TY - JOUR
AU - Lew, Roger R.
AB - Abstract Plasma membrane fluxes of the large unicellular model algal cell Eremosphaera viridis (De Bary) were measured under various light regimes to explore the role of plasma membrane fluxes during photosynthesis and high light-induced chloroplast translocation. Plasma membrane fluxes were measured directly and non-invasively with self-referencing ion-selective (H+, Ca2+, K+ and Cl−) potentiometric microelectrodes and oxygen amperometric microelectrodes. At light irradiances high enough to induce chloroplast migration from the cell periphery to its center, oxygen evolution declined to respiratory net O2 uptake prior to any significant chloroplast translocation, while net K+ and Cl− influx increased during the decline in photosynthetic activity (and the membrane potential depolarized). The results suggest that chloroplast translocation is not the cause of the cessation of O2 evolution at high irradiance. Rather, the chloroplast translocation may play a protective role: shielding the centrally located nucleus from damaging light intensities. At both high and low light intensities (similar to ambient growth conditions), there was a strong inverse correlation between H+ net fluxes and respiratory and photosynthetic net O2 fluxes. A similar inverse relationship was also observed for Ca2+ net fluxes, but only at higher light intensities. The net H+ fluxes are small relative to the buffering capacity of the cell, but are clearly related to both photosynthetic and respiratory activity. Introduction Fluxes at the plasma membrane are crucial to cellular life. Many obligatory cellular functions rely upon the uptake of nutrients from the surrounding extracellular environs and excretion of waste by-products. Osmotic balancing often requires the transport of ions and uncharged solutes into and/or out of the cell. Even cellular signaling may rely upon the availability of ions, such as Ca2+, in the extracellular solution. In all of these processes and others, the fluxes at the plasma membrane must be integrated with biochemical processes within the cell. Probably the best example of an integrative process in biological organisms is the interplay of photosynthesis and respiration with organellar, interorganellar and plasma membrane fluxes. To explore this integrative physiology, the unicellular Chlorophyte Eremosphaera viridis (De Bary) offers many advantages because of its large size and simple spherical geometry. It is amenable to intracellular measurements of electrical properties of the plasma membrane (see Kohler et al. 1983, Kohler et al. 1985, Kohler et al. 1986, Sauer et al. 1993, Bauer et al. 1999) and vacuolar membrane (see Linz and Köhler 1994, Bethmann et al. 1995), measurements of H2O transport (Lawaczeck, 1988), carbon transport (as CO2 and/or HCO3−) (Gimmler et al. 1990, Rotatore et al. 1992), excretion of small organic acids (Stabenau and Winkler, 2005), and turgor (Frey et al. 1988). It also allows in-depth exploration of pH regulation, which is of overriding importance to the long-term survival of cells that exist in solutions of varying acidity or alkalinity and under the pH changes caused by photosynthesis (Bethmann et al. 1998, Bethmann and Schönknecht, 2009). We undertook an integrated exploration of fluxes at the plasma membrane of E. viridis during photosynthesis using non-invasive self-referencing probes. This technique measures the external steady-state diffusive gradient created by solute transport across the plasma membrane by sampling the solute concentrations at two positions—one near to and one far from the cell—with a microprobe; the flux can be calculated from the difference in concentration (Smith et al. 1999). The potentiometric microprobe was used to measure net H+, K+, Ca2+ and Cl− fluxes. Oxygen was measured using an amperometric microprobe (Smith et al. 2007). Because of the non-invasive nature of the flux measurements, it is possible to measure multiple aspects of cellular physiology simultaneously without damaging the cell. We used this integrative approach to explore two light-related phenomena in E. viridis: plasma membrane fluxes during chloroplast translocations at high light irradiances, and net ion fluxes during photosynthesis at near ambient light intensities. Results Two basic questions were explored: the first was how O2 and ion fluxes are related to chloroplast translocation to the center of the cell in response to high irradiance. The second objective was to explore the relationship between H+ and Ca2+ fluxes and photosynthetic activity (O2 fluxes) at low light intensities, to confirm the tight correlations observed at high light intensities and to assess the role of plasma membrane net H+ fluxes in pH regulation during photosynthesis. Oxygen and ion fluxes during induction of chloroplast translocation Preliminary experiments characterized the light dependence of chloroplast translocation in E. viridis. This served to both confirm and extend previous reports by Weidinger (1982) and Weidinger and Ruppel (1985). A ×10 objective was used to maximize the number of cells that could be observed in the field of view. Actinic light intensity was varied from 1 to 10,000 μmol m−2 s−1; the duration of irradiations was 60 min. Both ‘incipient systrophe’ (in which some of the chloroplasts had moved towards the center of the cell) and ‘complete systrophe’ (in which all chloroplasts had moved from the cell periphery to the center) were scored every 2 min. ‘Incipient systrophe’ was observed at light intensities that ranged from 200 to 2,000 μmol m−2 s−1. ‘Complete systrophe’ was observed at light intensities >2,000 μmol m−2 s−1. Chloroplast translocation was not observed when red light (either 610 ± 5 nm or 650 ± 5 nm light) was used (at 200 μmol m−2 s−1), but blue light (467 ± 5 nm) was very effective (about 65% of the cells exhibited ‘incipient systrophe’ after 60 min at 200 μmol m−2 s−1). The time required to reach maximal chloroplast translocation was about 10–30 min, dependent on the intensity of light. Recovery—the return of chloroplasts to the cell periphery—at low light intensity required about 40 min. Examples of high-irradiance-induced chloroplast translocation are shown in Fig. 1. The response is limited to regions of irradiation (Fig. 1B). In dual imaging of mitochondria and chloroplasts (Fig. 2), close association of the organelles is commonly observed. However, only chloroplasts translocate from the outer peripheral shell of the cell to the center, surrounding the nucleus located at the center of the cell (Moore, 1901); mitochondria remain in the periphery of the cell. Fig. 1 View largeDownload slide Examples of chloroplast translocations after high light irradiation of Eremosphaera viridis cells. The actinic light irradiation was about 8,000 μmol m−2 s−1 for the durations shown. Images were captured with a color digital camera (Infinity2-1C, Lumenera Ltd.). (A) Two examples are shown that represent the range of response, from relatively little chloroplast translocation (upper panel) to the movement of the majority of chloroplasts to the center of the cell (lower panel). (B) To demonstrate the effect of localized irradiation, only half of the cell was irradiated [the light barrier was removed for the first (time 0) and last (48 min) irradiations to show the localized chloroplast translocation]. The nucleus is located in the center of the cell. Bars = 20 μm. Fig. 1 View largeDownload slide Examples of chloroplast translocations after high light irradiation of Eremosphaera viridis cells. The actinic light irradiation was about 8,000 μmol m−2 s−1 for the durations shown. Images were captured with a color digital camera (Infinity2-1C, Lumenera Ltd.). (A) Two examples are shown that represent the range of response, from relatively little chloroplast translocation (upper panel) to the movement of the majority of chloroplasts to the center of the cell (lower panel). (B) To demonstrate the effect of localized irradiation, only half of the cell was irradiated [the light barrier was removed for the first (time 0) and last (48 min) irradiations to show the localized chloroplast translocation]. The nucleus is located in the center of the cell. Bars = 20 μm. Fig. 2 View largeDownload slide Disparate response of chloroplasts and mitochondria during chloroplast translocation in Eremosphaera viridis. The left panels show imaging of mitochondria with MitoFluoGreen and the right panels show chloroplasts (Chl autofluorescence). In confocal imaging, it was common to observe some association of chloroplasts and mitochondria (since they co-localize to cytoplasmic strands and the peripheral cytoplasmic shell of the cell). Upon irradiation with high intensity light (about 8,000 μmol m−2 s−1, for the times shown), the chloroplasts translocate to the center of the cell (surrounding the nucleus), but mitochondria remain in the peripheral cytoplasmic shell. Bar = 40 μm. Fig. 2 View largeDownload slide Disparate response of chloroplasts and mitochondria during chloroplast translocation in Eremosphaera viridis. The left panels show imaging of mitochondria with MitoFluoGreen and the right panels show chloroplasts (Chl autofluorescence). In confocal imaging, it was common to observe some association of chloroplasts and mitochondria (since they co-localize to cytoplasmic strands and the peripheral cytoplasmic shell of the cell). Upon irradiation with high intensity light (about 8,000 μmol m−2 s−1, for the times shown), the chloroplasts translocate to the center of the cell (surrounding the nucleus), but mitochondria remain in the peripheral cytoplasmic shell. Bar = 40 μm. Preliminary measurements of O2 evolution from individual cells showed high rates of net O2 efflux at 320 μmol m−2 s−1. The magnitude of the net efflux was increased about 3-fold by the addition of KHCO3. This is evidence that inorganic carbon availability for carbon dioxide fixation limits the rate of the light reactions of photosynthesis. In the dark, there was a net influx of O2, consistent with respiratory activity. Subsequent high intensity irradiation (8,000 μmol m−2 s−1) inhibited net O2 efflux. Although some chloroplast translocation could be observed during high intensity irradiation (consistent with a scoring of ‘incipient systrophe’), ‘complete systrophe’ was not observed even though net O2 efflux was near zero, or even reversed to become net influx. At this time, the addition of KHCO3 was without effect, indicating that the inhibition of the light reactions was not due to depletion of carbon for carbon dioxide fixation. Respiration (net O2 influx) in the dark was unaffected by the high light intensity treatment (Fig. 3). Fig. 3 View largeDownload slide Example of oxygen evolution and its relationship to chloroplast translocation in Eremosphaera viridis. The extent of chloroplast translocation (upper panel) is shown at various times (min:s) (bar = 40 μm) during measurements of oxygen evolution (lower panel). In this experiment, KHCO3 was added to demonstrate that oxygen evolution is limited by carbon availability for CO2 fixation. At high irradiance (8,000 μmol m−2 s−1), oxygen evolution is arrested even though only limited chloroplast translocation has occurred. The inhibition of oxygen evolution was not relieved by KHCO3 addition. Fig. 3 View largeDownload slide Example of oxygen evolution and its relationship to chloroplast translocation in Eremosphaera viridis. The extent of chloroplast translocation (upper panel) is shown at various times (min:s) (bar = 40 μm) during measurements of oxygen evolution (lower panel). In this experiment, KHCO3 was added to demonstrate that oxygen evolution is limited by carbon availability for CO2 fixation. At high irradiance (8,000 μmol m−2 s−1), oxygen evolution is arrested even though only limited chloroplast translocation has occurred. The inhibition of oxygen evolution was not relieved by KHCO3 addition. Based upon these preliminary experiments of the relationship between O2 evolution and chloroplast translocation, a light protocol was designed that first examined the effect of light intensities about 3-fold greater than the normal light regime during cell culturing (160 μmol m−2 s−1) to determine ‘normal’ maximal O2 evolution rates. As an internal control, two identical light treatments were interspersed with a dark treatment to ensure that net O2 influx due to respiration was occurring. Then, step-wise increases in light intensity (with interspersed background measurements) were applied, culminating in a high intensity irradiation of 8,000 μmol m−2 s−1 to induce chloroplast translocation rapidly and completely. This was followed by irradiations at the original intensity of 160 μmol m−2 s−1 interrupted by a dark treatment to assess the competence of photosynthetic activity at normal light intensities expected to support maximal oxygen evolution, and the effect of the high intensity irradiation on respiratory activity, both after high intensity irradiation. The same protocol was performed during measurements of H+, Ca2+, K+ and Cl− fluxes, as well as measurements of the cell electrical potential. Digital images of the cell (usually 12) were captured throughout the protocol to assess the extent of chloroplast translocation. The flux data (and onset of chloroplast translocation) are compiled in Fig. 4. Fig. 4 View largeDownload slide Summary of oxygen (O2) and ion (H+, Ca2+, K+, Cl−) fluxes, and cell electrical potential (Em) in Eremosphaera viridis. The protocol was designed to examine fluxes under conditions of maximal oxygen evolution, and to examine the effect of high irradiance on inhibition of oxygen evolution and plasma membrane fluxes in the context of chloroplast translocation. Data are shown vs. time [with intervals added between each treatment to ease visualization and because measurements were interrupted for brief (but varying) periods of time to change neutral density filters]. Both individual experiments (gray lines) (except Em measurements) and means (open circles) are shown. To estimate ion concentration changes (nM min−1), the fluxes (nmol m−2 s−1) can be multiplied by 3,300 (assuming a cell radius of 55 μm and that cytoplasmic and organellar compartments contribute equally). Fig. 4 View largeDownload slide Summary of oxygen (O2) and ion (H+, Ca2+, K+, Cl−) fluxes, and cell electrical potential (Em) in Eremosphaera viridis. The protocol was designed to examine fluxes under conditions of maximal oxygen evolution, and to examine the effect of high irradiance on inhibition of oxygen evolution and plasma membrane fluxes in the context of chloroplast translocation. Data are shown vs. time [with intervals added between each treatment to ease visualization and because measurements were interrupted for brief (but varying) periods of time to change neutral density filters]. Both individual experiments (gray lines) (except Em measurements) and means (open circles) are shown. To estimate ion concentration changes (nM min−1), the fluxes (nmol m−2 s−1) can be multiplied by 3,300 (assuming a cell radius of 55 μm and that cytoplasmic and organellar compartments contribute equally). As noted in the preliminary characterization (Figs. 1, 2), the onset of chloroplast translocation occurs after the onset of the high intensity irradiation (Fig. 4), but usually does not progress very far, even though photosynthetic activity has become completely inhibited. The changes in net H+ and Ca2+ fluxes were a mirror image of O2 fluxes. That is, there was net influx of either ion when net O2 efflux was occurring, and net efflux of the ions when net O2 influx was occurring. The relative changes in fluxes were similar under all light treatments, and in the dark. Thus, net H+ and Ca2+ fluxes were tightly correlated with net O2 fluxes under all conditions. We were unable to resolve a fast transient change in Ca2+ flux, which may play a role in the signaling pathway for induction of chloroplast translocation. Although net K+ flux exhibited a similar mirroring with net O2 fluxes at low light intensities, when the cell was subjected to high intensity irradiation, there was an increase in net K+ influx (rather than the decline observed for net H+ and Ca2+ influx). Net Cl− influx was highest during the low light treatments at the end of the protocol. Using the same experimental protocol, the plasma membrane potential (Em) was measured. Digitized data were first averaged for successive 10 s intervals (n = 40), then averaged for all experiments (five independent experiments). At the initial light intensities (160 μmol m−2 s−1) interspersed with a dark treatment, the potential was hyperpolarized in the light compared with the dark. Subsequent high intensity irradiation (480 μmol m−2 s−1) caused a progressive depolarization of the potential, even before the maximal irradiation (8,000 μmol m−2 s−1) was applied. Respiration (net O2 influx in the dark) was unaffected by high light irradiation. The net O2 influx before (−56.4 ± 23.1 nmol m−2 s−1, n = 4) was the same as after high light irradiation (−59.8 ± 27.8 nmol m−2 s−1, n = 4) (two-tailed t-test P = 0.83). Dual flux measurements reveal tight correlations between H+ (but not Ca2+) and O2 fluxes in the dark and light The unexpectedly strong correlation between net H+ and Ca2+ fluxes and net O2 fluxes was explored in greater detail at low light intensities that were similar to the ambient light conditions of algal growth. To measure dual fluxes, the ion- selective probe was positioned on one side of the cell and the O2-selective probe on the other side (Fig. 5). Light intensities were adjusted by varying both the lamp output and the neutral density filters. Concurrent net fluxes were plotted vs. photon flux; best fits are with a hyperbolic function:   where Jphoton is the photon flux, Jmax is the maximal net ion or oxygen flux, K1/2 is the photon flux at which net ion or oxygen flux (J) is half-maximal, and the parameter ‘a’ accounts for the reversal of net fluxes. The inverse relationship of net O2 and H+ fluxes is very strong, exhibiting a very similar dependence on the photon flux. For net O2 flux, the K1/2 was 3.26 ± 1.816 μmol m−2 s−1; for net H+ flux, it was 4.08 ± 1.96 μmol m−2 s−1 (means ± SE) (Fig. 5). The photon flux at which the net O2 and H+ fluxes reversed was also very similar (mean ± SE): 0.803 ± 0.272 μmol m−2 s−1 for net O2 flux and 1.151 ± 0.267 μmol m−2 s−1 for net H+ flux. In the case of O2 fluxes, this is the point where respiratory O2 consumption and photosynthetic O2 production are equal. Finally, a linear regression analysis of the paired O2 and H+ flux measurements yielded a very high correlation coefficient (r2 = 0.759) (data not shown). In two experiments examining the effect of KHCO3 addition, the average net O2 efflux nearly doubled (from 348 to 569 nmol m−2 s−1), as did net H+ influx (from −0.017 to −0.027 nmol m−2 s−1). Therefore, there appears to be a strong relationship between oxygen utilization and production and net H+ fluxes at the plasma membrane in E. viridis in a complete growth mediun. Fig. 5 View largeDownload slide Light dependence of oxygen and H+ and Ca2+ fluxes in Eremosphaera viridis. Concurrent measurements of O2 and ion fluxes was made using dual probes as shown in the photograph (bar = 40 μm) overlaid with the measuring parameters for calculation of fluxes for the H+ microprobe (top panel) (see Materials and Methods). Best fits were to the equation: flux = a + [(Jphoton × Jmax)/(K1/2 + Jphoton)]. There is a strong correlation between net O2 flux and net H+ flux when fit with the hyperbolic function [the K1/2 values for photon flux (Jphoton) were very similar: 3.26 ± 1.81 and 4.08 ± 1.96 μmol m−2 s−1, respectively (mean ± SE)] (middle panels). In addition, when net O2 flux is inward (i.e. respiration exceeds photosynthesis), net H+ flux is outward. Conversely, when net O2 flux is outward (photosynthesis exceeds respiration), net H+ flux is inward. A similar correlation was not observed for Ca2+ fluxes at these relatively low light intensities (lower panels) (compared with the chloroplast translocation protocol, Fig. 4). Fig. 5 View largeDownload slide Light dependence of oxygen and H+ and Ca2+ fluxes in Eremosphaera viridis. Concurrent measurements of O2 and ion fluxes was made using dual probes as shown in the photograph (bar = 40 μm) overlaid with the measuring parameters for calculation of fluxes for the H+ microprobe (top panel) (see Materials and Methods). Best fits were to the equation: flux = a + [(Jphoton × Jmax)/(K1/2 + Jphoton)]. There is a strong correlation between net O2 flux and net H+ flux when fit with the hyperbolic function [the K1/2 values for photon flux (Jphoton) were very similar: 3.26 ± 1.81 and 4.08 ± 1.96 μmol m−2 s−1, respectively (mean ± SE)] (middle panels). In addition, when net O2 flux is inward (i.e. respiration exceeds photosynthesis), net H+ flux is outward. Conversely, when net O2 flux is outward (photosynthesis exceeds respiration), net H+ flux is inward. A similar correlation was not observed for Ca2+ fluxes at these relatively low light intensities (lower panels) (compared with the chloroplast translocation protocol, Fig. 4). In contrast, although there was an apparent relationship between O2 and Ca2+ net fluxes observed during the chloroplast translocation protocol (Fig. 4), the relationship was not strong enough to be seen at low light intensities (Fig. 5), for which the correlation coefficient was very low in linear regression analysis of the paired O2 and Ca2+ flux measurements (r2 = 0.015, data not shown). H+ fluxes in unbuffered media Measurements of H+ fluxes are complicated by the presence of buffers, which can transport H+ in their protonated state as a ‘hidden’ contribution to the total net H+ flux (Arif et al. 1995, Messerli et al. 2006). Bold's basal medium (BBM) is buffered with KPi at pH 6.8. At this pH, not only will phosphate act as a carrier of H+, but so will HCO3− in equilibrium with air levels of CO2. At pH 6.8, [HCO3−] is expected to be about 32 μM, based on a pKa of 6.351 (Goldberg et al. 2002). To test directly for an effect of buffering, experiments were performed in BBM lacking KPi and EDTA. The pH of the cell suspension in modified BBM was about 4.2. This was adjusted to 6.9 ± 0.3 by the addition of aliquots of 1 N NaOH (or 1 N HCl to adjust for overshoots). Dual O2 and H+ flux measurements were performed as described (see Fig. 5). Net O2 and H+ fluxes in the light were higher in buffered media (Fig. 6), suggesting that the beneficial effects of phosphate on O2 production (and the inversely correlated net H+ flux) outweighed any masking of the flux of H+ by buffers. Fig. 6 View largeDownload slide H+ and oxygen net fluxes in buffered and unbuffered solutions at pH 6.8. Concurrent measurements of net H+ and O2 fluxes were made in buffered (circles) and non-buffered solutions (without phosphate) (triangles) at pH 6.8. The net fluxes in normal buffered and non-buffered media were compared to test for attenuation of H+ fluxes by diffusion of H+ complexed with phosphate buffer. There was no indication that measurements of H+ flux were attenuated due to the presence of buffer. Instead, fluxes in the light were significantly higher; H+ fluxes in the dark were only slightly higher in unbuffered solution. Individual data, means ± SD and two-tailed t-tests comparisons are shown. Fig. 6 View largeDownload slide H+ and oxygen net fluxes in buffered and unbuffered solutions at pH 6.8. Concurrent measurements of net H+ and O2 fluxes were made in buffered (circles) and non-buffered solutions (without phosphate) (triangles) at pH 6.8. The net fluxes in normal buffered and non-buffered media were compared to test for attenuation of H+ fluxes by diffusion of H+ complexed with phosphate buffer. There was no indication that measurements of H+ flux were attenuated due to the presence of buffer. Instead, fluxes in the light were significantly higher; H+ fluxes in the dark were only slightly higher in unbuffered solution. Individual data, means ± SD and two-tailed t-tests comparisons are shown. Discussion The experimental approach was to create an integrated picture of physiological plasma membrane fluxes at the level of a single cell during light-mediated phenomena: (i) to explore in situ the process of chloroplast translocation; and (ii) to characterize correlative fluxes during photosynthesis at near ambient light levels. The self-referencing potentiometric and amperometric probes are very useful in this regard. They are non-invasive, and many of the measurements could be done concurrently, allowing for paired data analyses. The major result of our exploration of chloroplast translocation was unexpected. The translocation of chloroplasts to the center of the cell in response to high light irradiance occurred after photosynthetic activity has ceased, as measured by net O2 production. It is natural to assume that chloroplast translocation to the center of the cell functions to minimize light absorption and therefore to minimize photosynthetic activity (negative phototaxis) (Kasahara et al. 2002, Li et al. 2009). However, in retrospect, besides chloroplast movement, protection from photo-oxidative damage due to high light irradiance can involve a number of mechanisms that ‘quench’ the excessive number of excitons (Niyogi 1999, Müller et al. 2001). In E. viridis, when the chloroplasts translocate to the center of the cell, there are two effects on the efficiency of light absorption. One is the decrease in the cross-sectional target area resulting in lower absorption efficiency; the other is the increase in Chl density, resulting in higher absorption efficiency. The inter-relationship can be explored with a simplified model. Assuming all chloroplasts are located in a peripheral shell [ignoring chloroplasts that are located in cytoplasmic strands that extend from the center of the cell (where the nucleus is located; Moore 1901) to the periphery, see Figs. 1 and 2], when chloroplasts translocate to the center, the cross-sectional target area will decrease to about 0.40-fold. This estimate is based on the ratio of cross-sectional areas after and before translocation: [π(70 μm)2]/[π(110 μm)2]. This decrease in cross-sectional area is countered by the increase in absorptive efficiency. When the chloroplasts translocate to the center, their density is higher, resulting in a higher Chl concentration. The increase in light absorption depends on the Beer–Lambert law (A = εlc; where A is the absorbance, ε is the molar extinction coefficient, l is the path length and c is the concentration). Measuring the change in path length is not straightforward, although it probably increases when the chloroplasts translocate to the center (see Figs. 1 and 2), possibly 2-fold. Ignoring the path length for the sake of simplicity, and assuming that the chloroplasts reside in a shell of width 10 μm at the periphery or when they surround the nucleus in the center of the cell, the relative change in absorption efficiency can be estimated from the change in volume, which is directly proportional to the change in chlorophyll concentration:   The decrease in cross-sectional area (0.40-fold) and increase in Chl concentration (2.6-fold) results in virtually no change in the absorptive efficiency (1.05). So the effect on absorptive efficiency is slight, and O2 production declines before significant chloroplast translocation occurs anyway. Chloroplast translocation is a dramatic and clear cellular response to high light irradiance. If it plays only a small role in alleviating the effect of high photon flux on photosynthetic activity, what is its function? One possibility is to protect the nucleus from high light intensities, effectively shading it from UV light that would be present at high irradiances of normal sunlight. At low light intensities—closer to the ambient conditions used to culture cells—oxygen fluxes, either net production in photosynthesis or consumption in respiration, correlate with plasma membrane H+ fluxes under ionic conditions that are similar to those the alga would normally reside in. This raises the possibility that plasma membrane H+ fluxes may play a role in pH regulation. The effects of light/dark transitions (that would affect photosynthetic electron transport) on cytoplasmic and vacuolar pH have been well characterized by Bethmann et al. (1998) and pH regulation after intracellular acidification by Bethmann and Schönknecht (2009). The basic observation is a marked transient (2–4 min) cytoplasmic acidification (and slight vacuolar acidification) upon a transition from light to dark and transient cytoplasmic alkalinization in response to a dark/light transition (Bethmann et al. 1998). The initial light/dark/light treatments in the chloroplast translocation protocol (Fig. 4) and light- dependent fluxes in Fig. 5 have a similar time course. In the light when photosynthesis is active, there is a net H+ influx at the plasma membrane; in the dark, it changes to net efflux (fluxes were in the steady state, no transient was observed). Whether the net efflux would play a role in alleviating dark-induced cytoplasmic acidification can be assessed by calculating cellular [H+] changes. For the range of net H+ fluxes observed at moderate light irradiance (±0.015 nmol m−2 s−1, Fig. 5), assuming a cell radius of 55 μm, the concentration changes inside the cell would be ±49 nM min−1. Thus, the contribution of H+ fluxes at the plasma membrane to pH regulation would be small compared with the buffering capacity of the vacuole (10 mM per pH unit) (Bethmann et al. 1998) and cytoplasm (20–90 mM per pH unit) (Plieth et al. 1997, Bethmann and Schonknecht 2009). In addition to net H+ fluxes, other ion fluxes change during the light/dark/light transition. Both Ca2+ and K+ net influx declines in response to dark, and may contribute to the plasma membrane potential. If only CO2 is actively taken up by the cell (Rotatore et al. 1992), replenishment from extracellular HCO3− would result in extracellular H+ consumption (H+ + HCO3− → H2O + CO2) at the plasma membrane, contributing to the apparent net influx of H+ in the light. In the dark, CO2 produced by respiration would result in acidification (H2O + CO2 → HCO3− + H+) and apparent H+ net efflux from the cell. A 1 : 1 stoichiometry of H+ : O2 fluxes was reported in Chlamydomonas reinhardi (Neumann and Levine, 1971), but much lower ratios were observed in Spirogyra grevilleana (Porterfield and Smith, 2000). In E. viridis as well, the magnitude of the H+ fluxes is much lower than that of O2 net fluxes. It is unlikely that H+ net flux is ‘masked’ by extracellular buffers, since the removal of phosphate buffer causes lower H+ net fluxes in the light. Long-term, net H+ influx in the light and efflux in the dark are consistent with circadian changes in extracellular pH, exemplified by research on the model organism for biological rhythms, the dinoflagellate Gonyaulax polydra (Eisensamer and Roenneberg 2000). The plasma membrane H+-ATPase may play a role in the net H+ fluxes. However, the plasma membrane potential is relatively insensitive to external pH (Köhler et al. 1985, our unpublished data). Elevated CO2 (5%), which would increase photosynthetic activity, hyperpolarizes the potential in both the light and dark, indicating that its effects are independent of photosynthesis (Rotatore et al. 1992). No change in the potential is observed when CO2 treatments of ‘0’ and ambient air are used (also expected to affect photosynthetic activity) (Deveau et al. 2001). Thus, other transport mechanisms beside the plasma membrane H+-ATPase may contribute more to plasma membrane net H+ fluxes. In summary, a suite of solute fluxes has been measured in the context of ‘normal’ photosynthesis and high irradiance-induced chloroplast translocation. Oxygen production declines well before extensive chloroplast translocation has occurred. Net fluxes of all ions respond to the varying light treatments. Of these, H+ net fluxes exhibit the strongest correlation with photosynthetic activity but do not play a significant role in rapid regulation of cytoplasmic pH. Correlations between net ion fluxes and O2 net fluxes are weaker for Ca2+, but trend similarly to net H+ flux. Materials and Methods Strains Stock cultures of the alga E. viridis De Bary (CPCC 127) were obtained from the Canadian Phycological Culture Centre (University of Waterloo, Waterloo, Canada). The strain used in the present study was isolated from Plastic Lake, Ontario in 1987, a small lake that had a pH of 5.8 in 1981 and continued to acidify in subsequent years (at a rate of 0.035 pH units year−1) (Dillon et al. 1987). Other strains in culture collections are isolates from bogs (and soil) [Sammlung von Algenkulturen (SAG) Culture Collection of Algae, University of Gottingen, Germany]. Therefore, the species is commonly considered to be an acidophile; optimal growth of the CPCC 127 strain occurs at pH 5–7 (data not shown). The cultures were grown in 125 or 250 ml Ehrlenmeyer flasks mounted on an orbital shaker (125 r.p.m.) under 50 μmol m−2 s−1 photon flux from T8 fluorescent lamps (6,500 K color temperature) at room temperature with serial transfer every 5–10 d. The culture medium was BBM [http://www.phycol.ca/media (Nichols and Bold 1965)] supplemented with vitamins (thiamine-HCl, 0.5 μg ml−1; vitamin B12, 0.01 μg ml−1; and biotin, 0.005 μg ml−1). The major ions in BBM are (in meq l−1): Na+ (5.96), Cl− (3.17), NO3− (2.94), K+ (2.26), Pi (1.72), SO42− (0.32), Mg2+ (0.30) and Ca2+ (0.17); pH is ∼6.8. Culture preparation for experiments Cells from 1- to 2-week-old cultures were transferred to the lid of 35 mm culture dishes (1 ml of cells and 2 ml of fresh BBM). Cells were often observed as doublets, still attached by discarded cell walls; thus the cells were actively dividing and the cultures presumed to be in log phase [in separate growth experiments, the doubling time of this large alga was about 3 d at pH 5–7; very little growth occurred at more acidic (pH 3–4) or alkaline (pH 8–9) conditions]. For some experiments examining the effect of buffer on the H+ fluxes, 1 ml aliquots of cells were washed twice in 10–12 ml of unbuffered BBM (without K·PO4 or Na2·EDTA), before being transferred to the culture dish lid. Buffer was excluded to avoid the artifact of H+ transport by protonated buffer (Arif et al. 1995). Even in the absence of added buffer, H2O and carbonates from dissolved CO2 may transport H+. At pH <6.0, the effect of bicarbonate buffering is negligible (∼1%) (Arif et al. 1995). At pH 6.8, the effect of bicarbonate buffering should be small: [HCO3−] is about 38 μM, higher than [H+] (0.16 μM), but much lower than the buffering capacity in normal BBM with a phosphate concentration of 1.72 mM. For electrophysiology and flux measurements, cells of 100–120 μm were selected. For electrophysiology measurements, the cell was lifted from the bottom of the dish and held with a suction micropipet (inner diameter of ∼100 μm). Electrical measurement of the algal cells The algal cell was impaled with a double barrel micropipet. The fabrication and use of double barrel micropipets have been described by Lew (2006). Briefly, two borosilicate glass capillaries (OD 1.0 mm, ID 0.58 mm, with an internal filament; Friedrich and Dimmock Inc.) were placed in modified holders on a Sutter P-30 puller (Sutter Instruments Co.), heated, twisted 360° and then pulled. The twisted joint was strengthened with fast-setting epoxy, and one barrel was heated and pulled away to form a ‘y’ shape, separating the two barrels behind the micropipet tip. The micropipet barrels were filled with 3 M KCl. The reference electrode was a salt bridge containing 3 M KCl in 2% (w/v) agar connected to an Ag/AgCl electrode. Successful demonstration of impalement was confirmed by injecting current through one barrel, and observing a voltage deflection in the other barrel; no voltage deflection was observed before the cell was impaled, or after the micropipet was removed from the cell. Preliminary experiments were performed to determine the location of the micropipet tip after impalement, by imaging the distribution of Na-fluorescein after ionophoresis into the cell. The ionophoretic micropipet was loaded with 200 mg ml−1 Na-fluorescein in 100 mM KCl. Both micropipet barrels were backfilled with 3 M KCl. The confocal fluorescence imaging (Bio-Rad MRC-600) was performed on a Nikon microscope with a ×40 water immersion objective. Dual excitation (with a mixed gas krypton/argon laser) was used to image the fluorescein (488 nm excitation, bandpass emission, 500–560 nm) and Chl autofluorescence (568 nm excitation, longpass emission, 585 nm) of the chloroplasts. An example of an experiment is shown in Fig. 7. Fluorescence ‘cross-talk’ between the two excitation–emission channels was not observed. Ionophoretic injection (using a −1 nA current) of fluorescein through one micropipet caused a voltage deflection in the other micropipet, and movement of dye into the cell, which was distributed around the periphery of the cell near the impalement site, and around the chloroplasts (which are embedded in cytoplasm). Thus the micropipet tip is located in the cytoplasm. Fig. 7 View largeDownload slide Cytoplasmic location of the micropipet tip after impalement of Eremosphaera viridis cells. The upper panel (fluorescein ionophoresis) shows the electrical trace of the voltage-monitoring micropipet: impalement, followed by three ionophoretic injections (−1 nA) through the current-injecting micropipet containing 200 mM Na-fluorescein. The lower panel (Fluorescence imaging) shows medial section images taken at the times marked: (A) (90 s after impalement), before ionophoretic injection of the fluorescein; (B) (140 s), after the first ionophoretic injection; (C) (190 s), after the second ionophoretic injection; and (D) (250 s), after the third injection. The left column shows the fluorescein fluorescence and the right column shows Chl fluorescence. The middle column shows color merges (green, fluorescein; red, Chl) of the two images. Regions of co-localization of the fluorescein and Chl fluorescence are due to cytoplasm surrounding the chloroplasts. Fig. 7 View largeDownload slide Cytoplasmic location of the micropipet tip after impalement of Eremosphaera viridis cells. The upper panel (fluorescein ionophoresis) shows the electrical trace of the voltage-monitoring micropipet: impalement, followed by three ionophoretic injections (−1 nA) through the current-injecting micropipet containing 200 mM Na-fluorescein. The lower panel (Fluorescence imaging) shows medial section images taken at the times marked: (A) (90 s after impalement), before ionophoretic injection of the fluorescein; (B) (140 s), after the first ionophoretic injection; (C) (190 s), after the second ionophoretic injection; and (D) (250 s), after the third injection. The left column shows the fluorescein fluorescence and the right column shows Chl fluorescence. The middle column shows color merges (green, fluorescein; red, Chl) of the two images. Regions of co-localization of the fluorescein and Chl fluorescence are due to cytoplasm surrounding the chloroplasts. Data were recorded on a chart recorder (ionophoresis experiments) or with an NI USB-6009 data acquisition unit (electrical measurements) (National Instruments). Light treatments of the algal cells A variety of techniques were used to irradiate the cells on the specimen stage of the microscope. In all cases, irradiation was through the microscope condenser. Under Kohler illumination, the irradiated field was adjusted with the microscope diaphragm to fill the field of view under a ×10 objective, and the area calculated from the diameter of the field. The light intensity was measured with a radiometric probe [Model 268R with an internal radiometric filter (maximal light responsivity between 400 and 1,000 nm) attached to a Model S471 portable optometer; UDT Instruments]. The probe was placed above the condenser on the specimen stage where the cells were to be located. In all instances, the light source was a 50 W tungsten–halogen lamp, powered by an external DC power supply or with the internal power supply on the microscope. For some preliminary spectral experiments, interference filters were used (467 ± 4 nm, 610 ± 4 nm or 650 ± 4 nm; Optometrics Corporation). For ‘actinic’ light experiments, light output from the lamp was filtered through a 3.5% (w/v) cupric sulfate filter in a flat culture flask (depth 1.65 cm). This provided light >50% of maximal photon flux (at 565 nm) between 450 and 630 nm. The radiometer probe measures energy output in Watts. This was converted to a photon flux by assuming the average photon energy was 4.014 × 10−19 J photon−1. For light experiments characterizing chloroplast translocations (and O2 and ion fluxes), the light intensities were adjusted by insertion of neutral density filters into the light path, keeping the lamp output constant at 43 W. To explore the light intensity dependence of O2 evolution and ion fluxes (H+ and Ca2+), the light intensities were adjusted by modifying the lamp output in concert with the use of neutral density filters. The lamp wattages used were 7, 10, 12.75 and 17 W. This will result in a significant spectral shift of the light output. The different lamp outputs were attenuated with combinations of neutral density filters (attenuation factors of 4, 16 or 64). For different lamp outputs (7, 10 and 12.75 W) and neutral density attenuations (none, 4 or 16) that provided similar irradiances (6.6–16.9 μW), there was no statistically significant difference in O2 production. Since a 3.5% (w/v) cupric sulfate filter was used, it was assumed that the effect of spectral differences caused by varying the lamp output were insignificant compared with the effect of photon flux. Ion and oxygen flux meaurements The ion-selective probes used to measure net ion fluxes have been described in detail (Lew 1999, Lew et al. 2007). Fabrication and measuring protocols were those normally used at the BioCurrents Facility (Marine Biological Laboratory) (Messerli et al. 2006). The ion-selective ionophores (Sigma-Aldrich Corp.) and cocktails were: H+ [Hydrogen Ionophore I–Cocktail B, Cat. No. 95293 (pH 5.5–12.0)], Ca2+ (Calcium Ionophore I–Cocktail A, Cat. No. 21048), K+ {Potassium Ionophore I–Cocktail A, Cat. No. 60398 [modified by replacing 25% (w/w) 1,2-dimethyl-3-nitrobenzene with 25% (w/w) 2-nitrophenyl octyl ether (Messerli et al. 2006)]} and Cl− [Chloride Ionophore II, Cat. No. 24901 (2% w/w) and 0.03% (w/w) tridodecylmethylammonium chloride in ortho-nitrophenyloctylether (Messerli et al. 2008)]. In all cases, the ion-selective probes were calibrated before and after measurements at the cell in at least three concentrations of the ion that bracketed the normal concentration of the ion in BBM (calibrations were very similar before and after experiments). For the ion-selective microelectrodes, calibrations were H+ (59.9 ± 0.7 mV log[H+]−1, r2 = 0.997 ± 0.003, n = 13), Ca2+ (34.5 ± 2.9 mV log[Ca2+]−1, r2 = 0.999 ± 0.001, n = 6), Cl− (46.6 ± 5.4 mV log[Cl−]−1, r2 = 0.978 ± 0.007, n = 6) and K+ (55.9 ± 0.7 mV log[K+]−1, r2 = 0.999 ± 0.0003, n = 3). During measurement at the algal cell, the concentration of the selected ion was measured as near as possible to the algal cell wall and 20 μm away at a frequency of about 0.3 Hz (digital images of dual probe measurements are shown in Fig. 5). Background measurements were performed about 400 μm above the cell. Oxygen electrode design and fabrication at the BioCurrents Research Center has been described by Jung et al. (2000) and Smith et al. (2007); the measuring technique is amperometric. At a bias voltage of −0.6 V, the current generated from the reaction O2 + 4e− + 2H2O → 4 OH− is proportional to [O2]. Calibrations were performed in air-saturated BBM ([O2] of 280 μM) and N2-bubbled BBM ([O2] of 0 M). When dual measurements of either H+ or Ca2+ fluxes and O2 fluxes were performed, to avoid changes in the ion-selective probe output (μV) in response to simultaneous amperometric measurements of O2 evolution, two reference electrodes were used (both filled with 3 M KCl in 2% agar), one connected to the potentiometric probe amplifier, the other connected to the amperometric probe amplifier. To avoid interference between H+ sensing by the H+ probe and OH− generation by the O2 probe, the two probes were positioned on opposite sides of the cell (and thus separated by about 120 μm). Kunkel et al. (2006) have tested for interference between O2 and H+ probes using an artificial point source, and report no interference when the probes are positioned 40 μm apart. Furthermore, similar correlations between O2 and H+ net fluxes were observed when measurements were performed in separate experiments (see Fig. 4), or in dual measurements (see Fig. 5). Thus, interference between the two probes should not occur in the experimental set-up for net flux measurements in E. viridis. The time required to change the light treatment was <1 min; fluxes were at or near steady state during the flux measurements (duration of about 2.7 min). Ion fluxes were calculated from the ion concentration differences taking into account the spherical geometry of the algal cell, for which the flux, J (mol m−2 s−1), is:   where D is the diffusion coefficient (H+, 9.31 × 10−9 m2 s−1; Ca2+, 0.4 × 10−9 m2 s−1; K+, 1.96 × 10−9 m2 s−1; Cl−, 2.03 × 10−9 m2 s−1; O2, 2.51 × 10−9 m2 s−1), r is the cell radius, c2 and c1 are the concentrations at the two excursion points, r2 and r1 are the distances from the algal cell center to the two excursion points, and 4πr2 is the area of the cell. Outward net fluxes (efflux) are shown as positive fluxes; inward net fluxes (influx) are shown as negative fluxes. Microscopy and digital imaging Either a Nikon Optiphot or a Zeiss Axioskop microscope was used, normally with water immersion ×40 objectives. Most digital imaging was done with a color video camera (Model ZVS-47E, Carl Zeiss Inc.) and a screen-grabber controlled with the IonView software program developed at the BioCurrents Research Center. The resulting .avi files were imported into ImageJ (Rasband 2009) for further analysis. An Olympus Fluoview 300 confocal system with Fluoview software was used for confocal fluorescence imaging of chloroplasts (Chl autofluorescence) and mitochondria (MitoFluoGreen). Cells (1 ml) at log phase were added to 3 ml of BBM containing 2 μl of MitoFluoGreen (Molecular Probes) [from a 2 mM stock in dimethylsufoxide (DMSO); chloroplast distribution was unaffected by addition of the DMSO solvent alone]. Cells were mounted on a microscope slide in BBM, covered with a coverslip, and imaged with a ×60 oil immersion objective (infinity tube length). A multiargon laser (excitation line 488 nm) and He–Ne laser (excitation line 633 nm) were used for visualization of MitoFluoGreen (bandpass emission, 505–535 nm) and Chl autofluorescence (longpass emission, 660 nm), respectively. Z-sections (16 slices, 0.3 μm steps) were obtained, interspersed with brightfield illumination at high light intensity (about 8,000 μmol m−2 s−1) to induce chloroplast translocation. Z-projections were created from the Z-sections in ImageJ (Rasband 2009) and are shown without further processing. Statistical analysis Statistics are shown as mean ± SD (sample size) (SD was used as an estimator of population variance) unless stated otherwise. Independent two-tailed or one-tailed t-tests were performed in Excel (Microsoft). Non-linear regressions were performed in Kaleidagraph (Synergy Software). Funding This research was supported by the Natural Sciences and Engineering Research Council of Canada [Discovery Grant]; York University [sabbatical research support]. Acknowledgements Special thanks to Peter J. S. Smith, Director, and other members of the BioCurrents Research Facility, Marine Biological Laboratory, Woods Hole for their hospitality. Undergraduate Biophysics students performed some of the experimental characterization of chloroplast translocations [Robert Moscaritolo and Fidan (Dana) Gasumova], and pH dependence of growth and membrane potential experiments (Sandra Khine) as RAY (Research at York) summer research assistants. Abbreviations Abbreviations BBM Bold's basal medium DMSO dimethylsulfoxide. References Arif I,  Newman IA,  Keenlyside N.  Proton flux measurements from tissues in buffered solutions,  Plant Cell Environ. ,  1995, vol.  18 (pg.  1319- 1324) Google Scholar CrossRef Search ADS   Bauer CS,  Simonis W,  Schönknecht G.  Different xanthines cause membrane potential oscillations in a unicellular green alga pointing to a ryanodine/cADPR receptor Ca2+ channel,  Plant Cell Physiol. ,  1999, vol.  40 (pg.  453- 456) Google Scholar CrossRef Search ADS   Bethmann B,  Schönknecht G.  pH regulation in an acidophilic green alga—a quantitative analysis,  New Phytol. ,  2009, vol.  183 (pg.  327- 339) Google Scholar CrossRef Search ADS PubMed  Bethmann B,  Thaler M,  Simonis W,  Schönknecht G.  Electrochemical potential gradients of H+, K+, Ca2+ and Cl− across the tonoplast of the green alga Eremosphaera viridis,  Plant Physiol. ,  1995, vol.  109 (pg.  1317- 1326) Google Scholar CrossRef Search ADS PubMed  Bethmann B,  Simonis W,  Schönknecht G.  Light-induced changes of cytosolic pH in Eremosphaera viridis: recordings and kinetic analysis,  Plant Physiol. ,  1998, vol.  49 (pg.  1129- 1137) Deveau JST,  Lew RR,  Colman B.  Evidence for active CO2 uptake by a CO2-ATPase in the acidophilic green alga Eremosphaera viridis,  Can. J. Bot. ,  2001, vol.  79 (pg.  1274- 1281) Dillon PJ,  Reid RA,  de Grosbois E.  The rate of acidification of aquatic ecosystems in Ontario, Canada,  Nature ,  1987, vol.  329 (pg.  45- 48) Google Scholar CrossRef Search ADS   Eisensamer B,  Roenneberg T.  Extracellular pH is under circadian control in Gonyaulax polyedra and forms a metabolic feedback loop,  Chronobiol. Int. ,  2000, vol.  21 (pg.  27- 41) Google Scholar CrossRef Search ADS   Frey N,  Buchner K.-H,  Zimmerman U.  Water transport properties and regulatory processes in Eremosphaera viridis,  J. Membr. Biol. ,  1988, vol.  101 (pg.  151- 163) Google Scholar CrossRef Search ADS   Gimmler H,  Weiss C,  Baier M,  Hartung W.  The conductance of the plasmalemma for CO2,  J. Exp. Bot. ,  1990, vol.  41 (pg.  785- 795) Google Scholar CrossRef Search ADS   Goldberg RN,  Kishore N,  Lennen RM.  Thermodynamic quantities for the ionization reactions of buffers,  J. Phys. Chem. Ref. Data ,  2002, vol.  31 (pg.  231- 370) Google Scholar CrossRef Search ADS   Jung S.-K,  Hammar K,  Smith PJS.  Development of self-referencing oxygen micro-sensor and its application to single pancreatic HIT cells: effects of adenylate cyclase activator forskolin on oxygen consumption,  Biol. Bull. ,  2000, vol.  199 (pg.  197- 198) Google Scholar CrossRef Search ADS PubMed  Kasahara M,  Kagawa T,  Oikawa K,  Suetsugu N,  Miyao M.,  Wada M.  Chloroplast avoidance movement reduces photodamage in plants,  Nature ,  2002, vol.  420 (pg.  829- 832) Google Scholar CrossRef Search ADS PubMed  Köhler K,  Geisweid H-J,  Simonis W,  Urbach W.  Changes in membrane potential and resistance caused by transient increase of potassium conductance in the unicellular green alga Eremosphaera viridis,  Planta ,  1983, vol.  159 (pg.  165- 171) Google Scholar CrossRef Search ADS PubMed  Köhler K,  Steigner W,  Simonis W,  Urbach W.  Potassium channels in Eremosphaera viridis. I. Influence of cations and pH on resting membrane potential and on an action-potential-like response,  Planta ,  1985, vol.  166 (pg.  490- 499) Google Scholar CrossRef Search ADS PubMed  Köhler K,  Steigner W,  Kolbowski J,  Hansen U.-P,  Simonis W,  Urbach W.  Potassium channels in Eremosphaera viridis. II. Current- and voltage-clamp experiments,  Planta ,  1986, vol.  167 (pg.  66- 75) Google Scholar CrossRef Search ADS PubMed  Kunkel JG,  Cordeiro S,  Xu Y,  Shipley AM,  Feijo JA.  Volkov AG.  Use of non-invasive ion-selective microelectrode techniques for the study of plant development,  Plant Electrophysiology—Theory and Methods ,  2006 Berlin Springer(pg.  109- 137) Lawaczeck R.  First direct observation of water exchange across the membrane of a single-cell green alga on the cellular level,  Biochim. Biophys. Acta ,  1988, vol.  946 (pg.  165- 172) Google Scholar CrossRef Search ADS PubMed  Lew RR.  Comparative analysis of Ca2+ and H+ flux magnitude and location along growing hyphae of Saprolegnia ferax and Neurospora crassa,  Eur. J. Cell Biol. ,  1999, vol.  78 (pg.  892- 902) Google Scholar CrossRef Search ADS PubMed  Lew RR.  Volkov AG.  Use of double barrel micropipettes to voltage-clamp plant and fungal cells,  Plant Electrophysiology—Theory and Methods ,  2006 Berlin Springer(pg.  139- 154) Lew RR.  Ionic currents and ion fluxes in Neurospora crassa hyphae,  J. Exp. Bot. ,  2007, vol.  58 (pg.  3475- 3481) Google Scholar CrossRef Search ADS PubMed  Li Z,  Wakao S,  Fischer BB,  Niyogi KK.  Sensing and responding to excess light,  Annu. Rev. Plant Biol. ,  2009, vol.  60 (pg.  239- 260) Google Scholar CrossRef Search ADS PubMed  Linz KW,  Köhler K.  Vacuolar ion currents in the primitive green alga Eremosphaera viridis: the electrical properties are suggestive of both the Characeae and higher plants,  Protoplasma ,  1994, vol.  179 (pg.  34- 45) Google Scholar CrossRef Search ADS   Messerli MA,  Kurtz I,  Smith PJS.  Characterization of optimized Na+ and Cl− liquid membranes for use with extracellular, self-referencing microelectrodes,  Anal. Bioanal. Chem. ,  2008, vol.  390 (pg.  1355- 1359) Google Scholar CrossRef Search ADS PubMed  Messerli MA,  Robinson KR,  Smith PJS.  Volkov AG.  Electrochemical sensor applications to the study of molecular physiology and analyte flux in plants,  Plant Electrophysiology—Theory and Methods ,  2006 Berlin Springer(pg.  73- 107) Moore GT.  New or little known unicellular algae. II. Eremosphaera viridis and Excentrosphaera,  Bot. Gaz. ,  1901, vol.  32 (pg.  309- 324) Google Scholar CrossRef Search ADS   Müller P,  Li X.-P,  Niyogi KK.  Non-photochemical quenching: a response to excess light energy,  Plant Physiol. ,  2001, vol.  125 (pg.  1558- 1566) Google Scholar CrossRef Search ADS PubMed  Neumann J,  Levine RP.  Reversible pH changes in cells of Chlamydomonas reinhardi resulting from CO2 fixation in the light and its evolution in the dark,  Plant Physiol. ,  1971, vol.  47 (pg.  700- 704) Google Scholar CrossRef Search ADS PubMed  Nichols HW,  Bold HC.  Trichosarcina polymorpha Gen. et Sp. Nov.,  J. Phycol. ,  1965, vol.  1 (pg.  34- 38) Google Scholar CrossRef Search ADS   Niyogi KK.  Photoprotection revisited: genetic and molecular approaches,  Annu. Rev. Plant Physiol. Plant Mol. Biol. ,  1999, vol.  50 (pg.  333- 359) Google Scholar CrossRef Search ADS PubMed  Plieth C,  Sattelmacher B,  Hansen U.-P.  Cytoplasmic Ca2+–H+-exchange buffers in green algae,  Protoplasma ,  1997, vol.  198 (pg.  107- 124) Google Scholar CrossRef Search ADS   Porterfield DM,  Smith PJS.  Single-cell, real-time measurements of extracellular oxygen and proton fluxes from Spirogyra grevilleans,  Protoplasma ,  2000, vol.  212 (pg.  80- 88) Google Scholar CrossRef Search ADS   Rasband WS. ,  2009  ImageJ. U.S. National Institutes of Health, Bethesda, Maryland, USA, http://rsb.info.nih.gov/ij/, 1997–2009 Rotatore C,  Lew RR,  Colman B.  Active uptake of CO2 during photosynthesis in the green alga Eremosphaera viridis is mediated by a CO2-ATPase,  Planta ,  1992, vol.  188 (pg.  539- 545) Google Scholar CrossRef Search ADS PubMed  Sauer G,  Simonis W,  Schonknecht G.  An inwardly rectifying cation current across the plasma membrane of the green alga Eremosphaera viridis,  Plant Cell Physiol. ,  1993, vol.  34 (pg.  1275- 1282) Smith PJS,  Hammar K,  Porterfield DM,  Sanger RH,  Trimarchi JR.  Self-referencing, non-invasive, ion selective electrode for single cell detection of trans-plasma membrane calcium flux,  Microsc. Res. Tech. ,  1999, vol.  46 (pg.  398- 417) Google Scholar CrossRef Search ADS PubMed  Smith PJS,  Sanger RS,  Messerli MA.  Michael AC,  Borland LM.  Principles, development and applications of self-referencing electrochemical microelectrodes to the determination of fluxes at cell membranes,  Electrochemical Methods for Neuroscience ,  2007 Boca Raton, FL CRC Press(pg.  373- 405) Stabenau H,  Winkler U.  Glycolate metabolism in green algae,  Physiol. Plant ,  2005, vol.  123 (pg.  235- 245) Google Scholar CrossRef Search ADS   Weidinger M.  The inhibition of systrophe in different organisms,  Protoplasma ,  1982, vol.  110 (pg.  71- 74) Google Scholar CrossRef Search ADS   Weidinger M,  Ruppel HG.  Ca2+-requirement for a blue-light-induced chloroplast translocation in Eremosphaera viridis,  Protoplasma ,  1985, vol.  124 (pg.  184- 187) Google Scholar CrossRef Search ADS   © The Author 2010. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: journals.permissions@oxfordjournals.org
TI - Ion and Oxygen Fluxes in the Unicellular Alga Eremosphaera viridis
JF - Plant and Cell Physiology
DO - 10.1093/pcp/pcq149
DA - 2010-10-05
UR - https://www.deepdyve.com/lp/oxford-university-press/ion-and-oxygen-fluxes-in-the-unicellular-alga-eremosphaera-viridis-cATVi1h0GK
SP - 1889
EP - 1899
VL - 51
IS - 11
DP - DeepDyve
ER -