a “fluorescence switch” technique increases the sensitivity of proteomic detection and...

RESEARCH ARTICLE

A ‘‘fluorescence switch’’ technique increases

the sensitivity of proteomic detection and

identification of S-nitrosylated proteins

Daniel Tello1, Carlos Tarın2, Patricia Ahicart1, Rosa Breton-Romero3, Santiago Lamas3

and Antonio Martınez-Ruiz1

1 Servicio de Inmunologıa, Hospital de La Princesa, Madrid, Spain2 Centro Nacional de Investigaciones Cardiovasculares Carlos III (CNIC), Madrid, Spain3 Laboratorio Mixto CSIC-FRIAT de Fisiopatologıa Vascular y Renal, Centro de Investigaciones Biologicas (CSIC),

Madrid, Spain

Received: February 3, 2009

Revised: August 7, 2009

Accepted: September 1, 2009

Protein S-nitrosylation is a reversible post-translational modification of protein cysteines that is

increasingly being considered as a signal transduction mechanism. The ‘‘biotin switch’’ tech-

nique marked the beginning of the study of the S-nitrosoproteome, based on the specific

replacement of the labile S-nitrosylation by a more stable biotinylation that allowed further

detection and purification. However, its application for proteomic studies is limited by its rela-

tively low sensitivity. Thus, typical proteomic experiments require high quantities of protein

extracts, which precludes the use of this method in a number of biological settings. We have

developed a ‘‘fluorescence switch’’ technique that, when coupled to 2-DE proteomic methodol-

ogies, allows the detection and identification of S-nitrosylated proteins by using limited amounts

of starting material, thus significantly improving the sensitivity. We have applied this metho-

dology to detect proteins that become S-nitrosylated in endothelial cells when exposed to

S-nitroso-L-cysteine, a physiological S-nitrosothiol, identifying already known S-nitrosylation

targets, as well as proteins that are novel targets. This ‘‘fluorescence switch’’ approach also

allowed us to identify several proteins that are denitrosylated by thioredoxin in cytokine-activated

RAW264.7 (murine macrophage) cells. We believe that this method represents an improvement

in order to approach the identification of S-nitrosylated proteins in physiological conditions.

Keywords:

Cell biology / Endothelium / Macrophage activation / Post-translational modification /

Protein oxidation / S-nitrosylation

1 Introduction

Since the discovery of nitric oxide (NO) signaling more than 20

years ago, several pathways and mechanisms have arisen that

are expanding our understanding about how cells can use

different chemical mechanisms to respond to stimuli and

transduct these signals [1]. One of these mechanisms involves

the formation of post-translational modifications induced by

NO and related reactive nitrogen species [2]. Among them,

S-nitrosylation (also called S-nitrosation) has been implied in

NO signaling in several physiological and pathophysiological

contexts [3]. It can alter the functionality of different proteins,

and its particular characteristics (shared in part with other

oxidative post-translational modifications) have led to consider

that it might represent a new paradigm in signal transduction

[4, 5]. Its lability, which allows for its easy reversibility, also

represents a limitation for the direct detection of the modifi-

cation by proteomic methods; for example, MALDI ionization

has been shown to induce the breakage of the nitrosothiol S-N

bond, hampering its detection by one of the most widely used

techniques in proteomics [6]. The ‘‘biotin switch’’ technique

Abbreviations: Biotin-HPDP, N-[6-(Biotinamido)hexyl]-30-(20-

pyridyldithio)propionamide; CysSNO, S-nitroso-L-cysteine;

HUVEC, human umbilical vein endothelial cells; IFN-g, inter-

feron-g; iNOS, inducible nitric oxide synthase; LPS, bacterial

lipopolysaccharide; MMTS, methyl methanethiosulfonate; NEM,

N-ethylmaleimide; NO, nitric oxide; PDI, protein disulfide-

isomerase

Correspondence: Dr. Antonio Martınez-Ruiz, Servicio de Inmu-

nologıa, Hospital de La Princesa, C/Diego de Leon 62, E-28006

Madrid, Spain

E-mail: [email protected]

Fax: 134-915202374

& 2009 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com

Proteomics 2009, 9, 5359–5370 5359DOI 10.1002/pmic.200900070

overcomes some of these limitations by replacing the labile

nitrosothiol by a more stable biotinylation linked by a disulfide

bridge, by means of three successive chemical steps [7]. The

first step is the blocking of the protein free thiols, followed by

reduction of nitrosothiols by ascorbate to yield a new free thiol

that is subsequently labeled by the biotinylating reagent. In

order to maintain the specificity, both very effective blocking

and stringent conditions for reduction with ascorbate are

needed, and it has been described that only a fraction of the

initial S-nitrosylated protein is biotinylated [8]. Indeed, in many

physiological conditions, the amount of S-nitrosylated protein

is low, due to a tight control of the modification formation (by

regulated reactive nitrogen species production including its

subcellular localization) and its breakage (by denitrosylase

activities operating on both low molecular mass and protein

thiols) [9, 10]. Thus, it is assumed that there is a low proportion

of biotinylated proteins related to total protein, even in proteins

that could be preferentially S–nitrosylated. When the ‘‘biotin

switch’’ technique is used to assay the S-nitrosylation of

specific proteins, biotinylated protein capture (such as avidin

pull-down) can be coupled to antibody-based detection of the

specific proteins, which has allowed sensitive and specific

detection of S-nitrosylation in individual proteins in several

(patho)physiological contexts [10–13]. For proteomic studies,

biotinylated proteins purified through avidin capture are the

starting material for the detection and identification of the

proteins that were modified, either by electrophoretic (1-DE or

2-DE) protein separation or ‘‘second-generation proteomics’’

(LC-MS/MS) [14–16]. In this way, only the biotinylated fraction

of each protein is used for the identification. This may result in

an important limitation to study the S-nitrosoproteome in a

variety of (patho)physiological settings.

A different switch strategy, based on direct resin capture

(that replaces biotin labeling and avidin capture), has been

recently described for protein and peptide isolation, identi-

fication and even quantitation [17]. Although it seems to

increase the capture of high molecular mass proteins with

respect to the biotin switch, identification still relies on

purification of the modified fraction of the protein, so it still

bears the same limitation.

We herein report a ‘‘fluorescence switch’’ coupled to

2-DE that combines selective detection of fluorescent deri-

vatization of S-nitrosylated proteins with efficient identifi-

cation by digesting the total protein spot. This method

allows the use of only limited amounts of starting material,

thus making it more feasible for its application in a number

of physiological scenarios.

2 Materials and methods

2.1 Materials

Cell culture media and supplements were obtained from

Life Technologies, Invitrogen. SDS and other electrophor-

esis reagents were from Bio-Rad. Fluorescein-5-maleimide

was from Anaspec. PVDF ‘‘Immobilon P’’ membrane was

from Millipore. Streptavidin-peroxidase and 2-DE reagents

were from Amersham Biosciences. N-[6-(Biotinamido)-

hexyl]-30-(20-pyridyldithio)propionamide (biotin-HPDP) and

BCA reagent were from Pierce. Modified porcine trypsin,

sequencing grade, was from Promega. S-nitroso-L-cysteine

(CysSNO) was synthesized as previously described [18] and

quantified spectrophotometrically using an extinction coef-

ficient of 900 M�1 cm�1 at 338 nm [19], with yields of about

80%. Polyclonal antibody against inducible nitric oxide

synthase (iNOS) was from Biomol. Other reagents were

obtained from Sigma-Aldrich.

2.2 Cell culture and treatment

Cells were grown at 371C, 5% CO2. The EA.hy926 cell line

(kindly provided by Dr. Cora-Jean S. Edgell, UNC, Chapel

Hill, NC, USA) was cultured in DMEM with HAT supple-

ment, 20% FBS, 100 U/mL penicillin, 100mg/mL strepto-

mycin and 5mg/mL gentamicin. Human umbilical vein

endothelial cells (HUVEC) were isolated from umbilical

cord veins as previously described [20] and cultured in

medium 199 with 20% FBS, 1% ECGF, 100 U/mL penicillin

and 100 mg/mL streptomycin; umbilical cords were obtained

from Ruber International Hospital (Madrid, Spain) with the

approval of the donors and the ethics committee of the

institution. Unless otherwise stated, for cell treatments, cells

were washed with PBS and incubated in the dark in RPMI

without serum or phenol red at 371C for 15 min, and 1 mM

CysSNO was added or not added. The RAW264.7 cell line

was cultured in DMEM with 1 M HEPES supplement,

10% FBS, 100 U/mL penicillin, 100 mg/mL streptomycin.

RAW264.7 cells were incubated in RPMI medium without

phenol red, with or without 10 ng/mL interferon-g (IFN-g)

and 5 mg/mL bacterial lipopolysaccharide (LPS) for 18 h, and

treated or not with 1mM auranofin for 1 h at 371C. Nitrite

concentration in the supernatants was quantified using the

Griess assay, as previously described [21].

2.3 Fluorescence switch

All operations were carried out in the dark. Treated cells

were scraped and non-denaturing lysis solution (50 mM

Tris-HCl, pH 7.4, 300 mM NaCl, 5 mM EDTA. 0.1 mM

neocuproine, 1% Triton X-100 and 30 mM N-ethylmalei-

mide (NEM) plus protease inhibitors cocktail) was added,

incubated in ice for 15 min, and centrifuged at 10 000� g,

41C for 15 min. Supernatant was collected and protein was

quantified with BCA reagent (Pierce). Extracts were adjusted

to 0.5 mg/mL of protein and equal amounts were blocked

with 4 volumes of blocking buffer (225 mM HEPES, pH 7.2,

0.9 mM EDTA, 90 mM neocuproine, 2.5% SDS and 30 mM

NEM) at 371C for 30 min. After blocking, extracts were

precipitated with acetone and resuspended in HENS buffer

5360 D. Tello et al. Proteomics 2009, 9, 5359–5370


(250 mM HEPES, pH 7.2, 1 mM EDTA, 0.1 mM

neocuproine, 1% SDS) to which 100 mM ascorbate was

added (50 mL of buffer per 100mg of protein). Fluorescein-5-

maleimide was added in HENS buffer (40 mM, final

concentration), and incubated for 1 h at room temperature.

At this point NEM in HENS buffer was added to a final

4 mM concentration. Finally, proteins were acetone-preci-

pitated before separation by SDS- or 2-D-PAGE. As negative

controls, extracts were treated with 100 mM DTT for 10 min

at room temperature before blocking, or ascorbate was

omitted from the reduction step.

For some of the 2-DE gels, changes were made to the

protocol in order to reduce the saline content in the samples

and reactions with both conditions were compared before

running 2-DE gels: 25 M Tris replaced 250 mM HEPES,

0.25% SDS was used in the reduction step, after which

chloroform/methanol precipitation was performed as

previously described [22].

2.4 Standard and modified biotin switch

The standard biotin switch assay was performed essentially

as previously described [7, 14], with the improvements in the

reduction step that have been recently published [23]. The

process is essentially the same as the fluorescence switch

assay, except for the following differences. Protein extracts

were performed without NEM. Blocking was in pH 7.7

buffer with 20 mM methyl methanethiosulfonate (MMTS)

instead of NEM, for 20 min at 501C. Acetone-precipitated

samples were resuspended in pH 7.7 HENS buffer to which

100 mM ascorbate was added, and biotin-HPDP was added

to a final 1 mM concentration (from 4 mM stock in dime-

thylformamide). When NEM blocking or biotin-maleimide

labeling was used, these steps were performed in the same

conditions as in the fluorescence switch.

To detect biotinylated proteins by Western blot, samples

from the biotin switch assay were separated on 10% SDS-

PAGE gels, transferred to PVDF membranes, blocked with

non-fat dried milk, incubated with streptavidin-peroxidase,

and light was registered after addition of chemilumines-

cence reagents.

2.5 Protein electrophoresis and fluorescence

detection

For SDS-PAGE, samples from the fluorescence switch assay

were resuspended in SDS loading buffer and separated on

10% gels using standard protocols. Fluorescein detection

was performed using either a Kodak Image Station

4000MMPro with excitation/emission filters centred at 470/

535 nm, respectively, or a Typhoon scanner as stated below.

For 2-DE, samples from the fluorescence switch assay

were resuspended in lysis buffer (7 M urea, 2 M thiourea,

4% w/v CHAPS and 30 mM Tris-HCl, pH 8.5), diluted in

rehydration buffer (7 M urea, 2 M thiourea, 2% w/v CHAPS,

0.8% v/v Pharmalytes 3–10) to obtain a final volume of

350 mL and were applied in dry IPG strips 18 cm, 4–7 L for

reswelling. The first dimension was run at 75 mA per IPG

strip in the IPGphor IEF II System (GE Healthcare)

following a step-wise voltage increase: 30 V/h for 7 h, 60 V/h

for 7 h, 120 V/h for 1 h, 250 V/h for 1 h, 500 V/h for 1 h,

1000 V/h for 0.5 h, gradient was increased from 1000 to

8000 V/h for 0.5 h, and constant 8000 V until a total 68 000 V

was reached. After the first dimension, strips were equili-

brated with SDS equilibration buffer (6 M urea, 2% w/v

SDS, 30% v/v glycerol and 75 mM Tris-HCl, pH 8.8) and

separated on 10% Tris-glycine gels (0.05% SDS) using an

SE600 device (GE Healthcare) in the dark at 251C until the

tracking dye had migrated off the bottom of the gel.

After 2-DE, gels were scanned with a Typhoon 9400

scanner (GE Healthcare) at 100mm resolution using fluor-

escein conditions (blue laser at wavelength 488 nm for

excitation and 526BP emission filter). After scanning, gels

were detached from the glasses, stained with Lucy-565

fluorescent dye (Sigma-Aldrich) following the manu-

facturer’s instructions, and scanned with a Typhoon 9400

scanner for the fluorescein signal as before, and for Lucy

565 (green laser at wavelength 532 nm for excitation and

580BP30 emission filter). Finally, 2-DE gels were either

silver stained using a protocol compatible with MS [24], or

stained with Sypro Ruby Protein Gel Stain (Sigma) follow-

ing the manufacturer’s instructions and scanned with a

Typhoon 9400 scanner (green laser at wavelength 532 nm

for excitation and 610BP30 emission filter).

2.6 Protein digestion and identification

Protein spots from 2-DE gels were excised from the gels

manually and automatic digestion and MS analysis using an

Ultraflex MALDI-TOF/TOF mass spectrometer (Bruker

Daltonics) were performed as described [25].

Alternatively, protein spots from 2-DE gels were excised

automatically using an Ettan Spot Picker Workstation (GE),

deposited in 96-well plates and processed automatically in a

Proteineer DP (Bruker Daltonics). The digestion protocol

used was based on Shevchenko et al. [26] with minor

variations: gel plugs were washed first with 50 mM ammo-

nium bicarbonate and then with ACN prior to reduction

with 10 mM DTT in 25 mM ammonium bicarbonate solu-

tion, and alkylation was carried out with 55 mM IAA in

50 mM ammonium bicarbonate solution. Gel pieces were

then rinsed first with 50 mM ammonium bicarbonate and

then with ACN, and then were dried under a stream of

nitrogen. Modified porcine trypsin (sequencing grade;

Promega) at a final concentration of 16 ng/mL in 25% ACN/

50 mM ammonium bicarbonate solution was added and the

digestion took place at 371C for 6 h. The reaction was stop-

ped by adding 0.5% TFA for peptide extraction. The tryptic

eluted peptides were dried by speed-vacuum centrifugation

Proteomics 2009, 9, 5359–5370 5361


and were resuspended in 4 mL of MALDI solution (33%

2-propanol, 16.7% ACN and 0.5% TFA). A 0.8-mL aliquot of

each peptide mixture was deposited onto a 389-well Opti-

TOFTM Plate (Applied Biosystems, Framingham, MA, USA)

and allowed to dry at room temperature. A 0.8-mL aliquot of

matrix solution (3 mg/mL CHCA in MALDI solution) was

then deposited onto dried digest and allowed to dry at room

temperature. For MALDI-TOF/TOF analysis, samples were

automatically acquired in an ABi 4800 MALDI TOF/TOF

mass spectrometer (Applied Biosystems) in positive ion

reflector mode (the ion acceleration voltage was 25 kV to MS

acquisition and 1 kV to MSMS) and the obtained spectra

were stored into the ABi 4000 Series Explorer Spot Set

Manager. PMF and MSMS fragment ion spectra were

smoothed and corrected to zero baseline using routines

embedded in ABi 4000 Series Explorer Software v3.6. Each

PMF spectrum was internally calibrated with the mass

signals of two trypsin autolysis ions: [VATVSLPR1H]1

(m/z 5 842.510) and [LGEHNIDVLEGNEQFINAAK1H]1

(m/z 5 2211.105) to reach a typical mass measurement

accuracy of o25 ppm. Known trypsin and keratin mass

signals, as well as potential sodium and potassium adducts

(121 Da and 139 Da) were removed from the peak list. To

submit the combined PMF and MS/MS data to MASCOT

software v.2.1 (Matrix Science, London, UK), ABi GPS

Explorer v4.9 was used, searching in the Uniprot_SwissProt

protein database (v14.2_20080923; 39 181 sequences; 143

572 911 residues). Searches were submitted with a mass

tolerance of 100 ppm in MS mode and 0.8 Da in MS/MS

mode. One missed cleavage per peptide was allowed,

carbamidomethyl cysteine as fixed modification and oxida-

tion of methionine as variable modification were taken into

account. Searches were performed with Homo sapiens taxo-

nomic restriction and without constraining protein mole-

cular mass or pI. To achieve high confidence in the different

protein/peptide identifications, only high quality MS and

MS/MS spectra were accepted (MOWSE scores better than

65 and 35 for each spectrum, respectively), always with a

p-value greater than 0.05.

3 Results and discussion

3.1 Development of the fluorescence switch assay

We have developed a new derivatization technique called

‘‘fluorescence switch’’ that substitutes S–nitrosylation by a

fluorophore, based on the use of maleimide reagents.

Fluorescence switch, coupled to 2-DE separation of proteins,

enables specific detection of the fluorescence substituting

S-nitrosylation, while allowing for fluorescent total protein

detection in the same gel. Thus, the whole amount of a

protein that is detected as S-nitrosylated is used for proteo-

mic identification, allowing for a reduction of the starting

material. Indeed, this methodology allows for performing

‘‘differential post-translational modification proteomics,’’ i.e.

to detect variations in the levels of the post-translational

modification studied, instead of differential proteomics

centered on the comparison of protein amounts (Fig. 1B).

Conceptually, the steps of the fluorescence switch are

similar to the biotin switch: blocking of free thiols with NEM,

specific reduction with ascorbate and labeling of newly

formed thiols with fluorescent maleimide (Fig. 1A). The use

of fluorescent maleimides as labeling reagents allows for

selecting among a wide range of reagents that could be used

in different gel-based detection schemes. To reduce possible

artifacts we preferred to rely on the same chemistry for both

blocking and labeling steps, using NEM instead of MMTS for

blocking. Other groups have also used maleimide reagents

for modifying the biotin switch with good results [27–30].

SH

SH

SH

SH

SH

SNO

SNO SNO

SNO

SH

SH

SH

SNEM SNEM

SNEM SNEM

SNEM FS

FS FS

FS

SNEM SNEM

SNEM

Fluorescenceswitch

Labelledprotein

Labelledprotein

Totalprotein

Totalprotein MergeMerge

A

B

SH SNO

Protein

S S

S-NEM SNO

S S

Protein

S S

Ascorbate

NEM

S-NEM SH

S S

Protein

S S

Mal-Fluor

S-NEM S-

Protein

S S

Fluorescenceswitch

F

F

Figure 1. (A) Scheme of the fluorescence switch assay, showing

the three chemical steps involved: blocking with N-ethylmalei-

mide (NEM), reduction by ascorbate and labeling with fluor-

escent maleimide (adapted from reference [14] with permission).

(B) Scheme of the results expected in 2-DE gels for a protein

subjected to the fluorescence switch, depending on the degree

of S-nitrosylation. The squares show the signals corresponding

to the fluorescence switch derivatization in green, total protein

detection in red, and both signals merged. There is a displace-

ment of the fluorescence switch signal, as the electrophoretic

mobility changes due to the addition of the labeling molecule.



Indeed, to reduce the possible secondary reaction of malei-

mides with protein amine groups, we used a lower pH during

both blocking and labeling steps, and we used a limited

amount of labeling maleimide.

To assess the specificity of the reaction scheme, we

performed several controls of possible artifacts. We checked

that blocking with NEM was effective, comparing it with the

usual biotin switch blocking with MMTS by performing

blocking and labeling steps without a reducing agent

(Fig. 2A). Furthermore, we compared the usual biotin

switch (blocking with MMTS and labeling with biotin-

HPDP) with a biotin switch based on maleimide reagents

for both blocking and labeling steps; the results were simi-

lar, although a slightly stronger signal was obtained with the

usual biotin switch (Fig. 2B).

Once we had established the validity of the reaction

scheme based on maleimide reagents, we performed the

same reaction scheme comparing the labeling step

performed with either biotin or fluorescein (Fig. 2C),

thus establishing the basic protocol of the ‘‘fluorescence

switch.’’ We obtained similar results with both labels, and

even a stronger increase in the signal with the fluorescent

label.

We performed two kinds of negative controls for the

fluorescence switch. In the first one, samples were incu-

bated with DTT to reduce protein cysteines and eliminate

S-nitrosothiols before the blocking step (Fig. 2D). In this

case we observed a clear enhancement of the signal only in

the samples obtained from CysSNO-treated cells incubated

without DTT. In the second one, samples were treated or

not with ascorbate to assess the specificity of the reduction

step (Fig. 2E), which showed that the CysSNO-induced

signal increase was dependent on ascorbate addition.

3.2 Detection of S-nitrosylated proteins in

endothelial cells

We applied the fluorescence switch method coupled to non-

reducing 2-DE to endothelial cells treated with a permeable

natural nitrosothiol, CysSNO. In this system there is a clear

increase in protein S-nitrosylation and we have previously

described the proteomic identification of S-nitrosylated

proteins using the usual biotin switch method with avidin

capture [31]. Extracts of the EA.hy926 cell line (200 mg of

protein) were subjected to the fluorescence switch and

A B C

D E

Figure 2. The fluorescence switch, based

on maleimide chemistry, is efficient and

specific. Shown are representative images

of at least three experiments performed in

EA.hy926 cells. (A) Comparison of the

efficiency of blocking with MMTS and

N-ethylmaleimide. Endothelial cell extracts

were subjected to the same steps of the

biotin switch, without adding reducing

agent (ascorbate). Blocking agent was

none (--), N-ethylmaleimide (Mal.) or

MMTS. Labeling agent was biotin-HPDP or

biotin-maleimide. (B) Comparison of the

standard biotin switch with a maleimide-

based biotin switch. Extracts from either

untreated (--) or S-nitroso-L-cysteine-trea-

ted (CNO) endothelial cells were subjected

to the biotin switch process, with the indi-

cated reagents for blocking and labeling

steps, and the biotinylation profile was

assessed by avidin Western blot. (C)

Comparison of a maleimide-based biotin

switch with the fluorescence switch assay

in extracts from either untreated (--) or

S-nitroso-L-cysteine-treated (CNO) endo-

thelial cells. (D) Fluorescence switch assay

performed on untreated or DTT-treated

extracts from either untreated or S-nitroso-

L-cysteine-treated endothelial cells. (E)

Fluorescence switch assay performed on

untreated or ascorbate-treated extracts

from either untreated or S-nitroso-L-

cysteine-treated endothelial cells.

Proteomics 2009, 9, 5359–5370 5363


separated by 2-DE. The fluorescent signal corresponding

to the incorporated maleimide reagent was recorded

(Supporting Information Fig. 1A), showing a clear differ-

ence between both cell extracts. This result corresponds to

the difference in protein S-nitrosylation in both samples, as

it was assessed by control reactions without ascorbate

(Supporting Information Fig. 1A); and it is not due to

differences in protein levels, as the protein profile is very

similar between both gels, as detected by total protein

staining (Supporting Information Fig. 1B).

To correctly match the fluorescence switch signals with

the corresponding total protein signals, it is important to

couple both signals obtained in the same gel conditions.

Many of the commercially available fluorescent reagents

that detect total protein are used with acidic pH, which

causes a loss in the fluorescence switch signal of the fluor-

escein-maleimide reagent. Thus, a total protein fluorescent

reagent compatible with neutral pH (Lucy-565, Sigma-

Aldrich) was used and both fluorescent signals were recor-

ded simultaneously (Supporting Information Fig. 1C).

As this new methodology requires much less starting

protein amount when compared to the biotin switch

method, we were able to confirm the results obtained in

EA.hy926 cells using a primary cell culture. Thus, we treated

HUVEC with or without CysSNO and performed the

fluorescence switch assay, including negative controls

without ascorbate, obtaining an almost identical S-nitrosy-

lation signal pattern in 2-DE gels (Fig. 3).

Proteins that showed a differential increased signal in the

CysSNO-treated sample with respect to both the untreated

A B C

l asaB

l asaB

ON

SsyC

ON

SsyC

csA-

csA

+

Figure 3. Detection of S-

nitrosylated proteins in

primary endothelial cells

(HUVEC) after S-nitroso-L-

cysteine (CysSNO) treat-

ment, by fluorescence

switch and 2-DE (pI 4–7).

Images are representative of

two biological replicates. (A)

Fluorescence switch signal

from an assay performed on

untreated or ascorbate-trea-

ted extracts from either

untreated or S-nitroso-L-

cysteine-treated endothelial

cells. (B) Sypro staining. (C)

Simultaneous fluorescence

switch (green) and total

protein (red) signals.



sample and the control without ascorbate, and that were

assigned to a precise spot in the total protein signal, were

cut from the gels, digested with trypsin and identified by

combined peptide mass fingerprinting and MS/MS peptide

sequencing using MALDI-TOF/TOF instruments (Fig. 4

and Table 1). In our previous reports using the usual biotin

switch on this system [31, 32], we were able to identify ten

proteins starting from 5 mg of protein extract. Now, we have

been able to identify 21 proteins that are S-nitrosylated in

intact endothelial cells, starting from only 200 mg of protein

extract in each gel and using a limited pI range in 2-DE,

thus significantly increasing sensitivity of the analysis.

An analysis of the proteins identified reveals that most of

them have already been detected as S-nitrosylation targets,

either in biotin-switch proteomic assays or in individual

protein studies. Hsp90, protein-glutamine gamma-gluta-

myltransferase 2 (tissue transglutaminase), vimentin, 14-3-3

proteins and beta-actin (actin, cytoplasmic 1) have been

already identified to be targets of S-nitrosylation in endo-

thelial cells [31–34]. Other proteins have been described as

S-nitrosylation targets in proteomic studies in other cell

systems: Grp75 (stress-70 protein, mitochondrial), lactate

dehydrogenase B chain, Hsp 27 (HSP beta-1), heat shock

70 kDa protein 4, elongation factor 2, alpha actinin and

tubulin alpha and beta [7, 15, 16, 29, 35–39]. A different

subunit (zeta) of the T-complex protein 1, another tropo-

myosin and other annexins have also been described to

be S-nitrosylated [16, 31, 37, 40]. Glyceraldehye-3-phosphate

dehydrogenase, a typical protein which is repeatedly iden-

tified as a target for S-nitrosylation in endothelial cells, was

not detected, most likely due to the limited pI range chosen

for 2-DE.

However, the method has some limitations that can be

considered. It relies on co-detection of the modified and

non-modified fractions of the protein and identification of

the whole protein amount, and thus some false positives

could arise. Indeed, some results can be ambiguous, as

S-nitrosylation detected in a particular 2-DE spot where two

proteins co-migrate cannot be undoubtedly assigned to any

of them. In our results, this can be the case for transitional

endoplasmic reticulum ATPase and 26S proteasome non-

ATPase regulatory subunit 2. It can be the case of protein

isoforms that comigrate, as in the cases of Hsp90 and

tubulin beta and alpha chains, although it is plausible that

the modified cysteines could be homologous in the different

isoforms, as it happens in Hsp90 [32].

Two more limitations can be considered. One, shared by

the ‘‘classic’’ biotin switch and circumvented by peptide-

based approaches [15–17], is that the modified residue is not

easily identified. The other one is that the fluorescence

switch assay cannot be coupled to individual protein detec-

tion by Western blot as it happens with the biotin switch

(see scheme in [14]) because the modified protein is not

separated from the non-modified one.

As it happens in other proteomic methods, including the

biotin switch, this new method can be considered a

screening technique that points out new putative targets that

can be confirmed individually. Actually, for studying

S-nitrosylation, it is recommended to employ several

orthogonal techniques (based on different chemical

approaches) to confirm the modification [41]. The particular

usefulness of this method would be for screening

scarce samples, for example, in animal models or clinical

studies.

3.3 Detection of endogenously S-nitrosylated

proteins in activated macrophages

To detect changes induced by NO endogenously

produced by nitric oxide synthase enzyme, we applied the

fluorescence switch technique to RAW264.7 (murine

macrophage) cells treated with LPS and IFN-g.

Although iNOS expression was induced and nitrite was

21

4 3

56

7 89

1011

12

1314

18

1 34

67

9

13

14

A B

Figure 4. Fluorescence switch

detection of 2-DE gels from

CysSNO-treated endothelial

cells, with indication of the

spots identified in each gel. (A)

EA.hy926, representative of

four biological replicates. (B)

HUVEC, representative of two

biological replicates.

Proteomics 2009, 9, 5359–5370 5365


Tab

le1.

Pro

tein

sid

en

tifi

ed

fro

mC

ysS

NO

-tre

ate

den

do

theli

al

cell

s

Sp

ot

no

.

Acc

ess

ion

cod

e

Pro

tein

defi

nit

ion

Tim

es

iden

tifi

ed

a)

Masc

ot

sco

reb

)M

asc

ot

exp

ect

b)

MM

(Da),

theo

r.

pI

theo

r.M

atc

hed

pep

tid

esb

)C

over.

%b

)

HU

VE

A

1c)

HS

90A

_HU

MA

NH

eat

sho

ckp

rote

inH

SP

90-a

23

1090

2e-1

05

85

006

4.9

434

48

HS

90B

_HU

MA

NH

eat

sho

ckp

rote

inH

SP

90-b

22

572

1.3

e-0

53

83

554

4.9

715

23

2H

SP

74_H

UM

AN

Heat

sho

ck70

kDa

pro

tein

41

2512

1.3

e-0

47

95

127

5.1

120

30

3c)

PS

MD

2_H

UM

AN

26S

pro

teaso

me

no

n-A

TP

ase

reg

ula

tory

sub

un

it2

3185

6.5

e-0

15

100

877

5.0

87

10

TE

RA

_HU

MA

NT

ran

siti

on

al

en

do

pla

smic

reti

culu

mA

TP

ase

22

417

4.1

e-0

38

89

950

5.1

425

40

4T

GM

2_H

UM

AN

Pro

tein

-glu

tam

ine

gam

ma-g

luta

mylt

ran

sfera

se2

23

624

8.1

e-0

59

78

420

5.1

118

37

5H

NR

PK

_HU

MA

NH

ete

rog

en

eo

us

nu

clear

rib

on

ucl

eo

pro

tein

K1

2290

2e-0

25

51

230

5.3

910

30

6V

IME

_HU

MA

NV

imen

tin

11

1090

2e-1

05

53

676

5.0

639

74

7c)

TB

B5_H

UM

AN

d)

Tu

bu

linb

chain

13

682

1.3

e-0

64

50

095

4.7

823

58

TB

A1C

_HU

MA

Nd

)T

ub

uli

na-

1C

chain

13

1020

2e-0

98

50

548

4.9

618

53

8IF

4A

1_H

UM

AN

Eu

kary

oti

cin

itia

tio

nfa

cto

r4A

-I1

373

1e-0

33

46

353

5.3

214

38

9A

CT

B_H

UM

AN

Act

in,

cyto

pla

smic

12

3710

2e-0

67

42

052

5.2

916

57

10

TM

OD

3_H

UM

AN

Tro

po

mo

du

lin

-31

69

0.0

044

39

741

5.0

83

14

11

HN

RP

C_H

UM

AN

Hete

rog

en

eo

us

nu

clear

rib

on

ucl

eo

pro

tein

sC

1/C

21

144

8.1

e-0

11

33

707

4.9

54

15

12

NP

M_H

UM

AN

Nu

cleo

ph

osm

in2

166

5.1

e-0

13

32

726

4.6

44

21

13

TP

M3_H

UM

AN

Tro

po

myo

sina-

3ch

ain

12

297

4.1

e-0

26

32

856

4.6

87

16

14

c)

1433Z

_HU

MA

N14-3

-3p

rote

inz/d

11

366

5.1

e-0

33

27

899

4.7

36

33

1433E

_HU

MA

N14-3

-3p

rote

ine

1152

1.3

e-0

11

29

326

4.6

34

18

1433B

_HU

MA

N14-3

-3p

rote

inb/a

192

1.4

e-0

05

28

179

4.7

62

13

15

c)

AC

TN

1_H

UM

AN

a-A

ctin

in-1

11

418

3.2

e-0

38

103

563

5.2

531

46

AC

TN

4_H

UM

AN

a-A

ctin

in-4

2210

2.0

e-0

17

105

245

5.2

717

23

16

GR

P75_H

UM

AN

Str

ess

-70

pro

tein

,m

ito

cho

nd

rial

1207

1.2

e-0

14

73

920

5.8

77

13

17

TC

PQ

_HU

MA

NT

-co

mp

lex

pro

tein

1su

bu

nit

theta

1123

1.0

e-0

08

60

153

5.4

23

6

18

LD

HB

_HU

MA

NL-l

act

ate

deh

yd

rog

en

ase

Bch

ain

2309

2.6

e-0

27

36

900

5.7

17

26

19

HS

PB

1_H

UM

AN

Heat

sho

ckp

rote

inb-

11

121

1.6

e-0

08

22

826

5.9

82

13

20

AN

XA

1_H

UM

AN

An

nexin

A1

1122

1.3

e-0

08

38

918

6.5

78

31

21

EF2_H

UM

AN

Elo

ng

ati

on

fact

or

21

159

2.6

e-0

12

96

246

6.4

112

13

Th

eta

ble

sum

mari

zes

the

resu

lts

fro

mfo

ur

ind

ep

en

den

texp

eri

men

tso

nE

A.h

y926

cell

san

dtw

oin

dep

en

den

texp

eri

men

tso

nH

UV

EC

.a)

Nu

mb

er

of

rep

lica

tes

inw

hic

heach

pro

tein

was

iden

tifi

ed

inH

UV

EC

(HU

V)

or

EA

.hy926

cell

s(E

A).

b)

Data

fro

mth

eid

en

tifi

cati

on

wit

hh

igh

est

Masc

ot

sco

re.

c)M

ixtu

reo

fp

rote

ins

dete

cted

inth

esa

me

spo

ts.

d)

Severa

lis

ofo

rms

were

iden

tifi

ed

.



accumulated extracellularly (Fig. 5A), we did not observe an

increment in the S-nitrosothiol signal (Fig. 5C). There are

reports that show a difference in signal using either

biotin switch or a similar fluorescence technique [30, 36].

However, we have repeatedly observed this result using

both fluorescence switch and biotin switch (Fig. 5B).

Recently, in accordance with our results, Forrester et al.have reported that only one protein was clearly differen-

tially labeled using a novel switch methodology, SNO-RAC

[17].

This could be explained by a protection mechanism

exerted by the nitric-oxide producing macrophage in

1 2

35

6

7 8

9

10 11

4

C

A

iNOS

-+

B

Figure 5. Detection of S-nitrosylated proteins in RAW264.7 cell cultures, treated or not with LPS and IFN-g or auranofin. (A) Nitrite

detection by Griess reaction, and Western blot detection of iNOS. (B) Avidin Western blot of cell extracts subjected to the biotin switch

assay and separated by SDS-PAGE. (C) Fluorescence switch signal of 2-DE gels of extracts from RAW264.7 cells, indicating the spots that

were cut and digested for MS identification. Images are representative of two biological replicates.

Table 2. Proteins identified from auranofin-treated activated macrophages

Spot no. Accession code Protein definition Mascot

score

Mascot

expect

MW (Da),

theor.

pI theor. Matched

peptides

Cover. %

1 HSP74_MOUSE Heat shock 70 kDa protein 4 158 2.50E�12 94 872 5.15 7 11

2 HSP7C_MOUSE Heat shock cognate 71 kDa protein 159 2.00E�12 71 055 5.37 5 10

3 UBA1_MOUSE Ubiquitin-like modifier-activating

enzyme

166 4.00E�13 118 931 5.43 2 2

4 UBP5_MOUSE Ubiquitin carboxyl-terminal hydrolase 5 149 2.00E�11 96 685 4.89 5 8

5 PSMD2_MOUSE 26S proteasome non-ATPase regulatory

subunit 2

122 1.00E�08 100 937 5.06 2 3

6 GRP78_MOUSE 78 kDa glucose-regulated protein 210 1.60E�17 72 492 5.07 5 12

7 PDIA1_MOUSE Protein disulfide-isomerase 134 6.30E�10 57 507 4.79 9 20

8 PDIA3_MOUSE Protein disulfide-isomerase A3 88 2.60E�05 57 099 5.88 6 11

9 RINI_MOUSE Ribonuclease inhibitor 132 1.00E�09 51 495 4.69 4 11

10 CO3_MOUSE Complement C3 316 4.00E�28 187 904 6.39 6 5

11 RSSA_MOUSE 40S ribosomal protein SA 216 4.00E�18 32 931 4.8 4 17

Proteomics 2009, 9, 5359–5370 5367


order to maintain the intracellular protein redox

status. Several systems involved in the homeostasis of

S-nitrosylation by acting as ‘‘denitrosylases’’ have been

described and they could be responsible for this protection.

Among them, the role of thioredoxin in protein

denitrosylation, as well as the impairment of this

activity with thioredoxin reductase inhibitors such as

auranofin, has been recently described [10, 42]. Thus, we

performed the fluorescence switch assay in activated

RAW264.7 cells treated with auranofin for 1 h, and we were

able to detect several proteins that had an increased signal

(Fig. 5C).

Among them, we were able to identify 11 proteins that

showed an increased fluorescence switch signal when trea-

ted with auranofin, and that were matched to total protein

spots in 2-DE gels (Fig. 5C and Table 2). It is noteworthy

that there are targets from different subcellular locations

(endoplasmic reticulum (Grp78 and protein disulfide-

isomerases), ribosome and cytoplasm). This finding

suggests that S-nitrosylation regulation by thioredoxins

could be a mechanism exerted throughout several subcel-

lular environments.

As far as we know, some of the proteins identified are

novel targets for S-nitrosylation. 26S proteasome non-

ATPase regulatory subunit 2, found in CysSNO treatment,

has also been found in this setting. Two proteins related

with ubiquitination have been found, an ubiquitin-activating

E1 protein (ubiquitin-like modifier enzyme) and a protease

(ubiquitin carboxyl-terminal hydrolase 5), which reinforce

the role of S-nitrosylation in regulating ubiquitination, as it

has been shown for a E3 protein, parkin [12, 43]. Ribonu-

clease inhibitor has a high number of redox-sensitive

cysteine residues whose oxidation results in functional

changes [44]. It also plays a role in cell redox homeostasis

[45] and thus enzymes such as thioredoxin could be

implied in the maintenance of its redox status, including

S-nitrosylation.

4 Concluding remarks

We have developed a methodology to selectively label

S-nitrosylated proteins with maleimide-based fluorescent

reagents. This ‘‘fluorescence switch,’’ coupled to 2-DE,

allows detection of variations in protein S-nitrosylation, and

an easier identification of modified proteins, while using a

limited amount of protein extract. This represents an

increase in sensitivity compared to the commonly used

‘‘biotin switch’’ technique.

This approach would allow to study the S-nitrosopro-

teomes in several (patho)physiological settings where the

starting material might be limiting, such as in vivo samples

of small tissues. Another potential advantage is the use of

different fluorophores, in a similar fashion as the DIGE

technology, which would allow to more precisely quantify

variations in S-nitrosylation of individual proteins.

Tight regulation of protein S-nitrosylation (both the

build-up and breakdown) is considered to be highly relevant

in order to accept this modification as a fully physiological

mechanism. By using the ‘‘fluorescence switch’’ we have

been able to detect several proteins that became S-nitrosy-

lated when activated macrophages were treated with an

inhibitor of the thioredoxin/thioredoxin reductase pathway.

This suggests the existence of novel targets that could be

regulated by this post-translational modification under

(patho)physiological stimuli.

This work was supported by grants from the SpanishGovernment (FIS CP03/00025 and CP07/00143 to A. M.-R.,SAF2006-02410 to S. L. and CSD2007-00020 to A. M.-R. andS. L.). We want to thank Juan Miguel Redondo and ManuelOrtiz de Landazuri and all the people in their labs for the helpand support received, Silvia Vazquez for technical assistance, andJesus Jorrın-Novo for his helpful comments on the manuscript. Wethank the following proteomics facilities and persons involved in2-DE and protein identification, for their expert service: JuanAntonio Lopez and Emilio Camafeita from the Unidad deProteomica, CNIC (Madrid, Spain); Marisol Fernandez from theLaboratorio de Proteomica, CNB (Madrid, Spain) and Unidadde Proteomica, CIMA (Pamplona, Spain); CNB and CIMAfacilities are members of ProteoRed.

The authors have declared no conflict of interest.

5 References

[1] Martınez-Ruiz, A., Lamas, S., Two decades of new concepts

in nitric oxide signaling: From the discovery of a gas

messenger to the mediation of nonenzymatic posttransla-

tional modifications. IUBMB Life 2009, 61, 91–98.

[2] Martınez-Ruiz, A., Lamas, S., Signalling by NO-induced

protein S-nitrosylation and S-glutathionylation: Conver-

gences and divergences. Cardiovasc. Res. 2007, 75,

220–228.

[3] Foster, M. W., McMahon, T. J., Stamler, J. S., S-nitrosyla-

tion in health and disease. Trends Mol. Med. 2003, 9,

160–168.

[4] Stamler, J. S., Lamas, S., Fang, F. C., Nitrosylation: the

prototypic redox-based signaling mechanism. Cell 2001,

106, 675–683.

[5] Martınez-Ruiz, A., Lamas, S., S-nitrosylation: a potential

new paradigm in signal transduction. Cardiovasc. Res.

2004, 62, 43–52.

[6] Kaneko, R., Wada, Y., Decomposition of protein nitro-

sothiols in matrix-assisted laser desorption/ionization and

electrospray ionization mass spectrometry. J. Mass Spec-

trom. 2003, 38, 526–530.

[7] Jaffrey, S. R., Erdjument-Bromage, H., Ferris, C. D., Tempst, P.,

Snyder, S. H., Protein S-nitrosylation: a physiological

signal for neuronal nitric oxide. Nat. Cell Biol. 2001, 3,

193–197.



[8] Zhang, Y., Keszler, A., Broniowska, K. A., Hogg, N., Char-

acterization and application of the biotin-switch assay for

the identification of S-nitrosated proteins. Free Radic. Biol.

Med. 2005, 38, 874–881.

[9] Liu, L., Hausladen, A., Zeng, M., Que, L. et al., A metabolic

enzyme for S-nitrosothiol conserved from bacteria to

humans. Nature 2001, 410, 490–494.

[10] Benhar, M., Forrester, M. T., Hess, D. T., Stamler, J. S.,

Regulated protein denitrosylation by cytosolic and mito-

chondrial thioredoxins. Science 2008, 320, 1050–1054.

[11] Uehara, T., Nakamura, T., Yao, D., Shi, Z.-Q. et al.,

S-Nitrosylated protein-disulphide isomerase links protein

misfolding to neurodegeneration. Nature 2006, 441,

513–517.

[12] Chung, K. K. K., Thomas, B., Li, X., Pletnikova, O. et al.,

S-Nitrosylation of parkin regulates ubiquitination and

compromises parkin’s protective function. Science 2004,

304, 1328–1331.

[13] Ibiza, S., Perez-Rodrıguez, A., Ortega, A ., Martınez-Ruiz, A.

et al., Endothelial nitric oxide synthase regulates

N-Ras activation on the Golgi complex of antigen-stimu-

lated T cells. Proc. Natl. Acad. Sci. USA 2008, 105,

10507–10512.

[14] Martınez-Ruiz, A., Lamas, S., Detection and identification of

S-nitrosylated proteins in endothelial cells. Methods Enzy-

mol. 2005, 396, 131–139.

[15] Hao, G., Derakhshan, B., Shi, L., Campagne, F., Gross, S. S.,

SNOSID, a proteomic method for identification of cysteine

S-nitrosylation sites in complex protein mixtures. Proc.

Natl. Acad. Sci. USA 2006, 103, 1012–1017.

[16] Greco, T. M., Hodara, R., Parastatidis, I., Heijnen, H. F. G.

et al., Identification of S-nitrosylation motifs by site-specific

mapping of the S-nitrosocysteine proteome in human

vascular smooth muscle cells. Proc. Natl. Acad. Sci. USA

2006, 103, 7420–7425.

[17] Forrester, M. T., Thompson, J. W., Foster, M. W., Nogueira, L.

et al., Proteomic analysis of S-nitrosylation and denitrosyla-

tion by resin-assisted capture. Nat. Biotechnol. 2009, 27,

557–559.

[18] Jourd’heuil, D., Gray, L., Grisham, M. B., S-nitrosothiol

formation in blood of lipopolysaccharide-treated rats.

Biochem. Biophys. Res. Commun. 2000, 273, 22–26.

[19] DeMaster, E. G., Quast, B. J., Redfern, B., Nagasawa, H. T.,

Reaction of nitric oxide with the free sulfhydryl group of

human serum albumin yields a sulfenic acid and nitrous

oxide. Biochemistry 1995, 34, 11494–11499.

[20] Munoz, C., Castellanos, M. C., Alfranca, A., Vara, A. et al.,

Transcriptional up-regulation of intracellular adhesion

molecule-1 in human endothelial cells by the antioxidant

pyrrolidine dithiocarbamate involves the activation of acti-

vating protein-1. J. Immunol. 1996, 157, 3587–3597.

[21] Zaragoza, C., Lopez-Rivera, E., Garcıa-Rama, C., Saura, M.

et al., Cbfa-1 mediates nitric oxide regulation of MMP-13 in

osteoblasts. J. Cell Sci. 2006, 119, 1896–1902.

[22] Wessel, D., Fl .ugge, U. I., A method for the quantitative

recovery of protein in dilute solution in the presence of

detergents and lipids. Anal. Biochem. 1984, 138, 141–143.

[23] Forrester, M. T., Foster, M. W., Stamler, J. S., Assessment

and application of the biotin switch technique for examin-

ing protein s-nitrosylation under conditions of pharmaco-

logically induced oxidative stress. J. Biol. Chem. 2007, 282,

13977–13983.

[24] Yan, J. X., Wait, R., Berkelman, T., Harry, R. A. et al.,

A modified silver staining protocol for visualization of

proteins compatible with matrix-assisted laser desorption/

ionization and electrospray ionization-mass spectrometry.

Electrophoresis 2000, 21, 3666–3672.

[25] Corton, M., Botella-Carretero, J. I., Lopez, J. A., Camafeita, E.

et al., Proteomic analysis of human omental adipose tissue

in the polycystic ovary syndrome using two-dimensional

difference gel electrophoresis and mass spectrometry. Hum.

Reprod. 2008, 23, 651–661.

[26] Shevchenko, A., Tomas, H., Havlis, J., Olsen, J. V., Mann, M.,

In-gel digestion for mass spectrometric characterization of

proteins and proteomes. Nat. Protoc. 2007, 1, 2856–2860.

[27] Ckless, K., Reynaert, N. L., Taatjes, D. J., Lounsbury, K. M.

et al., In situ detection and visualization of S-nitrosylated

proteins following chemical derivatization: identification of

Ran GTPase as a target for S-nitrosylation. Nitric Oxide

2004, 11, 216–227.

[28] Kettenhofen, N. J., Broniowska, K. A., Keszler, A., Zhang, Y.,

Hogg, N., Proteomic methods for analysis of S-nitrosation.

J. Chromatogr. B 2007, 851, 152–159.

[29] Moon, K.-H., Hood, B. L., Mukhopadhyay, P., Rajesh, M.

et al., Oxidative inactivation of key mitochondrial proteins

leads to dysfunction and injury in hepatic ischemia reper-

fusion. Gastroenterology 2008, 135, 1344–1357.

[30] Santhanam, L., Gucek, M., Brown, T. R., Mansharamani, M.

et al., Selective fluorescent labeling of S-nitrosothiols (S-

FLOS): A novel method for studying S-nitrosation. Nitric

Oxide 2008, 19, 295–302.

[31] Martınez-Ruiz, A., Lamas, S., Detection and proteomic

identification of S-nitrosylated proteins in endothelial cells.

Arch. Biochem. Biophys. 2004, 423, 192–199.

[32] Martınez-Ruiz, A., Villanueva, L., Gonzalez de Orduna, C.,

Lopez-Ferrer, D. et al., S-nitrosylation of Hsp90 promotes

the inhibition of its ATPase and endothelial nitric oxide

synthase regulatory activities. Proc. Natl. Acad. Sci. USA

2005, 102, 8525–8530.

[33] Lai, T. S., Hausladen, A., Slaughter, T. F., Eu, J. P. et al.,

Calcium regulates S-nitrosylation, denitrosylation, and

activity of tissue transglutaminase. Biochemistry 2001, 40,

4904–4910.

[34] Yang, Y., Loscalzo, J., S-nitrosoprotein formation and

localization in endothelial cells. Proc. Natl. Acad. Sci. USA

2005, 102, 117–122.

[35] Lindermayr, C., Saalbach, G., D .urner, J., Proteomic Identi-

fication of S-Nitrosylated Proteins in Arabidopsis. Plant

Physiol. 2005, 137, 921–930.

[36] Gao, C., Guo, H., Wei, J., Mi, Z. et al., Identification of

S-nitrosylated proteins in endotoxin-stimulated RAW264.7

murine macrophages. Nitric Oxide 2005, 12, 121–126.

[37] Dall’Agnol, M., Bernstein, C., Bernstein, H., Garewal, H.,

Payne, C. M., Identification of S-nitrosylated proteins after

Proteomics 2009, 9, 5359–5370 5369


chronic exposure of colon epithelial cells to deoxycholate.

Proteomics 2006, 6, 1654–1662.

[38] Lopez-Sanchez, L. M., Corrales, F. J., Gonzalez, R., Ferrın, G.

et al., Alteration of S-nitrosothiol homeostasis and targets

for protein S-nitrosation in human hepatocytes. Proteomics

2008, 8, 4709–4720.

[39] Han, P., Chen, C., Detergent-free biotin switch combined

with liquid chromatography/tandem mass spectrometry in

the analysis of S-nitrosylated proteins. Rapid Commun.

Mass Spectrom. 2008, 22, 1137–1145.

[40] Kuncewicz, T., Sheta, E. A., Goldknopf, I. L., Kone, B. C.,

Proteomic analysis of S-nitrosylated proteins in mesangial

cells. Mol. Cell Proteomics 2003, 2, 156–163.

[41] Lancaster, J. R., Jr., Gaston, B., NO and nitrosothiols:

spatial confinement and free diffusion. Am. J. Physiol. Lung

Cell Mol. Physiol. 2004, 287, L465–466.

[42] Sengupta, R., Ryter, S. W., Zuckerbraun, B. S., Tzeng, E.

et al., Thioredoxin catalyzes the denitrosation of low-

molecular mass and protein S-nitrosothiols. Biochemistry

2007, 46, 8472–8483.

[43] Yao, D., Gu, Z., Nakamura, T., Shi, Z.-Q. et al., Nitrosative

stress linked to sporadic Parkinson’s disease: S-nitrosyla-

tion of parkin regulates its E3 ubiquitin ligase activity. Proc.

Natl. Acad. Sci. USA 2004, 101, 10810–10814.

[44] Ferreras, M., Gavilanes, J. G., Lopez-Otın, C., Garcıa-

Segura, J. M., Thiol-disulfide exchange of ribonuclease

inhibitor bound to ribonuclease A. J. Biol. Chem. 1995, 270,

28570–28578.

[45] Monti, D. M., Montesano Gesualdi, N., Matousek, J., Espo-

sito, F., D’Alessio, G., The cytosolic ribonuclease inhibitor

contributes to intracellular redox homeostasis. FEBS Lett.

2007, 581, 930–934.



a “fluorescence switch” technique increases the sensitivity of proteomic detection and...

Documents