Ion and Oxygen Fluxes in the Unicellular Alga Eremosphaera
viridis
Roger R. Lew*
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 lightinduced chloroplast translocation. Plasma membrane
fluxes were measured directly and non-invasively with selfreferencing 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.
Keywords: Chloroplast translocation • Eremosphaera viridis •
Ion fluxes • Oxygen fluxes • Photosynthesis • Unicellular alga.
Abbreviations: BBM,
dimethylsulfoxide.
Bold's
basal
medium;
DMSO,
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.
Regular Paper
Department of Biology, York University, Toronto, Ontario Canada
*Corresponding author: E-mail, [email protected]; Fax, +1-416-736-5698
(Received August 19, 2010; Accepted September 27, 2010)
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149, available online at www.pcp.oxfordjournals.org
© The Author 2010. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
All rights reserved. For permissions, please email: [email protected]
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
1889
R. R. Lew
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.
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
0 minutes
11 minutes
16 minutes
A
B
0
6
12
18
24
30
36
42
48 min
Fig. 1 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.
1890
Fig. 2 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.
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
Shining light on ion flux and chloroplast function
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).
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.
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
Chloroplast Translocation
nmole m–2 sec–1
1000
net
efflux
Oxygen Evolution
plus 0.2 mM KHCO3
8000 mmol
600
320 mmol m–2 s–1
200
-200
m–2 s–1
plus 0.2 mM KHCO3
dark
dark
4
net
influx
8
12
16
20
24
28
32
36
40
Time (min)
Fig. 3 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.
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
1891
R. R. Lew
800
160 mmol
160
480
600
onset of chlt
translocation
(−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).
O2
2160
8000
400
200
160
0
-200
1.0
600
bckgnd
Dark
160 Dark
bckgnd
H+
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 ionselective 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:
0.0
Flux (nmoles m–2 s–1)
-1.0
-2.0
10
Ca2+
0
J=a+
-10
-20
100
0
net
efflux
100
K+
200
700
-100
-200
-300
2500
net
influx
Cl–
1500
500
-500
-1500
10
-2500
20
30
40
50
160
Dark
Time (min)
Voltage (mV)
–10
–30
–50
Dark
–70
160 mmol
480
160
–90
2160 8000
Em
Fig. 4 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).
1892
Dual flux measurements reveal tight correlations
between H+ (but not Ca2+) and O2 fluxes in the
dark and light
J photon • J max
K 1 / 2 + J photon
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.
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
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
Shining light on ion flux and chloroplast function
light
dark
r1
1.0
r2
150
100
50
0
-50
Vmax = 137
K1/2 = 3.26
-100
-0.5
-1.0
buffered
(pH 6.8 ± 0.1)
non-buffered
P = 0.0482
-1.5
0.01
Vmax = –0.02
0.005
K1/2 = 4.08
-2.0
0
700
-0.005
600
-0.01
-0.015
-0.02
O2 Flux (nmoles m–2 s–1)
0.0
0.015
0
40
80
120
Photon Flux (µmoles m–2 s–1)
160
500
400
300
O2 Flux (nmoles m–2 s–1)
H+ Flux (nmoles m–2 s–1)
-150
buffered
net
influx
buffered
net
efflux
500
400
300
P = 0.4546
non-buffered
200
100
buffered
non-buffered
0
200
100
Vmax = 440
K1/2 = 3.20
0
-100
Ca2+ Flux (nmoles m–2 s–1)
P = 0.0045
0.5
H+ Flux (nmoles m–2 s–1)
O2 Flux (nmoles m–2 s–1)
r
net
efflux
non-buffered
(pH 6.9 ± 0.3)
0.6
0.4
0.2
0
-0.2
-0.4
0
40
80
120
160
Photon Flux (µmoles m–2 s–1)
200
Fig. 5 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).
-100
-200
net
influx
P = 0.0060
Fig. 6 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.
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
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
1893
R. R. Lew
effects of phosphate on O2 production (and the inversely
correlated net H+ flux) outweighed any masking of the flux
of H+ by buffers.
which is directly proportional to the change in chlorophyll
concentration:
r =110 mm
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 crosssectional 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,
1894
∫
r =100 mm
r = 70 mm
∫
4π r 2dr
= 2.6
2
4π r dr
r = 60 mm
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 lightdependent 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
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
Shining light on ion flux and chloroplast function
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 irradianceinduced 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 fastsetting 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
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
1895
R. R. Lew
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.
Data were recorded on a chart recorder (ionophoresis
experiments) or with an NI USB-6009 data acquisition unit
(electrical measurements) (National Instruments).
Fluorescein Ionophoresis
100 mV
A
B
C
D
5 minutes
Fluorescence Imaging
A
B
C
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
1896
D
Flouorescein
fluorescence
Color
merge
Chlorophyll
fluorescence
Fig. 7 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 Nafluorescein. 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.
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
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
Shining light on ion flux and chloroplast function
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-dimethyl3-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:
⎛ rr ⎞
4π D ⎜ 1 2 ⎟ (c2 − c 1)
⎝ ( r2 − r1 ) ⎠
J=
4π r 2
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.
Plant Cell Physiol. 51(11): 1889–1899 (2010) doi:10.1093/pcp/pcq149 © The Author 2010.
1897
R. R. Lew
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.
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