Reyer A et al. (2020), Channelrhodopsin-mediated optogenetics highligh...

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Proc Natl Acad Sci U S A 2020 Aug 25;11734:20920-20925. doi: 10.1073/pnas.2005626117.

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Channelrhodopsin-mediated optogenetics highlights a central role of depolarization-dependent plant proton pumps.

Reyer A , Häßler M , Scherzer S , Huang S , Pedersen JT , Al-Rascheid KAS , Bamberg E , Palmgren M , Dreyer I , Nagel G , Hedrich R , Becker D .

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In plants, environmental stressors trigger plasma membrane depolarizations. Being electrically interconnected via plasmodesmata, proper functional dissection of electrical signaling by electrophysiology is basically impossible. The green alga Chlamydomonas reinhardtii evolved blue light-excited channelrhodopsins (ChR1, 2) to navigate. When expressed in excitable nerve and muscle cells, ChRs can be used to control the membrane potential via illumination. In Arabidopsis plants, we used the algal ChR2-light switches as tools to stimulate plasmodesmata-interconnected photosynthetic cell networks by blue light and monitor the subsequent plasma membrane electrical responses. Blue-dependent stimulations of ChR2 expressing mesophyll cells, resting around -160 to -180 mV, reproducibly depolarized the membrane potential by 95 mV on average. Following excitation, mesophyll cells recovered their prestimulus potential not without transiently passing a hyperpolarization state. By combining optogenetics with voltage-sensing microelectrodes, we demonstrate that plant plasma membrane AHA-type H+-ATPase governs the gross repolarization process. AHA2 protein biochemistry and functional expression analysis in Xenopus oocytes indicates that the capacity of this H+ pump to recharge the membrane potential is rooted in its voltage- and pH-dependent functional anatomy. Thus, ChR2 optogenetics appears well suited to noninvasively expose plant cells to signal specific depolarization signatures. From the responses we learn about the molecular processes, plants employ to channel stress-associated membrane excitations into physiological responses.

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	Fig. 1. Blue light induces transient membrane potential depolarizations in Arabidopsis thaliana mesophyll cells. (A) Diagram showing the experimental setup for recording blue light-induced membrane potential changes in A. thaliana mesophyll cells. Note: Leaves were mounted with the adaxial side on the microscope slide and the abaxial epidermis up, facing the bath medium. (B) Representative trace of membrane potential recordings using microelectrode impalement of mesophyll cells in intact leaves in response to a 50-s blue light pulse. Upon impalement, typical membrane potentials of ca. −160 to −180 mV were recorded. Upon 50 s of illumination with blue light, fast membrane potential depolarizations to about −60 mV were triggered as a singular event. Membrane potential depolarizations were followed by fast repolarization and transient hyperpolarization, before cells returned to prestimulus membrane potential levels. The mean depolarization (ΔV) triggered by extended blue light pulses was about 101.2 ± 3.3 mV, while the observed hyperpolarization, relative to the recorded resting membrane potential was about −32.5 ± 2.3 mV (Inset). (C) No difference in kinetics is observed when comparing the blue light-induced depolarization under the same experimental conditions as in B between A. thaliana WT (Col-0) and the gork1-2 mutant, indicating that the mesophyll K+ efflux channel does not contribute to blue light-induced membrane potential changes. External solution was composed of 1 mM KCl, 1 mM CaCl2, 10 mM MES/BTP, pH 6.0. Blue light was applied via fiber optics at an intensity density of 17.5 mW·mm−2.
	Fig. 2. Blue light activates ChR2-XXL in tobacco mesophyll cells and Arabidopsis guard cells. (A) Representative voltage traces of blue light-induced depolarization in N. benthamiana mesophyll cells expressing YFP (control) or different ChR2 variants. A statistical analysis of the properties of selected ChR2 variants is presented in SI Appendix, Table S1 (n ≥ 4; mean ± SE). Note that blue light did not evoke depolarizations in control cells and that ChR2-mediated depolarizations depended on retinal substitution (SI Appendix, Fig. S3). (B) Guard cells in isolated epidermal strips were impaled with double-barreled electrodes, filled with 300 mM KOAc (pH 7.0). The plasma membrane potential was clamped at −100 mV and superimposed with a 10-s blue light stimulus. Blue light-stimulated photocurrents were recorded in guard cells expressing ChR2-XXL (n = 5) but not in controls (control, n = 6). The bath solution was composed of 1 mM KCl, 1 mM CaCl2, and 10 mM MES/BTP, pH 6. (C and D) External pH affects the ChR2-XXL current. Photocurrents of ChR2-XXL were recorded in guard cells under identical conditions as in B but at different pH values as indicated in the figure (mean ± SE, n ≥ 5).
	Fig. 3. ChR2-XXL provides for reproducible, blue light-triggered membrane potential depolarizations in Arabidopsis mesophyll cells. (A) Representative membrane potential recordings in response to 5-s blue light pulses using microelectrode impalement of mesophyll cells in intact Arabidopsis leaves expressing ChR2-XXL. Upon impalement, typical membrane potentials of ca. −160 to −190 mV were recorded. Repeated 5-s blue light-pulses triggered fast, reproducible depolarizations of the membrane potential. Depolarizations typically peaked at about −105 mV followed by fast repolarization and weak transient hyperpolarization before cells returned to prestimulus membrane potential levels. (B) Mean standardized membrane potential depolarizations of A. thaliana mesophyll cells expressing fully reconstituted (100 µM retinal) ChR2-XXL were 92 ± 2.9 mV, (n = 8, Inset) in response to a 5-s blue light-pulse (black curve). In the absence of retinal, mean depolarizations were only 14 ± 2.4 mV (red curve; n = 7, Inset). (C) Statistic analysis of the standardized membrane potential characteristics from 16 individual plants revealed a mean depolarization of 95 ± 2.3 mV with a minimum of 74 mV, a maximum of 108 mV and a median of 98 mV. Following ChR2-XXL–mediated depolarization, mesophyll cells repolarized toward their resting potential within 83 ± 4.5 s (minimum 51 s, maximum 107 s, and median 82 s). The associated repolarization speed was calculated to −1.2 ± 0.08 mV·s−1 (minimum −0.78 mV·s−1, maximum −1.69 mV·s−1, median −1.1 mV·s−1, and an outlier of −2.06 mV·s−1). (D) Representative membrane potential recordings of mesophyll cells in intact leaves in response to consecutive, extending blue light pulses (5, 10, 20, 2 × 50 s). Fast membrane potential depolarizations were observed for each pulse. 50-s blue light pulses, however, caused an additional depolarization (Fig. 1) on top of the ChR2-XXL–based depolarization followed by a pronounced hyperpolarization that was typically absent in response to short, 5-s blue light illumination.
	Fig. 4. Proton pump modulators affect repolarization kinetics. (A) Representative membrane potential recordings of impaled mesophyll cells preincubated with 1 mM Na3VO4 (vanadate, red curve). Traces represent individual depolarizations from a continuous recording cut out and superimposed for comparison. In comparison to control cells (black curve), vanadate inhibition of the PM H+-ATPase resulted in a positive shift of the prestimulus membrane potential (resting potential, see SI Appendix, Fig. S5C) and further delayed the repolarization phase by clamping cells in a depolarized state after blue light-induced ChR2-XXL activation. (B) The fungal toxin fusicoccin activates the PM H+-ATPase. Representative membrane potential recordings from ChR2-XXL–expressing mesophyll cells before (black line) and in response to 4 µM fusicoccin (blue line). The repolarization speed of ChR2-XXL–mediated membrane potential transients is significantly accelerated due to hyperstimulated PM H+-ATPase activity (see also SI Appendix, Fig. S6). (C) Functional expression of the PM H+-ATPase in Xenopus oocytes. H+ flux across the Xenopus oocyte plasma membrane was measured using the SISE technique. Oocytes were injected with water (black), AtAHA2 WT (light blue), or the aha2Δ73 mutant RNA (blue). After 5 d of expression, oocytes were voltage-clamped to −40 mV and H+ flux was recorded. Control oocytes exhibited proton influx under tested conditions. Expression of AHA2 WT resulted in significant reduction of net H+ flux and expression of the constitutively active mutant aha2Δ73 resulted in significant H+ efflux. Preincubation of aha2Δ73-expressing oocytes for 30 min in 10 mM vanadate resulted in significant reduction of H+ efflux (red). The bottom and top of the boxes denote the first and third quartiles, respectively, the middle line represents the median, whiskers mark the most extreme values within 1.5 times the interquartile distance below the first or above the third quartile, crosses indicate the first and 99th percentiles, and different letters indicate significantly different values: control-WT, P = 6.7 × 10−8; control-aha2Δ73, P = 6.7 × 10−8; control-aha2Δ73+vanadate, P = 9.2 × 10−8; WT-aha2Δ73, P = 8.8 × 10−8; WT-aha2Δ73+vanadate, P = 0.4; aha2Δ73-aha2Δ73+vanadate, P = 8.7 × 10−8; F3,48 = 502.8, one-way ANOVA, n ≥ 9 oocytes). (D) To determine voltage dependence of H+-pumping activity, H+ fluxes were measured in aha2Δ73-expressing oocytes and membrane potential was varied between −210 and +30 mV. Fitting the normalized fluxes with equation (A3; red line) resulted in half-maximal pump activity at −140 mV (mean ± SD, n ≥ 4).

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