DTT may reduce
other oxidation species (e.g., disulfides), potentially leading to
false positives. However, control experiments substituting GSH
or GSSG for GSNO showed no time-dependent increase in oxidation on the peptides containing the non-active-site Cys (Cys62,
Cys69, or Cys73), though the active-site Cys-containing peptide
oxidized upon treatment with GSSG. GSNO treatment showed
a clear, time-dependent increase of the NEM to d5 -NEM ratio
on the peptides containing Cys62, Cys69, and Cys73 (Fig. S6),
indicating the analysis accurately tracks the nitrosation of Trx.
Furthermore, the nitrosation rates calculated with this method
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Fig. 4. Reductive-switch assay for nitrosation. (A) Rates of nitrosation for
rTrx and oTrx on the peptides containing Cys73 (gray) and Cys62/Cys69
(white). C62S+C73S (mix) is a 1∶1 mixture of the two mutant proteins (final
concentration 1 μM). All data points (0.1, 3, 4, and 5 min) were collected in
triplicate from independently prepared samples. All samples were subjected
to the reductive-switch protocol, and nitrosation at each data point was
determined as the ratio of heavy labeled to light-labeled peptide. Rates
were calculated assuming pseudo-first-order kinetics, as the negative of the
linear slopes of the plot of ln (Trx consumption) versus time. (B) Table of the
rates shown in A.
Barglow et al.
Discussion
In the studies reported here we have undertaken a kinetic analysis
of the redox dependence on Cys selectivity of Trx following
exposure to GSNO. To this end, two previously undescribed
methods for the analysis of Trx nitrosation were developed. The
first, using a dual-valve rapid desalting device, followed by TOFMS, allowed for the direct detection of Trx nitrosation and redox
state at short time intervals. By analyzing the intact protein without derivatization, SNO loss and intra- or intermolecular scrambling of the nitrosation site was minimized. This technique
allowed for analysis at time points as short as 1 min, allowed many
replicates in a single experiment, was highly sensitive (approximately 1 pmol protein could be detected), and was highly repro-
Fig. 5. Model of nitrosation selectivity for oTrx and rTrx. (A) rTrx structure
(based on PDB ID code 1ERT) (39) showing the location of the 5 Cys. (B) Proposed model for interconversion and nitrosation of rTrx and oTrx.
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ducible. The second methodology, the reductive switch, was
complementary to the online-desalting/TOF-MS intact protein
experiments. Analysis of peptides following the reductive-switch
procedure allowed the assignment of nitrosation sites to single
peptides via TOF-MS. Taken together, these two techniques
revealed the kinetics and selectivity of Trx nitrosation by GSNO.
Of the five cysteine residues in human Trx1 (Fig. 5A), only
two were selectively nitrosated by GSNO, Cys62 and Cys73. rTrx
nitrosated rapidly and primarily at Cys62, whereas oTrx nitrosated more slowly overall with selectivity toward Cys73 (Fig. 5B).
Cys69, Cys32, and Cys35 did not appear to be selective targets for
nitrosation; nitrosation occurred only at high concentrations of
GSNO (1 mM) . At biologically relevant concentrations, Trx was
nitrosated at only a single Cys (Cys62 or Cys73). This striking
specificity preference in Cys reactivity between oTrx and rTrx is
what would be expected for a modification involved in signal
transduction. The differences in rates of nitrosation (with rTrx
nitrosation >2-fold faster than oTrx) imply that, in the case of a
mixture of rTrx and oTrx, rTrx nitrosation would predominate
under kinetically controlled conditions.
The concentrations of protein and GSNO had dramatic effects
not only on the rates but also on the selectivity of Trx nitrosation.
At the high concentrations of GSNO often used for in vitro
experiments (100 μM–1 mM), nucleophile–electrophile reactivity
drives the reaction leading to full nitrosation of all cysteines in
the protein. This intrinsic reactivity, and the high concentrations
needed for clear signal when using many standard nitrosation
assays, provides a basis for the conflicting data in the literature
regarding Trx nitrosation and highlights the importance of physiological conditions and concentrations for studying the selectivity of transnitrosation reactions. In sharp contrast, the results
when the reaction is carried out at concentrations comparable to
measured physiological concentrations of Trx [2–12 μM (38) and
GSNO (1–10 μM (16, 17, 39)] showed clear selectivity for specific
Cys and was redox-dependent.
We hypothesize that the selectivity of nitrosation is controlled
by the conformation of the protein, either by conformational
changes affecting GSNO binding (binding near Cys73 in oTrx and
Cys62 in rTrx) or by different protein microenvironments in rTrx
and oTrx that either activate the nucleophilicity of the respective
Cys (Cys62 and Cys73) or that stabilize the corresponding SNOs.
Although rTrx and oTrx are structurally similar (40) according to
crystallographic measurements, the local dynamics of the protein
may well be different in the two redox states, because the activesite disulfide bond of oTrx is predicted to reduce movement of
the loops surrounding the active site. Cys73, the Cys nearest the
active site (Fig. 5A), would be predicted to be particularly susceptible to conformational changes around the active site.
The peptide-level analysis using the reductive-switch assay
supports the nitrosation of rTrx on Cys62 and of oTrx on Cys73;
however, these data revealed an additional mechanistic insight:
Cys62 is required for rTrx nitrosation on Cys73, but not for oTrx
nitrosation on Cys73. oTrx appears to react with GSNO directly
at Cys73, whereas rTrx appears to react with GSNO directly at
Cys62, making rTrx-Cys62-SNO the direct kinetic product of the
reaction of rTrx with GSNO. However, nitrosation was observed
on Cys73 of wild-type rTrx, but not C62S-rTrx, and the reaction of
a mixture of C62S-rTrx and C73S-rTrx showed wild-type nitrosation rates on Cys73, implying that intermolecular transnitrosation
is possible. It appears that rTrx-Cys62-SNO may subsequently
react with another molecule of rTrx to form rTrx-Cys73-SNO
(Fig. 5B, Lower Left), and thus rTrx itself can be considered a
target for transnitrosation reactions with rTrx-Cys62S. The in
vivo significance of this in unknown, as it would be dependent
on transient Trx-Trx interactions in areas of high localized Trx
concentration. It is possible that for rTrx, Cys62-SNO dominates
at shorter time points (at 5 min or less) and is the kinetically
favored product, whereas Cys73-SNO is the more stable, thermo-
BIOCHEMISTRY
for rTrx-C62S (1.29∕s). Furthermore, the rTrx-C73S mutant protein nitrosated slightly faster than wild-type rTrx on the Cys62/
Cys69-containing peptide. Taken together with the intact protein
results, these data indicate that Cys62 is key to the nitrosation
of rTrx, but that downstream reactions of rTrx-Cys62-SNO can
occur, possibly via inter- or intra-Trx transnitrosation from Cys62
to Cys73, explaining the nitrosation observed on Cys73 of wildtype rTrx. If intermolecular transnitrosation was occurring, we
would expect that Cys62 on one molecule of Trx would compensate for the mutation in rTrx-C62S, allowing faster nitrosation
on Cys73; whereas if intramolecular transnitrosation from Cys62
to Cys73 was occurring, the addition of Cys62 on another Trx
molecule would not change the rate of nitrosation on Cys73. To
determine which of these possibilities were most likely, a 1∶1
mixture of the C62S and C73S Trx proteins was assayed. Both
the reduced and oxidized forms of this mixture (rC62S + rC73S
and oC62 + oC73S, respectively) had rates identical to wild type
on both the Cys73-containing peptide and the Cys62/Cys69containing peptides (Fig. 4), consistent with intermolecular transnitrosation, where rTrx-Cys62-SNO transfers NO to Cys73 on
another rTrx molecule (Fig. 5, Lower Left). The in vivo relevance
of this SNO-Trx mediated transnitrosation of Trx remains unclear.
dynamically favored product. rTrx-Cys62-SNO could then be considered a signaling intermediate for transnitrosation reactions,
whereas Trx-C73-SNO could serve a dual role as both a transnitrosation agent (for example, with Casp3) and a thermodynamic
sink for “NO storage.”
The potential stability of Trx-C73-SNO is intriguing in light of
data indicating that GSNO is unstable in the presence of GSH
(41). Attempted nitrosation of Trx by a mixture of GSNO and
GSH showed no nitrosation, likely due to the reaction of GSNO
with 1 mM GSH. In contrast, Trx-SNO is fairly stable to GSH
at concentrations of 1 mM (60% SNO-rTrx remained, Fig. S7).
Thus, if the Trx-GSNO reaction were to take place in areas of
high local concentrations of GSNO, the stable Trx-SNO species
could be available for downstream transnitrosation reactions
in the presence of cytoplasmic GSH. Trx-SNO is thus an ideal
candidate as an intermediate for redox and nitrosation-dependent signal transduction.
In vivo, cellular redox conditions control the ratio of rTrx
to oTrx. Glutathione reductase (GR) (42, 43) likely minimizes
concentrations of GSSG, at least in healthy (nonoxidatively
stressed) cells, and TrxR (26) also drives oTrx toward rTrx. High
intracellular GSH concentrations (1–5 mM) would suggest that
rTrx (and rTrx-SNO) likely predominate in vivo under normal
conditions. Because both GR and TrxR are NADPH-dependent,
it is reasonable to assume that oTrx and oTrx-SNO may accumulate under conditions of oxidative stress or during depletion of
NADPH. The differences in Cys selectivity for nitrosation between rTrx and oTrx may represent a redox-dependent “switch”
for signal transduction via cascading transnitrosation reactions.
Because Cys73 and Cys62 are on opposite faces of the protein
(Fig. 5A), it is possible that oTrx-SNO and rTrx-SNO have differ-
Fig. 6. Hypothetical model for redox-dependent signal transduction via Trx
nitrosation. Under cellular reducing conditions, rTrx (Left) nitrosates on Cys62
and transfers that nitroso group to protein partners (blue). Under conditions
of oxidative stress, oTrx (Right) nitrosates on Cys73 and transfers that nitroso
group to different protein partners (red), such as Casp3.
6 of 7 ∣
www.pnas.org/cgi/doi/10.1073/pnas.1110736108
ent binding partners and thus different downstream transnitrosation targets (Fig. 6). This would provide an alternate mechanism
of signal transduction during periods of nitrosative or oxidative
stress.
Materials and Methods
Plasmids, Protein, and Reagents. All reagents and chemicals were purchased
from Sigma unless otherwise noted. Human Trx1 was expressed as an N-terminal his10 -tagged construct from the expression vector pET16b, in Escherichia
coli BL21(DE3) as described previously (4). The [15 N]-labeled Trx was expressed
in BL21(DE3) cells grown in minimal media (M9) supplemented with 14 g∕L
[15 N]-ammonium sulfate (Cambridge Isotopes) and purified as previously
described. Cys mutants were generated by QuikChange (Stratagene) mutagenesis using the manufacturer’s protocol. Trx Cys variants were confirmed
using TOF-MS of the intact proteins and the tryptic-digested peptides. Protein purity was assayed by SDS-PAGE, and concentration was determined by
the Bradford assay using BSA as the standard. To generate rTrx, 50 μM Trx was
treated with 1 mM TCEP or DTT for 5 min at room temperature, then desalted
over a Bio-Spin 6 column (Bio-rad). To generate oTrx, 50 μM Trx was treated
with TCEP or DTT and desalted as above, followed by treatment with 60 μM
(1.2 equivalents) bovine insulin, and desalted over a Bio-Spin 6 column.
Removal of insulin was confirmed by TOF-MS.
Direct Analysis of Protein Nitrosation. rTrx and oTrx were prepared as described above, then diluted to 2 μM in 50 mM potassium phosphate, pH 7.4
(PB). A fresh solution of GSNO (Cayman Chemicals) was prepared in PB and
the concentration determined spectrophotometrically (ϵ ¼ 0.92 at 335 nm).
GSNO stocks were diluted to 20 μM and stored protected from light for no
longer than 6 h. All reactions were carried out at room temperature. Trx and
GSNO were loaded into separate 1-mL syringes and connected to a dual
syringe pump attached to a 3-μL mixing tee. After mixing, Trx and GSNO
were allowed to react in the tubing for a set time (based on flow rate and
dead volume) before application to the RapidFire™ DS Module (Agilent Technologies). The system was equilibrated with water (containing 0.1% formic
acid, mobile phase A) in the first valve state. During the second valve state,
the sample loop (10 μL) was filled with the reaction mixture. In the third valve
state, the sample was applied to the trapping cartridge (C18) and washed
with mobile phase A. In the fourth valve state, the sample was eluted from
the trapping cartridge with mobile base B (acetonitrile containing 0.1%
formic acid) and injected directly onto an Agilent TOF mass spectrometer
(MSD TOF, Model 1969A). Following injection the device returned to valve
state 1. The cycle (equilibration, loop filling, washing/desalting, and elution)
was repeated 5 times for each time point. Syringe pump flow rates were
changed for each time point to increase reaction time. The following instrument parameters were used for the TOF-MS: gas temperature, 350 °C; gas
flow, 10 L∕ min; nebulizer, 30 psi; capillary voltage, 3,500 V; fragmentor,
90 V. The total ion chromatogram consisted of periodic elutions for each
replicate. The extracted protein envelopes were manually examined to
ensure no dimer or insulin remained. Deconvolution of the protein envelopes
was performed using MassHunter Software (Agilent Technologies) in the
13,000- to 16,000-MW range, using the extraction ion m∕z range of 600–
1,600. The resulting deconvoluted spectra intensities were used to calculate
the relative ratio of Trx to SNO-Trx for quantitative analysis of reaction rates.
For denitrosation experiments, 1 μM oTrx was treated with 10 μM GSNO
or 1 mm GSH for 2 min, after which 10 μM GSNO or 1 mm GSH was added
via a second syringe pump and mixing tee, and the mixture incubated for
1 min before rapid desalting and TOF-MS analysis as above.
Reductive-Switch Assay. rTrx and oTrx were prepared as described above,
then diluted to 1 μM in 50 mM potassium phosphate, pH 7.4, with
200 mM NaCl (PBS). Fresh GSNO stocks were prepared as above in PBS. All
reactions were performed in triplicate. For each reaction, the final volume
was 500 μL containing 1 μM Trx and 10 μM GSNO. Reactions were quenched
(Cys alkylated) by the addition of 1 mM (final concentration) d 5 -labeled
N-methyl-maleimide (d 5 -NEM, CDN Isotopes) at 37 °C for 1 h. For the 0.1-min
reactions, GSNO was added, tubes vortexed, then d 5 -NEM was immediately
added. Samples were then treated with 2 mM DTT for 10 min at room temperature to reduce nitrosothiols. NEM (5 mM) was added as a counterlabel
and samples were reacted for 1 h at 37 °C. Samples were concentrated over
5-kDa cutoff spin concentrators (Vivaspin) to 50 μL, followed by addition
of trypsin (4 μg∕mL) (Trypsin Gold, Promega). Samples were digested for
1 h at 37 °C, then analyzed via LC-MS using a Zorbax C18 (2.1 × 50 mm)
column and a Waters LCT Premier XE TOF-MS. The heavy- and light-labeled
peptides containing Cys32/Cys35, Cys62/Cys69, and Cys73 were quantified
using MassLynx software (Waters).
Barglow et al.
1. Hess DT, Matsumoto A, Kim SO, Marshall HE, Stamler JS (2005) Protein S-nitrosylation:
Purview and parameters. Nat Rev Mol Cell Biol 6:150–166.
2. Benhar M, Forrester MT, Hess DT, Stamler JS (2008) Regulated protein denitrosylation
by cytosolic and mitochondrial thioredoxins. Science 320:1050–1054.
3. Hashemy SI, Holmgren A (2008) Regulation of the catalytic activity and structure of
human thioredoxin 1 via oxidation and S-nitrosylation of cysteine residues. J Biol Chem
283:21890–21898.
4. Mitchell DA, Marletta MA (2005) Thioredoxin catalyzes the S-nitrosation of the
caspase-3 active site cysteine. Nat Chem Biol 1:154–158.
5. Mitchell DA, Morton SU, Fernhoff NB, Marletta MA (2007) Thioredoxin is required for
S-nitrosation of procaspase-3 and the inhibition of apoptosis in Jurkat cells. Proc Natl
Acad Sci USA 104:11609–11614.
6. Sengupta R, et al. (2007) Thioredoxin catalyzes the denitrosation of low-molecular
mass and protein S-nitrosothiols. Biochemistry 46:8472–8483.
7. Wu C, et al. (2010) Redox regulatory mechanism of transnitrosylation by thioredoxin.
Mol Cell Proteomics 9:2262–2275.
8. Stoyanovsky DA, et al. (2005) Thioredoxin and lipoic acid catalyze the denitrosation of
low molecular weight and protein S-nitrosothiols. J Am Chem Soc 127:15815–15823.
9. Powis G, Montfort WR (2001) Properties and biological activities of thioredoxins. Annu
Rev Biophys Biomol Struct 30:421–455.
10. Madej E, Folkes LK, Wardman P, Czapski G, Goldstein S (2008) Thiyl radicals react with
nitric oxide to form S-nitrosothiols with rate constants near the diffusion-controlled
limit. Free Radic Biol Med 44:2013–2018.
11. Keszler A, Zhang Y, Hogg N (2010) Reaction between nitric oxide, glutathione, and
oxygen in the presence and absence of protein: How are S-nitrosothiols formed? Free
Radic Biol Med 48:55–64.
12. Keshive M, Singh S, Wishnok JS, Tannenbaum SR, Deen WM (1996) Kinetics of S-nitrosation of thiols in nitric oxide solutions. Chem Res Toxicol 9:988–993.
13. Dickinson DA, Forman HJ (2002) Cellular glutathione and thiols metabolism. Biochem
Pharmacol 64:1019–1026.
14. Filomeni G, Rotilio G, Ciriolo MR (2002) Cell signalling and the glutathione redox
system. Biochem Pharmacol 64:1057–1064.
15. Gaston B, et al. (1993) Endogenous nitrogen oxides and bronchodilator S-nitrosothiols
in human airways. Proc Natl Acad Sci USA 90:10957–10961.
16. Kluge I, Gutteck-Amsler U, Zollinger M, Do KQ (1997) S-nitrosoglutathione in rat
cerebellum: Identification and quantification by liquid chromatography-mass spectrometry. J Neurochem 69:2599–2607.
17. Yap LP, et al. (2010) Determination of GSH, GSSG, and GSNO using HPLC with electrochemical detection. Methods Enzymol 473:137–147.
18. Xu L, Eu JP, Meissner G, Stamler JS (1998) Activation of the cardiac calcium release
channel (ryanodine receptor) by poly-S-nitrosylation. Science 279:234–237.
19. Yang Z, et al. (2010) Lymphocyte development requires S-nitrosoglutathione reductase. J Immunol 185:6664–6669.
20. Weichsel A, Brailey JL, Montfort WR (2007) Buried S-nitrosocysteine revealed in crystal
structures of human thioredoxin. Biochemistry 46:1219–1227.
21. Weichsel A, Kem M, Montfort WR (2010) Crystal structure of human thioredoxin revealing an unraveled helix and exposed S-nitrosation site. Protein Sci 19:1801–1806.
22. Haendeler J, et al. (2002) Redox regulatory and anti-apoptotic functions of thioredoxin depend on S-nitrosylation at cysteine 69. Nat Cell Biol 4:743–749.
23. Bao R, Zhang Y, Zhou CZ, Chen Y (2009) Structural and mechanistic analyses of
yeast mitochondrial thioredoxin Trx3 reveal putative function of its additional cysteine
residues. Biochim Biophys Acta 1794:716–721.
24. Wang Y, Liu T, Wu C, Li H (2008) A strategy for direct identification of protein
S-nitrosylation sites by quadrupole time-of-flight mass spectrometry. J Am Soc Mass
Spectrom 19:1353–1360.
25. Benhar M, Thompson JW, Moseley MA, Stamler JS (2010) Identification of S-nitrosylated targets of thioredoxin using a quantitative proteomic approach. Biochemistry
49:6963–6969.
26. Holmgren A, Lu J (2010) Thioredoxin and thioredoxin reductase: Current research
with special reference to human disease. Biochem Biophys Res Commun 396:120–124.
27. Jaffrey SR, Snyder SH (2001) The biotin switch method for the detection of S-nitrosylated proteins. Sci STKE 2001(86):pl1.
28. Forrester MT, Foster MW, Benhar M, Stamler JS (2009) Detection of protein S-nitrosylation with the biotin-switch technique. Free Radic Biol Med 46:119–126.
29. Forrester MT, Foster MW, Stamler JS (2007) Assessment and application of the
biotin switch technique for examining protein S-nitrosylation under conditions of
pharmacologically induced oxidative stress. J Biol Chem 282:13977–13983.
30. Forrester MT, et al. (2009) Proteomic analysis of S-nitrosylation and denitrosylation by
resin-assisted capture. Nat Biotechnol 27:557–559.
31. Sinha V, et al. (2010) Proteomic and mass spectroscopic quantitation of protein
S-nitrosation differentiates NO-donors. ACS Chem Biol 5:667–680.
32. Zhang Y, Keszler A, Broniowska KA, Hogg N (2005) Characterization and application
of the biotin-switch assay for the identification of S-nitrosated proteins. Free Radic
Biol Med 38:874–881.
33. Han P, Chen C (2008) Detergent-free biotin switch combined with liquid chromatography/tandem mass spectrometry in the analysis of S-nitrosylated proteins. Rapid
Commun Mass Spectrom 22:1137–1145.
34. Zhou X, et al. (2010) ESNOQ, proteomic quantification of endogenous S-nitrosation.
PLoS One 5:e10015.
35. Gladwin MT, Wang X, Hogg N (2006) Methodological vexation about thiol oxidation
versus S-nitrosation—A commentary on “An ascorbate-dependent artifact that
interferes with the interpretation of the biotin-switch assay”. Free Radic Biol Med
41:557–561.
36. Huang B, Chen C (2006) An ascorbate-dependent artifact that interferes with the
interpretation of the biotin switch assay. Free Radic Biol Med 41:562–567.
37. Tao L, English AM (2004) Protein S-glutathiolation triggered by decomposed S-nitrosoglutathione. Biochemistry 43:4028–4038.
38. Holmgren A, Luthman M (1978) Tissue distrubution and subcellular localization of
bovine thioredoxin determined by radioimmunoassay. Biochemistry 17:4071–4077.
39. Bramanti E, et al. Determination of S-nitrosoglutathione in plasma: Comparison of
two methods. Talanta 81:1295–1299.
40. Weichsel A, Gasdaska JR, Powis G, Montfort WR (1996) Crystal structures of reduced,
oxidized, and mutated human thioredoxins: Evidence for a regulatory homodimer.
Structure 4:735–751.
41. Singh SP, Wishnok JS, Keshive M, Deen WM, Tannenbaum SR (1996) The chemistry of
the S-nitrosoglutathione/glutathione system. Proc Natl Acad Sci USA 93:14428–14433.
42. De Vega L, et al. (2002) Glutathione determination and a study of the activity of
glutathione-peroxidase, glutathione-transferase, and glutathione-reductase in renal
transplants. Ren Fail 24:421–432.
43. De Vega L, et al. (2003) Study of the activity of glutathione-peroxidase, glutathionetransferase, and glutathione-reductase in renal transplants. Transplant Proc
35:1346–1350.
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PNAS Early Edition ∣
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PNAS PLUS
Institute of Technology (MIT)-Merck fellowship, and K.T.B. by an American
Cancer Society Postdoctoral Fellowship. C.G.K., S.R.T., and J.S.W. are
supported by the National Institutes of Health (NIH) (CA26731) and by the
MIT Center for Environmental Health Sciences (ES002109). K.T.B. and
M.A.M. are supported by the NIH (GM080272).
BIOCHEMISTRY
ACKNOWLEDGMENTS. We acknowledge Agilent Technologies for access to
the UHPLC 1290 binary pump, autosampler, degasser, and the RapidFire
DS Module™. Special thanks go to Can Ozbal, Michelle Romm, and Kari
Schlicht (Agilent Technologies) for technical assistance with the RapidFire
DS Module™. We thank Eric Underbakke and Brian Smith for helpful
assistance editing this manuscript. C.G.K. is supported by the Massachusetts