Genetically encoded reactive oxygen species (ROS) and redox indicators.

Biotechnology Journal

Biotechnol. J. 2014, 9, 282–293

DOI 10.1002/biot.201300199

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Review

Genetically encoded reactive oxygen species (ROS) and redox indicators Sandrine Pouvreau1,2 1 University 2 CNRS,

of Bordeaux, Interdisciplinary Institute for Neuroscience, UMR 5297, Bordeaux, France Interdisciplinary Institute for Neuroscience, UMR 5297, Bordeaux, France

Redox processes are increasingly being recognized as key elements in the regulation of cellular signaling cascades. They are frequently encountered at the frontier between physiological functions and pathological events. The biological relevance of intracellular redox changes depends on the subcellular origin, the spatio-temporal distribution and the redox couple involved. Thus, a key task in the elucidation of the role of redox reactions is the specific and quantitative measurement of redox conditions with high spatio-temporal resolution. Unfortunately, until recently, our ability to perform such measurements was limited by the lack of adequate technology. Over the last 10 years, promising imaging tools have been developed from fluorescent proteins. Genetically encoded reactive oxygen species (ROS) and redox indicators (GERRIs) have the potential to allow real-time and pseudo-quantitative monitoring of specific ROS and thiol redox state in subcellular compartments or live organisms. Redox-sensitive yellow fluorescent proteins (rxYFP family), redox-sensitive green fluorescent proteins (roGFP family), HyPer (a probe designed to measure H2O2), circularly permuted YFP and others have been used in several models and sufficient information has been collected to highlight their main characteristics. This review is intended to be a tour guide of the main types of GERRIs, their origins, properties, advantages and pitfalls.

Received 04 JUL 2013 Revised 10 SEP 2013 Accepted 06 NOV 2013

Keywords: Fluorescent protein sensors · Live cell imaging · Reactive oxygen species · Redox processes · Subcellular compartments imaging

1 Introduction: Intracellular redox signaling Redox signaling occurs whenever a variation in the concentration of reactive oxygen species (ROS) or a shift in the redox state of a redox couple alters a biological system. An increasing number of evidences suggest that redox processes regulate diverse physiological functions

Correspondence: Dr. Sandrine Pouvreau, Interdisciplinary Institute for Neuroscience, CNRS/University Victor Segalen Bordeaux 2, 146 rue Leo Saignat, 33077 Bordeaux cedex, France E-mail: [email protected] Abbreviations: cpYFP, circularly permuted YFP; E°’, midpoint redox potential; EGFP, enhanced GFP; ER, endoplasmic reticulum; GERRI, genetically encoded ROS and redox indicator; GFP, green fluorescence protein; Grx, glutaredoxin; GSH, reduced glutathione; GSSG, glutathione disulfide; roGFP, redox-sensitive GFP; ROS, reactive oxygen species; rxYFP, redoxsensitive YFP; wtYFP, wild-type YFP; YFP, yellow fluorescent protein

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such as responses to growth factor stimulation, differentiation, cell cycle, metabolism, cell migration, gene transcription, intracellular calcium release [1], communication between organelles [2, 3] immune responses, and stress responses (for review, see [4]).

1.1 Redox-sensitive proteins To act as signaling messengers, redox-active molecules have to induce a reversible change in the activity of target proteins. Generally, this involves modification of thiol (SH) groups on specific cysteine residues located in redox-sensitive proteins. Thiols can be modified by a direct reaction with oxidants, independently of the average redox state of the cell compartment, or through reactions with another thiol–disulfide redox couple. Certain thiols are particularly sensitive to oxidation. These thiols are known as “functional switches” (for review, see [5]).

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Biotechnol. J. 2014, 9, 282–293


The redox sensitivity of a thiol is mainly determined by its local environment. One important factor is accessibility since the redox molecule has to be able to reach the reactive thiol in the protein. The pH of the subcellular compartment in which the target protein is located is also a determinant for the regulation process. For instance, the oxidant hydrogen peroxide (H2O2) reacts only with thiolates (-S–). Although the pKa of thiol groups on free cysteines is typically around 8, in proteins the amino acids surrounding the cysteine residue can substantially lower the pKa value to as low as 4–5. As a consequence, these residues are mostly deprotonated under physiological conditions. Deprotonated cysteine residues are vulnerable to oxidation to sulfenic acids (RSOH). Sulfenic acids are unstable and can be irreversibly oxidized to sulfinic (RSO2H) or sulfonic (RSO3H) acids under stronger oxidative stress. Post-translational cysteine redox modifications also include S-glutathionylation (RSSG), S-nitrosylation (RSNO), or the formation of an intra- or intermolecular disulfide bond (RSSR). Oxidized cysteines can be reduced through thiol disulfide exchanges with other redox couples. These exchanges can be catalyzed by thioredoxins or glutaredoxins. Numerous proteins are modulated by these redox modifications under physiological conditions, including protein tyrosine phosphatases, members of the Ras family, proteases, transcription factors, and ion channels (for reviews, see [6–9]).

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Glutathione is with few exceptions the most abundant nonprotein thiol in the cell, and is considered to be the major player in maintaining intracellular redox equilibrium [11]. Intracellular glutathione concentrations are reported to be as high as 5–10 mM, and ratios of GSH to GSSG to range from 30 to 100:1 [12].

1.3 Specificity of the reaction between redox species and target proteins In order to understand the regulation of signaling pathways by redox processes, we need to know what species are involved under different circumstances and their reactivities. Indeed, not only are some species much more reactive or strongly oxidizing than others, but also the electron transfer is context dependent. For instance, O2·–, despite its reduction potential of 0.9 V, is a poor oxidant in vivo as it is rapidly dismuted to H2O2, and also because its anionic charge limits reactivity with electron-rich centers. H2O2 also reacts poorly with most biological molecules as its reactions with typical thiols are slow. Its biological effects are mainly due to the secondary products of its main reaction with seleno-, thiol or heme peroxidases or other transition metals, as well as hydroxyl radicals formed by Fenton chemistry (for review, see [7]). Consequently, it is necessary to analyze the various redox species specifically and separately.

1.4 Redox signaling: Location, location, location 1.2 Intracellular production and clearance mechanisms of ROS The term ROS generally includes the primary species generated by oxygen reduction (superoxide or H2O2) as well as their secondary reactive products. The mitochondrial electron transport chain and NADPH oxidases are the main sources of ROS within the cell. The superoxide anion O2·– is the first species to be produced, but it is rapidly converted to H2O2 by spontaneous dismutation or superoxide dismutases that are present within the mitochondrial matrix and the cytosol [2, 6]. H2O2 can be converted through the Fenton reaction into a highly reactive product, the hydroxyl radical (OH·). Additional intracellular sources of ROS include peroxisomes and numerous other enzymes, such as nitric oxide synthases, xanthine oxidase, cyclooxygenases, cytochrome p450 enzymes, the redox enzyme p66Shc and lipoxygenases (for reviews, see [6, 7, 10]). Oxidants are also generated in the endoplasmic reticulum (ER) during the process of oxidative protein folding. Decomposition of H2O2 is catalyzed by catalases, glutathione peroxidases, and peroxiredoxins. Glutathione peroxidases reduce free H2O2 to water using reduced glutathione (GSH) as the reducing substrate. To complete the cycle, the resulting glutathione disulfide (GSSG) is then reduced to GSH by glutathione reductases, using NADPH.


The vast existing literature about redox signaling often appears very confusing, probably due to the fact that dynamic changes in redox species and their pharmacology are reported as “redox state”, “oxidative stress”, “antioxidant therapy”, with no further precision about the species involved, or the spatial or temporal resolution of the process. Such a terminology implies that redox states are global properties of the cell, and are not compartment or species dependent. This is evidently highly incorrect as cells contain numerous redox species that are kinetically controlled, far from equilibrium, and which activities are compartment, and sometimes tissue, dependent. Accordingly, a gradient of H2O2 levels within tissues has been recently identified in the zebrafish [13]. In the same way, Albrecht et al. [14] found, using genetically encoded redox probes for GSH/GSSG and H2O2 in Drosophila, that redox changes are associated with development and aging in this insect. These changes are redox species specific, as the two oxidant species have been shown to vary independently, as well as subcellular compartment and tissue specific. To understand redox processes, knowing their location and intracellular compartmentalization is essential, as oxidants, due to their reactive nature, must be generated close to their targets. Moreover, while certain ROS such as H2O2 can cross biological membranes, others, such as O2·–, cannot (for reviews,

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see [2, 4, 7, 8]), so the target protein has to be located in the same compartment as the source of O2·–. In conclusion, in the same way that we talk about “increase in cytosolic calcium level” and not “change in cellular ionic state” while reporting calcium signaling events, we should always be specific about the species involved, the dynamics and the specific subcellular compartment while studying redox processes.

2 Measuring redox species: Challenges and limitations As discussed above, redox signaling is species – subcellular compartment – and time dependent. Therefore, unraveling redox mechanisms requires the ability to look at redox processes with high specificity, spatial and temporal resolution. Hence the ideal technique for redox species measurement should fulfill the following requirements: (i) be highly sensitive to physiologically relevant changes in redox species level; (ii) provide specificity towards a particular molecule; (iii) allow temporal resolution of redox signaling; (iv) allow spatial resolution of redox signaling; (v) be able to detect changes in oxidized or reduced compartments; (vi) be quantitative; (vii) be insensitive to other environmental parameters such as pH; (viii) should not interfere with the system, e.g. ROS sensors should not behave as antioxidants; and (ix) be non invasive and ultimately allow ROS detection in vivo. The initial redox measurements in cell or animal models have relied either on disruptive methods, such as cell or tissue extracts, or on indirect evidence, such as latestage markers of cellular damage [11]. Measurement of the ratio between the oxidized and reduced forms of a redox couple in tissue or cell extracts does not provide adequate spatial resolution, as it averages over different cell types or subcellular compartments. In the same way, oxidative damage markers are not reliable as they might reflect changes in repair and turnover rather than increase in oxidant levels [15]. In addition, these techniques are invasive and do not provide the adequate temporal resolution and specificity. In this context, synthetic fluorescent sensors such as dichlorodihydrofluorescein, hydroethidine (and its mitochondrial targeted version known as MitoSox), dihydrorhodamine, coumarin boronate and Amplex red appeared promising, as they allow the visualization of changes in redox species in a direct and non-invasive manner using fluorescence microscopy (for review, see [16]). Unfortunately, apart from MitoSox, these sensors cannot be targeted to subcellular compartments. Furthermore, most of them are not specific and/or not reversible, hence impairing the temporal resolution of the dynamic events. To date, the most advanced and promising tools for real-time monitoring of ROS and thiol redox state in living cells and tissues are genetically encoded ROS and redox

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Box 1: The ideal probe… 1. is highly sensitive to relevant changes in redox species level 2. is specific to a particular molecule 3. exhibits a high dynamic range 4. has fast kinetics compared to the time course of the redox signal 5. is reversible 6. allows subcellular resolution of redox signaling 7. is able to detect redox shifts in oxidized or reduced compartments 8. is ratiometric 9. does not interfere with the recorded biological system 10. is not sensitive to other environmental parameters such as pH fluctuations 11. can be expressed in live organisms

indicators (GERRIs) derived from the green fluorescent protein (GFP). These biosensors exhibit most of the qualities required for an ideal probe (see Box 1). As genetically encoded probes, they can be specifically targeted to different subcellular locations. Some are highly specific to certain redox couples. Several are ratiometric, allowing quantitative measurements with proper calibration. They are reversible, by in large using the same reducing mechanisms as endogenous redox-sensitive proteins. In addition, redox biosensor transgenic model organisms open the possibility of dynamic in vivo imaging of redox processes. Finally, GERRIs can be upgraded with targeted mutations. Within a few years since their initial introduction, several redox-sensitive fluorescent proteins have been developed to monitor dithiol/disulfide ratio, GSH/GSSG, H2O2, and O2·–. The chemistry of these probes is reviewed in the excellent article by Meyer and Dick [17]. The goal of the present review is to discuss the most relevant characteristics for cell biology of the main redox-sensitive fluorescent probes, and how those characteristics can affect the outcome of our experiments.

3 Genetically encoded sensors for dithiol/disulfide equilibrium Redox-sensitive YFP (rxYFP) and redox-sensitive GFP (roGFP) were created by two different laboratories by introduction of redox-reactive cysteines into fluorescent proteins at locations at which a change in the oxidation state of these redox groups would change the fluorescent properties. Both fluorescent probes allow non-invasive imaging of dithiol/disulfide equilibrium.



Biotechnol. J. 2014, 9, 282–293


3.1 rxYFP rxYFP was engineered by the group of Winther [18] by introducing pairs of cysteines into the wild-type YFP (wtYFP) in appropriate locations so that they form disulfide bridges in an oxidizing environment (see Fig. 1A). As the cysteine residues are located on β-strands 7 and 10 (amino acids 199–208), the formation of a disulfide bridge induces a conformational change in the YFP, and a change in its fluorescent properties. Only one cysteine pair (N149C/S202C) among the four tested provoked a significant redox-dependent change in fluorescence intensity. Importantly, the disulfide bond was shown to be fully reversible, which allows dynamic recording of the cell dithiol/disulfide ratio. Cysteine 48 of the wtYFP was also mutated to valine to avoid interfering side reactions. The midpoint redox potential E°’ of the cysteine couple was determined as –261 mV at pH 7 [18], which is within the physiological range for redox-active cysteines.


As an example, the E°’ of thioredoxin is –270 mV (see e.g. [19]), and of the GSSG/GSH couple –240 mV [20]. This makes rxYFP a good candidate for redox measurements in reducing compartments, and, in fact, several studies have reported detection of thiol/disulfide variations with rxYFP in different subcellular locations of eukaryotic cells: mitochondrial matrix and intermembrane space [21], cytosol [22], and nucleus and cytosol [23]. Just like wtYFP, rxYFP exhibits two absorption peaks (392 and 512 nm), and an emission peak at 523 nm [18]. However, due to fluorescence quenching, the protonated form of wtYFP and rxYFP (λmax 392 nm) is not fluorescent. Hence, formation of a disulfide bond between the two cysteines in rxYFP results in a 2.2-fold decrease in the signal fluorescence at pH 7 without a significant shift in the excitation wavelength [18]. Spectroscopic studies revealed that the decrease in the rxYFP fluorescence upon oxidation was due to a change in the protonation state of the chromophore

Figure 1. Family trees of the four main families of genetically encoded ROS and redox indicators: (A) roGFP2, (B) rxYFP, (C) Hyper, (D) cpYFP. The mutations leading to the next generation of the protein are indicated on the side of the arrows. The number in superscript next to each sensor refers to the original description of the protein.


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Table 1. The properties of genetically encoded ROS and redox indicators

Probe

Specificity

Disulfide bond

Eo’ V (pH 7, 30°C)

λ ex

λ em

Dynamic range

Cell compartments [ref]

roGFP1

Thiol/disulfide equilibrium

C147-C204

–0.288

405/475

around 505

6.1

Cytosol, mitochondria, endosomes, lysosomes, nucleus [27–29].

roGFP2


C147-C204

–0.272

400/490

around 505

5.8

Cytosol, mitochondria, endosomes, lysosomes, nucleus [27–29].

roGFP1-RX


C147-C204

–263 to –0.284

395/475

around 505

5.4 to 7.5

R12: mitochondria [27]

roGFP1-iX


C147-C204

–0.229 to –0.287

395/465

505

2.4 to 7.2

Endoplasmic reticulum [31, 64–67]

Grx1-roGFP2

2 GSH/GSSG

C147-C204

400/490

around 505

4.4

Cytosol, mitochondria [36]

roGFP2-Orp1

H2O2

C147-C204 roGFP2 C36-C82 Orp1

400/490

around 505

4.8

Cytosol, mitochondria [15]

rxYFP


C149-C202

–0.261

512

523

2.2

Cytosol, nucleus, mitochondrial matrix, intermembrane space [22–24]

rxYFP 3R


C149-C202

ND

512

523

ND

rxYFP-Grx1p

2 GSH/GSSG

C149-C202

–0.267

512

523

2.1

cpYFP

Superoxide

C171-C193

405/490

515

4.2

Mitochondria [46, 55, 58–60]

Hyper

H2O2

OxyR-RD: C199-C208

420/500

516

3-4

Cytosol, mitochondria, peroxisomes, nucleus, ER [39–43]

Hyper 2

H2O2

OxyR-RD: C199-C208

420/500

516

6 to 7

Cytosol [50]

Hyper 3

H2O2

OxyR-RD: C199-C208

420/500

516

6

Cytosol [51]

–0.185

towards the protonated (non fluorescent) state, and to a 1.5-fold decrease in the molar extinction coefficient of the deprotonated (fluorescent) state [18]. Moreover, the apparent pKas of the redox sensor chromophore, measured by fluorescence pH titration, were 6.05 for the reduced sensor and 6.76 for the oxidized sensor. These results confirm that redox detection by rxYFP is pH sensitive. As additional evidence, the authors showed that at pH 6.5, the dynamic range of the probe was 2.9 as compared to 1.75 at pH 7.5. This might as well alter redox detection by the probe in alkaline compartments such as mitochondria. The pH sensitivity of rxYFP is therefore a major pitfall of the probe, and has to be considered while performing experiments, especially as the chromophore pKa renders the probe very sensitive to pH changes under near-neu-

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tral conditions. Other limitations include the following points: the probe is not ratiometric so quantitative measurements are almost impossible; just like wtYFP, rxYFP might be quenched by low concentrations of certain small anions [24]; rxYFP does not exhibit clear specificity toward a specific redox couple. We should note that an improved version of rxYFP has been implemented by the same group [25] through the introduction of three positively charged residues in the proximity of the two cysteines involved in the disulfide bond. This mutation decreases the pKa value of the cysteines, which results in a stabilization of the reactive thiolate form under physiological pH conditions. The new version of the sensor, named rxYFP 3R (for rxYFP200R/ 204R/227R) exhibits a 13-fold increase in reactivity towards glutathione.





All the properties of rxYFP and derivatives are summarized in Table 1.

3.2

roGFP

The roGFP family is probably the most extended family of the GERRIs, as well as the most represented in the literature.

3.2.1 roGFP1 and roGFP2 roGFP1 and roGFP2 were developed by the group of Remington by inserting reactive cysteine residues into the positions S147 and Q204 located on the β-strands 7 and 10 of enhanced GFP (EGFP) [26]. Interestingly, these positions are in close proximity to the ones mutated in rxYFP. In addition, for roGFP1, the S65T mutation encountered in EGFP was reversed (T65S) (see Fig. 1). The authors also tried to insert the disulfide bond at position C149=C202 (roGFP3 and roGFP4), or to introduce four cysteines residues into GFP (roGFP5 and roGFP6) [26]. However, the resulting probes exhibited a lower dynamic range and were rapidly overshined by their siblings. roGFP1 and roGFP2 report changes in the global thiol/disulfide equilibrium rather than in the ratio of a specific redox couple. It is important to note that roGFPs are not ROS sensors as the active cysteine pair of these sensors is protonated at physiological pH. Indeed, only cysteine residues with low pKa that are deprotonated at physiological pH react significantly with ROS. Importantly, as for rxYFP, the oxidation of the cysteine couples of roGFPs is fully reversible, allowing the recording of dynamic redox changes in living cells. The E°’s of the cysteine couples at pH 7 were determined as –288 mV for roGFP1 and –272 mV for roGFP2 [26], which is more negative than the midpoint potential of rxYFP (Table 1). These indicators are thus best suited for use in reducing subcellular environments such as mitochondria or the cytosol. As an example, using roGFPs, the dithiol/disulfide equilibrium within the mitochondria and cytosol was reported at (EpH) 8 ≈–360 mV and (EpH) 7 ≈–320 mV, respectively [26, 27]. roGFP1 and 2 have also been utilized to report redox changes in the nucleus and peroxisomes [27, 28]. Of note, roGFPs targeted to endosomes, lysosomes and the ER were found to be fully oxidized, illustrating the need of redox sensors with less negative midpoint potential [29]. In addition to the midpoint redox potential, another parameter determining the usefulness of a probe is the kinetics. In their 2004 study, Dooley et al. [27] show that redox-mediated changes in roGFP fluorescence can require several minutes. Hence, roGFPs might be better suited for detecting steady-state redox conditions and long-lasting redox changes than for reporting rapid fluctuation in thiol/disulfide equilibrium. The two absorption peaks of the EGFP (400–405 nm for the protonated form and 475–490 nm for the deprotonated form), and the emission peak (505 nm) are maintained


in the two roGFPs variants [26]. Similar to rxYFP, oxidation of the cysteine couple C147=C204 favors the protonation of the chromophore. However, the protonated form of roGFP being fluorescent, this results in an enhancement of the 400–405 nm excitation peak and a decrease of the 475–490 nm peak. Thus, contrary to rxYFP, roGFP sensors are ratiometric in excitation. Oxidation of the chromophore results in an increase in the 400/475 ratio of 6.2 for roGFP1, and in the 400/490 ratio of 5.8 for roGFP2 (see Table 1). It should be noted that, in contrast to roGFP1, roGFP2 fluorescence emission depends on pH as well as redox potential. Fluorescence pH titrations of roGFP2 gave apparent chromophore pKa values of 5.6 for the reduced protein and 6.76 for the oxidized protein. However, pHinduced changes in fluorescence are corrected by the ratio 400/490 nm; in other words, the ratiometric readout of roGFP2 is pH insensitive. Unfortunately, the midpoint potential for roGFP2 was also found to vary linearly with pH in the range of 6.0–8.0, following the equation E°’pH= E°’ – 65.5 mV × (pH-7) [26]. Hence, as with rxYFP, special care should be taken when using the sensor in acidic or alkaline compartments.

3.2.2 roGFP1-RX roGFPX (roGFPR1 through roGFPR14) are a newer version of roGFP1 with higher response rate, created by Cannon and Remington in 2006 [30]. The strategy underlying the development of these probes resembles the one implemented for rxYFP 3R. Indeed, the pKa values of the two reactive cysteine residues of roGFPs are between 8.9 and 9.5 [26]. Thus, at neutral pH, most of the residues are protonated, which results in a low reactivity. Introduction of three positively charged amino acids next to the cysteines can lead to stabilization of the thiolate form and increase in the reactivity. The rate constants approximately double with each additional positively charged substitution. However, the midpoint potentials are not dramatically affected by the mutation. Among the roGFP-RX, roGFP1-R12 (obtained by insertion of two lysines close to the two cysteines plus an arginine in position 223) is the most suitable for use in live cells, due to the significantly increased reaction rate (sixfold higher than the one of roGFP1).

3.2.3 roGFP1-iX roGFP1-iX were developed by Lohman and Remington in 2008 [31] to fulfill the need for redox probes with higher midpoint potential, suitable for oxidizing compartments such as the ER (estimated reduction potential around –180 mV [12]). In these probes, the thermodynamic stability of the disulfide is substantially lowered by insertion of an amino acid into the β-strand 7 of the roGFP1, adjacent to cysteine 147. This insertion, in conjunction with the mutation H148S, caused an increased geometric strain in the disulfide, which resulted in indicators with

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higher reaction kinetics and midpoint potentials. Several amino acid were tested and the highest E°’ values were achieved with the insertion of leucine (–229 mV) and glutamate (–236 mV). To identify members of this family, the notation roGFP1-iX, was adopted, where X denotes the amino acid at insertion position 147. The excitation spectra of oxidized and reduced roGFP1-iX are pH sensitive [31]. However, the 465/395 ratio is essentially constant between pH 6.0 and 8.0. Hence, roGFP-iX should be used in a ratiometric way to avoid pH artifacts.

4 Genetically encoded sensors for 2GSH/GSSG rxYFP-Grx1p and Grx1-roGFP are chimeric sensors based on a redox relay between redox-active enzymes (glutaredoxin) and redox-sensitive proteins (rxYFP and roGFP, respectively). The idea behind this approach is that the redox enzyme will lend specificity and faster kinetics to the redox sensing process.

4.1 rxYFP-Grx1p rxYFP-Grx1P was developed by Bjornberg et al. [32] by fusing the enzyme glutaredoxin-1 from yeast (Grx1p) with the redox probe rxYFP (see Fig. 1). The strategy was based on the following observations: (i) rxYFP contains a dithiol/disulfide pair whose midpoint potential of –261 mV is suitable for monitoring intracellular glutathione redox potential [18]; (ii) purified rxYFP equilibrates very slowly with oxidized/reduced glutathione; and (iii) addition of glutaredoxin significantly accelerates the reaction [33]. As expected, in comparison to isolated rxYFP, the rate of GSSG-mediated oxidation of the chimera was improved by a factor of 3300. In addition, reactivity towards other oxidants such as H2O2, hydroxyethyl disulfide or cystine remained low, giving a high glutathione specificity. Characteristics of the probe are detailed in Table 1.

4.2 Grx1-roGFP2 Like the rxYFPs, conventional roGFPs exhibit undefined specificity and slow response to changes in redox potential [26]. In particular, roGFPs have been shown to equilibrate very slowly with oxidized/reduced glutathione, the rate-limiting factor being the availability of glutaredoxin [34]. As mentioned above, the issue of glutaredoxin availability has been solved for rxYFP by fusing the sensor with Grx1p [32]. The same strategy was adopted by Gutscher et al. [35] with roGFP 2. The N terminus of the sensor was linked to the human Grx1 by a 30-amino acid spacer, (GlyGly-Ser-Gly-Gly)×6, allowing flexible interactions between the two moieties. This fusion facilitates specific and rela-

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tively fast equilibration between roGFP2 and GSH/GSSG. Consequently, the resulting chimera Grx1-roGFP2 detects nanomolar variations in GSSG against a background of millimolar concentration of GSH on a scale of seconds to minutes, whereas the classical roGFP2 is not reactive under these conditions [35]. Moreover, Grx1roGFP2 is sensitive to even small traces of GSSG in commercial samples of GSH. However, the probe is insensitive to H2O2 [35]. Thus, Grx1-roGFP2 appears to be suitable for dynamically measuring the GSH/GSSG redox state in highly reduced compartments with high sensitivity and temporal resolution. It was successfully used in HeLa cell cytosol and mitochondria [35], as well as in Drosophila tissues and whole body [14]. Grx1-roGFP2 exhibited the same fluorescence excitation spectrum as roGFP2, and the excitation ratio of the sensor was also found to be insensitive to pH changes between 5.5 and 8.5 [35]. Full protocol and considerations for measuring the GSH/GSSG redox state using Grx1roGFP2 can be found in Morgan et al. [36]. It should be noted that another chimera resulting from the fusion of roGFP2 and thioredoxin-1 in place of glutaredoxin-1 has been tested. However, this attempt was not successful as Trx1-roGFP2 did not respond to changes in oxidized glutathione concentration [35].

5 Genetically encoded sensors for H2O2 5.1 HyPer HyPer has been developed by the group of Lukyanov [37] using an approach similar to the genetically encoded calcium indicators Pericams: the fluorescent protein, circularly permuted YFP (cpYFP), was inserted into a sensing protein domain (H2O2 sensing domain for HyPer and calmodulin/peptide M13 for Pericam) [38]. As a result, conformational changes in the sensing domain are translated into conformational changes in the fluorescent protein, coupling the process of interest to a change in fluorescence. The redox sensing protein included in HyPer is OxyRRD, the H2O2-sensing regulatory domain (RD) of the transcription factor OxyR found in Escherichia coli. The key residues of OxyR for H2O2 sensing are the reactive cysteines 199 and 208. C199 reacts with H2O2 to form an intramolecular disulfide bridge with C208. Consequently, an important conformational change occurs within the flexible region of the regulatory domain of OxyR, located between the residues 205 and 222. Hence, to create HyPer, a mutated form of the cpYFP (for optimized folding and maturation) was inserted between residues 205 and 206 of OxyR-RD [37]. In addition, the Y203F mutation was introduced in the cpYFP chromophore to allow the visualization of its protonated form (see Fig. 1). Importantly, all





mutants in which serine replaces one or both of the reactive cysteine residues of OxyR-RD showed no change in fluorescence upon exposure to H2O2. HyPer displays a high sensitivity to H2O2 (low-to-middle nanomolar range), faster kinetics than roGFP2-Orp1 (in agreement with the different mechanisms of probe oxidation: direct reaction for HyPer and Orp1-mediated oxidation for roGFP2-Orp1), and has been shown to be fully reversible in eukaryotes [37, 39]. It is likely that the reduction of HyPer in eukaryotes is mediated by the glutathione system, just like the reduction of OxyR in E. coli [39]. Therefore, HyPer could be a probe reporting the balance between 2H2O/H2O2 and 2GSH/GSSG rather than a pure H2O2 sensor. The cysteine pair of the OxyR-RD part of the HyPer has a midpoint potential of -185 mV [37]. Thus, HyPer can be used only in reducing compartments such as cytosol, nucleus, mitochondria and peroxisomes (see e.g. [37, 40]). HyPer was shown to be completely oxidized when expressed in the ER [40], although some studies have reported detection of the production of H2O2 within the ER lumen using Hyper [41, 42]. HyPer has also been used to monitor changes in H2O2 level in vivo in the zebrafish, the Xenopus laevis, and the nematode Caenorhabditis elegans (see e.g. [43–45]). Since the Y203F mutation was introduced into the cpYFP of HyPer, the protonated form of the chromophore is fluorescent. Hence, HyPer displays two excitation peaks at 420 (protonated chromophore) and 500 nm (deprotonated chromophore) and a single emission peak at 516 nm. Oxidation of HyPer by H2O2 results in a shift in the equilibrium between the two protonation states of the chromophore, causing a decrease in the 420-nm excitation peak and a proportional increase in the 500-nm excitation peak [37]. Therefore, the sensor is ratiometric. The maximal ratiometric dynamic range in vitro was reported to be three- to fourfold, but was smaller in living cells [37]. As the fluorescent moiety of HyPer, cpYFP, is very sensitive to the environmental pH [45, 46], it is not surprising that the probe also exhibits a strong pH sensitivity. Acidification of the environment leads to an increase in the 420-nm excitation peak (protonated chromophore) and a decrease in the 500-nm peak (deprotonated chromophore), which can be mistaken for a reduction of the probe [37, 47].Conversely, alkalinization mimics oxidation of HyPer. Hence, pH changes have to be monitored while investigating H2O2 variations using HyPer. The best control is provided by HyPer-C199S, the mutated version insensitive to H2O2 [37], also known as SypHer [47]. How specific is HyPer for H2O2? Contrary to rxYFPs and roGFPs, the redox-sensitive cysteine pair in HyPer is not located on the chromophore but on a redox sensing domain, OxyR-RD. While there is little doubt that OxyRRD is specific for H2O2, recent work suggests that cpYFP is sensitive to O2·– [46]. Although in vitro study of HyPer sensitivity towards various oxidants suggests that HyPer


does not detect O2·– [37], transient increases in fluorescence of mitochondrial targeted HyPer and its mutated version SypHer have been detected in skeletal muscle, which very much resemble the superoxide flashes described previously [46, 48]. Strikingly, these flashes occurred simultaneously with increases in MitoSox fluorescence intensity [49], challenging the lack of sensitivity of HyPer for O2·–. Lukyanov and Belousov [39] proposed an alternative explanation: pH increase would decrease the spontaneous dismutation rate of O2·–. Thus, flashes would embody transient alkalinization of the mitochondria matrix as well as increase in O2·– level. However, a decrease in spontaneous dismutation of O2·– should also affect H2O2 level, but no differences were observed between flashes detected with HyPer or SypHer. Further experiments are needed to assess the possible sensitivity of HyPer to O2·–. Calibration of the probe at alkaline pH (the mitochondrial pH being around 8) could be necessary as redox potentials depend on pH. For instance, midpoint potential of roGFPs has been shown to vary by –65 mV for every pH unit [26].

5.2 HyPer-2 and HyPer-3 Two improved versions of HyPer have been recently developed by mutating the OxyR-RD: HyPer-2 [50] and HyPer-3 [51] (see Fig. 1). HyPer-2 was derived by random mutation (A406V) of HyPer [50]. It has an expanded dynamic range (6–7 compared to 3 in HyPer) and higher brightness upon expression in eukaryotic cells, but slower kinetics in vitro [50] (see Table 1). HyPer-3 was obtained after H34Y mutation of HyPer-2, resulting in a sensor with high dynamic range and faster kinetics [51]. HyPer-3 was successfully used for in vivo imaging of tissue-scale H2O2 gradients in zebrafish larvae.

5.3 roGFP2-Orp1 Recently, another H2O2 sensor has been developed by Dick and colleagues [52], which consist of a fusion protein between a peroxidase, the yeast peroxidase Orp1, and a redox fluorescent probe, roGFP2 (Fig. 1). The yeast peroxidase Orp1 is endogenously part of an oxidation relay with the transcription factor Yap1. In other words, in the yeast, H2O2 oxidizes Orp1, which in turn oxidizes the transcription factor Yap1. Thus, Orp1 is a good candidate for the peroxidase moiety of fusion proteins sensors. The same relay is found in the resulting probe, roGFP2-Orp1, with roGFP2 being the reducing substrate of Orp1. Orp1 catalyzes a near stoichiometric conversion of H2O2 to disulfides in roGFP2. As a result, the roGFP2Orp1 redox relay converts physiological H2O2 signals into measurable fluorescent signals in living cells. roGFP2-Orp1 responds to low micromolar concentrations of H2O2 and exhibits a dynamic range of 4.8 in living cells, with spectral properties similar to roGFP2 (Table 1).

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In addition, the ratiometric readout of roGFP2-Orp1 is also pH stable [52]. roGFP2-Orp1 was successfully used to study age- and condition-related alterations of H2O2 in cytosol and mitochondria of living larval and fixed adult Drosophila melanogaster [14].

6 Genetically encoded sensors for O2·–: The case of cpYFP As discussed above, cpYFPs were previously developed by the group of Miyawaki to be used as the core structure of the genetically encoded calcium indicator Pericam [38]. In cpYFP, the original amino and carboxyl moiety of the fluorescent protein are inverted, and connected with a short linker. The sensitivity of cpYFP to O2·– was discovered serendipitously by the groups of Dirksen, Cheng and Sheu, while trying to measure calcium in mitochondria using mitochondria-targeted ratiometric Pericam. Ratiometric Pericam as well as the isolated cpYFP exhibited transient increases in fluorescence intensity (flashes), which were identified as increases in superoxide level – superoxide flashes [46, 48]. The two cysteine residues involved in O2·– sensing have been identified as C171 and C193, as mutation of both cysteines to either alanine or methionine alter the sensitivity of the probe to aldrithiol, a strong oxidant [46]. However, this result is debated, as the double mutation also largely reduces fluorescence of the entire molecule [17]. Just like the cpYFP used in HyPer, and independently developed by the group of Lukyanov [37], the cpYFP isolated from the ratiometric pericam bears the mutation Y203F, which renders the protonated form of the chromophore fluorescent. Thus, cpYFP exhibits two excitation peaks at 405 nm (protonated chromophore) and 490 nm (deprotonated chromophore), and a single emission peak at 515 nm. cpYFP is therefore ratiometric in excitation. It should be noted that, in terms of redox sensing, the 405-nm excitation wavelength behaves as an isoemissive point: no changes in fluorescence intensity are detected for λex 405 nm, while it increases at λex 490 nm when the chromophore is fully oxidized with aldrithiol [46]. In the same way, fluorescence intensity at λex 405 nm is unchanged during a flash. This point is discussed below. cpYFP specificity towards O2·– was demonstrated in vitro using the following experiments [46]. First, the fluorescence emission of purified cpYFP was found to be five times brighter under strong oxidizing conditions (1 mM aldrithiol) compared to strong reducing conditions (10 mM reduced dithiothreitol). The authors then showed that cpYFP responded to O2·– (generated by 2 mM xanthine and 20 mU xanthine oxidase under aerobic conditions) but not to H2O2 or peroxinitrite. Hydroxyl radicals and NO only caused a slight decrease of fluorescence intensity. Similarly, no significant changes in cpYFP fluo-

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rescence was found after application of Ca2+, ATP, ADP, NAD(P)+, and NAD(P)H. Finally, unlike redox biosensors [18, 26], the fluorescence emission of cpYFP was unaltered by variation of redox potential between -319 and 7.5 mV (produced by different mixes of reduced and oxidized dithiothreitol). However, the specificity of cpYFP towards O2·– is currently a matter of debate. The cpYFP, as a circularly permuted fluorescent protein, is very sensitive to pH. Circular permutation results in disturbances of the β-barrel that may allow easy access of H+ to the chromophore [17]. Furthermore, the pKa of cpYFP is around 8.5 [46], which makes cpYFP very sensitive to minor changes in pH in the alkaline matrix of mitochondria. Since 1) the pharmacology of superoxide flashes is quite unusual compared to what is known about superoxide production in mitochondria, and 2) Schwarzlander et al. [53, 54] reported that cpYFP fluorescence intensity in isolated mitochondria from plants was not affected by genetic and pharmacological manipulation of superoxide level but responded to changes in matrix pH, it has been proposed that superoxide flashes were in fact pH flashes. However, a change in the protonation state of the chromophore should induce an increase in the fluorescent intensity measured with the 488-nm excitation wavelength as well as a decrease with the 405-nm excitation wavelength. Yet, the flashes were detected at 488 nm with no change in the fluorescence intensity for λex= 405 nm. Furthermore, Wang et al. [46] and Wei-Lapierre et al. [55] reported changes in flashes amplitude and frequency upon manipulation of mitochondrial matrix level of O2·– in mammalian cells. Finally, Pouvreau [48] showed that cpYFP flashes were simultaneous with an increase in fluorescence intensity of a redshifted synthetic O2·– sensor, MitoSox. In addition, flashes have also been recently indentified in cardiac cells loaded with MitoSox or 2,7-dichlorodihydrofluorescein diacetate (with no cpYFP expression), ruling out the possibility of an energy transfer between cpYFP and MitoSox [56]. Hence, cpYFP flashes seem to be effectively reporting transient increase in O2·– fluorescence. Another argument in favor of the pH nature of cpYFP flashes is that transient increases in fluorescence resembling the flashes have been detected with mitochondrialtargeted SypHer, a pH probe derived from HyPer [57]. However, SypHer and HyPer are based on cpYFP. An increase in MitoSox fluorescence intensity simultaneously to HyPer of SypHer flashes has been reported [49, 57]. Thus, it is not clear whether cpYFP flashes are reporting pH or superoxide changes. Let’s try to untangle the story. The same pH dependency and spectral properties apply to the four protein sensors that have been used to detect flashes: Ratiometric Pericam [48], cpYFP [46], HyPer [49] and SypHer [49, 57]. This means: deprotonation of the chromophore is reported by an increase in fluorescence intensity at λex= 490 nm and a concomitant decrease at λex= 405 nm. This has





been verified during alkalinization of the mitochondrial matrix from intact cells for cpYFP [55], ratiometric Pericam, HyPer and SypHer (Pouvreau, unpublished results). However, during flashes detected by either cpYFP or ratiometric Pericam, the intensity at λex= 405 nm does not change [46, 48]. In addition, strong oxidation of cpYFP with aldrithiol does not affect the intensity of fluorescence recorded at λex= 405 nm [46]. This result shows that cpYFP flashes are not reporting changes in pH, and also that detection of superoxide does not involve a shift in the protonation state of the chromophore. The mechanism could be an alteration of the molar extinction coefficient as has been observed for rxYFP [18]. In conclusion, although the O2·– nature of cpYFP flashes is still disputed, it appears unlikely that it is reporting changes in pH. However, the final answer to this debate will require unraveling the chemistry of the sensor. Crystallization of the protein in oxidized and reduced state would help. In addition, supplemental experiments using probes with different structures, such as roGFP2-Orp1 for ROS or mito AlpHi (pKa 8.5) for pH, would bring essential information. It should be noted that, although cpYFP has been used in vivo and in numerous different cell types [46, 48, 58–60], to my knowledge no publications refer to detection of O2·– using cpYFP in other cell compartments. It would be interesting to know how the biosensor behaves in nonalkaline cellular compartments.

7 Future directions Compared to the more established genetically encoded calcium indicators, the GERRIs are still in their infancy. This review covers only the most popular, but new indicators based on other strategies or specific for other redox molecules are being developed every year. Examples are förster resonance energy transfer (FRET)-based H2O2 sensors (PerFRET and OxyFRET, [61]), FRET-based redox sensor (Redoxfluor, [62]) or NAD+/NADH sensors (Peredox, [63]). No doubt some of them will become as common as their older siblings over the next few years. Currently existing sensors still have limitations, such as kinetics, pH dependency, and lack of specificity. Furthermore, all the existing sensors are imaged in the green channel. Red-shifted sensors would be very useful to perform dual color imaging, e.g. of redox and calcium signaling, or of redox signaling in two cellular compartments. There is therefore still a long way to go before being able to image most of the redox species with the accuracy that currently exists for calcium ions. But seeing how fast the field moved for genetically encoded calcium sensors, we can predict that it will not take long to cover the distance.


Sandrine Pouvreau is a senior researcher at the French National Centre for Scientific Research (CNRS). She received her PhD in Physiology from the University of Lyon (France) in 2005 for her work on intracellular calcium dynamics. She then joined the Department of Biophysics at Rush University (Chicago, USA) for her postdoctoral research. In 2008 she was assigned as a CNRS researcher in Lyon working on the imaging of ROS production in the mitochondria of living cells. She currently works at the Interdisciplinary Institute for Neurosciences in Bordeaux (France). Her research interests lie in the involvement of mitochondria in synaptic plasticity.

This work was supported by grants from the Centre National de la Recherche Scientifique. The author declares no commercial or financial conflict of interest.

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ISSN 1860-6768 · BJIOAM 9 (2) 171–310 (2014) · Vol. 9 · February 2014

Systems & Synthetic Biology · Nanobiotech · Medicine

2/2014 FRET imaging Synthetic probes Live-cell imaging

Fluorescent Biosensors www.biotechnology-journal.com

The Fluorescent Biosensor special issue of Biotechnology Journal is edited by Dr. May Morris and Prof. Marc Blondel. The cover image is an artistic interpretation of how fluorescent biosensors function as molecular beacons for scientists navigating a sea of molecules. Image courtesy of L. Divita and R. Wintergerst.

Biotechnology Journal – list of articles published in the February 2014 issue. Editorial: Fluorescent biosensors May C. Morris and Marc Blondel http://dx.doi.org/10.1002/biot.201400008 Review Newly engineered cyan fluorescent proteins with enhanced performances for live cell FRET imaging

Review Fluorescent biosensors for high throughput screening of protein kinase inhibitors Camille Prével, Morgan Pellerano, Thi Nhu Ngoc Van and May C. Morris

http://dx.doi.org/10.1002/biot.201300196

Fabienne Mérola, Asma Fredj, Dahdjim-Benoît Betolngar, Cornelia Ziegler, Marie Erard and Hélène Pasquier

Review FRET-based and other fluorescent proteinase probes


Hai-Yu Hu, Stefanie Gehrig, Gregor Reither, Devaraj Subramanian, Marcus A. Mall, Oliver Plettenburg and Carsten Schultz

Review Decoding spatial and temporal features of neuronal cAMP/PKA signaling with FRET biosensors Liliana R. V. Castro, Elvire Guiot, Marina Polito, Danièle Paupardin-Tritsch and Pierre Vincent

http://dx.doi.org/10.1002/biot.201300202 Review Imaging early signaling events in T lymphocytes with fluorescent biosensors Clotilde Randriamampita and Annemarie C. Lellouch

http://dx.doi.org/10.1002/biot.201300195 Review Deciphering the spatio-temporal regulation of entry and progression through mitosis Lilia Gheghiani and Olivier Gavet

http://dx.doi.org/10.1002/biot.201300194 Review Shining light on cell death processes – a novel biosensor for necroptosis, a newly described cell death program François Sipieter, Maria Ladik, Peter Vandenabeele and Franck Riquet

http://dx.doi.org/10.1002/biot.201300201 Review Genetically encoded reactive oxygen species (ROS) and redox indicators Sandrine Pouvreau

http://dx.doi.org/10.1002/biot.201300199 Technical Report Time-resolved microfluorimetry: An alternative method for free radical and metabolic rate detection in microalgae Amadine Bijoux and Anne-Cécile Ribou

http://dx.doi.org/10.1002/biot.201300217 Research Article Acri-2,7-Py, a bright red-emitting DNA probe identified through screening of a distyryl dye library Delphine Naud-Martin, Xavier Martin-Benlloch, Florent Poyer, Florence Mahuteau-Betzer and Marie-Paule Teulade-Fichou


http://dx.doi.org/10.1002/biot.201300200 Review Advances in lanthanide-based luminescent peptide probes for monitoring the activity of kinase and phosphatase Elena Pazos and M. Eugenio Vázquez




Mitochondrial reactive oxygen species (ROS) and ROS-induced ROS release.

Reactive oxygen species and redox compartmentalization.

Redox imaging using genetically encoded redox indicators in zebrafish and mice.

Reactive oxygen species in redox cancer therapy.

Genetically encoded fluorescent redox probes.

Genetically Encoded Voltage Indicators: Opportunities and Challenges.

Substituent effects on reactive oxygen species (ROS) generation by hydroquinones.

Genetically encoded indicators of neuronal activity.

Reactive Oxygen Species (ROS): Beneficial Companions of Plants' Developmental Processes.

Genetically encoded Ca2+ indicators in cardiac myocytes.

Genetically Encoded Voltage Indicators in Circulation Research.

Redox Roles of Reactive Oxygen Species in Cardiovascular Diseases.

Tuning of Redox Regulatory Mechanisms, Reactive Oxygen Species and Redox Homeostasis under Salinity Stress.

The evolving capabilities of rhodopsin-based genetically encoded voltage indicators.

Photoactivatable genetically encoded calcium indicators for targeted neuronal imaging.

Genetically Encoded Fluorescent Indicators for Organellar Calcium Imaging.

Calcium imaging with genetically encoded indicators in behaving primates.

Generation of transgenic marmosets expressing genetically encoded calcium indicators.

Recent progress in the development of genetically encoded Ca2+ indicators.

Optogenetic Monitoring of Synaptic Activity with Genetically Encoded Voltage Indicators.

Mitohormesis: Promoting Health and Lifespan by Increased Levels of Reactive Oxygen Species (ROS).

Decitabine induces delayed reactive oxygen species (ROS) accumulation in leukemia cells and induces the expression of ROS generating enzymes.

Detection and quantification of reactive oxygen species (ROS) in indoor air.

Estrogen potentiates reactive oxygen species (ROS) tolerance to initiate carcinogenesis and promote cancer malignant transformation.