Genetically encoded fluorescent sensors are becoming extremely powerful tools for functional imaging. For instance, the classic calcium sensor introduced by Roger Tsien's lab, cameleon, is now extensively used in the plant field (Miyawaki et al., 1997; Romoser et al., 1997). Such calcium sensors have allowed major breakthroughs regarding, for instance, the molecular dialog between plant roots and symbiotic microorganisms (Sieberer et al., 2012; Singh and Parniske, 2012). More recently, dedicated strategies have led to the development of very promising biosensors, which allow to probe the local concentration of disaccharides, phosphate, amino acid, or ammonium (reviewed in Gjetting et al., 2013; Jones et al., 2013). In this perspective issue, we will focus on a specific type of sensors optimized for pH measurement.

## **PROPERTIES OF GENETICALLY ENCODED pH SENSORS**

Classical fluorescent dyes have been used to estimate pH in plants. For example, fluorescein coupled to dextran allows pH recording in the close vicinity of roots (Monshausen et al., 2009, 2011) or shoots (Geilfus and Mühling, 2012). The 2- ,7- -Bis(2 carboxyethyl)-5(6)-carboxyfluorescein (BCECF), which was first introduced by Rink et al. (1982), is also classically used to measure plant vacuolar pH, especially in its membrane permeable form as acetomethyl ester (BCECF-AM; Krebs et al., 2010). Nevertheless, such approaches are invasive and do not allow measuring pH in other intracellular cell structures. Genetically encoded pH sensors have been designed to overcome these limitations.

The chromophore of green fluorescent protein (GFP) is formed by two autocatalytic reactions (cyclization and oxidation) between Ser65, Tyr66, and Gly67 (review in Wachter, 2007). It absorbs light at two maxima: 395 nm when protonated and 475 nm when depronated (Bizzarri et al., 2009). This shift in absorption has been extensively used to record pH in living organisms (Kneen et al., 1998; Llopis et al., 1998), and optimized versions of GFP for pH measurement have been generated. Ecliptic and superecliptic pHluorins display almost no fluorescence when protonated, allowing sensitive detection of biological processes associated with pH increase (**Table 1**; Miesenböck et al., 1998; Sankaranarayanan and Ryan, 2001). However, the equilibrium between the protonated and deprotonated states of these proteins is also affected by temperature and ionic strength. Furthermore, the light emitted by such sensor protein is dependent on its own concentration. Thus a variation in fluorescence signal can result either from a change in pH or in concentration of the sensor. To circumvent this issue, ratiometric pH sensors have been developed providing a pH readout independent of the probe concentration. Among such sensors, one of the most popular is pHluorin. It was generated by introducing point mutations in *Aequorea victoria* GFP (AvGFP) sequence, especially S202H, which conferred ratiometric behavior to the sensor (Miesenböck et al., 1998; Bizzarri et al., 2009). For example, pHluorin displays a strong increase in absorption at 475 nm

**Table 1 | Selected genetically encoded pH sensors available for** *in vivo* **pH measurements.**

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*\*Eightfold brighter version of pHluorin (Mahon, 2011)*

*\*\*pHluorin with optimize codon usage for Aspergillus (Bagar et al., 2009)*

*\*\*\*pHluorin with optimize codon usage for Arabidopsis (Shen et al., 2013)*

*\*\*\*\*pHGFP, chimera between smGFP (Davis and Vierstra, 1998) and pHluorin to prevent splicing (Moseyko and Feldman, 2001)*

and, simultaneously, a strong decrease in absorption at 395 nm when the pH is shifted from 7.5 to 5.5. The pHluorin sensor has a pKa of 6.9 and the dynamic range of fluorescent signal is from pH 5.4 to 8.4 (**Table 1**). This pH range is suitable for most plant subcellular compartments, with the exception of the apoplast, which is too acidic, and the vacuole, where GFP is degraded (Tamura et al., 2003). Later, derivatives of this protein were engineered to improve the level of expression. This is the case for ratiometric Venus Citrine (RaVC), whose codon usage has been optimized for expression in *Aspergillus* (Bagar et al., 2009), and the plant-solubility-modified ratiometric pHluorin (*PRpHluorin*), optimized for *Arabidopsis* (Shen et al., 2013). To avoid erroneous pH measurement due to chromatic aberrations, an alternative type of ratiometric sensor was generated from S65T GFP, where the emission is yellow-shifted upon alkalization (**Table 1**). This dual emission GFP (deGFP) contains the point mutations T203C and H148G or H148C (Hanson et al., 2002). In addition, some sensors such as E2GFP and E1GFP combine dual emission and dual excitation features (Bizzarri et al., 2006; Arosio et al., 2007).

The major limitation of using pHluorin and its derivates, especially in plant tissues, is their acidic quenching at an apparent pH near 4.5. Two strategies were used to improve pH sensing in acidic environments. The first strategy is to use a fluorescent protein (FP), *Pt-*GFP, from orange sea pen, *Ptilosarcus gurneyi* (Schulte et al., 2006). Although *Pt*-GFP has a pKa similar to that of pHluorin (pKPtGFP: 7.3), this protein has a broader pH response range and is more stable in acidic conditions (Schulte et al., 2006). The second strategy is to make a tandem of FPs with highly different ranges of pH responsiveness. The measure of pH is then either given by Förster resonance energy transfer (FRET) between these two proteins, as for pHlameleon (Esposito et al., 2008) or by the ratio of fluorescence of the proteins. In particular, the tandem of enhanced GFP (eGFP) and monomeric red FP (mRFP1) is suitable for measuring pH in the apoplasm (Gjetting et al., 2012). Finally, a new pH biosensor was developed recently based on bioluminescence resonance energy transfer (BRET). This pHlash sensor is composed of a luciferase as a donor, fused to pH-sensitive Venus as acceptor (Zhang et al., 2012). Use of bioluminescence significantly improves the signal-to-noise ratio making BRET particularly useful for experimentation at the whole tissue or organism scale.

Regardless of the exact type of genetically encoded pH sensors, estimation of the local pH value from the fluorescence data is not straightforward. This requires calibration to characterize the relationship between the fluorescence properties of the sensor and pH. The calibration can be carried out either *in vitro* or *in vivo* (Gao et al., 2004; Schulte et al., 2006; Shen et al., 2013). *In vitro* calibration uses heterologously expressed sensors in buffers of different pH. Such a calibration does not take into account the ionic strength and buffering capacity of the cell and thus can lead to substantial artifacts (Schulte et al., 2006). *In vivo* calibration uses sensor-expressing plant cells that are treated with an ionophore, such as nigericin, in presence of a solution with sufficient buffering capacity to clamp the pH of the cellular environment containing the sensor. This approach should be favored whenever possible, but is still limited for several reasons. First, there is no objective way to ensure that the equilibrium has reached between the bathing media and the cell compartment where the sensor is located. This is especially true when the sensor is in the lumen of intracellular structures, meaning that the ionophore has to be effective on two successive membranes. Secondly, the ionophore modifies the ion content of the cell and consequently the native buffering capacity and ionic strength. Such calibration issues may explain the ongoing debate on the feasibility of measuring absolute pH values with sensors *in vivo* (Schulte et al., 2006; Bizzarri et al., 2009).

## **ANALYSIS OF EXOCYTOSIS EVENTS WITH pH SENSORS**

Genetically encoded pH sensors can be used to investigate processes that give rise to, or rely on, changes in pH. During exocytosis, vesicle lumen undergoes a rapid alkalinization upon fusion to the plasma membrane. Since the pioneering study by Miesenböck et al. (1998), ecliptic pHluorin has been extensively used as an exocytosis indicator (Sankaranarayanan et al., 2000; Gandhi and Stevens, 2003; Taylor et al., 2011). For instance, in the synaptic cleft, Gandhi and Stevens (2003) were able to distinguish three types of vesicle behaviors at the plasma membrane. In another study, ecliptic pHluorin used to describe the dependency of vesicle fusion on actin filaments and cdc42, a protein controlling cell division (Alberts et al., 2006). Similar approaches would appear useful in the plant field. According to the classical view of ligand–receptor trafficking, a difference in pH is expected between the lumen of the *trans*-Golgi network (TGN) and the apoplastic face of the plasma membrane (Robinson et al., 2012). Hence, the luminal pH of exocytic vesicles is predicted to be different before and after fusion with the plasma membrane. In recent reports, TGN compartments have been shown to have the most acidic lumen of the secretory pathway, with a pH around 6 (Martinière et al., 2013; Shen et al., 2013; **Table 2**). On the other side, apoplastic pH is likely to be highly variable and dependent on tissue type and development stage (Gao et al., 2004; Staal et al., 2011; Geilfus and Mühling, 2012; Gjetting et al., 2012). Unfortunately, studies that concomitantly measure the pH in the TGN and in the apoplasm are still lacking.

## **pH WITHIN THE APOPLAST**

In plants, the plasma membrane is energized by P-type proton pump ATPases. Direct evidence has been obtained that proton diffusion within the apoplast away from the membrane can be ratelimiting when compared to proton excretion by the proton pumps. Using weak acid influx for assaying pH, it has been shown that the local pH in the unstirred aqueous layers next to the plasmalemma can be much lower than that of the external bathing solution, by up to 2 pH units under conditions that maximize the rate of net proton excretion (Sentenac and Grignon, 1987; Grignon and Sentenac, 1991). A few studies on the dependency of membrane transport in roots on external pH have quantitatively assessed this steady state pH shift at the membrane surface due to proton excretion by using weak acid influx assays under the same conditions as those used for the transport measurements (Thibaud et al., 1998). Clearly, the results indicated that this shift in pH strongly contributed to energization of the cell membrane and had to be taken into account to explain, for instance, kinetic and thermodynamic features of nitrate, phosphate, or potassium transport (Thibaud et al., 1988), or the effect of HCO− 3 , a buffer that can be naturally present in the external solution (in calcareous soils), on the transport rate of these nutriments (Toulon et al., 1989) or on iron reduction at the cell surface (Toulon et al., 1992).

Different methods have been developed to estimate local pH values, using proton-selective micro-electrodes (Felle, 1998) or impermeable pH-sensitive dyes (Taylor et al., 1996; Monshausen et al., 2009, 2011), or collecting apoplasmic fluids (Mühling and Sattelmacher, 1995). However, such approaches provide estimates of the pH conditions at the organ surface or, at best, of the mean pH within the apoplast, and not of the local pH in close vicinity of the membrane, which is the actual pH sensed by the ion transporters. Another source of difficulties is the possibility of heterogeneities within the apoplast. For example, in the root cortex and for the inner cortical cells, the diffusion barrier that

*The average pH is indicated for each subcellular compartment the pH sensor used for the measurement being indicated between parenthesis. ST, sialyltransferase; ManI, mannosidase 1; AtVSR2, At2g14720; trunc, truncated AtVSR2 with no luminal domain; BCECF-AM, 2* - *,7* - *-Bis(2-carboxyethyl)-5(6)-carboxyfluorescein; tetrakis(acetoxymethyl) ester mito, transit peptide of the* β*-subunit of NpF1-ATP synthase; NLS, nuclear localization signal.*

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slows proton migration toward the external solution comprises the entire apoplastic continuum, from cortical cells surrounding the endodermis to the root epidermis surface. Furthermore, longitudinal heterogeneities in surface pH have been observed between the root apex, the elongation region and the fully differentiated zone, due to differences in the rate and direction of proton transport. The physiological consequences and roles of such radial and longitudinal differences, in terms of membrane energization and transport activity, have been poorly investigated. Clearly, genetically encoded pH sensors open novel perspectives in this domain of the root structure–function relationship. In this respect, Moseyko and Feldman (2001) and Gao et al. (2004) have developed pioneering approaches. For example, large pH variations were observed in response to salt stress. When challenged with 100 mM NaCl, root cells displayed a more alkaline apoplasm and a more acidic cytosol. These changes in pH might result from specific activation of H+:Na+ antiport activity mediated by the plasma membrane Salt-Overly-Sensitive1 (SOS1) antiporter in response to salinity constraint (Quintero et al., 2011). Such analyses clearly indicate that expression of pH sensor proteins in mutant plants could allow rapid progress in investigating the root structure–function relationship and its role in membrane energization, ion transport and plant adaptation to environmental stresses.

## **CELL WALL pH AND GROWTH**

Cell capacity to control the apoplastic pH plays a major role also in plant growth and development. Indeed, according to the acid growth theory, cell wall "loosening" favored by low pH allows cell elongation (Rayle and Cleland, 1992). In a typical sequence of events, auxin stimulates proton secretion by plasmalemma proton pump ATPases, resulting in cell wall acidification, causing activation of expansins, which loosen cellulose microfibrils. As a result, cell walls become more extensible favoring turgor-driven cell elongation. In support to the acid growth theory, the elongation zone of the root generally displays a more acidic apoplasm than the fully differentiated zone (Staal et al., 2011). Intriguingly, a close-up analysis using a pH sensor reveals highly dynamic fluctuations of root surface pH with alkalinization and acidification periods and alkalinization bursts along the root longitudinal axis or alternating between opposing flanks (Monshausen et al., 2011). Such results indicate that a net influx of protons can occur in epidermal cells, as previously reported in other studies (O'Neill and Scott, 1983). Intriguingly, a recent report shows that treatment of root tips with indole-3-acetic acid (IAA) induces an increase in pH by about 0.5–0.8 pH units (Gjetting et al., 2012), which might be related to the periodical alkalinization previously reported in root tips (Monshausen et al., 2011).

#### **pH MAPPING OF ORGANELLES AND PROTEIN TARGETING**

Taking advantage of growing knowledge in protein targeting mechanisms, appropriate signal sequences can be fused to the coding sequence of the pH sensor so that this is targeted later to various cell membranes or compartments. Using such an approach, pH in various intracellular compartments was measured *in vivo* in mammalian and, more recently, in *Arabidopsis* protoplasts (Shen et al., 2013), tobacco leaf epidermal cells or *Arabidopsis* root tip cells (Martinière et al., 2013). These studies directly demonstrate a gradual acidification of 1.5–2 pH units in the lumen of the endomembrane system with ER displaying the most alkaline pH and vacuole the most acidic one (see **Table 2**). Interestingly, some pH differences exist for a given compartment between plant species (cytosol, endoplasmic reticulum and vacuole; **Table 2**). These variations could be the result of biological variation between the systems tested (intact cells versus protoplast) or could be due to differences in growing condition. For instance, pH homeostasis is known to be sensitive to light intensity (Martinière et al., 2013). Most compartment markers cycle at least between two subcellular localizations and there is still no way to target a sensor solely to either the lumen of the TGN, the prevacuolar compartment (PVC) or the late PVC. Fusion of pH sensors to the vacuolar sorting receptor (VSR) mutants Y165A and IMAA allowed to partially solve such localization issue, the former mutant being preferentially retained in the TGN and the latter localized in the LPVC. This allowed differentiation between TGN and the PVC and getting pH estimates for these two compartments (Martinière et al., 2013; Shen et al., 2013). However, it should be noted that, in addition to TGN, the Y165A VSR mutant is very likely to also label recycling endosomes and plasma membranes (daSilva et al., 2006; Saint-Jean et al., 2010). Furthermore, accumulation of the IMAA mutant in the LPVC is only transient (Saint-Jean et al., 2010).

In average, the population of VSR labeled compartments has a pH intermediate between the *trans-*Golgi and the vacuole in all the systems tested. But surprisingly, the TGN is actually more acidic, with a pH 6, than the PVC (pH 6.8) or even the late PVC (pH 7.1, see **Table 2**; Martinière et al., 2013). This result fits well with the localization of V-type ATPase involved in pumping protons. Indeed, VHA-a1 protein is preferentially accumulated in the TGN (Dettmer et al., 2006) and is not detected in the multivesicular bodies (MVB; Robinson et al., 2012) or PVC. Accordingly to the proton leak model, while V-type ATPase acidifies a compartment, antiporters could be involved in proton extrusion (Demaurex, 2002). For instance, NHX5 and NHX6 Na+/H+ antiports have been proposed to be involved in such a mechanism in plant endosomes (Bassil et al., 2011). Therefore, the relative amount of VHA-a1 and proton antiporters such as NHX5/6, in compartment on the way to the vacuole could have a preponderant role in proton homeostasis and substantial consequence in terms of protein sorting. In support to this hypothesis, is the mistargeting of a soluble vacuolar marker in *nhx5 nhx6* double knockout (Bassil et al., 2011) and the fact that the pH measured in the TGN and the PVC fits well with the distribution of VHA-a1 and NHX5/6 in tobacco leaf epidermal cells (Martinière et al., 2013).

The fact that the PVC is more alkaline than TGN is surprising and in contradiction with the VSR-mediated vacuolar sorting model where acidification in the PVC is thought to be responsible for a release of the ligand by the receptor (Kirsch et al., 1994; Ahmed et al., 2000; Shimada et al., 2003). In fact, the *in vitro* binding curve between VSR and the ligand is bell shaped with an optimum pH at 6 (Kirsch et al., 1994). This value corresponds to the pH measured *in vivo* in the TGN (Martinière et al., 2013)

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suggesting that binding of VSR to its ligand would be optimal in the TGN but significantly lower in more acidic and alkaline compartments. The use of ratiometric sensors to measure the endomembrane luminal pH therefore suggests that alkalinization rather than acidification would trigger ligand release from the receptor VSR in the PVC (Martinière et al., 2013). Of course this model does not take into account other environmental parameters in TGN and PVC such as the calcium concentration that is also known to regulate ligand binding for a least some members of the VSR family (Watanabe et al., 2002, 2004). Finally, it is also important to point out that the pH reported for the PVC or the TGN is an average from a large population of independent organelles with some variation of pH values. This difference is either due to technical limitations in imaging or may reflect pH variability between compartments of same identity, e.g., TGN or PVC. A model for VSR-mediated transport is still a matter of debate (Robinson and Pimpl, 2013).

It is likely that pH ratiometric sensors will open new perspectives in our understanding of the mobility, functional heterogeneity and remodeling capacities of organelles. For instance, the TGN has been assumed to undergo maturation as its morphology changes over time (Scheuring et al., 2011). Using pH sensors along with dynamic studies could provide evidence for these maturation events.

## **CONCLUDING REMARKS**

Genetically encoded pH sensors have opened avenues for cell biology. They will provide valuable functional data with respect to cell compartmentation, protein trafficking, acidic growth, or ion diffusion within plant tissue. One of the future challenges will be to improve the definition of pH information provided by the sensor as cell nano-environments are anticipated. For instance, protein and lipid composition of membranes have a substantial effect on proton diffusion. As well, it is reasonable to anticipate a higher proton concentration in the vicinity of ATPase pumps and therefore heterogeneity on membrane surfaces with respect to pH.

## **REFERENCES**

emission pH indicator for intracellular studies. *Biophys. J.* 90, 3300–3314. doi: 10.1529/biophysj.105.074708

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 30 August 2013; accepted: 04 December 2013; published online: 18 December 2013.*

*Citation: Martinière A, Desbrosses G, Sentenac H and Paris N (2013) Development and properties of genetically encoded pH sensors in plants. Front. Plant Sci. 4:523. doi: 10.3389/fpls.2013.00523*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2013 Martinière, Desbrosses, Sentenac and Paris. This is an openaccess article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

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## Development of roGFP2-derived redox probes for measurement of the glutathione redox potential in the cytosol of severely glutathione-deficient *rml1* seedlings

#### *Isabel Aller 1, Nicolas Rouhier 2 and Andreas J. Meyer 1 \**

*1 INRES-Chemical Signalling, University of Bonn, Bonn, Germany*

*2 Interactions Arbres Microorganismes, IFR 110 EFABA, Faculté des sciences, Université de Lorraine, UMR 1136 Université de Lorraine/INRA, Vandoeuvre, lès-Nancy, France*

#### *Edited by:*

*Alex Costa, University of Milan, Italy*

*Reviewed by: Mirko Zaffagnini, University of Bologna, Italy Frank E. Schleifenbaum, University of Tuebingen, Germany Taras P. Pasternak, Albert-Ludwigs-University of Freiburg, Germany*

#### *\*Correspondence:*

*Andreas J. Meyer, INRES-Chemical Signalling, University of Bonn, Friedrich-Ebert-Allee 144, 53113 Bonn, Germany e-mail<EMAIL_ADDRESS>Glutathione is important for detoxification, as a cofactor in biochemical reactions and as a thiol-redox buffer. The cytosolic glutathione buffer is normally highly reduced with glutathione redox potentials (*EGSH*) of more negative than −310 mV. Maintenance of such negative redox potential is achieved through continuous reduction of glutathione disulfide by glutathione reductase (GR). Deviations from steady state glutathione redox homeostasis have been discussed as a possible mean to alter the activity of redox-sensitive proteins through switching of critical thiol residues. To better understand such signaling mechanisms it is essential to be able to measure *EGSH* over a wide range from highly negative redox potentials down to potentials found in mutants that show already severe phenotypes. With the advent of redox-sensitive GFPs (roGFPs), understanding the *in vivo* dynamics of the thiol-based redox buffer system became within reach. The original roGFP versions, roGFP1 and roGFP2, however, have midpoint potentials between −280 and −290 mV rendering them fully oxidized in the ER and almost fully reduced in the cytosol, plastids, mitochondria, and peroxisomes. To extend the range of suitable probes we have engineered a roGFP2 derivative, roGFP2-iL, with a midpoint potential of about −238 mV. This value is within the range of redox potentials reported for homologous roGFP1-iX probes, albeit with different excitation properties. To allow rapid and specific equilibration with the glutathione pool, fusion constructs with human glutaredoxin 1 (GRX1) were generated and characterized *in vitro*. GRX1-roGFP2-iL proved to be suitable for *in vivo* redox potential measurements and extends the range of *EGSH* values that can be measured *in vivo* with roGFP2-based probes from about −320 mV for GRX1-roGFP2 down to about −210 mV for GRX1-roGFP2-iL. Using both probes in the cytosol of severely glutathione-deficient *rml1* seedlings revealed an *EGSH* of about −260 mV in this mutant.

#### **Keywords: glutathione, glutathione redox potential, GRX1-roGFP2,** *rml1***, redox imaging**

## **INTRODUCTION**

Thiol redox biochemistry is considered to play a fundamental role in cellular processes including signaling and cell fate decisions. The ability to dynamically and quantitatively measure such cellular processes *in vivo* is key to understand the underlying principles and the coordination of redox processes in the context of intact cells. Redox-sensitive GFP (roGFP) allows a direct read-out of the glutathione redox potential (*EGSH*) particularly in reducing compartments (Meyer and Dick, 2010). However, current variants of roGFPs are largely inadequate in mutants with very low glutathione levels, generally oxidizing conditions like in the ER, or oxidizing conditions triggered by pathological processes. These limitations demand the development of further roGFP variants with less negative midpoint potentials.

The tripeptide glutathione (γ-L-glutamyl-L-cysteinylglycine) constitutes the major low molecular weight thiol in most prokaryotic and virtually all eukaryotic organisms. Glutathione is synthesized in two sequential steps that are catalyzed by two enzymes, glutamate-cysteine ligase (GSH1) and glutathione synthase (GSH2). In plants, glutathione fulfills a broad range of essential functions including detoxification of heavy metals and xenobiotics and serving as an electron donating cofactor in biochemical reactions (Cobbett and Goldsbrough, 2002; Noctor et al., 2011). Moreover glutathione constitutes one of the most important redox buffer systems in the cell. The capacity to act as a redox buffer relies on the reversible convertibility of glutathione between the reduced form of glutathione (GSH) and the oxidized form glutathione disulfide (GSSG). While the glutathione pool in the cytosol, mitochondria, plastids, and peroxisomes is maintained in a highly reduced state by NADPH and glutathione reductase (GR) (Schwarzländer et al., 2008; Marty et al., 2009), the glutathione redox buffer in the endoplasmic reticulum (ER) is highly oxidized (Hwang et al., 1992; Brach et al., 2009). *EGSH* depends on the absolute glutathione concentration and the ratio of [GSH]:[GSSG] (Meyer and Hell, 2005). The local *EGSH* is assumed to affect the glutathionylation status of proteins which is an important means of regulating protein activity (Michelet et al., 2005; Noctor et al., 2011; Zaffagnini et al., 2012).

Conditions of environmental challenge are frequently considered to cause the generation of oxidants in the form of reactive oxygen species (ROS) (Apel and Hirt, 2004; Torres et al., 2006). Apart from potential toxic effects of ROS, the oxidants are increasingly recognized as vital messengers in cellular signaling (Finkel, 2011). Oxidant-dependent signaling may occur either directly by specific recognition of oxidants (Forman et al., 2010) or indirectly through detoxification of oxidants and resulting changes in the cellular redox buffer system providing an opportunity for readout of these changes. Stress-dependent alterations of *EGSH* caused by a transient draw of electrons from the glutathione redox buffer for oxidant detoxification and coupling of *EGSH* to target proteins thus may result in altered protein function and hence pronounced metabolic and developmental consequences for individual cells and the whole organism.

Depletion of total GSH also directly impacts on *EGSH* and thus mutants with defects in GSH biosynthesis may cause constitutive activation or inactivation of respective signaling pathways. Indeed, Arabidopsis *rax1* mutants carrying a mutation in the GSH1 enzyme leading to only 20–50% of wild-type GSH are affected in stress signaling and have induced defense pathways (Ball et al., 2004). Another partially GSH-deficient mutant, *pad2*, is not capable of activating appropriate defense mechanisms against biotic stressors (Parisy et al., 2007; Schlaeppi et al., 2008). The most severe, yet viable, mutant affected in GSH1 is *rml1*, which contains only 2–10% of wild-type glutathione (Vernoux et al., 2000; Cairns et al., 2006). Homozygous *rml1* mutant seedlings show impaired root development, which is assumed to be caused by redox-dependent inhibition of cell cycle components (Vernoux et al., 2000).

Better understanding of redox-regulatory processes mediated by glutathione demands experimental approaches enabling quantitative monitoring of *EGSH*. The development of roGFPs enables ratiometric thiol redox imaging at subcellular level (Dooley et al., 2004; Hanson et al., 2004). The two excitation maxima of the parental GFP arise from the protonation state of the chromophore and excited state proton transfer that converts the neutral form of the chromophore into the green emitting anionic form (Brejc et al., 1997; Palm et al., 1997). Engineering of two surface-exposed cysteines into the GFP barrel on the two adjacent β-strands 7 and 10 in positions allowing reversible disulfide formation exploits a structure-dependent shift in the protonation status of the chromophore for ratiometric measurements (Dooley et al., 2004; Hanson et al., 2004). The degree of chromophore protonation is also dependent on the type of chromophore: while roGFP1 derived from the wild-type chromophore with the amino acid S65 in the chromophore displays a dominant excitation peak in the UV range, the roGFP2 variant derived from EGFP containing the S65T mutation has a main excitation peak at 490 nm (Hanson et al., 2004). The chromophore type also slightly affects the redox potential of the engineered disulfide with roGFP1 being slightly more negative than roGFP2. It has been shown that glutaredoxin (GRX) catalyzes the oxidation and reduction of roGFP in the presence of glutathione (Meyer et al., 2007; Gutscher et al., 2008). Fusion of GRX to roGFP overcomes kinetic limitations of the GRX/roGFP interaction and possible limitations caused by the absence of appropriate GRXs in some organisms or subcellular compartments (Gutscher et al., 2008; Meyer and Dick, 2010). In addition to kinetic properties, the thermodynamic properties of roGFP probes are similarly important for dynamic measurements and ideally redox sensors with redox potentials adapted to the desired measuring range should be selected. It has been shown that the redox potentials of roGFP variants, roGFP3 and roGFP4, which both contain the engineered disulfide C149/C202, are more negative than the redox potentials of their respective counterparts roGFP1 and roGFP2, which both contain the disulfide C147/C204 (Hanson et al., 2004). The redox potential can also be shifted to less negative values by engineering basic amino acids next to the disulfide-forming cysteines. As a consequence, such probes show an increased pI and hence a significantly enhanced response rate for reductive processes (Cannon and Remington, 2006). In the oxidizing ER lumen and under highly severe stress conditions, however, these probes would still be fully oxidized. To overcome this limitation, Lohman and Remington (2008) further engineered roGFP1 by replacing the chromophore-interacting H148 by serine and inserting an additional amino acid between the disulfide forming C147 and S148. Depending on the amino acid inserted behind C147 the respective members of the socalled roGFP1-iX family had less negative midpoint potentials between −229 and −246 mV (Lohman and Remington, 2008). However, even these less reducing probes appear almost fully oxidized in the ER at steady state of HeLa cells (Birk et al., 2013).

While under highly reducing conditions roGFP1 may have some advantages, roGFP2 has advantages in less reducing conditions as they occur in partially glutathione-deficient mutants (Meyer et al., 2007; Schwarzländer et al., 2008). Specifically, roGFP2 has a larger dynamic range than roGFP1 and it avoids potential illumination-dependent photoisomerization artifacts associated with roGFP1 (Schwarzländer et al., 2008). Because roGFP2 also is less reducing than roGFP1 we asked whether roGFP2-iX probes can be engineered and whether such probes would have an even less negative redox potential than their respective roGFP1-iX counterparts to enable redox imaging in compartments with a less negative *EGSH*. Inhibition of GSH biosynthesis in wild-type roots by BSO leads to severe oxidation in the cytosol (Meyer et al., 2007). Similarly, *EGSH* in *rml1* roots has been shown to be far less negative than in wild-type plants (Meyer and Dick, 2010), but the exact value of cytosolic *EGSH* in this mutant has not been determined because the redox potential is close to the edge or even beyond the usable measuring range of roGFP2. After detailed characterization the novel roGFP2-iL probe was thus used to determine *EGSH* in the cytosol of *rml1* seedlings in order to define the lower limit of glutathione redox potentials under which viability can be maintained.

## **MATERIALS AND METHODS**

## **GENE CONSTRUCTION, PROTEIN EXPRESSION AND PURIFICATION**

roGFP2 (C48S/S65T/Q80R/F99S/S147C/Q204C, (Dooley et al., 2004; Hanson et al., 2004) was used as a template to generate roGFP2-iL (C48S/S65T/Q80R/F99S/S147CL/H148S/Q204C) in which a leucine was added after C147. For site-specific insertion of C147CL and substitution of H148S into the roGFP2 sequence the primers 5- -AACTACAACTGC CTGAGCAACGTCTATATCATGGCC-3 and 5- -GCTCAGGCA GTTGTAGTTGTACTCCAGCTTGTG-3 were used. N-terminal fusion of human GRX1 and roGFP2-iL was done by PCR using gene-specific primers 5- -TCAGGAGGAGTGAGCAAGGGCGA-3 and 5- -TCGCCCTTGCTCACTCCTCCTGA-3- . Amplification of full-length product was done with the primer pair 5- -AC CATGATGGCTCAAGAGTTTGTGAA-3 and 5- -TCTAGACT TGTACAGCTCGTCCATG-3 generating GRX1-TS(GGSGG)6 roGFP2-iL (GRX1-roGFP2-iL). Both PCR products were cloned into pCAP*s* vector (Roche, www.roche-applied-science.com) for sequence confirmation.

The sequence of free roGFP2-iL and GRX1-roGFP2-iL were cloned into the BamHI and NcoI restriction sites of the protein expression vector pQE-30 (Qiagen, www.qiagen.com) for the production of a recombinant protein containing an N-terminal His6-tag. The restriction sites were introduced by PCR using the primer pairs 5- -TGGATCCGCTCAAGAGTTTGTGAACTG-3- and 5- -CTAAGCTTTTACTTGTACAGCTCGTCC-3 for GRX1 roGFP2iL and 5- -AAGGATCCGTGAGCAAGGGCGAGGAGC-3- and 5- -CTAAGCTTTTACTTGTACAGCTCGTCC-3 for roGFP2 iL. Recombinant roGFP1 was produced as described earlier (Schwarzländer et al., 2008). pQE-30 plasmids for expression of roGFP1-iL and roGFP1-iE were kindly provided by Dr. J. Remington (Univ. Oregon) and Dr. C. Appenzeller-Herzog (Univ. Basel), respectively.

For stable expression in plants, a modified version of pBinAR vector (Höfgen and Willmitzer, 1990) containing UBQ10 promoter instead of 35S promoter was used. The coding sequence of GRX1-roGFP2iL was amplified by 5- -AG GTACCATGGCTCAAGAGTTTGTGAAC-3 and 5- -TATGTCG ACTTACTTGTACAGCTCGTCCAT-3 to add KpnI and SalI restriction sites used for cloning.

## **ISOLATION OF RECOMBINANT PROTEINS**

After transformation of the *E. coli* strain Origami (DE3) (Novagen, www.merckmillipore.de), a pre-culture of 10 ml was grown over night at 37◦C. Five mL of the pre-culture were added to 450 ml LB medium and grown at room temperature to an OD600 of 0.5–0.8. Expression of the different roGFP variants was induced by addition of isopropyl-β-D-thiogalactopyranoside (IPTG) to a final concentration of 1 mM. Protein expression was performed at room temperature for 24 h. The cells were harvested and resuspended in protein extraction buffer (50 mM Tris-HCl pH 8.0, 250 mM NaCl). The cells were sonicated for 10 min. Cell lysate was centrifuged and the soluble roGFP proteins purified via a Ni2+ loaded HiTrap™ Chelating HP Column (GE Healthcare, www.gelifesciences.com).

## **SPECTROSCOPY**

Fluorescence excitation spectra were collected using a LS55 fluorescence spectrophotometer (Perkin Elmer Life Sciences, http:// www.perkinelmer.com). Five hundred microliter protein solution was placed in a 1 ml quartz cuvette. Samples contained 0.2μM protein in reaction buffer (100 mM K2HPO4/KH2PO4 pH 7.4, 1 mM EDTA) and 10 mM total dithiothreitol (DTT) for full reduction or 10 mM total H2O2 for full oxidation of the sensor, respectively. The spectra were collected from 350 to 520 nm with a bandwidth of 10 nm and a scan speed of 500 nm min−1. Fluorescence was detected at 540 nm.

The fluorescence quantum yield (QY) of roGFPs was determined by comparison to Rhodamine 6G (Sigma-Aldrich, www. sigmaaldrich.com) which has a QY of 0.9 when dissolved in water (Magde et al., 2002). Six different dilutions of the respective proteins with absorbances between 0.01 and 0.1 were prepared in aqueous solution. Both, standard and roGFP samples were excited at 488 nm with a bandwidth of 5 nm. Total emission was collected from 505 to 590 nm. Fluorescence quantum yields for roGFPs were calculated from the integrated fluorescence intensities of the spectra after correction for wavelength-dependent photomultiplier sensitivity.

## *IN VITRO* **CHARACTERIZATION OF roGFP VARIANTS**

*In vitro* characterization of roGFP2-iL and GRX1-roGFP2-iL fusions by ratiometric time-course measurements with isolated proteins was performed on a fluorescence plate reader (POLARstar Omega; BMG Labtech, www.bmglabtech.com) with filter-based excitation at 390 and 480 nm and detection of emitted light at 520 nm. Phosphate buffer (100 mM K2HPO4/KH2PO4, 1 mM EDTA, pH 7.4) with 1μM of the respective roGFP according to the information given in the text were pipetted into the wells of a 96-well plate with a clear bottom (NUNC™ 96, www.thermoscientific.com). Reduced glutathione (in 100 mM K2HPO4/KH2PO4 buffer, pH 7.0) was automatically injected to the indicated final concentration using the built-in injectors. For maximum achievable reduction of the glutathione buffer, 0.1 μM recombinant *Arabidopsis thaliana* GR1 (*At*GR1) and 100μM NADPH were added to each well. To allow comparable reduction kinetics, the proteins were pre-oxidized with 10 mM H2O2 for 30 min. The remaining H2O2 was removed by desalting spin columns according to the manufacturer's manual (Zeba™ Spin Desalting Columns, www.thermoscientific.com). H2O2 and DTT to a final concentration of 10 mM were separately used to define maximum oxidation and maximum reduction of the sensors. In the comparative reduction assays with free roGFP1, roGFP1-iL, roGFP2, and roGFP2iL, purified *At*GRX2 to a final concentration of 2μM was included in the reaction mix.

## **DETERMINATION OF MIDPOINT POTENTIALS OF roGFP VARIANTS**

Both roGFP2-iL and GRX-roGFP2-iL (1μM final concentration) were allowed to equilibrate (2–3 h) with lipoic acid buffers [reduced form dihydrolipoic acid (DHLA); oxidized form lipoic acid (LA)] (Equation 1). DHLA/LA was used at a total concentration of 2.5 mM in degassed HEPES buffer (100 mM HEPES, 300 mM NaCl, 1 mM EDTA, pH 7.0). The appropriate concentrations of DHLA and LA to set distinct redox potentials were calculated from the Nernst equation based on the standard reduction potential of lipoic acid (*E*◦- *LA*) of −290 mV at pH 7.0 (Lees and Whitesides, 1993).

$$\text{roGFP}\_{\text{ox}} + \text{DHLA} \Leftrightarrow \text{roGFP}\_{\text{red}} + \text{LA} \tag{1}$$

The redox potential for each of the two redox pairs is defined by the Nernst equation using the respective midpoint potential and ratios (*Q*) of oxidized to reduced roGFP and LA to DHLA, respectively (Equation 2).

$$E = E^{\circ \prime} - \frac{RT}{nF} \ln Q \tag{2}$$

In this equation, *R* is the gas constant (8.314 J K−1 mol−1), *T* is the temperature (298.15 K), *n* is the number of transferred electrons, and *F* is Faraday's constant (96,485 C mol−1). After redox equilibration of both redox couples it is possible to equate the two redox potentials (Equation 3):

$$E\_{LA} = E\_{LA}^{\circ\prime} - \frac{0.0592V}{2} \log \frac{[\text{DHLA}]}{[\text{LA}]} = E\_{roGFP}^{\circ\prime}$$

$$-\frac{0.0592V}{2} \log \frac{[\text{roGFP}\_{\text{red}}]}{[\text{roGFP}\_{\text{ox}}]} = E\_{roGFP} \tag{3}$$

Because *E*◦- LA is known this leaves two unknowns, *E*◦- roGFP and [roGFPred]/[roGFPox]. The redox potential of roGFP is dependent on the degree of oxidation (*OxDroGFP*) of the redox pair roGFPred/roGFPoxaccording to Equations 4 and 5:

$$\text{OxD}\_{\text{roGFP}} = \frac{\text{[roGFP}\_{\text{ox}}\text{]}}{\text{[roGFP}\_{\text{ox}}\text{]} + \text{[roGFP}\_{\text{red}}\text{]}} \tag{4}$$

$$\frac{[\text{roGFP}\_{\text{red}}]}{[\text{roGFP}\_{\text{ox}}]} = \frac{1 - \text{OxD}\_{\text{roGFP}}}{\text{OxD}\_{\text{roGFP}}} \tag{5}$$

Substitution of Equation 5 into Equation 3 allows transformation of Equation 3 into Equation 6:

$$E\_{LA} = E\_{LA}^{\circ \prime} - \frac{0.0592V}{2} \log \frac{[\text{DHLA}]}{[\text{LA}]} = E\_{roGFP}^{\circ \prime}$$

$$-\frac{0.0592V}{2} \log \frac{1 - \text{OxD}\_{roGFP}}{\text{OxD}\_{roGFP}} = E\_{roGFP} \tag{6}$$

*OxDroGFP* can be measured *in vitro* by monitoring the fluorescence emission intensities (*I*) at 520 nm for excitation at 390 and 480 nm. After equilibration in an environment of a defined redox potential, the respective values were used to determine *OxDroGFP* according to Equation 7:

$$\text{OxD}\_{roGFP} = \frac{R - R\_{red}}{\left(\frac{I480\_{\text{ox}}}{I480\_{\text{red}}}\right)(R\_{\text{ox}} - R) + (R - R\_{red})} \tag{7}$$

Here, *R* denotes the ratio of the fluorescence intensities measured at 390 and 480 nm. *Rred* and *Rox* represent the fluorescence ratios of fully reduced and fully oxidized roGFP, respectively. The raw values of *I* were always corrected by subtracting the respective blank values.

For determination of the midpoint potentials *E*◦ of roGFP variants *OxDroGFP* or, alternatively, the degree of reduction *ReDroGFP* = 1 − *OxDroGFP* was plotted against the calculated redox potentials of the respective lipoic acid redox buffers and all data points were fitted to a sigmoidal dose-response curve using GraphPadPrism5 (GraphPad Software, www.graphpad. com). The titration of each protein was carried out three times with 4 technical replicates. Titration of GRX1-roGFP2 and roGFP2 was done with DTT, which has a more negative reduction potential than lipoic acid (*E*◦- DTT = −323 mV, Shaked et al., 1980).

#### **PLANT MATERIAL AND GROWTH CONDITIONS**

Heterozygous Arabidopsis *rml1* mutants were selected by genotyping for the mutant allele and exploiting the fact that the point mutation (Vernoux et al., 2000) generates a new ApoI restriction site. After transformation of heterozygous plants with the respective roGFP constructs transformed plants were first screened for uniform fluorescence on a stereomicroscope equipped with fluorescence optics and a GFP filter. In a subsequent molecular screen heterozygous *rml1* seedlings were selected for further propagation. Seeds from transgenic plants expressing either GRX1-roGFP2 of GRX1-roGFP2-iL were surface sterilized with 70% ethanol twice and resuspended in sterile deionized water. Seeds were plated on nutrient medium [5 mM KNO3, 2.5 mM KH2PO4, 2 mM MgSO4, 2 mM Ca(NO3)2, 10μM Fe-EDTA, 0.1% (v/v) micronutrient mix (Somerville and Ogren, 1982), pH 5.8 solidified with 0.8% phytagel]. Plants were kept at 4◦C for 1 day before placing them in vertical orientation in a growth chamber with a diurnal cycle of 16 h light at 22◦C and 8 h dark at 18◦C. The light intensity was 75μmol photons m−2 s −1. Plants were grown for 3 days until the characteristic dwarf phenotype of *rml1* became visible.

#### **CLSM IMAGING**

Pre-selected *rml1* seedlings were mounted on a slide in a drop of water and immediately transferred to a Zeiss confocal microscope LSM780 (Carl Zeiss Microscopy, www.zeiss.de/mikro). Images were collected with a 40× lens (Zeiss Objective C-Apochromat 40×/1.2 W Corr M27) in multi-track mode with line switching between 488 nm excitation and 405 nm excitation and taking an average of four readings in case of GRX1-roGFP2 and two readings for GRX1-roGFP2-iL, respectively. The roGFP fluorescence was collected with a 505–530 nm emission band-pass filter. Autofluorescence excited at 405 nm was collected from 430 to 480 nm and values were used to subtract autofluorescence bleeding into the roGFP channel as described previously (Schwarzländer et al., 2008; Samalova et al., 2013).

#### **RATIOMETRIC IMAGE ANALYSIS**

Images were imported into a custom written MatLab (The MathWorks, www.mathworks.de) analysis suite (M.D. Fricker, Dept. Plant Sciences, Oxford). The ratio analysis was performed on a pixel-by-pixel basis as *I*405/*I*488 following spatial averaging in (*x*,*y*) using a 3 × 3 kernel. Correction of *I*405 for autofluorescence bleeding into the 405 nm channel and subtraction of background signals for each channel was performed. The average background signal was typically measured from the vacuole of one of the cells. For pseudocolor display, the ratio was coded on a spectral color scale ranging from blue (fully reduced) to red (fully oxidized), with limits set by the *in situ* calibration. The calibration was done by incubation of *rml1* seedlings expressing GRX1 roGFP2 or GRX1-roGFP2-iL in 10 mM DTT and 25 mM H2O2, respectively, to drive the roGFPs to their fully reduced and fully oxidized forms. *OxD* of GRX1-roGFP2 and GRX1-roGFP2-iL expressed in the cytosol of *rml1* mutants was calculated according to Equation 7 albeit with wavelengths 405 nm and 488 nm used for excitation of roGFPs *in vivo*.

## **RESULTS**

## **DEVELOPMENT AND CHARACTERIZATION OF roGFP2-iL**

To investigate whether roGFP2 can be modified to generate less reducing probe variants, the roGFP2 sequence was further engineered by introducing site-specific mutations thus generating roGFP2-iL (**Figures 1A,B**). Subsequently, roGFP2-iL was fused behind human GRX1 to generate GRX1-roGFP2-iL (**Figure 1C**) to ensure specificity of the novel probe for *EGSH* similar to other roGFP variants used before (Gutscher et al., 2008; Meyer and Dick, 2010; Albrecht et al., 2013; Birk et al., 2013). IPTGinduction of *E. coli* cultures transformed with roGFP2-iL and GRX1-roGFP2-iL already resulted in bright green cultures suggesting that the introduced mutations did not abolish the fluorescence (not shown). Comparison of roGFP2-iL and roGFP1-iE with the standard Rhodamine 6G resulted in a QY of 0.6 for roGFP2-iL and a QY of 0.4 for roGFP1-iE. This initial observation was further confirmed through side-by-side fluorescence scans of recombinant roGFP2 and roGFP2-iL in buffers containing either 10 mM DTT for full reduction or 10 mM H2O2 for full oxidation of the probes, respectively (**Figure 2**). As previously described (Dooley et al., 2004; Hanson et al., 2004), formation of the disulfide upon oxidation of roGFP2 favors the protonated, neutral form of the chromophore causing an increase in fluorescence at 395 nm while the 488 nm peak is decreased (red curve). Conversely, full reduction of roGFP2 leads to a decrease in excitation at 395 nm while the peak at 488 nm is increased (blue curve; **Figure 2A**).

Modification of roGFP2 through the C147CL insertion and H148S exchange does alter the general spectral properties but does not abolish the dual excitation behavior in roGFP2-iL (**Figure 2B**). Full oxidation of the sensor causes an increase of fluorescence at 395 nm and a corresponding decrease in 488 nm excitation while full reduction of roGFP2-iL leads to decreased fluorescence excitation at 395 nm and an increase at 488 nm. A clear isosbestic point at 425 nm separates the two peaks further indicating two equilibrating molecular species (**Figure 2B**). Thus, all spectral properties of roGFP2 are fully retained in the newly-generated roGFP2-iL. In contrast to roGFP2, however, the maximum change in fluorescence intensity between fully oxidized and fully reduced sensor states was significantly lower in roGFP2-iL. The dynamic range (δ) for the maximum change of the fluorescence ratio between the fully oxidized and fully reduced state of the chromophore was calculated from the fluorescence excitation at the wavelengths used for ratiometric imaging (i.e., 405 and 488 nm). While roGFP2 has a δ405/488 of 8.18 the δ405/488 for roGFP2-iL was found to be 2.55 (**Figure 2**; **Table 1**). Further equilibration of roGFP2-iL in redox buffers with varying pH revealed that the fluorescence ratio of roGFP2-iL is pH-insensitive in the physiological range (**Figure 3**).

close proximity so that they can facilitate disulfide formation. **(B)** The introduction of a leucine next to Cys147 (Leu147\*) and the substitution of His148 by serine increase the relative distance between Cys147 and Cys204 that causes alterations in the thermodynamic stability of the disulfide. **(C)** Schematic representation of the GRX1-roGFP2-iL fusion protein. GRX1 was fused to the N-terminus of roGFP2-iL by a TS(GGSGG)6 linker.

The standard redox potential (*E*◦-) strongly depends on the thermodynamic stability between C147 and C204 that is influenced by the nature of the surrounding amino acid residues. It has been described for roGFP1-iL that the insertion C147CL and the substitution H148S lower the thermodynamic stability of the inter-strand disulfide C147–C204 resulting in a less negative *E*◦ of the sensor (Lohman and Remington, 2008). To test whether this effect on thermodynamic stability is retained in roGFP2-iL, *E*◦- *roGFP*2-*iL* was determined through titration with LA/DHLA. Plotting the degree of reduction of the respective sensor variants against the redox potential of the ambient LA/DHLA buffer allowed to deduce *E*◦ values of −237.7 ± 2 mV for roGFP2-iL (**Figure 4A**) and 243.2 ± 5 mV for GRX1-roGFP2-iL (**Figure 4B**). The difference in *E*◦of roGFP2-iL and GRX1-roGFP2-iL suggests that

excitation ratios for fully reduced and fully oxidized probes. Both excitation wavelengths are indicated by vertical lines.

N-terminal fusion of GRX1 influences the thermodynamic properties of the C147–C204 disulfide. To test this hypothesis, the midpoint potential of roGFP2 and GRX1-roGFP2 was determined side-by-side by titration against DTT redox buffers. As depicted in **Figure 5**, free roGFP2 shows an *E*◦ of −277.5 ± 1 mV (**Figure 5A**) while *E*◦ of GRX1-roGFP2 is shifted to −290.2 ± 3 mV (**Figure 5B**).

To test whether the reduction kinetics of roGFP2-iL is affected by GRX, roGFP2-iL was fused to the C-terminus of *Hs*GRX1 and the catalytic efficiency for the reduction of the resulting fusion protein by GSH was compared to free roGFP2-iL in *in vitro* experiments (**Figure 6**). In the absence of GRX, pre-oxidized roGFP2 iL responds only slowly to injection of 2 mM GSH while the GRX1-roGFP2-iL fusion protein shows a dramatically increased reduction rate.

To further confirm the less negative *E*◦ of roGFP2-iL, the sensor was used side-by-side with other roGFP variants in reduction experiments with GSH at physiological concentrations as reducing agent. Since not all compared roGFPs were available as fusion proteins with GRX1, recombinant free GRX was added to the reaction mix. Pre-oxidized roGFP1, roGFP2, roGFP1 iL and roGFP2-iL were all reduced in the presence of GSH which was injected into the roGFP solutions to a final concentration of 5 mM. Subsequently, the reduction kinetics of the sensors were followed over time (**Figure 7A**). Addition of GSH resulted in complete reduction of roGFP1-iL and roGFP2-iL within 3 min while the reduction of roGFP2 was clearly delayed by several minutes and reached only 95% within 21 min. The reduction of the most negative probe roGFP1 was even slower and reached only 81% within 21 min. The rapid change in the degree of reduction of sensors with significantly less negative *E*◦ values than *E*◦ values of roGFP2 and roGFP1 (**Table 1**) can be further illustrated by the first derivative *dReD/dt* of the degree of reduction. *dRed/dt* of the four different probes plotted against time after injection of GSH shows the most rapid change in the degree of reduction for roGFP1-iL and roGFP2 iL while the values for roGFP2 and roGFP1 are much lower (**Figure 7B**).

*aDynamic range determined with the excitation wavelengths given in parenthesis. 405 nm and 488 nm are the excitation wavelengths used for confocal imaging. bHanson et al., 2004.*

*cDooley et al., 2004.*

*d Lohman and Remington, 2008.*

*eSchwarzländer et al., 2008.*

*f Dihydrolipoic acid (DHLA, reduced form), Lipoic acid (LA, oxidized form).*

**FIGURE 3 | pH-dependence of roGFP2-iL fluorescence.** Recombinant roGFP2-iL was incubated in potassium-phosphate buffer at different pH values. The changes in roGFP2-iL fluorescence intensity of the 390 nm (black) and 480 nm channel (white) in buffers of different pH values under full oxidation with 10 mM H2O2 **(A)** and full reduction by 10 mM DTT **(B)** is shown. Increasing pH values beyond pH 7.0 equally increase fluorescence intensity in both channels thus the 390/480 nm ratio remains unaffected. Conversely, decreasing pH below pH 7.0 has the opposite effect for fully oxidized and fully reduced roGFP2-iL. Similar pH-dependent effects have been reported for wild-type GFP (Patterson et al., 1997). **(C)** The excitation ratios (390/480 nm) in 10 mM H2O2 (-) and in 10 mM DTT () of roGFP2-iL are plotted against the respective pH solution. For several data points, standard error is smaller than data marker (*n* = 3 technical replicates).

## **MEASUREMENT OF THE CYTOSOLIC** *EGSH* **IN SEVERELY GLUTATHIONE-DEFICIENT** *rml1* **MUTANTS**

Conventional roGFP1 and roGFP2 are highly reduced when expressed in the cytosol of non-stressed cells (Meyer et al., 2007; Schwarzländer et al., 2008). Therefore, roGFP-iX probes with a less negative midpoint potential would obviously be expected to

be also fully reduced, particularly when fused GRX1 ensures specific equilibration with the local *EGSH*. Severe stress situations and pathological conditions, however, can lead to pronounced oxidation in the cytosol and thus probes with a less negative midpoint potential may be advantageous for redox imaging in the cytosol to fully resolve the dynamic changes in *EGSH*. Severe depletion of GSH in the Arabidopsis mutant *rml1* has been reported to result in growth inhibition and it has been speculated that this effect may be caused by severe oxidation of cytosolic *EGSH* and redox-dependent inhibition of the cell cycle (Vernoux et al., 2000). Indeed, roGFP2 has been shown to be largely oxidized in the cytosol of *rml1* seedlings (Meyer and Dick, 2010). To further investigate the effect of severe GSH depletion on the cytosolic *EGSH*, both probes, GRX1-roGFP2 and GRX1-roGFP2 iL were expressed in the cytosol of *rml1* and used to measure *EGSH* in root epidermal cells. As expected, the conventional GRX1-roGFP2 was largely oxidized (**Figures 8A,B**). Incubation of *rml1* seedlings with 25 mM H2O2 resulted only in minor additional increase in the fluorescence ratio of GRX1-roGFP2 while incubation of seedlings in 10 mM DTT caused a pronounced drop in fluorescence ratio. Expression of GRX1-roGFP2-iL in the cytosol of *rml1* mutants, on the other hand, resulted in a largely reduced probe (**Figures 8A,C**). Full reduction of the probe through incubation of seedlings with 10 mM DTT caused only a small drop in the detected fluorescence ratio. In contrast, incubation with 25 mM H2O2 resulted in a pronounced increase in the 405/488 nm fluorescence ratio. The dynamic range of the

of 2 mM reduced glutathione. The reaction buffer contained 100μM NADPH and glutathione reductase at pH 7.4 to maintain full reduction of glutathione over the measured time (*n* = 4).

**FIGURE 7 | GSH-dependent reduction of roGFP variants.** The kinetics for the reduction of roGFP by reduced glutathione strongly depends on redox potential of the disulfide forming cysteines of the respective roGFP sensors. **(A)** The excitation ratio 390/480 nm with emission at 520 nm was followed over time. Fully reduced glutathione (final concentration 5 mM) was added to a solution of pre-oxidized roGFP1-iL (), roGFP2-iL (♦), roGFP2 (), and roGFP1 (◦) 3.75 min after start of the measurement. The reaction buffer contained 2μM GRX, 100μM NADPH, and glutathione reductase at pH 7.4 to maintain maximum reduction of glutathione over the measured time course (*n* = 4). **(B)** *dRed/dt* further illustrates the rapid change in reduction of roGFP1-iL () and roGFP2-iL (♦) compared to roGFP2 () and roGFP1 (◦).

probe in the cytosol calculated from the respective *min* and *max* ratios was about 3-fold (**Figure 8C**), which is similar to the predictions from the spectral analysis of recombinant roGFP2-iL (**Figure 2**).

Calculation of the degree of oxidation (*OxD*) of both probes from the ratiometric imaging data presented in **Figure 8** showed a low *OxD* of only 14% for GRX1-roGFP2-iL and a corresponding high *OxD* of 93% for GRX1-roGFP2 (**Figure 9A**). The measured *OxD* values for both probes can be compared to titration curves for GRX1-roGFP2 (blue) and GRX1-roGFP2-iL (red) that were calculated from the Nernst-Equation using *E*◦ values determined earlier (**Figure 9B**). The interception points between the titration curves and the *OxD* for GRX1-roGFP2 and GRX1 roGFP2-iL, respectively, suggest an *EGSH* in the cytosol of *rml1* of about −260 mV.

**FIGURE 8 | The GRX1-roGFP2-iL probe expressed in the cytosol of homozygous** *rml1* **mutants is almost completely reduced while GRX1-roGFP2 is highly oxidized. (A)** Ratiometric images **(a–f)** show the redox state of GRX1-roGFP2 and GRX-roGFP2-iL in the cytosol of root epidermal cells of *rml1* mutants. Treatment with 10 mM DTT results in full reduction of GRX1-roGFP2 **(a)** and GRX-roGFP2-iL **(d)**, while under resting conditions GRX1-roGFP2 is highly oxidized **(b)** whereas GRX1-roGFP2-iL is in a reduced state **(e)**. Treatment with 25 mM H2O2results in complete oxidation of GRX1-roGFP2 **(c)** and GRX-roGFP2-iL **(f)**. Scale bar 10μm. (**B,C**) Fluorescence ratio of cytosolic GRX1-roGFP2 **(B)** and GRX1-roGFP2-iL **(C)** show complete reduction and oxidation of GRX-roGFP2 and GRX1-roGFP2-iL after treatment with 10 mM DTT and 25 mM H2O2, respectively. Under resting conditions in H2O, GRX1-roGFP2 is highly oxidized while GRX1-roGFP2-iL is highly reduced. Values are calculated as the mean ± SD (*n* = 5 − 10).

**FIGURE 9 | Deduction of the redox potential of the roGFP probes from the Degree of Oxidation (***OxD***) in the cytosol of** *rml1* **seedlings. (A)** GRX1-roGFP2-iL and GRX1-roGFP2 expressed in the cytosol of *rml1* plants as calculated according to equations given in Materials and Methods from the mean fluorescence data presented in **Figure 7**. While GRX1-roGFP2 is highly oxidized only a small fraction of GRX1-roGFP2-iL is in the oxidized state. **(B)** Titration curves drawn for GRX1-roGFP2 (blue) and GRX1-roGFP2-iL (red) were calculated from the Nernst-Equation using the midpoint potentials determined in **Figures 3**, **4**. Dotted horizontal lines refer to *OxD* of GRX1-roGFP2 and GRX1-roGFP2-iL from *rml1* measurements shown in panel **(A)**. The vertical dotted line indicates the interception points for GRX1-roGFP2 (blue) and GRX-roGFP2-iL (red) which suggests an *EGSH* of about −260 mV in the cytosol of *rml1*.

## **DISCUSSION**

The glutathione redox potential in life cells varies depending on the compartment and on environmental conditions imposing stress situations for the plant. While cytosol, peroxisomes, mitochondria, and plastids in Arabidopsis wild-type plants maintain a highly reduced glutathione buffer with redox potentials of less than −310 mV, the ER is far less reducing with *EGSH* values of less than −240 mV (Meyer et al., 2007; Schwarzländer et al., 2008). In nominally reducing compartments, severe stress can trigger physiological conditions frequently described as oxidative stress. Better understanding of the underlying molecular processes leading to gradual oxidation and also the downstream processes involved in signal transduction cascades enabling the plant to adapt to a stress situation requires the ability to measure individual components of the cellular redox buffer system in a dynamic way. The roGFPs, and particularly GRX1-roGFP2 fusion proteins, have been shown to sense the local *EGSH* with high specificity due to selective mediation of thiol/disulfide exchange reaction between glutathione and roGFP by GRXs (Meyer et al., 2007; Gutscher et al., 2008). The originally introduced variants roGFP1 and roGFP2, however, are particularly suitable only in the reducing compartments of wild-type plants and mutants with limited effects on the amount of glutathione (Meyer et al., 2007; Maughan et al., 2010), or mutants affected in the GSH/GSSG redox equilibrium due to lack of GR (Marty et al., 2009). In more oxidizing compartments or more severe mutants affected in glutathione redox homeostasis, *EGSH* is outside the measuring range of these probes thus limiting their usability. The need for less reducing probe variants has been partially fulfilled through the introduction of roGFP1-iX variants derived from roGFP1 (Lohman and Remington, 2008). However, even the least reducing variant roGFP1-iL is still largely oxidized in the ER of human cells and thus hardly capable of detecting oxidative processes in the ER. In addition, the most oxidizing roGFP1-iX variant roGFP1-iL has been reported as being relatively dim compared to the parent roGFP1 (Avezov et al., 2013; Birk et al., 2013). Similar to the continuous extension of the range of differentially colored fluorescent proteins (Shaner et al., 2005) there is also a demand for redox reporter proteins with midpoint potentials matching the *EGSH* in distinct compartments or in mutants with targeted alterations in glutathione homeostasis.

The insertion of amino acids in the GFP backbone may lead to changes in the spectroscopic properties of GFP but still yield functional fluorescent proteins (Topell et al., 1999; Cannon and Remington, 2006; Lohman and Remington, 2008). Insertion of single amino acid residues within the roGFP backbone, adjacent to the reactive cysteine 147, leads to a destabilization of the disulfide formed between Cys147 and Cys204. As a consequence, the resulting roGFP1-iX variants show a less negative midpoint potential compared to the parental roGFP1. However, the substitution of the chromophore phenol(ate) interacting His148 by Ser148 was required to maintain ratiometric behavior (Lohman and Remington, 2008). A roGFP1-iL carrying the mutations S147CL and H148S has been reported to have the least negative midpoint potential (−229 ± 5 mV) of a whole series or roGFP1 iX variants (Lohman and Remington, 2008). Introduction of the same changes into roGFP2 lead to roGFP2-iL. This new variant maintained the general ratiometric behavior of roGFP2 with the excitation peak for the anionic form of the chromophore at 488 nm being more pronounced than the UV excitation peak for the protonated chromophore. However, the dynamic range δ for the maximum change in peak excitation ratio was significantly decreased compared to roGFP2 (see **Figure 2** and **Table 1**). A similar reduction of the dynamic range due to amino acid insertion next to Cys147 has already been reported for several roGFP1-iX variants but not for the roGFP1-iL version (Lohman and Remington, 2008). Similarly to roGFP1-iL, roGFP2-iL is also lower in fluorescence than the respective parent roGFP. Nevertheless, the maintenance of ratiometric properties and a sufficiently large dynamic range to resolve biologically relevant redox changes suggest that roGFP2-iL is generally suitable for *in vivo* measurements.

The roGFP1-iL and roGFP2-iL only differ in the S65T substitution introducing an additional methyl group in roGFP2. This minor change not only affects the protonation of the chromophore leading to pronounced stabilization of the anionic form of the chromophore (Elsliger et al., 1999; Jung et al., 2005) but also shifts the midpoint potential of roGFP2 −10 mV less negative compared to roGFP1. The shift in *E*◦ from −280 mV for roGFP2 to −238 mV for roGFP2-iL is lower than the shift for roGFP1-iL which at −229 mV is 62 mV less negative than its parent roGFP1. Different from the original expectation, the modification of roGFP2 did thus not result in a linear additive effect in which the resulting roGFP2-iL was expected to be slightly more negative than roGFP1-iL. The deviation from linearity is probably due to geometric constraints in the roGFP barrel similar to those described for roGFP1-iR which had been expected to be less reducing than roGFP1-iL due to stabilization of the thiolate anion by the introduced adjacent basic arginine residue (Lohman and Remington, 2008). The roGFP1-iR, however, turned out to be more reducing than roGFP1-iL. Whether introduction of basic amino acids in the vicinity of the redox active cysteines may cause a stabilization of thiolates and thus render the redox potential of roGFP2-iL less negative is not known at this stage.

For live cell measurements biosensors should ideally exhibit a strong preference for one specific analyte. By definition, roGFPs undergo thiol/disulfide exchange reactions and it has been shown that specificity for glutathione is achieved through specific interaction with GRXs (Meyer et al., 2007). Fusion of GRX1 to roGFP2 leads to a permanent increase of the local GRX concentration around roGFP and hence this kinetic coupling further increases the likelihood of interaction between roGFP and the fused GRX1 compared to other non-specific interactions with other oxidoreductases (Gutscher et al., 2008; Albrecht et al., 2013). Human GRX1 does also interact with roGFP2-iL and the fusion protein GRX1-roGFP2-iL responds much faster to changes in *EGSH* than free roGFP2-iL. Interestingly, fusion of GRX1 to the N-termini of different roGFP variants consistently leads to a slight shift of 5–13 mV in the midpoint potentials toward more negative values. A similar shift of 6 mV has also been reported for GRX1-roGFP1 iE (Birk et al., 2013). Consistent with the respective order of redox potentials, Birk et al. (2013) also observed a higher degree of oxidation of GRX1-roGFP1-iE than for free roGFP1-iE expressed in the ER. A possible explanation for the effects of N-terminally fused GRX on the *E*◦ of roGFP probes might be a mechanical strain that acts on the GFP barrel slightly affecting the thermodynamic stability of the disulfide. Despite the slightly less negative midpoint potential of roGFP1-iL compared with roGFP2-iL, the kinetic properties of both probes for reduction by GSH are very similar. This suggests that both probes may be equally suitable in compartments with oxidizing conditions. RoGFP2-iL thus extends the range of suitable probes for oxidizing compartments by offering additional spectral features. The dominant excitation peak at 488 nm may be of particular advantage for lifetime imaging of roGFPs with pulsed blue excitation. This imaging approach has been successfully used for the original roGFP2 probe and different roGFP1-iX variants (Wierer et al., 2012; Avezov et al., 2013). For roGFP2-iL, excitation with blue light would maximize the excitation and hence minimize the integration time required to sample sufficient photons for analysis. Furthermore, it has been reported that roGFP1 under conditions of intense illumination undergoes an irreversible photoswitch reaction that would foster the anionic form of the chromophore and thus may artificially report reducing conditions even though the local environment of the probe is oxidizing (Schwarzländer et al., 2008).

Generally, roGFP probes are appropriate for measuring the thiol/disulfide equilibrium within a linear range of about ±30 mV from their midpoint potential equivalent to an *OxD* between 10 and 90%. RoGFP2-iL may thus be suitable in the range from ∼ −205 to ∼ −275 mV. Recently it was shown that GRX1-roGFP1 iE is still oxidized to more than 90% when expressed in the ER of HeLa cells (Birk et al., 2013). This high degree of oxidation clearly limits the use of the probe in that it would not allow investigating processes leading to increasing oxidation in the ER. Based on the even slightly more negative midpoint potential a very similar response can be expected for GRX1-roGFP2-iL.

Stable expression of GRX1-roGFP2-iL and GRX1-roGFP2 in the cytosol of homozygous *rml1* plants allowed ratiometric measurements of the yet undefined *EGSH* with both sensors. In both cases the measured values are close to the end of the linear range of the respective probes with GRX1-roGFP2 being even slightly beyond its useful linear measurement range with an oxidation of 93%. GRX1-roGFP2-iL with *OxD* = 14% is still in the linear range. Importantly, *OxD* for both sensor variants can be converted to the respective redox potential and in both cases the redox potential is about −260 mV. A combination of both probes thus effectively doubles the useful dynamic range without leaving a gap between the two probes. The concentration of cytosolic glutathione in wild-type Arabidopsis root tips has been shown to be between 2 and 3 mM (Fricker et al., 2000). Assuming a medium concentration of 2.5 mM cytosolic GSH and an *OxDGSH* of 0.002% (Meyer et al., 2007) the Nernst equation predicts an *EGSH* of about −310 mV. A remaining GSH level of only 2% in *rml1* would then lead to an *EGSH* of −260 mV. This calculation assumes a constant *OxDGSH* which is not necessarily given. Obviously, the *rml1* mutant seedlings are stressed and supply with redox equivalents in form of NADPH may be restricted under these severe metabolic constraints. If indeed *OxDGSH* was higher in *rml1* than in the wild-type then the total glutathione level in the cytosol would have to be assumed to be slightly above 2% in order to reach an *EGSH* of −260 mV. This, however, is still well within the measured range of glutathione concentrations in *rml1*.

In conclusion, GRX1-roGFP2-iL further extends the range of roGFP-based probes for measurements of *EGSH* far less reducing than typical *EGSH* values normally found in the cytosol. This applies similarly to cell-type specific developmental differences as well as to mutants with low glutathione levels and to generally oxidizing compartments like the ER. GRX1-roGFP2-iL may still be largely oxidized in the ER of non-stressed plants as it has been shown for GRX1-roGFP1-iE in HeLa cells (Birk et al., 2013). The oxidizing redox potential in the lumen supports protein folding whereas deviations from steady-state redox conditions induce an unfolded protein response (Merksamer et al., 2008). Vice versa, malfunction in oxidative protein folding is assumed to affect the redox potential of the lumenal glutathione pool. Under stress situations and in mutants causing a shift in the ER redox potential toward more negative values, GRX1-roGFP2-iL will likely allow dynamic measurement of the glutathione-related redox processes in the ER lumen. Conversely, severe stress situations can cause very strong oxidation in the cytosol, chloroplasts, mitochondria, and peroxisomes and the extent of these reactions may also be dependent on the developmental state of particular cells. Darkinduced senescence has been shown to cause an oxidation of mitochondrial roGFP1 and roGFP2 up to the end of their linear range within only 3 days (Rosenwasser et al., 2010). Under such pathological conditions GRX1-roGFP2-iL will allow to further investigate the most oxidizing phases of the responses.

## **ACKNOWLEDGMENTS**

We thank James Remington for providing pQE-30 with roGFP1 iL and Christian Appenzeller-Herzog for pQE-30 with roGFP1 iE. The UBQ10 promoter was kindly provided by Karin Schumacher. This work was supported by a grant from the Deutsche Forschungsgemeinschaft (DFG) to Andreas J. Meyer (grant ME1567/5-1).

## **REFERENCES**

Finkel, T. (2011). Signal transduction by reactive oxygen species. *J. Cell Biol.* 194, 7–15. doi: 10.1083/jcb.201102095

the importance of glutathione in disease resistance of Arabidopsis. *Plant J.* 49, 159–172. doi: 10.1111/j.1365-313X.2006.02938.x

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 03 September 2013; accepted: 26 November 2013; published online: 16 December 2013.*

*Citation: Aller I, Rouhier N and Meyer AJ (2013) Development of roGFP2-derived redox probes for measurement of the glutathione redox potential in the cytosol of severely glutathione-deficient rml1 seedlings. Front. Plant Sci. 4:506. doi: 10.3389/fpls. 2013.00506*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2013 Aller, Rouhier and Meyer. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## *Tou Cheu Xiong1,2,3,4\*, Elsa Ronzier 1,2,3,4, Frédéric Sanchez 1,2,3,4, Claire Corratgé-Faillie1,2,3,4, Christian Mazars 5,6 and Jean-Baptiste Thibaud1,2,3,4*

*1 Biochimie et Physiologie Moléculaire des Plantes, Institut National de la Recherche Agronomique, UMR 386, Montpellier, France*

*2 Biochimie et Physiologie Moléculaire des Plantes, Centre National de la Recherche Scientifique, UMR 5004, Montpellier, France*

#### *Edited by:*

*Alex Costa, University of Milan, Italy*

#### *Reviewed by:*

*Zhi Qi, Inner Mongolia University-Hohhot, China Nathalie Leborgne-Castel, Université de Bourgogne, France Markus Schwarzländer, University of Bonn, Germany*

#### *\*Correspondence:*

*Tou Cheu Xiong, Biochimie et Physiologie Moléculaire des Plantes, Institut National de la Recherche Agronomique, UMR 386/Centre National de la Recherche Scientifique, UMR 5004/Montpellier SupAgro/Université Montpellier 2, Campus INRA-SupAgro, Place Pierre Viala, F34060 Montpellier Cedex 2, France*

*e-mail<EMAIL_ADDRESS>
Calcium (Ca2+) is a second messenger involved in many plant signaling processes. Biotic and abiotic stimuli induce Ca2+ signals within plant cells, which, when decoded, enable these cells to adapt in response to environmental stresses. Multiple examples of Ca2+ signals from plants containing the fluorescent yellow cameleon sensor (YC) have contributed to the definition of the Ca2+ signature in some cell types such as root hairs, pollen tubes and guard cells. YC is, however, of limited use in highly autofluorescent plant tissues, in particular mesophyll cells. Alternatively, the bioluminescent reporter aequorin enables Ca2+ imaging in the whole plant, including mesophyll cells, but this requires specific devices capable of detecting the low amounts of emitted light. Another type of Ca2+ sensor, referred to as GFP-aequorin (G5A), has been engineered as a chimeric protein, which combines the two photoactive proteins from the jellyfish *Aequorea victoria*, the green fluorescent protein (GFP) and the bioluminescent protein aequorin. The Ca2+-dependent light-emitting property of G5A is based on a bioluminescence resonance energy transfer (BRET) between aequorin and GFP. G5A has been used for over 10 years for enhanced *in vivo* detection of Ca2+ signals in animal tissues. Here, we apply G5A in Arabidopsis and show that G5A greatly improves the imaging of Ca2+ dynamics in intact plants. We describe a simple method to image Ca2+ signals in autofluorescent leaves of plants with a cooled charge-coupled device (cooled CCD) camera. We present data demonstrating how plants expressing the G5A probe can be powerful tools for imaging of Ca2+ signals. It is shown that Ca2+ signals propagating over long distances can be visualized in intact plant leaves and are visible mainly in the veins.

**Keywords:** *Arabidopsis thaliana***, calcium imaging, cooled CCD camera, GFP-aequorin, leaf, long distance calcium signaling, calcium waves, salt stress**

## **INTRODUCTION**

Calcium (Ca2+) has long been established as a second messenger. Transgenic expression of fluorescence resonance energy transfer (FRET)-based fluorescent Ca2+ reporters such as the popular yellow cameleon (YC) or of the bioluminescent aequorin has permitted non-invasive monitoring of free Ca2+ levels and enabled real-time imaging of Ca2+ levels in different cell-types and organisms, including plants (Knight et al., 1991; Perez Koldenkova and Nagai, 2013). The YC has been used extensively for imaging Ca2+ signals in specific plant cell types such as guard cells (Allen et al., 1999), germinating pollen tubes (Iwano et al., 2012), and root hairs (Miwa et al., 2006; Monshausen et al., 2008). YC is also well suitable for Ca2+ sensing in subcellular compartments (Krebs et al., 2012; Bonza et al., 2013). However, YC requires excitation by exogenous light, which limits its relevance in plant photosynthetic tissues due to high background emission from auto-fluorescent cell walls, chlorophyll, and secondary metabolites. Indeed, wide autofluorescent spectrum of plant leaf pigments that overlap YC emission limits visualization of changes in intensity of YC fluorescence emission upon Ca2+ elevation. Moreover, Ca2+ imaging at plant tissue level requires strong and long excitation to detect fluorescence signals. Long term Ca2+ measurements would result in some YC photo-bleaching and/or tissue damage, this limiting long term Ca2+ measurements, over 24 h for example. On the other hand, the bioluminescent Ca2+ reporter aequorin does not require exogenous excitation light and very little background signal is produced resulting in a high signal-to-noise ratio throughout long acquisition periods. Aequorin has the largest dynamic range among Ca2+ reporters, allowing the monitoring of Ca2+ signals over several days and over a wide range of Ca2+ concentrations (Alonso and Garcia-Sancho, 2011). Aequorin has been introduced into several plant species (Knight et al., 1991; Webb et al., 2010) and has enabled photon counting based monitoring of Ca2+ in intact plant leaves. Many reports of aequorin application in plants have been published, where photon counting with luminometers was used to

**METHODS ARTICLE** published: 18 February 2014 doi: 10.3389/fpls.2014.00043

*3 Biochimie et Physiologie Moléculaire des Plantes, SupAgro, Montpellier, France*

describe Ca2+ signaling under several stress conditions. However, to image photons emitted by aequorin with good resolution in both space and time requires sophisticated detection devices such as image photon detectors (IPDs) (Webb et al., 2010) or cameras fitted with an Intensified Charge-Coupled Device (ICCD) (Webb et al., 2010) or Electron Multiplying Charge-Coupled Device (EMCCD) (Rogers et al., 2008; Webb et al., 2010). This is a significant limitation to *in planta* Ca2+ imaging which could be overcome by using the G5A probe, an engineered fusion between the green fluorescent protein (GFP) and aequorin (**Figures 1A,C**) initially developed for Ca2+ imaging in animal cells (Baubet et al., 2000; Rogers et al., 2005). Through a bioluminescence resonance energy transfer (BRET) from aequorin to GFP, the wavelength of the emitted photon is 510 nm, instead of 470 nm and detection yield by CCD is found optimized, compared to aequorin, with a better signal/noise ratio (Baubet et al., 2000; Rogers et al., 2005, 2008).

Here, (i) we applied G5A in Arabidopsis, (ii) we show that, in comparison to aequorin, G5A enhances *in vitro* and *in vivo* detection of weak Ca2+ events in intact plants, including in photosynthetic tissues and (iii) we describe a simple method that only

requires a cooled-CCD camera to visualize Ca2+ signals in plant leaves as, for example, Ca2+ waves propagating along leaf veins of intact plants after imposing a salt stress to roots of these plants. It is concluded that G5A reporter is an interesting alternative to aequorin.

## **MATERIALS AND METHODS**

## **CLONING G5A AND ENGINEERING G5A-EXPRESSING PLANTS**

The original vector harboring the *G5A* construct (Baubet et al., 2000) was kindly provided by Dr. Philippe Brûlet's group (CNRS, Gif-sur-Yvette, France). The G5A coding sequence was cloned into the Gateway® entry vector pDONR™ by two sequential PCRs amplification using a *G5A* forward primer 5 -GGAGATAGAACC**ATGAGCAAGGGCGAGGAGCTGTTCA**-3 and a *G5A* reverse primer 5 -TCCACCTCCGG**ATCAGGGGAC AGCTCCACCGTAG-**3 , followed by a second PCR using a U5 forward primer 5 -GGGGACAAGTTTGTACAAAAA AGCAGGCTTCGAAGGAGAT-AGAACCATG-3 and a U3 reverse primer 5 -AGATTGGGGACCACTTTGTACAAGAAA GC-TGGGTCTCCACCTCCGGATC-3 . Next step was a transfer, by LR Gateway® recombination, of the *G5A* construct into the expression vector pGWB502-(Nakagawa et al., 2007).

The pGWB502--*G5A* construct was introduced in *Agrobacterium tumefaciens* (GV3101), for transformation of *Arabidopsis thaliana* ecotype Col-0 by the floral dip method (Clough and Bent, 1998). G5A expressing transgenic plants were selected using hygromycin selective media and checked for GFP fluorescence emission under direct excitation of GFP at 488 nm (see **Figure 2**). Homozygous G5A expressing T3 and T4 plants (here below denoted *G5A* plants) were used and compared to transgenic plants expressing aequorin in the cytoplasm (Col-0 ecotype, denoted below *Aeq* plants) obtained from Prof. Marc Knight (Durham, UK).

## **PLANT MATERIAL AND GROWTH CONDITIONS**

Seeds from *Aeq* and *G5A* plants were surface-sterilized and placed on half-strength Murashige and Skoog plate medium supplemented with sucrose 1% (w/v) and with hygromycin (15μg/mL)(*G5A*) or kanamycin (50μg/mL)(*Aeq*), and stratified at 4◦C for 2 days in the dark. Seedlings were subsequently grown in a growth chamber at 22◦C with a 70% relative humidity, in long-day conditions (150μE/m2/s light for 16 h a day) for 7 days. These 7-day old seedlings were either used directly or further grown in soil under short day conditions (200μE/m2/s light for 8 h a day) for 3–7 weeks, as indicated.

## **LUMINESCENCE MEASUREMENT AND IMAGING** *Seedlings*

*In vivo* reconstitution of functional aequorin was performed by incubating 7 day-old seedlings for 4 h at 22◦C in the dark with a 2.5μM aqueous solution of coelenterazine HCP (Interchim). For imaging, coelenterazine-treated seedlings were placed within a dark chamber over a gelosed layer (water with 1% agar) in large Petri dishes. A cooled-CCD camera (Hamamatsu 4880-30), fitted at the top of the chamber, collected photons. Sequential image acquisition was carried out using the Hipic 5.1.0 software with an exposure time per image in the 15–60 s range (as indicated in the

plants of the *G5A* line were checked for reporter expression by excitation of GFP (at λex = 488 nm) under a stereo microscope **(A,B)** or a confocal microscope **(C–J)**. Fluorescence emission by GFP is shown in **(A,C,E,G,I)** and corresponding bright field images are shown respectively in **(B,D,F,H,J)**. microscopy observation of leaf epidermal cells **(C,D)** and mesophyll cells **(E,F)** showed good expression of G5A in leaf tissues. G5A fluorescence was observed in both the cytosol and the nucleus (arrows) of root cells **(G,H)** and of stomatal guard cells **(I,J)**. Scale bar = 20μm.

Figure legends). Images were analyzed in ImageJ (Schneider et al., 2012). The first 5 min in each sequence were discarded because of chlorophyll autofluorescence decay.

## *Older soil-grown plants*

At the end of the day, the soil was gently removed from the roots and the whole plants were incubated for 4 h at 22◦C in the dark in a 2.5μM aqueous solution of coelenterazine HCP. Plants were then placed with the roots in Qualibact® (CEB) tubes filled with water through a hole in the cap of the tube to separate leaves from roots. Plants were left for 1 h in the dark at room temperature for recovery. Images of the rosette were acquired as described above for seedlings, with a 30 s integration time. NaCl at a final concentration of 200 mM was injected with a remotely controlled syringe at the root level after 25 min of acquisition.

## *Excised mature leaves*

Mature leaves were excised from 6–7-week old plants grown as described above and incubated at 22◦C in the dark for 4 h in a 2.5μM aqueous solution of coelenterazine HCP. The treated excised leaf was then transferred to a dark chamber under the cooled-CCD camera for G5A imaging as described above with 15 s exposure time by frame. For salt stress, 100μL of 200 mM NaCl was pipetted onto the excised leaf petiole (see **Figure 8A** arrow) and subsequent light emission was acquired.

## **IMAGE ANALYSIS**

All images were analyzed in ImageJ. Shading correction of all images was performed by subtraction of a dark field image (acquisition without sample) acquired with the same exposure time. Backgrounds of each image were normalized by subtraction of ROIs of non-plant pixels. ROIs of plant pixels were then quantified and the average values are plotted over the time. Background noise after chlorophyll fluorescence decay was determined by imaging light emission from wild-type plant leaves and then subtracted for data obtained from *G5A* or *Aeq* plants. Ca2+ signal velocity was determined with the MtrackJ plugin of ImageJ and the localization (x,y) of each velocity value was plotted with Matlab® software (R2006a).

## **AEQUORIN IMMUNOBLOTTING**

Soluble protein was extracted from 50 pooled seedlings (Mithofer and Mazars, 2002) and separated by SDS-PAGE. Immunoblotting was carried out using an anti-aequorin rabbit polyclonal antibody (Novus Biological, NB100-1877) as described by the manufacturer. Signals of immunodetection were acquired with LAS-3000 imager (Fujifilm) and quantified with Multi-Gauge v3.2 (Fujifilm).

### **CALIBRATION OF THE TWO PROBES G5A AND AEQUORIN**

Soluble proteins were extracted from *G5A* and *Aeq* plants as described by Mithofer and Mazars (2002) and were diluted in a buffer (Tris-HCl 200 mM, pH 7.4, EGTA 5 mM, NaCl 0.5 M, β-mercaptoethanol 5 mM) containing the coelenterazine HCP cofactor for 2 h in the dark at 4◦C. Relative amounts of G5A and aequorin reporters in these crude extracts were estimated by immunoblotting (**Figure 3A**). So-called *G5A* and *Aeq* buffers were prepared by diluting soluble protein crude extracts from *G5A* and *Aeq* plants in Tris-HCl 200 mM, pH 7.0, EGTA 5 mM at a protein content of 0.1μg/μL and 0.15μg/μL respectively (to ensure that subsequent *in vitro* comparison of both reporters was made with equal quantities of them). Wells of 96-well plates were filled with 50μL of different Ca2+ solutions, of which the free Ca2+ concentration was estimated by MaxChelator Software (http://www.stanford.edu/∼cpatton/downloads.htm). To start probe calibration, 50μL of either *G5A* or *Aeq* buffer was dispensed into each well and maximum light emitted per second (L) was measured. In a second step, 100μL of a 2 M CaCl2 solution was dispensed into each well for discharging the remaining reconstituted G5A or aequorin reporters. Light emitted at this time (Ltotal) allowed the total amount of functional Ca2+ reporter to be estimated. All light measurements were made with a plate-spectrophotometer Victor2 (Perkin Elmer). Collected photons were integrated over 1 s lapses during 180 s. Results are expressed as the ratio ±SE of maximum light over total light (L/Ltotal).

## **RESULTS**

Effective transformation using the pGWB502--*G5A* construct was expected to yield a broad constitutive expression pattern of the G5A probe. This was checked in a Ca2+-independent manner by direct excitation, at 488 nm, of the GFP moiety of the chimera probe (**Figure 2**).

A strong ubiquitous GFP signal was observed in 7 day-old seedlings. The subcellular pattern of the GFP signal suggested cytosolic and nuclear localization (arrows in **Figures 2G,I**).

An anti-aequorin polyclonal antibody (Novus Biological, USA) was used to evaluate the amount of G5A and aequorin proteins in the soluble protein fraction (**Figure 3A**). This antibody revealed strong bands at 22 kDa and 52 kDa in both protein extracts from *Aeq* and *G5A* plants. The G5A/aequorin ratio (protein level) was estimated at 1.48 ± 0.14.

Calibration curves were performed with soluble protein extracts from *G5A* or *Aeq* plants. Equal amounts of G5A and aequorin reporters were used for *in vitro* calibration curves. Data are expressed as maximum light emitted per second (denoted "L") (Fricker et al., 1999) over total light (denoted "Ltotal") ratio (**Figure 3B** see "Methods" section). G5A and aequorin showed similar responses to free Ca2+ concentration and calibration curves in **Figure 3B** do not differ significantly over the 10−8 to 10−6 M free Ca2+ range. Linear regression between 10−8 to 10−6 M free Ca2+ reveals a straight line with a slope of 1.065 ± 0.016 (*R*2 = 0.988) and 0.888 ± 0.015 (*R*2 = 0.987) for

**FIGURE 3 |** *In vitro* **calibration of G5A and aequorin as Ca2+ reporters. (A)** Immunoblotting of soluble protein fractions from fifty 14 day-old plants of the *G5A* and *Aeq* lines with an anti-aequorin polyclonal antibody. Signals of immunoblotting were quantified and indicated as a G5A/aequorin ratio of protein accumulation of 1.48 ± 0.14 (average of one hundred seedlings obtained in two independent experiments ± SE, *n* = 2). **(B)** *In vitro* calibration curves of G5A and aequorin in soluble protein extracts from *G5A* and *Aeq* plants. Equivalent amounts of Ca2+ reporters were used for the comparison (based on assay reported in **A**) and incubated in a buffer containing 2.5μM coelenterazine HCP cofactor for 2 h in the dark at 4◦C prior light emission assay in the presence of various free Ca2+ amounts (see Materials and Methods). Results are expressed as the ratio ± SE (*n* = 6 in two independent experiments) of maximum light over total light (L/Ltotal). The linear range of L/Ltotal ratio as a function of free Ca2+ concentration is shown for G5A (full line) and aequorin (dashed line).

G5A and aequorin respectively (**Figure 3B**). The reciprocal relationship, i.e., between free Ca2+ concentration and the rate of consumption of G5A or aequorin, can be represented by the equation:

$$-\log([Ca^{2+}]) = a\* - \log(\frac{L}{L\_{\text{Total}}}) + b \text{ (Fricker et al., 1999)}$$

Coefficients *a* and *b* in the above equation are 0.93919 and 5.61289 for G5A, and 1.13646 and 5.26608 for aequorin.

The bioluminescent reporter aequorin has very low noise and high signal/noise ratio (Brini, 2008; Webb et al., 2010). No signal from G5A and aequorin was detected under *in vitro conditions*, in the absence of coelenterazine HCP. In the presence of coelenterazine HCP and without Ca2+, G5A and aequorin noise levels were respectively 271.16 ± 10.76 and 198.67 ± 11.52 RLU (Relative Light Unit). This difference is, however, negligible compared to signal after injection of free Ca2+. At the basal level of cytosolic free Ca2+ (0.1μM), maximum light level was increased to 6.61 × 103 ± 0.32 × 103 and 1.72 × 103 ± 0.29 × 103 RLU for G5A and aequorin respectively. At 1 μM free Ca2+, light levels increased up to 90.14 × 103 ± 7.44 × 103 and 20.25 × 103 ± 1.65 × 103 RLU for G5A and aequorin respectively. Data expressed as signal/noise ratio for the two reporters (**Figure 4A**) show that G5A is approximately 3–5 times better than aequorin. For instance, the signal/noise ratio of aequorin with 1μM free Ca2+ is reached with only 300 nM free Ca2+ with G5A reporter.

In parallel, comparison of the two Ca2+ reporters was performed *in planta*. Different time lapses were tested for collecting photons emitted by *G5A* and *Aeq* plants (**Figure 4B**). The threshold for signal detection for basal level of free Ca2+ was approximately 10 s and 30 s with G5A and aequorin respectively. A 30 s time lapse allowed sufficient light to be collected from *G5A* plants while 1 min was hardly sufficient in the case of *Aeq* plants.

Sudden light-dark transition has been reported to induce weak Ca2+ signals in photosynthetic tissues (Johnson et al., 1995; Sai and Johnson, 2002; Dodd et al., 2006). To assess the capability of G5A to detect weak Ca2+ events in intact plant tissues, we challenged Arabidopsis plants with darkness: the reactions of *G5A* plants upon light-dark transition were compared to those of *Aeq* plants (**Figure 5**). Significantly more photons could be collected from *G5A* plants than from the *Aeq* plants over this period (**Figure 5B**). Successive integrations of photons over 1 min time lapses provided an overview of the Ca2+ signal kinetics (**Figure 5C**). Dark-induced Ca2+ signals displayed by G5A and aequorin had parallel kinetics (**Figure 5C** and inset), with a maximal light emission between 40 and 60 min. However, approximately five times more photons were detected from plants of the *G5A* line.

This interesting G5A feature allowed us to follow the dynamics of free Ca2+ in leaves triggered by a salt stress applied to roots of intact plants (**Figure 6** and **Supplementary videos S1**, **S2**). The stress was sensed by roots and propagated to leaves, suggesting that Ca2+ waves might contribute to plant adaptation to salt stress. A time series of representative results (from video S1) is presented in **Figure 6A**. It shows that a 30 s delay after the

**and aequorin (A)** *In vitro* assay of light emission from equal amounts of Ca2+ reporters G5A and aequorin is plotted against buffer free Ca2+ concentration. Results (mean ± SE, *n* = 6) are expressed as the ratio of signal (maximum emitted light) over noise. **(B)** Representative *in vivo*

over different exposition time lapses (10, 30, and 60 s). Left panel shows bright field view (scale bar = 1 cm) of the imaged plants. The other panels display cumulative Ca2+ responses in false colors (color scale in the 10 s-labeled panel).

application of NaCl (at time = 0) was required before Ca2+ levels increased in the petioles. Elevated calcium levels then propagated to the rest of the leaves. It is interesting to note that the Ca2+ responses of mature and young leaves differed in terms of kinetics. Mature leaves responded by an initial rapid, transient, Ca2+ peak (**Figure 6A**, time = 1–2 min) followed by a second very slow, wave-like, increase and subsequent decrease of free Ca2+ level lasting more than 50 min (from 6 to 60 min) with a maximum at 12–13 min. The young leaves displayed a single rapid Ca2+ transient peak (**Figure 6A** red arrows), similar to that observed in mature leaves although slightly later (3.5–4.5 min after salt stress application). Defining the whole plant as an ROI and plotting the time course of the signal summed over each 30 s lapse (in RLU/30 s) for 60 min after salt stress application yielded a dynamic view of these Ca2+ events at leaf level (**Figure 6B**). It was found that both the Ca2+ peaks (observed for mature and young leaves) and the Ca2+ wave had a maximum at 19–20 RLU/30 s (**Figure 6B**). Despite inevitable variations from a plant to another one, an analogous pattern of distribution in space and time of Ca2+ events was observed when challenging a plant with a salt stress at the root level (**Supplementary video S2**).

Light emitted by G5A in intact plants facing a salt stress therefore appeared to be sufficiently intense to image the propagation of Ca2+ signals in leaves with good time resolution. We performed a simple analysis of Ca2+ waves on each leaf of plants subjected to a salt stress applied to roots. Ca2+ signal velocities were then calculated for each leaf of the plant shown in **Figure 6** (red dashed arrows, **Figure 7**). This shows that velocity was not constant within a given leaf (it decreased at leaf tip) and differed depending on the leaf. Detailed numerical values are given (**Table 1**): maximum and minimum of velocities were 0.52 and 0.03 mm/s respectively.

Further applications of G5A were investigated by imposing similar salt stress on mature leaves excised from 7–8 week-old plants (**Figure 8**). Application of 100μL of 200 mM NaCl onto the petiole end of an excised leaf (**Figure 8A**, white arrow) was enough to generate detectable elevations in free calcium after 1 min. They started immediately after the exposure to the NaCl solution at the site of application before they spread throughout the rest of the leaf (**Figure 8A** and **Supplementary videos S3**, **S4**). The propagation of free calcium elevation seems to be different in the basal third part of the leaf (denoted by ∗ in **Figure 8C**) as compared to the rest of the leaf (denoted by ∗∗ in **Figure 8C**). In the early phase of response to the applied stress, increase in free Ca2+ were observed in the peripheral regions of the basal third part of the leaf and not in the middle vein. Subsequently, Ca2+ responses seem to propagate throughout the leaf, firstly, along the vascular tissues (primary and secondary veins) and later in the mesophyll tissue. Quantification of increase in free Ca2+ from the entire leaf shows that salt stress induced two different peaks of Ca2+, which correspond to these two successive episodes of Ca2+ increase, firstly in the basal part of the leaf, subsequently in the rest of the leaf (as indicated by asterisks in **Figure 8B** as described for **Figure 8C**).

In this example (representative of five independent leaves), elevation of free Ca2+ induced by NaCl needed 315 s (image at 1:00 to 6:15) to travel through a 57 mm-long leaf. Thus, in this example the average velocity was 0.181 mm/s. A further analysis was performed on the primary vein of this leaf (**Figure 8C**, red dashed line X–Y), a kymographic representation of velocity value on the axis X–Y shows that there were three different Ca2+ response velocities (**Figure 8D**). Two of them (red dashed and orange dashed arrows) spread acropetally (from X to Y) whereas one propagated the opposite way (Y–X, cyan dashed arrow). Interestingly, between the first third and the second third of the leaf, was observed a region where no Ca2+ signals were detected with G5A. Despite this gap, Ca2+ signal propagation was observed all along the XY axis (**Figure 8C** and **Supplementary video S3**).

Analyses of Ca2+ waves on this excised leaf (representative of five leaves) show that Ca2+ signal velocities along different veins (**Figure 9A** dashed arrows) were different. Local velocity values were plotted on the image (as spots in false-color scale, **Figure 9B**). Higher velocities in the center of the leaf and slower velocities at leaf borders were found (**Figure 9B**). Details of velocities of Ca2+ signals in leaf veins, including on X–Y axis (#1, #8, and #16) are presented in **Table 2** below. Despite inevitable variation from a leaf excised from a plant to another leaf excised from another plant, the nature and pattern (both in space and time) was essentially reproducible (see **Supplemental video S4**).

## **DISCUSSION**

Since the successful use of aequorin as a Ca2+-signaling reporter in plant tissues (Knight et al., 1991), examples of aequorin imaging in plants have relied on the use of ultra-sensitive camera devices (intensified-CCD or electron-multiplying-CCD) to detect the few photons emitted by aequorin. They have revealed, for example, that Ca2+ oscillations occur during diurnal rhythms in plant leaves (Johnson et al., 1995; Sai and Johnson, 2002; Dodd et al., 2006). Optimized imaging of aequorin signals in plants has been reported recently that, with integration time

**high salt stimulus applied to roots. (A)** Free Ca2+ elevations on each leaf were analyzed with ImageJ. Velocities (mm/s) of Ca2+ responses were determined for each leaf along paths figured by red dashed arrows (values of Ca2+ response velocity are presented on **Table 1**). **(B)** Propagation speeds along the main leaf vain are indicated for selected points (same space scale as in **A**). False color scale is in mm/s.

down to 40 s, showed stimulus- and tissue-specific Ca2+ signatures in seedlings (Zhu et al., 2013). In these examples of aequorin imaging in plants, however, low light detection relied upon sophisticated and costly equipment. Another interesting development of aequorin-based approach to *in planta* Ca2+ signaling has recently been reported: expression of aequorin in specific cell types of Arabidopsis was used to determine which cell types release calcium into the cytosol in response to a given stimulus (Marti et al., 2013). Photometry (with high time resolution) of aequorin emission from plants expressing the Ca2+-sensor in specific leaf cell–types (mesophyll cells, guard cells, [peri-]vascular cells, epidermal cells, and trichomes) may be used to follow various stresses. These experimental conditions allowed the collected information to be ascribed to a given cell-type, without spatial localization of the measured signals.

The G5A-based method we report here provides an interesting complement to these recent improvements of aequorin-based methods. In the *Aequoria victoria* jellyfish, a naturally evolved BRET phenomenon between aequorin and GFP occurs. Several artificial proteins assembling aequorin with GFP-derived proteins have been engineered to mimic the natural BRET observed in *Aequoria victoria*. Performance of these artificial Ca2+ reporters depends on the linker motif placed in between the BRET partner proteins (Baubet et al., 2000; Gorokhovatsky et al., 2004). Comparison of different linkers in GFP-aequorin protein fusions has demonstrated that the five repeat motifs used here in the so-called G5A artificial reporter (see **Figure 1**) allows a high BRET efficiency (Baubet et al., 2000). Subsequently, the G5A reporter has been successfully used to monitor Ca2+ elevation at cellular and organ levels in animals (Baubet et al., 2000; Chiesa et al., 2001; Cassidy and Radda, 2005; Rogers et al., 2005, 2008; Martin et al., 2007; Naumann et al., 2010). To date, however, no G5A application in plants has been reported.

Although our *in vitro* assays show similar calibration curves for both reporters (**Figure 3B**), there was a significantly better signal/noise ratio for G5A than for aequorin (**Figure 4A**) corresponding to a 3–5 fold increase in light collected from the former compared to the latter. Such an amplification is consistent with the data obtained *in planta* (**Figures 4B**, **5**) where small changes of free Ca2+ concentration were more easily detected with G5A than with aequorin. The (up to) 5-fold amplification of light in plants expressing G5A is similar to the 5.7-fold increase

*\*Time resolution is limited by the time acquisition (30 s).*

**the vascular tissue. (A)** Time series of images (time of capture indicated in the right top corner of each image in min:s) showing propagation of Ca2+ responses in a detached leaf. First image of the series is a bright-field view of dispensed onto the petiole (white arrow) and light subsequently emitted from leaves was integrated over 7 min (each image shows cumulative light over *(Continued)*

#### **FIGURE 8 | Continued**

a 15 s time lapse). Ca2+ responses were immediately detected on petioles and propagation of Ca2+ responses to the end of leaf was observed. Scale bar = 1 cm. Calibration bar in gray scale is shown in the last image (RLU). Representative result of five independent experiments. **(B)** Quantification of light signals from ROI for the whole leaf (see **C**) over 8 min (in RLU per 15 s). At time = 1 min, NaCl was applied onto the petiole (black arrow). The two peaks denoted by ∗ and ∗∗ correspond to a Ca2+ increase in the

in the light signal resulting from BRET between luciferin and GFP in *Renilla reniformis* (Ward and Cormier, 1976). It is interesting to note however that the improved detection of light emission from aequorin through BRET is still not completely understood (Webb et al., 2010). In the jellyfish *Aequorea victoria*, aequorin is associated with the GFP that allows the amplification of light by BRET phenomena. *In vitro*, the binding between aequorin and GFP does not occur, even at high concentration and no BRET events was observed (Baubet et al., 2000). Fusion of aequorin and GFP is a prerequisite to observe such natural light amplification seen in jellyfish. Baubet et al. have designed an optimized linker that results in sufficient high quantum yield to visualize Ca2+ induced light emission from G5A (Baubet et al., 2000).

An issue for plant cell physiologists is the autofluorescence of chlorophyll and other organic pigments which hinders the use of fluorescent probes with overlapping spectroscopic properties. For instance chlorophyll *b* and carotenoids (lutein, neoxanthin, and violaxanthin) have major absorption peaks between 469 and 490 nm (Rivadossi et al., 2004; Taylor et al., 2006) and may absorb photons emitted at ∼470 nm by aequorin in leaf cells. As a consequence of the efficient BRET between its aequorin and GFP moieties, G5A emits photons at∼510 nm (Baubet et al., 2000; Rogers et al., 2005, 2008), thus reducing absorbance by plant tissues and resulting in better detection by cameras or luminometers. This also could contribute to the better performance of G5A compared to aequorin in the present *in planta* experiments (**Figure 5C**).

In addition to these considerations, the G5A-based results that we report here exemplify the possibilities that this Ca2+ reporter holds for plant biologists. Comparison of *Aeq* and *G5A* plants subjected to darkness shows that similar responses could be visualized with both Ca2+ reporters, but five times more light was emitted from *G5A* plants (**Figure 5**). This suggests that G5A is an alternative tool to aequorin for Ca2+ imaging when signals are either too low or if ultra-sensitive camera is not available. Moreover, further examples of potential application of G5A are introduced here, showing the analysis of longdistance Ca2+ signaling (potentially involved in the coordination of the integrative responses of plant to stresses, here salt stress, **Figures 6**–**9**).

Long distance propagation of Ca2+ signals ("Ca2+ waves") is attracting increasing interest in the plant biology community (Steinhorst and Kudla, 2013) and in this context, G5A seems to be a complementary tool, along with fluorescent Ca2+ reporters, to investigate complex cell to cell communication within plants. Most of Ca2+ imaging experiments in intact tissue with a fluorescent reporter have been carried out in roots (Fasano basal third and the rest of the leaf respectively. **(C)** Localization of Ca2+ responses is represented by a z-stack of standard deviation of Ca2+ signals over the 8 min of measurement. Ca2+ signals seem to take place in leaf veins. The basal third of the leaf is indicated by ∗ and the rest of the leaf by ∗∗. Line scan of primary vein (red dashed line X–Y) is analyzed in **(D)**. **(D)** Kymographic representation of Ca2+ signals in the primary vein (X–Y from **C**). Three different velocities of Ca2+ signals could be measured and are indicated.

et al., 2001; Monshausen et al., 2011; Gjetting et al., 2012), where auto-fluorescence is much less of an obstacle than in leaves. It is by using FRET-confocal laser scanning microscopy that Ca2+ imaging on leaves with YC has recently been reported (Benikhlef et al., 2013; Verrillo et al., 2014). Aequorin imaging requires ultrasensitive cameras (intensified-CCD or electron-multiplied-CCD) while G5A imaging does not. The high dynamics and intensity of G5A light emission upon Ca2+ events open opportunities to detect low Ca2+ signals and to analyse their propagation with good time resolution, using a "regular" cooled CCD camera. In practical terms, time resolution is the time required to make an image (a frame) with an acceptably high signal/noise ratio. Under the present experimental conditions, integration time for a frame was well under 1 min: depending on amplitude of the Ca2+ signal this integration time ranged from 30 s (**Figure 6**) down to 15 s (**Figure 8**) for whole seedlings or mature leaves and even down to 5 s in the most favorable case (wounding stress, data not shown). Thus, values of Ca2+ wave propagation speed as fast as 0.5–0.6 mm/s can be resolved (**Figure 9**, **Tables 1**, **2**).

We consider that G5A reporter opens exciting perspectives for the study of cell-to-cell communication in plants. The physiological meaning of Ca2+ waves observed within vascular tissues (but not solely there) is one of the interesting aspects which could be investigated. Velocity values observed for these Ca2+ signals are of the same order of magnitude than those of "fast" electrical signals (i.e., "action potentials") reported to travel leaf tissues in Arabidopsis (Favre et al., 2011). This substantiates the hypothesis of an interplay between Ca2+ and electrical signaling (Król et al., 2011). Recently, glutamate receptor-like putative Ca2+ channels were reported to play a role in leaf to leaf signaling after wounding (Mousavi et al., 2013), together with electrical signals. In this context, combining G5A-based imaging of Ca2+ with electrophysiological recording of electrical signals might be a powerful method to decipher the molecular basis of electrical signaling in plants upon different types of stress, including wounding.

In conclusion, G5A allows to image free Ca2+ elevation in intact plant leaves, making this probe a promising addition in the toolbox of plant cell physiology. Ca2+ imaging in intact plant leaves with widely affordable imaging equipment has the potential to boost the investigation of Ca2+ signaling in plants.

## **ACKNOWLEDGMENTS**

We thank Dr. T. Nakagawa (Nagoya, Shimane University, Japan) for the pGWB502 plasmid, Prof. M. Knight (Durham University, UK) for the *Aeq* line (pMAQ2, Col-0 ecotype), the

along paths represented by dashed arrows (details of Ca2+ signal velocity,

*\*Time resolution is limited by the time acquisition (15 s).*

late Prof. Philippe Brûlet (CNRS, Gif-sur-Yvette, France) who kindly provided us with the *G5A* construct that we used to build the *G5A* line. This work was supported by a joint CNRS-INRA fellowship to Elsa Ronzier, by the Agropolis Fondation (Grant 0803-022), and by the European Commission's FP7 (Grant IRG 268393). We are grateful to Drs. Cliona Hann and Siobhán Staunton for their helpful comments on the manuscript.

for selected points (same space scale as in **A**). False color scale is in mm/s.

## **SUPPLEMENTARY MATERIAL**

The Supplementary Material for this article can be found online at: http://www.frontiersin.org/journal/10.3389/fpls.2014.00043/ abstract

**Supplementary video S1 | Light emission from 4 week-old G5A-expressing leaves upon a high salt stimulus applied to roots of the plant presented in Figures 5, 6.** The left part **(A)** shows the bright field view of the plant (scale bar is 2 cm). Video in the right part **(B)** is made of frames, each corresponding to photons collected over a 30 s time lapse, displayed at a rate of three frames per second (video rate is ×90). At time zero of the video, the plant has been adapted to dark for 20 min and no Ca2+ signals were detected (data not shown). Application of 200 mM NaCl to roots at *t* = 5 min (3.33 s, video time) induced propagation of Ca2+ elevation over long distances from roots to leaves over 77 min. NaCl-induced Ca2+ waves were different in expanded leaves and in young ones. Propagation of strong Ca2+ elevations to fully expanded leaves can be visualized just after the NaCl stress and is followed by slow Ca2+ variations in leaves over 77 min. Small young leaves emit strong and transient Ca2+ elevation only 3.5–4.5 min after the NaCl stress (5.66–6.66 s, video time).

**Supplementary video S2 | Second example of light emission from 4 week-old G5A-expressing leaves upon a high salt stimulus applied to roots of the same plant.** The left part **(A)** shows the bright field view of this plant (scale bar is 2 cm). Video in the right part **(B)** is made of frames, each corresponding to photons collected over a 30 s time lapse, displayed at a rate of three frames per second (video rate is ×90). At time zero of the video, the plant has been adapted to dark for 20 min and no Ca2+ signals were detected (data not shown). Application of 200 mM NaCl to roots at *t* = 5 min (3.33 s, video time) induced propagation of a Ca2+ elevation over long distances from roots to leaves over 150 min. As in video S1, NaCl-induced Ca2+ waves were different in expanded leaves and in young ones. In this other plant, however, propagation rates and kinetics of Ca2+ changes were different from those in the plant featured in video S1, exemplifying the complex nature of Ca2+ responses in intact plants.

**Supplementary video S3 | Light emission by a leaf detached from a** *G5A* **plant upon a high-salt stimulus applied to the petiole.** The left part **(A)** shows the bright field view of the leaf (scale bar is 1 cm). Video in the right part **(B)** is made of frames, each corresponding to photons collected over a 15 s time lapse, displayed at a rate of six frames per second (video rate is ×90). At time zero no light is detected. Application of 200 mM NaCl to petiole at *t* = 1 min (0.66 s, video time) induced Ca2+ waves from petiole to leaf tips over 7 min.

**Supplementary video S4 | Second example of light emission from a detached G5A-expressing leaf upon a high salt stimulus applied to the petiole.** The left part **(A)** shows the bright field view of the leaf (scale bar is 1 cm). Video in the right part **(B)** is made of frames, each corresponding to photons collected over a 15 s time lapse, displayed at a rate of six frames per second (video rate is ×90). At time zero no light is detected. Application of 200 mM NaCl to petiole at *t* = 1 min (0.66 s, video time) induced Ca2+ waves from the petiole to leaf tips over 17 min. In this leaf from another plant, however, propagation rates and kinetics of Ca2+ changes were different from those in the leaf featured in video S3, exemplifying the complex nature of Ca2+ responses in excised leaves.

## **REFERENCES**

bioluminescent reporters. *Eur. J. Neurosci.* 21, 597–610. doi: 10.1111/j.1460- 9568.2005.03871.x

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 12 December 2013; accepted: 29 January 2014; published online: 18 February 2014.*

*Citation: Xiong TC, Ronzier E, Sanchez F, Corratgé-Faillie C, Mazars C and Thibaud J-B (2014) Imaging long distance propagating calcium signals in intact plant leaves with the BRET-based GFP-aequorin reporter. Front. Plant Sci. 5:43. doi: 10.3389/fpls. 2014.00043*

*This article was submitted to Plant Cell Biology, a section of the journal Frontiers in Plant Science.*

*Copyright © 2014 Xiong, Ronzier, Sanchez, Corratgé-Faillie, Mazars and Thibaud. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

#### *Sébastien Peter 1, Martina B. Zell 2, Christian Blum3, Alexander Stuhl 4, Kirstin Elgass 1, Marcus Sackrow4†, Vinod Subramaniam , Alfred J. Meixner 4, Klaus Harter 1, Veronica G. Maurino2,6 and Frank E. Schleifenbaum1,4\* 3,5*

#### *Edited by:*

*George R. Littlejohn, University of Exeter, UK*

#### *Reviewed by:*

*Suleyman I. Allakhverdiev, Russian Academy of Sciences, Russia Tomas Morosinotto, Università di Padova, Italy Thomas Paul Howard, University of Exeter, UK Cornelia Spetea, University of Gothenburg, Sweden*

#### *\*Correspondence:*

*Frank E. Schleifenbaum, University of Tübingen, Auf der Morgenstelle 18,72076 Tübingen, Germany e-mail: frank.schleifenbaum@ uni-tuebingen.de*

During photosynthesis, energy conversion at the two photosystems is controlled by highly complex and dynamic adaptation processes triggered by external factors such as light quality, intensity, and duration, or internal cues such as carbon availability. These dynamics have remained largely concealed so far, because current analytical techniques are based on the investigation of isolated chloroplasts lacking full adaptation ability and are performed at non-physiologically low temperatures. Here, we use non-invasive *in planta* spectro-microscopic approaches to investigate living chloroplasts in their native environment at ambient temperatures. This is a valuable approach to study the complex function of these systems, because an intrinsic property—the fluorescence emission—is exploited and no additional external perturbations are introduced. Our analysis demonstrates a dynamic adjustment of not only the photosystemI/photosystemII (PSI/PSII) intensity ratio in the chloroplasts but also of the capacity of the LHCs for energy transfer in response to environmental and internal cues.

**Keywords: fluorescence, spectroscopy, photochemistry, chloroplast, photosystem, spectromicroscopy**

#### *†Present address:*

*Marcus Sackrow, Picoquant GmbH, Rudower Chaussee 29, 12489 Berlin, Germany*

## **INTRODUCTION**

Photosynthesis of plants still sets the benchmark in light-toenergy conversion regarding efficiency, stability and the ability to adapt to dynamic surrounding conditions. Photosynthesis is a highly complex process, involving multi-step energy migration within and between the two photosystems (PS) I and II, which are located in adaptable ratios in the grana and the stroma thylakoids of chloroplasts. The PSs are highly ordered functional and structural units that perform the primary photochemistry of photosynthesis: the absorption of light and the conversion of energy by charge separation. Especially the light harvesting complexes (LHCs) are responsible for the high efficiency of the whole process of photosynthesis due to the broad spectral absorption of ambient light by the LHCs, which is directed to the reaction center via a complex and efficient energy migration chain.

To optimize the use of available light-energy, higher plants undergo highly complex adaptation processes, which are mainly concerned with a structural rearrangement of the chloroplasts (Anderson et al., 1988, 2008; Anderson, 1999).The thylakoid membranes of chloroplasts, which are organized in stacked (grana thylakoids) and non-stacked (stromathylakoids) membranes, comprise different amounts of PSI and PSII. Ultrastructural studies of chloroplasts reveal an accumulation of PSII predominantly in the grana thylakoids, whereas PSI is found mainly in the stroma thylakoids (Anderson, 1986; Allen and Forsberg, 2001). Plants that experience changes in ambient light conditions offer an altered photosynthesis efficiency, which most likely is believed to be connected to a reorganization of the chloroplasts and, hence, a change in the PSI/PSII ratio (Anderson et al., 1988).

Accordingly, it is desirable to determine this ratio *in vivo* at ambient conditions as a measure of the plant ability to adapt its photosynthesic apparatus to the changing light environment. Non-invasive analyses of chloroplasts use the inherent autofluorescence of these compartments, which generally makes them accessible to optical studies. The emission spectrum of chloroplasts exhibits a maximum at 680 nm, which can be assigned to the fluorescence of the reaction center of PSII. Additionally, a spectral shoulder between 700 and 750 nm in the emission spectrum arising from the PSI reaction center is observed (Krause and Weis, 1991). However, a straightforward determination of the PSI/PSII ratio is hampered using conventional fluorescence microscopy due to the strong overlap of the spectral signatures of PSI and PSII. Hence, fluorescence spectroscopic measurements up to now have mostly been carried out at low temperatures (77 K and below) using isolated chloroplasts, where a differentiation between PSI and PSII is possible (Krause and Weis, 1991). However, similar to ultrastructural studies, plant adaptation processes in a living cell context remain obscured with these methods, since the composition of the photosynthetic apparatus reacts highly sensitive to sample treatments (Andersen et al., 1972). Furthermore, the influence of environmental and endogenous cues on the photosynthetic capacity cannot be explored using isolated chloroplasts. Here, we present two strategies using optical spectro-microscopy to overcome these limitations. These non-invasive techniques provide access to the composition of the photosynthesis apparatus and its adaptation to environmental changes at ambient conditions. To this end, we analyzed the PSI/PSII intensity ratio in chloroplasts at room temperature and in their cellular context by applying high spatial resolution fluorescence spectroscopy. We show that in a high-resolution fluorescence emission spectrum, the contributions of the distinct photosystems can be derived from the spectral shape, which depends on the relative contributions of PSI and PSII, respectively, and hence, the two subpopulations can be identified and compared. A careful statistical analysis of the room temperature emission spectra (SART) enables for monitoring global adaptations in the chloroplasts (i.e., the PSI/PS II intensity ratio).

Using conventional fluorescence-based techniques, however, only the final chromophore of an energy transfer cascade in the photosystems (**Figure 1A**) can be monitored, while the individual steps and composition of the energy transfer systems are not accessible (**Figure 1B**). This is because the energy absorbed by the LHCs is very efficiently funneled to chlorophyll *a* in the reaction centers as the last component of the energy migration chain (Butler and Kitajima, 1975) (**Figure 1A**). Accordingly, any detected fluorescence light originates from the emission of the reaction centers and information about the upstream energy cascade is not available (**Figure 1C**). The crystal structures of LHCs suggest a rigid, uniform assembly of these complexes (Liu et al., 2004).Variations are known to involve different supramolecular organizations of PSs and LHCs detected by electron microscopy (Yakushevska et al., 2001) as well as alterations in the total pigment amount and ratio in the chloroplasts (Bailey et al., 2004). Pigment analysis is usually based on the isolation of pigments using organic solvents followed by chromatographic separation. However, a technique that enables the observation of the photosynthetic efficiency *in vivo* has been missing so far. To overcome this restriction and get access to the composition of the energy transfer complexes *in vivo*, we established a novel *in planta* approach by disentangling the distinct energy transfer steps at ambient temperature. This was achieved by recording the fluorescence excitation spectra (FExS) in the diffraction limited focal spot by confocal fluorescence microscopy.