Effect of Redox‐Modifying Agents on the Activity of Channelrhodopsin‐2

Summary

Background

The algal protein Channelrhodopsin‐2 (ChR2) has been widely used in recent years in optogenetic technique to investigate the functions of complex neuronal networks through minimally invasive and temporally precise photostimulation of genetically defined neurons. However, as with any other new technique, current optogentic approaches have various limitations. In addition, how ChR2 may behave in response to complex biochemical changes associated with various physiological/pathological conditions is largely unknown.

Aim

In this study, we investigated whether a change in redox state of the cell affects the activity of ChR2 channels.

Methods

Whole‐cell patch‐clamp recordings were used to examine the effect of reducing and oxidizing agents on ChR2 currents activated by blue light.

Results

We show that the reducing agent dithiothreitol (DTT) dramatically potentiates the ChR2 currents in a reversible and concentration‐dependent manner. Glutathione, an endogenous reducing agent, shows a similar effect on ChR2 currents. The oxidizing agent 5,5′‐dithio‐bis‐(2‐nitrobenzoic acid) (DTNB) has no effect on ChR2 currents by itself; however, it completely reverses the potentiating effect of DTT. DTT also causes a shift in the current–voltage relationship by 23 ± 4.31 mV, suggesting a change in ion selectivity.

Conclusion

Taken together, these data suggest that redox modification of ChR2 plays an important role in its sensitivity to the light stimulation. Our findings not only help for a better understanding of how ChR2 may behave in physiological/pathological conditions where changes in redox state are common, but also provide a new direction for further optimization of this important opsin.

Introduction

Optogenetics is a recently developed novel technique in neuroscience field which combines optics and genetics to modulate the activity of specific populations of neurons 1. Channelrhodopsin‐2 (ChR2), a light‐activated cation‐selective channel, is the most widely used opsin and the critical component in the optogenetic technique since its first introduction into neurons 10 years ago 2, 3, 4. Stimulation of ChR2‐expressing neurons with blue light (bandwidth 450–490 nM) evokes membrane depolarization and action potential firing with high temporal precision on millisecond timescales 2. The optogenetic technique offers a promising approach to enable minimally invasive, genetically defined and temporally precise control of electrical activity of specific cell populations 5. It is a powerful technique for delineating complex neuronal network and for future treatment of various neuropsychiatric and neurological disorders.

However, as with any other new technique, current optogentic approaches have various limitations. In addition, how ChR2 may behave in response to complex biochemical changes associated with various physiological/pathological conditions is largely unknown. Further understanding the properties and modulations of this channel will help for future optimization of the system and increase the likelihood of using this technique for future clinical applications.

Changes in the redox state are associated with various physiological and pathological processes 6, 7. In various neurological disorders, for example, an increase in oxidative stress is a common feature 6, 8. Changes in the redox states have been shown to affect the properties and functions of some ion channels such as NMDA and ASICs 9, 10. In this regard, we determined whether a change in the redox state affects the activity/function of ChR2 channels.

We show that reducing agent DTT dramatically potentiates the ChR2 currents expressed in Chinese hamster ovary (CHO) while the addition of oxidizing agent DTNB reverses the effect of DTT. These findings suggest that redox state plays an important role in determining the activity/function of ChR2. The current studies will help for a better understanding of how ChR2 may behave in physiological/pathological conditions accompanied by changes in the redox state. Our findings also suggest a new direction for further optimization of this important opsin.

Methods

Culture of CHO Cells and ChR2 Transfection

Chinese hamster ovary (CHO) cells were cultured and seeded onto culture dishes (35 mm in diameter) at a density of 1.25 × 10⁵ cells/mL, as described previously 11. Cells were used for transfection 24 h after plating. pCAGGS‐ChR2‐Venus, a gift from Karel Svoboda 12 (Addgene plasmid # 15753), was used. Transient transfection was performed using 1 μg pCAGGS‐ChR2‐Venus, 100 μL Opti‐MEM, 3 μL Fugene 6 Transfection Reagent (Promega, Madison, WI, USA) for each dish (2 mL media) and cultured for 36–48 h before experiments. GFP‐positive cells were chosen for recording.

Electrophysiology

Whole‐cell voltage‐clamp recordings were performed as described 13, 14. Patch electrodes were fabricated from borosilicate capillary tubing of 1.5 mm in diameter (WPI) using a vertical puller (PP‐83, Narishige, Amityville, NY, USA) and had resistances of 3–4 MΩ when filled with electrode solution (see below). Currents were recorded using Axopatch 200B amplifier with pCLAMP 10 software (Molecular Devices, Sunnyvale, CA, USA). They were filtered at 2 kHz and digitized at 5 kHz using Digidata 1322A. Data were eliminated from statistical analysis when access resistance was >10 MΩ or leak current was >100 pA at −60 mV. A multibarrel perfusion system (SF‐77B, Warner Instruments, Hamden, CT, USA) was used to achieve a rapid exchange of external solutions. All experiments were performed at room temperature.

Solutions and Chemicals

Standard ECF contained (in mM): 140 NaCl, 5.4 KCl, 2 CaCl₂, 1 MgCl₂, 20 HEPES, 10 glucose (pH 7.4 adjusted with NaOH; 320–335 mOsm). Patch electrodes contained (in mM): 140 CsF, 10 HEPES, 1 CaCl₂, 11 EGTA, 2 TEA, 4 Mg‐ATP (pH 7.3 adjusted with CsOH, 290–300 mOsm). L‐Glutathione (reduced form, GSH) DL‐Dithiothreitol (DTT) and 5,5′‐dithiobis(2‐nitrobenzoic acid) (DTNB) were purchased from Sigma‐Aldrich (St. Louis, MO, USA). A stock solution of 1M DTT was prepared in distilled water before dilution to the bath solutions. GSH and DTNB were directly dissolved in bath solutions. All the solutions were prepared freshly and used within 1 day.

Light Stimulation

ChR2 currents were activated by blue light generated with the optogenetic kit (Thor Labs). The tip of a fiber optic was placed within 0.3 mm from the cells of interest. The intensity of light was detected with a digital handheld optical power meter (Thor Labs, Newton, NJ, USA).

Statistical Analysis

Data are presented as mean ± SEM. Student's t‐test and ANOVA were used for statistical analysis where appropriate. P‐value of <0.05 was considered significant.

Results

Reducing Agents Potentiate ChR2 Currents

To determine whether redox state modulates the activity of ChR2, we first examined the effect of reducing agent DTT on ChR2 currents. 36–48 h following transfection, whole‐cell patch‐clamp recording was performed in GFP‐positive CHO cells. Following the establishment of whole‐cell configuration and recording of stable ChR2 currents by repeated light stimulations, bath solution containing 1 mM DTT was perfused to the cell for 5–10 min. As shown in Fig. 1A, a large potentiation of both the peak and sustained current components was observed after perfusion of DTT. At 1 mM, DTT increased the peak current to 1.88 ± 0.57 fold of control (n = 10, Fig. 1B). The maximal potentiation can be reached after perfusion of DTT for ~5 min (Fig. 1C). However, it took up to 20 min for the effect of DTT to be completely reversed after washout (n = 6; Fig. 1C).

Figure 1.

Potentiation of ChR2 currents by DTT. (A) Representative traces and (B) bar graphs showing the potentiation of ChR2 currents by 1 mM DTT in a reversible manner. n = 10, **P < 0.01 compared with control, paired t‐test. (C) Time course of the potentiation by DTT (1 mM). DTT was applied as indicated after at least three stable current traces were obtained (n = 6).

We next studied the dose–response relationship. We examined the effects of different concentrations of DTT (0.03–10 mM) on ChR2 currents. As indicated in Fig. 2A and 2B, ChR2 currents were potentiated by DTT in a concentration‐dependent manner (DTT, 0.1 mM, 1.14 ± 0.46; 0.3 mM, 1.37 ± 0.43; 1 mM 1.55 ± 0.18; 3 mM, 1.91 ± 0.22; 10 mM, 2.20 ± 0.27; n = 6). A small but significant potentiation was observed when 0.1 mM DTT was applied to the cell for ~5 min (Fig. 2A). At 10 mM, DTT increased the peak amplitude of ChR2 currents to 2.20 ± 0.27 fold of control (n = 6; Fig. 2B). As the concentration above 10 mM caused unstable in membrane seal, we were unable to record the maximal effect of DTT (data not shown).

Figure 2.

Concentration‐dependent potentiation of ChR2 currents by DTT. (A) Representative current traces and (B) bar graphs showing the concentration‐dependent potentiation of ChR2 currents by DTT; n = 6, *P < 0.05 and **P < 0.01 compared with control.

We next tested the effect of endogenous reducing agents GSH. Perfusion of 1 mM GSH for 5 min increased the peak amplitude of the ChR2 current to 1.34 ± 0.04 of control value, and this effect was also reversible (n = 6; Fig. 3A and 3B). Together, these data suggest that there exist redox modulatory sites on ChR2 or closely associated protein, which can be modulated by reducing agents, resulting in increased ChR2 currents.

Figure 3.

Potentiation of ChR2 currents by GSH. (A) Representative traces and (B) bar graphs showing the potentiation of ChR2 currents by GSH (1 mM). n = 6, **P < 0.01.

Oxidizing Agent DTNB Abolishes the Potentiation of ChR2 Currents by DTT

We further examined whether oxidizing agents have an opposite effect on ChR2 currents. After recording a stable ChR2 current, cells were perfused with DTNB (1 mM) for as long as 10 min. However, no significant change in ChR2 currents by DTNB was observed (the relative peak amplitude of the ChR2 current after 1 mM DTNB was 0.99 ± 0.02 of control, n = 7; Fig. 4A and 4B). Interestingly, the addition of DTNB completely reversed the potentiating effect of DTT on ChR2 currents within 10 min of DTNB addition (the relative peak amplitude was reduced from 1.78 ± 0.18 to 1.01 ± 0.14‐fold of control; n = 6, Fig. 4C–E). These data suggest that, in the resting condition before DTT, ChR2 or its closely associated protein might be already in an oxidized state.

Figure 4.

Oxidizing agent DTNB reverses the potentiation of ChR2 currents by DTT. (A) Representative traces and (B) bar graph showing the effect of 1 mM DTNB on ChR2 currents (n = 7). (C) Representative traces and (D) bar graph showing that the potentiation of ChR2 currents by DTT was reversed by addition of DTNB (1 mM). (E) Time course showing that the persistent potentiation by DTT was quickly reversed by addition of DTNB (1 mM). n = 6, *P < 0.05.

Effect of DTT on the Light Intensity—Current Response Curve of ChR2

We then investigated whether DTT changes the light intensity–current response curve of ChR2. As shown in Fig. 5, the amplitude of ChR2 currents gradually increases as the light intensity was increased from 0.46 to 20.65 μW/mm². In the presence of 1 mM DTT, a significant potentiation of ChR2 currents was observed at all light intensities tested. There was no significant change in EC₅₀ value (1.01 ± 0.15 μW/mm² in the absence of DTT and 1.24 ± 0.18 μW/mm² in the presence of DTT, n = 8, P = 0.32). The relative degree of potentiation was independent of the light intensity (n = 8; Fig. 5C).

Figure 5.

The effect of DTT on light intensity–current response curve of ChR2. (A) Representative traces and bar graphs showing ChR2 currents activated by different intensity of light ranging from 0.46 to 20.65 μW/mm², in the absence or presence of 1 mM DTT. (B) Light intensity–response curves inferred from A (n = 8). The peak current amplitude was normalized to the one activated by the maximal intensity of light. (C) The ratios show the potentiation of ChR2 peak currents by 1 mM DTT under variable light intensities.

DTT Shifts the Current–Voltage Relationship of ChR2

Finally, we examined whether DTT has any effect on the current‐voltage relationship of ChR2. ChR2 currents were recorded at variable holding potentials from −80 mV to +60 mV (Fig. 6A). In the absence of DTT, the reversal potential of ChR2 currents was at −0.44 ± 3.81 mV, consistent with a nonselective cation channel. Application of 1 mM DTT significantly shifted the reversal potential to 23 ± 4.31 mV (Fig 6B, n = 5), suggesting that DTT may alter the ion selectivity of ChR2.

Figure 6.

The effect of DTT on the current–voltage relationship of ChR2. (A) Representative traces and (B) summary data showing current–voltage relationships in the absence or presence of 1 mM DTT. ChR2 currents were activated at various holding potentials ranging from −80 mV to 60 mV, in the absence or presence of 1 mM DTT. The peak amplitude of all the currents was normalized to the one at −80 mV of the control. (n = 5, P < 0.01).

Discussion

In the present study, we provided the evidence suggesting that the activity of ChR2 channels can be dramatically influenced by the redox state of the cell.

Since optogenetics was introduced into the field of neuroscience 10 years ago 2, it has been widely employed to achieve a better understanding of the neural circuits mediating normal behavior as well as dysfunctions in the neural circuit underlying neurological/psychiatric disorders such as mood disorders, addiction, and Parkinson's disease4, 15.

As with any other new technique, limitations with optogenetics exist. Despite that the employment of wild‐type ChR2 in optogenetic experiments is satisfactory in some studies, limitations of the technique such as insufficient level of expression, potential phototoxicity, and heating of the tissues are considered 16. In addition, how ChR2 may behave in response to complex biochemical changes associated with physiological and pathological conditions is largely unclear. Therefore, understanding the properties and modulations of these channels and development of more light‐sensitive ChR2 variants will be beneficial for further optimizing the optogenetic system and improving chances of using the technique for successful clinical applications 4. Our current finding that ChR2 currents are potentiated by reducing agent suggests a potential new direction for improving the light sensitivity of ChR2, that is, by modification of redox state of the protein.

ChR2 is a light‐activated nonselective cation channel, which has permeability for H⁺, Na⁺, and Ca²⁺ 3, 17. An increase in the amplitude of ChR2 current along with a positive shift in reversal potential by DTT suggests that there might be an increase in the permeability of ChR2 to Na⁺ or Ca²⁺ by reducing agents. Further experiment with ion substitution assay is needed to identify the change in permeability to specific ions.

Multiple membrane proteins including voltage‐ and ligand‐gated ion channels are modulated by redox status 9, 18, 19. For instance, N‐methyl‐d‐aspartate receptor (NMDA) and acid sensing ion channels (ASICs) are potentiated by reducing agents but inhibited by oxidizing agents 9, 19. Similar to their effects on NMDA receptors and ASICs, reducing agents potentiate the activity of ChR2. Unlike NMDA receptors and ASICs, however, oxidizing agent (DTNB) does not have a clear effect on ChR2. It is likely that the cysteines on ChR2 or its closely associated proteins involved in redox modulation are already in the oxidized state under our recording condition. Our studies suggest that under neurological conditions where there is a shift in the redox states toward more oxidized states, the activity of ChR2 will be largely suppressed thus a higher light intensity will be required to activate the channels.

Variable membrane proteins are known to be modulated by reducing/oxidizing agents, and the sulfur moiety in cysteine residues in those proteins is considered to be a primary target for redox modification 18, 19, although other residues including methionine, tyrosine, phenylalanine, histidine, and lysine may also be involved 9. Redox‐modulating sites may exist in the extracellular, transmembrane, or intracellular domains of membrane proteins. Our finding that the membrane‐impermeable reducing agent GSH has a clear effect on ChR2 suggests that at least some redox‐modulating sites might be located on the extracellular domains of the channel. Further experiment with mutation of the extracellular cysteines may help identify the site(s) involved in redox modulation of ChR2.

In summary, we demonstrate, for the first time, that redox state plays an important role in modulating the activity of ChR2 channels. The current findings not only help for a better understanding of how ChR2 may behave in vivo where redox states change with different physiological and pathological conditions, but also suggest a new direction for future development of more light‐sensitive and efficient opsins.

Conflict of Interest

The authors declare no conflicts of interest.