
==== Front
Plant Physiol
Plant Physiol
plphys
Plant Physiology
0032-0889
1532-2548
Oxford University Press US

38701198
10.1093/plphys/kiae241
kiae241
Research Article
Membranes, Transport, and Bioenergetics
AcademicSubjects/SCI01270
AcademicSubjects/SCI01280
AcademicSubjects/SCI02286
AcademicSubjects/SCI02287
AcademicSubjects/SCI02288
Genetically manipulated chloroplast stromal phosphate levels alter photosynthetic efficiency
https://orcid.org/0000-0002-5497-608X
Raju Aditi Subramani Department of Biology, Texas A&M University, College Station, TX 77843, USA

https://orcid.org/0000-0003-2181-6888
Kramer David M Department of Energy Plant Research Laboratory, Michigan State University, East Lansing, MI 48824, USA
Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 48824, USA
Jan IngenHousz Institute, Bornsesteeg 48A, 6708 PE Wageningen, The Netherlands

https://orcid.org/0000-0002-3725-1036
Versaw Wayne K Department of Biology, Texas A&M University, College Station, TX 77843, USA

Author for correspondence: wversaw@tamu.edu
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (https://academic.oup.com/plphys/pages/General-Instructions) is: Wayne K. Versaw (wversaw@tamu.edu).

Conflict of interest statement. None declared.

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© The Author(s) 2024. Published by Oxford University Press on behalf of American Society of Plant Biologists.
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Abstract

The concentration of inorganic phosphate (Pi) in the chloroplast stroma must be maintained within narrow limits to sustain photosynthesis and to direct the partitioning of fixed carbon. However, it is unknown if these limits or the underlying contributions of different chloroplastic Pi transporters vary throughout the photoperiod or between chloroplasts in different leaf tissues. To address these questions, we applied live Pi imaging to Arabidopsis (Arabidopsis thaliana) wild-type plants and 2 loss-of-function transporter mutants: triose phosphate/phosphate translocator (tpt), phosphate transporter 2;1 (pht2;1), and tpt pht2;1. Our analyses revealed that stromal Pi varies spatially and temporally, and that TPT and PHT2;1 contribute to Pi import with overlapping tissue specificities. Further, the series of progressively diminished steady-state stromal Pi levels in these mutants provided the means to examine the effects of Pi on photosynthetic efficiency without imposing nutritional deprivation. ΦPSII and nonphotochemical quenching (NPQ) correlated with stromal Pi levels. However, the proton efflux activity of the ATP synthase (gH+) and the thylakoid proton motive force (pmf) were unaltered under growth conditions, but were suppressed transiently after a dark to light transition with return to wild-type levels within 2 min. These results argue against a simple substrate-level limitation of ATP synthase by depletion of stromal Pi, favoring more integrated regulatory models, which include rapid acclimation of thylakoid ATP synthase activity to reduced Pi levels.

Mutational manipulation of chloroplast stromal phosphate content implicates integrated regulation of ATP synthase activity and photoprotective mechanisms.

U.S. Department of Energy 10.13039/100000015 Office of Science 10.13039/100006132 Bioimaging Technology Program DE–SC0014037 Basic Energy Sciences 10.13039/100006151 DE-FG02-04ER15559 Hagler Institute for Advanced Study 10.13039/100017306 Texas A&M University 10.13039/100007904
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pmcIntroduction

During photosynthesis, the concentration of inorganic phosphate (Pi) in the chloroplast stroma must be high enough to sustain ATP synthesis (Selman and Selman-Reimer 1981; Aflalo and Shavit 1983; Junge 1987; Strotmann et al. 1990; Pänke and Rumberg 1996; Grotjohann and Graber 2002) but below levels that inhibit starch synthesis and reactions of the Calvin cycle (Cockburn et al. 1968; Furbank et al. 1987; Ballicora et al. 2004; Marcus et al. 2005). Substrate-level and allosteric effects of Pi within this concentration range contribute to the regulation of photosynthetic efficiency and carbon partitioning throughout the photoperiod (Marschner et al. 1996; Mengin et al. 2017). This regulation must be coordinated with environmental and metabolic conditions since high CO2 concentrations, cold temperatures, and high light can all induce a feedback-inhibition of photosynthesis that has been attributed to low stromal Pi concentrations and a resulting decline in ATP synthesis (Sharkey and Vanderveer 1989; Takizawa et al. 2008; McClain and Sharkey 2019; Bouain et al. 2022).

Three major processes determine stromal Pi concentration: assimilation of Pi to ATP via photophosphorylation (Grotjohann and Graber 2002), metabolic recycling of Pi from organophosphate compounds in the Calvin cycle and during starch synthesis (Geigenberger 2011; McClain and Sharkey 2019), and transport of Pi between the stroma and cytosol. However, the contribution of transport to the fine tuning of stromal Pi levels and, in turn, to photosynthesis, is poorly understood.

Chloroplastic Pi transport has been attributed to members of the plastidic phosphate translocator (pPT), PHOSPHATE TRANSPORTER 2 (PHT2), and PHT4 transporter families that localize to the chloroplast inner envelope membrane and catalyze Pi transport when reconstituted in artificial membranes or when expressed in heterologous systems. All pPTs catalyze stoichiometric exchange of Pi with phosphorylated C3, C5, or C6 compounds (Flugge 1999). For example, phosphoenolpyruvate/Pi translocators (PPT), glucose 6-phosphate/Pi translocators (GPT), and xylulose 5-phosphate/Pi translocators (XPT) mediate the export of stromal Pi in exchange for cytosolic metabolites that serve as precursors for biosynthetic activities within the chloroplast (Knappe et al. 2003). In contrast, the triose phosphate/Pi translocator (TPT) is the only member of the pPT family that catalyzes the import of Pi from the cytosol during the forward assimilatory reactions and couples it with the export of triose phosphates (TPs), the end products of photosynthesis (Fliege et al. 1978; Flügge et al. 1989). This exchange is the primary mechanism for carbon allocation to the cytosol and is expected to be a key route for Pi entry to the chloroplast during the day. However, loss-of-function tpt mutants are viable and exhibit no obvious growth phenotype, indicating the presence of additional carbon and Pi transport mechanisms (Walters et al. 2004). Indeed, increased starch synthesis and turnover throughout the day coupled with export of carbon from the chloroplast in the form of maltose and glucose compensate for the tpt defect in carbon export (Riesmeier et al. 1993; Walters et al. 2004). It is plausible that members of the PHT2 and PHT4 transporter families compensate for the tpt defect in Pi import via H+- or Na+-coupled Pi transport (Versaw and Harrison 2002; Guo et al. 2008; Pavon et al. 2008). Loss-of-function mutants for PHT2;1, PHT4;1, and PHT4;4 have modest phenotypes (Versaw and Harrison 2002; Karlsson et al. 2015; Miyaji et al. 2015), suggesting at least partially redundant and/or unrelated activities. However, distinct effects of these mutations on stromal Pi are unknown.

Live Pi imaging allows for nondestructive monitoring of Pi with subcellular spatial resolution. We previously demonstrated the utility of a genetically-encoded Förster resonance energy transfer (FRET)-based Pi sensor to monitor Pi levels in subcellular compartments within Arabidopsis (Arabidopsis thaliana) roots and leaves and Brachypodium distachyon mycorrhizal roots (Mukherjee et al. 2015; Banerjee et al. 2016; Sahu et al. 2020; Zhang et al. 2022). These studies revealed variations in cytosolic Pi concentration during root development and in response to Pi supply, and also confirmed a physiological role for PHT4;2 in Pi export from root plastids. In this study, we used live ratiometric imaging of a FRET-based Pi sensor to evaluate the contributions of Arabidopsis chloroplastic Pi transporters TPT, PHT2;1, and PHT4;4 toward the control of stromal Pi concentration in different leaf tissues and over time in the photoperiod. We found no evidence of Pi transport (import or export) for PHT4;4, which is consistent with results indicating that this protein functions as an ascorbate transporter (Miyaji et al. 2015; Nam et al. 2021). However, our results indicated that TPT and PHT2;1 both contribute to the transport of Pi into chloroplasts with overlapping but distinct tissue-dependent activities. We also utilized tpt and pht2;1 mutants to evaluate the effect of altered stromal Pi concentration on photosynthetic efficiency and thylakoid photoprotective measures.

Results

Effects of chloroplastic Pi transporter mutations on stromal Pi concentration

We hypothesized that the TPT, PHT2;1, and PHT4;4 chloroplastic Pi transporters have distinct roles in the modulation of stromal Pi concentration. To explore this possibility, we first needed to compare stromal Pi levels in wild-type plants with those in tpt, pht2;1, and pht4;4 mutants. To image stromal Pi in each of these genetic backgrounds, we generated transgenic lines that constitutively express stroma-targeted versions of a Pi sensor (cpFLIPPi-5.3m), single fluorescent protein controls, and a Pi-insensitive control sensor (cpFLIPPi-Null) (Mukherjee et al. 2015; Banerjee et al. 2016). Plants were grown for 21 d and then adaxial mesophyll (ADM, palisade) cells in the third or fourth rosette leaves were imaged 4 h into the photoperiod using confocal microscopy.

Ratiometric imaging (FRET-derived cpVenus emission: CFP emission), hereafter referred to as FRET ratio, was used to assess Pi levels. Raw FRET emission values were corrected for tissue-specific spectral bleed-through and cross-excitation of fluorescence emission from the donor (CFP) and acceptor (cpVenus) proteins, respectively, using location-matched single fluorescent protein controls to yield sensitized FRET values. The same imaging conditions were applied to the Pi-insensitive control, which differs from the Pi sensor in 6 amino acids in a predicted Pi-binding domain (Banerjee et al. 2016). At least 6 independent plants and 50 to 60 chloroplasts were imaged for each genotype. Linear regression was applied to each genotype data set to determine FRET ratios, i.e. slope values of sensitized FRET vs. CFP (Banerjee et al. 2016). This analysis also confirmed that FRET ratio values were not influenced by variation in sensor protein abundance. As an example, data for wild-type stroma are shown in Supplementary Fig. S1. FRET ratio values were constant with image exposure times ranging from 0.3 to 2 s, indicating that imaging conditions did not alter steady-state stromal Pi levels. We found that FRET ratios were significantly higher for the tpt and pht2;1 mutants than wildtype (Fig. 1A), but there were no significant changes for the Pi-insensitive control (Fig. 1B). Because FRET ratio decreases with increasing Pi, elevated FRET ratio values for tpt and pht2;1 indicate that both have reduced levels of stromal Pi, consistent with defects in Pi import. To facilitate interpretation of these results, y axes in Fig. 1 and subsequent figures are reversed so “up” corresponds to higher Pi concentration. In contrast to tpt and pht2;1, we detected no change in stromal Pi for the pht4;4 mutant. It was possible that PHT4;4 has a small contribution to stromal Pi that would be more readily revealed in a sensitized background, but we detected no measurable effect when the pht4;4 mutation was combined with tpt or pht2;1, whereas combining the tpt and pht2;1 mutations resulted in lower stromal Pi levels than in the single mutants (Supplementary Fig. S2). These results support previous findings suggesting that PHT4;4 catalyzes ascorbate transport rather than Pi transport in Arabidopsis (Miyaji et al. 2015; Nam et al. 2021). Given the lack of any detectable effect on stromal Pi concentration, the pht4;4 mutant was not included in subsequent experiments.

Figure 1. Effect of chloroplastic Pi transporter mutations on stromal Pi. Relative stromal Pi levels were measured in wildtype (WT) and the indicated chloroplastic Pi transporter mutants in adaxial mesophyll tissue. Plotted data are Förster resonance energy transfer (FRET) emission ratios ± SE for Pi sensor (top) and control sensor (bottom). y axes are reversed to emphasize that higher FRET ratios correspond to lower Pi levels. Fifty to 60 chloroplasts from 6 independent plants of each genotype were examined at Hour 4 of the photoperiod. A) Pi levels in tpt and pht2;1 were significantly lower than WT, but levels in pht4;4 were equivalent to WT. * indicates significant difference from WT, Student's t-test, P < 0.05. B) No significant differences (n.s.) were detected with the control sensor.

Although we previously used microinjections to develop a calibration curve to estimate absolute Pi concentrations from FRET ratios in roots (Sahu et al. 2020), FRET ratios measured in leaves were consistently outside the root calibration limits. This difference was observed previously and attributed to unequal quenching of sensor fluorophores by leaf pigments rather than difference in Pi concentration since FRET ratios for the control sensor were affected similarly (Banerjee et al. 2016). Microinjection of chloroplasts is not possible so results here are reported as relative Pi concentrations.

Spatial and temporal variation of stromal Pi

To determine if chloroplasts in different leaf tissues have different stromal Pi contents, if these Pi levels change during the photoperiod, and if TPT and PHT2;1 have spatial and/or temporal specificities, we imaged each of the 4 main leaf tissues, adaxial epidermis (ADE), adaxial mesophyll (ADM, palisade), abaxial mesophyll (ABM, spongy), and abaxial epidermis (ABE) at 6 different time points chosen to span the 16-h photoperiod. Figure 2A illustrates the arrangement of these tissues in a leaf. In wild-type leaves, relative stromal Pi concentration differed in each tissue and in their temporal profiles (Fig. 2B). Stromal Pi levels were most similar and least variable over time in epidermal tissues (ADE and ABE). In ADM, Pi levels decreased during the first half of the photoperiod and then increased back to starting levels by the end of the photoperiod. In contrast, Pi levels in ABM rose over the first 4 h then declined over the remainder of the photoperiod.

Figure 2. Spatial and temporal variation of stromal Pi concentration in wild-type leaves. A) Graphical illustration of the main tissue types of a dicot leaf—adaxial epidermis (ADE), adaxial mesophyll (ADM), abaxial mesophyll (ABM), and abaxial epidermis (ADE), created with BioRender.com. B) Stromal Pi contents differed significantly between tissues and over time as determined by ANOVA (P < 0.05). Plotted values are Förster resonance energy transfer (FRET) ratio ± SE obtained from 50 to 60 chloroplasts in 6 independent plants of each genotype at each time point. Data were normalized using tissue-specific FRET ratios obtained from control sensors in wild-type plants.

To determine the contributions of TPT and PHT2;1 toward the spatial and temporal stromal Pi profiles observed in wildtype, we imaged stromal Pi in the tpt and pht2;1 mutants as well as a tpt pht2;1 double mutant. As shown in Fig. 3, A and B, tpt and pht2;1 mutations reduced stromal Pi in both mesophyll tissues (ADM, ABM) with time of day effects that largely mirrored changes in wildtype. Stromal Pi content was further diminished in the tpt pht2;1 double mutant suggesting that TPT and PHT2;1 have additive contributions. These contributions were less obvious in epidermal cells because stromal Pi levels in the mutants were either equivalent to, greater than, or less than wildtype at different times in the photoperiod (Fig. 3, C and D), suggesting that chloroplasts in these tissues control Pi through additional mechanisms, which could include compensatory changes in Pi homeostatic processes. It is also possible that TPT and PHT2;1 influence Pi levels in these chloroplasts under other environmental conditions. No significant spatial or temporal changes in FRET ratio in any of these genotypes were detected with the Pi-insensitive control sensor (Supplementary Fig. S3), supporting the assertion that differences detected with the Pi sensor reflect relative Pi concentrations.

Figure 3. Spatial and temporal variation in relative stromal Pi levels. Pi-dependent Förster resonance energy transfer (FRET) ratios were measured in wildtype (WT), pht2;1, tpt, and tpt pht2;1 in A) adaxial mesophyll (ADM), B) abaxial mesophyll (ABM), C) adaxial epidermis (ADE), and D) abaxial epidermis (ABE) tissues. Plotted values are FRET ratio ± SE obtained from 50 to 60 chloroplasts in 6 independent plants of each genotype at each time point. * indicates significant genotypic differences from WT, ANOVA (P < 0.05); not significant (n.s.).

Although tpt and pht2;1 mutations reduced stromal Pi in both mesophyll tissues, the effects on ADM and ABM were unequal and therefore suggest a level of specificity. That is, in ADM, the tpt mutation had a greater effect on stromal Pi than pht2;1 and this relative effect was reversed in ABM (Fig. 3, A and B). To visualize this difference more readily, FRET ratio data for the mutants were normalized to wild-type values and replotted as percent of wildtype (Supplementary Fig. S4). These results indicate that TPT and PHT2;1 both contribute to chloroplastic Pi import in ABM and ADM, but TPT has the greater role in ADM whereas PHT2;1 has the greater role in ABM.

Complementation of tpt and pht2;1 mutations

To test for effects of secondary mutations, we asked if wild-type alleles of the Pi transporter genes complement the mutations. We assembled genomic TPT and PHT2;1 constructs with C-terminal translational fusions to mCherry to aid detection, and then introduced these constructs into the respective mutant backgrounds by transformation. Pi imaging of mesophyll tissues (ADM and ABM) showed that stromal Pi levels were restored to wild-type levels in both complementation lines (Fig. 4, A and B). These results support our initial hypothesis that tpt and pht2;1 mutations are causal for low stromal Pi phenotypes.

Figure 4. Complementation of tpt and pht2;1 mutations. Reduced stromal Pi phenotypes of tpt and pht2;1 were both rescued to wild-type (WT) levels in the respective complement lines in ADM A) and ABM B) tissues as indicated. Plotted values are Förster resonance energy transfer (FRET) ratio ± SE for the stroma-localized Pi sensor obtained from 50 to 60 chloroplasts in 6 independent plants of each genotype at each time point. * indicates significant genotypic differences from WT, ANOVA (P < 0.05); not significant (n.s.).

Tissue-specific abundance of TPT and PHT2;1

To test if changes in TPT and PHT2;1 protein levels could explain the results in Fig. 3, we produced lines in which a mCherry fusion tag was incorporated into the complementation constructs. We monitored mCherry fluorescence as a readout of transporter protein abundance in each tissue and used emission from a stroma-targeted cpVenus expressed from the constitutive UBQ10 promoter in the same plants and tissues for normalization. As shown in Supplementary Fig. S5, no spatial or temporal variation in the abundance of either TPT or PHT2;1 was detected.

The lack of variation in TPT and PHT2;1 protein abundance was not an artifact of mCherry stability because changes in relative mCherry fluorescence that closely mimic those of native PHT2;1 transcripts (Versaw and Harrison 2002; Guo et al. 2008) were detected when plants were subject to light/dark transitions (Supplementary Fig. S6). As a result, we conclude that the abundances of TPT and PHT2;1 did not vary with location or time during the photoperiod.

Tpt and pht2;1 mutations have opposite effects on cytosolic Pi concentration

Given that the tpt and pht2;1 mutants have defects in the import of cytosolic Pi into the chloroplast stroma, we hypothesized that Pi would accumulate in the cytosol of these mutants. To test this idea, we expressed a cytosol-localized Pi sensor in wild-type plants as well as each of the single mutants and the tpt pht2;1 double mutant, and then measured Pi-dependent FRET ratios in ADM (palisade) cells to evaluate relative cytosolic Pi concentrations. In wild-type cells, cytosolic Pi levels varied over the photoperiod (Fig. 5A). As predicted, cytosolic Pi levels were elevated in pht2;1, but only between 7 and 12 h into the photoperiod. Surprisingly, cytosolic Pi levels were reduced in tpt during this same time interval. These opposite effects on cytosolic Pi accumulation were partially balanced in the tpt pht2;1 double mutant (Fig. 5A). No significant variations in FRET ratio were detected for the control sensor in these genotypes (Fig. 5B), validating the changes observed with the Pi sensor.

Figure 5. Tpt and pht2;1 mutations have opposite effects on cytosolic Pi concentration. A) Comparison of relative cytosolic Pi levels in adaxial mesophyll (palisade) cells of the indicated genotypes. B) Förster resonance energy transfer (FRET) ratios for the control sensor in the same tissue and genotypes. Plotted values are FRET ratio ± SE obtained from 50 to 60 chloroplasts in 6 independent plants of each genotype at each time point. * indicates significant genotypic differences from wildtype (WT), ANOVA (P < 0.05); not significant (n.s.). x indicates significant differences in cytosolic Pi levels in opposite directions relative to WT (Student’s t-test, P < 0.05).

Effect of tpt and pht2;1 mutations on total Pi content of leaves

Although stromal Pi was reduced in ADM tissues in both tpt and pht2;1 mutants, the opposite effects of these mutations on cytosolic Pi levels raised the question of whether the total Pi content of leaves was altered and if so, if it correlated with the contents of either of these subcellular pools. We therefore measured total Pi contents in leaves of 21-d-old wild-type, tpt, pht2;1, and tpt pht2;1 plants using a colorimetric assay. As shown in Fig. 6A, total Pi content was significantly reduced in both of the single mutants and was further reduced in the tpt pht2;1 double mutant. This trend correlated (R2 = 0.93) with relative stromal Pi levels in ADM tissue (Fig. 6B), which likely reflected the large number of chloroplasts per ADM cell and therefore substantial portion of total leaf volume. This interpretation was supported by 3D rendering (Supplementary Fig. S7A) and voxel counting, which showed that chloroplast stroma occupied 30% to 40% of palisade mesophyll cell volume, whereas cytosol was only ∼10% of cell volume (Supplementary Fig. S7B).

Figure 6. Total leaf Pi contents correlate with stromal Pi levels. A) Total Pi content (nmol Pi/mg fresh weight) was determined from 3 independent samples for each genotype. Values shown are means ± SD. * indicates significant difference from wildtype (WT), Student’s t-test (P < 0.05). B) Correlation between relative stromal Pi concentration (data from Fig. 4A, Hour 4) and total leaf Pi content.

Tpt and pht2;1 mutants have reduced photosynthetic efficiency

Previous studies showed that photosynthesis is reduced when leaf Pi content is diminished by either nutritional deprivation during plant growth or by infiltration of leaves with non-metabolizable hexokinase substrates like mannose to sequester cytosolic Pi (Fredeen et al. 1989; Takizawa et al. 2008; Carstensen et al. 2018). Although this correlation was attributed to effects of reduced stromal Pi concentration, it is difficult to disentangle the myriad pleiotropic effects of Pi deprivation on cellular metabolism (Plaxton and Tran 2011) without direct measures of stromal Pi. The effects of tpt, pht2;1, and tpt pht2;1 mutations on stromal Pi levels under the same Pi-replete growth conditions (Fig. 3) therefore present a unique opportunity to evaluate the relationship between stromal Pi and photosynthesis, and to discern the effects of individual chloroplastic Pi transport processes.

We used the chloroplastic Pi transporter mutants to examine the effect of reduced stromal Pi on quantum yield of photosystem II (ΦPSII) as a measure of photosynthetic efficiency. As shown in Fig. 7A, ΦPSII did not vary significantly over the photoperiod in wild-type plants. However, ΦPSII was clearly reduced in each of the mutants with greatest effect in the tpt pht2;1 double mutant. This trend was mirrored by increases in nonphotochemical quenching (NPQt, Fig. 7B), which represents the dissipation of excess absorbed light energy in the photosynthetic antenna complexes as heat (Butler 1978; Genty et al. 1989; Tietz et al. 2017). Fv/Fm measured immediately before the onset of photosynthesis was unaffected in these mutants (Supplementary Fig. S8) indicating that the decline in ΦPSII observed under growth conditions was not attributable to lower maximal PSII quantum yield. Moreover, chlorophyll contents of these mutants were indistinguishable (Supplementary Fig. S9) from wild-type plants so differences in ΦPSII and NPQt align with stromal Pi levels in leaf mesophyll tissues (Fig. 3).

Figure 7. Effect of chloroplastic Pi transporter mutations on ΦPSII and NPQt. A) ΦPSII and B) NPQt for the indicated genotypes were measured at 4 different time points in the photoperiod. Each data point represents mean +/SE for 6 independent plants. * indicates significant genotypic differences from wildtype (WT), ANOVA (P < 0.05); not significant (n.s.).

The qE form of NPQ is triggered by acidification of the thylakoid lumen via protonation of the PsbS protein (Li et al. 2004) and through activation of violaxanthin de-epoxidase (Demmig-Adams 1990; Crofts and Yerkes 1994; Horton et al. 1996; Müller et al. 2001; Jahns and Holzwarth 2012). Pi-dependent acidification of the lumen could occur if stromal Pi availability limited the ATP synthase conductivity of protons (gH+), which would concomitantly increase the thylakoid proton motive force (pmf) (Takizawa et al. 2008). To evaluate gH+ and relative changes in pmf, we analyzed the decay kinetics of the electrochromic shift (ECS) signal during brief dark intervals, also referred to as the dark interval relaxation kinetics (DIRK) analysis (Sacksteder and Kramer 2000). The ECS signal is a change in the absorbance of chloroplast pigments when the electric field formed across the thylakoid membrane is altered (Junge and Witt 1968; Witt 1979). Despite the observed changes in NPQt (Fig. 7B), we found no significant differences between mutants and wild-type plants for either gH+ or pmf as estimated by the ECSt parameter under steady-state conditions (Supplementary Fig. S10).

We next asked if effects observed under standard growth conditions would be exacerbated by rapid increases in light intensity. Specifically, we subjected wild-type, tpt, pht2;1, and tpt pht2;1 plants to sequentially increasing light intensities with 1-min intervals. As shown in Fig. 8, A and B, all genotypes showed the expected increase in NPQt and decrease in ΦPSII in response to increasing light intensity, but relative genotypic differences from wildtype were not exacerbated. For example, for pht2;1, ΦPSII and NPQt differed from wildtype over the experimental range but magnitudes were nevertheless not statistically significant (Fig. 8B). However, a contribution of PHT2;1 was inferred from significantly greater responses for the tpt pht2;1 double mutant than the tpt single mutant (Fig. 8, A and B).

Figure 8. Effect of rapid changes in light intensity on photosynthetic parameters. Plants were subjected to the indicated light intensities for 1-min intervals and the following photosynthetic parameters were measured: A) ΦPSII, B) NPQt, C) pmf (ECSt(×103)), D) gH+. Each data point represents values +/SD for 6 independent plants. * indicates significant genotypic differences from wildtype (WT), ANOVA (P < 0.05); not significant (n.s.).

Under these dynamic light conditions, pmf was unaffected in the mutants at low light intensities but was significantly less than wildtype at higher intensities with greatest difference in the double mutant (Fig. 8C), suggesting that 1 or both the ΔpH and Δψ components of pmf were affected. Similarly, linear electron flow (LEF) increased with light intensity in all genotypes but as observed for pmf, differences were significant only at higher light intensities (Supplementary Fig. S11A). Further, LEF is expected to translocate three H+/e−, so that with LEF alone, a plot of flux of protons estimated by the ECS decay (vH+) against LEF should be a straight line (Baker et al. 2007). However, if cyclic electron flow (CEF) contributes substantially to proton translocation, vH+ should increase above this line (Avenson et al. 2005; Joliot and Joliot 2005). As shown in Supplementary Figure S11B, the relationship between vH+ and LEF was not significantly different between genotypes (ANOVA, P > 0.05, n = 7). The apparent linear relationship between LEF and thylakoid proton flux (vH+) (Supplementary Fig. S11B) indicates that, within the noise and uncertainty levels, altered stromal Pi did not have substantial effects on the contributions from cyclic electron flow (CEF) to pmf. However, we cannot rule out the possibility that small changes in CEF could contribute to ATP/NADPH balancing. Differences in gH+ were still not apparent (Fig. 8D). In wildtype, gH+ values initially fell with increasing light intensity and then leveled at higher light intensities as previously reported for Arabidopsis (Kanazawa and Kramer 2002). The Pi transporter mutants exhibited similar patterns.

Pi-dependent acclimation of ATP synthase activity

In dark-adapted wild-type plants, illumination generates a transient high-magnitude pmf that decreases to steady-state levels within minutes after onset of actinic light, and changes in gH+ follow this pattern with a brief initial delay (Fig. 9, A and B). Chloroplastic Pi transporter mutants also showed a transient increase in pmf with onset of light followed by a decline to steady-state levels within 2 min (Fig. 9A). The magnitude of the initial transient pmf was slightly reduced in the mutants, although this difference was statistically significant only for the tpt pht2;1 double mutant. In contrast, induction of gH+ was significantly reduced in both tpt and pht2;1, and further reduced in the tpt pht2;1 double mutant (Fig. 9B). Nevertheless, gH+ reset to wild-type levels within minutes. Taken together, these results suggest that thylakoid pmf and ATP synthase activity acclimate to reduced stromal Pi levels within minutes, although the effects of this acclimation on NPQt and ΦPSII persist under both standard growth and dynamic light conditions. The basis for this acclimation may encompass alterations in the rate of the reaction, influenced by adaptive modifications in either the affinity toward the Pi substrate or the fraction of the ATP synthase pool that is active (Rott et al. 2011)

Figure 9. Reduced stromal Pi transiently limits ATP synthase activity. Plants were dark adapted for 30 min then illuminated with low intensity light (∼75 μmol m−2 s−1). Changes in A) pmf (ECSt(×103)) and B) gH+ were measured simultaneously before, during and up to 5 min after dark-light transition. Plotted data are mean +/SE for 6 independent plants of each genotype. Values for gH+, but not pmf, differed significantly from wildtype (WT) as determined by ANOVA (P < 0.05). x indicates significant differences in gH+ relative to WT (Student’s t-test, P < 0.05).

Discussion

The roles of stromal Pi as a substrate for ATP synthesis and as an effector for carbon partitioning have been well established (Marschner et al. 1996). However, until now, it was unclear if stromal Pi concentration varies between chloroplasts in different tissues or varies over the course of a day. Live Pi imaging revealed that chloroplast Pi levels display striking variations both spatially and temporally. It is likely that tissue-dependent differences reflect specialized metabolism, which may include CO2 capture or partitioning of carbon to different metabolites. Proteomic studies suggest that such specialization may also include sensing of environmental conditions because profiles of plastids isolated from vasculature and epidermal tissues have environment sensory functions that are underrepresented in the photosynthesis-specialized chloroplasts in the leaf mesophyll tissue (Beltran et al. 2018).

Temporal changes in stromal Pi concentration within a tissue suggest metabolic shifts in the rates of 1 or more of the processes that govern stromal Pi, namely Pi assimilation, recycling, and transport. For example, in wild-type plants, the concentration of stromal Pi in ADM (palisade) tissue decreased during the first half of the photoperiod and then returned to starting levels by the end of the photoperiod (Fig. 3A). The decrease suggests that the rate of Pi assimilation required to sustain photosynthesis is greater than the rates of Pi recycling and/or Pi import resulting in a net decline. The subsequent rise in stromal Pi levels in the latter half of the photoperiod may then reflect a reduced sink demand for photosynthate (Brauner et al. 2018) coupled with high rates of metabolic recycling from starch production (Lu et al. 2005).

Chloroplastic Pi transporter mutants allowed visualization of the relative contributions of Pi transport to the spatial and temporal control of stromal Pi (Supplementary Fig. S4). Stromal Pi was reduced in tpt and pht2;1 mutants and was further reduced in a tpt pht2;1 double mutant (Fig. 3A), indicating that TPT and PHT2;1 both catalyze Pi import and that these activities are not redundant. Comparisons of the relative effects of these mutations on chloroplasts in different tissues indicated that PHT2;1 has a greater contribution to stromal Pi than TPT in ABM (spongy) tissue, whereas TPT has the greater role in ADM (palisade) tissue (Supplementary Fig. S4). The physiological relevance of this tissue specificity is unclear but may be related to the coupled transport of triose phosphates by TPT. Interestingly, although Pi transport by TPT and PHT2;1 varied between tissues and over time, as inferred from the respective mutants, protein abundance did not, suggesting that both transporters are regulated by unknown post-translational mechanisms.

If chloroplasts functioned as a closed system, independent of other cellular compartments, we would expect defects of Pi import in tpt and pht2;1 to result in greater accumulation of Pi in the cytosol. However, it is generally considered that the large vacuolar Pi pool buffers cytosol Pi concentration against substantial change (Mimura 1999). It was therefore surprising when we found that cytosolic Pi levels were altered in both tpt and pht2;1, and even more so that the effects were in the opposite direction (Fig. 5A). Elevated cytosolic Pi in the pht2;1 mutant is consistent with its defect in chloroplast Pi import and indicates that excess Pi in the cytosol, for at least a portion of the photoperiod, is tolerated and is not transferred to the vacuole. Reduced cytosolic Pi levels in the tpt mutant further illustrates the plasticity of cytosolic Pi and that a reduced Pi level is also not readily countered by vacuolar Pi stores.

The diminished cytosolic Pi concentration in tpt is not consistent with its defect in chloroplast Pi import (Fig. 5A). We therefore hypothesize that reduced cytosolic Pi levels in tpt reflect compensation for its coupled defect in the export of phosphorylated carbon (triose phosphates). That is, fixed carbon in this mutant follows the night path for carbon mobilization with starch synthesis/hydrolysis to yield maltose and glucose that are exported to the cytosol (Riesmeier et al. 1993; Walters et al. 2004). These sugars then undergo phosphorylation in the cytosol, which would drive down cytosolic Pi, at least transiently. Consistent with this hypothesis, a tpt pht2;1 double mutant has an intermediate cytosolic Pi level that is comparable to that in wild-type plants.

The series of progressively diminished steady-state stromal Pi levels in wild-type, tpt, pht2;1, and tpt pht2;1 plants provided an opportunity to evaluate the effect of stromal Pi concentration on photosynthesis independent of pleiotropic effects associated with nutritional Pi deprivation. Our central hypothesis was that diminished Pi levels would impose a substrate-level limitation on ATP synthase activity and the resulting reduction in proton conductivity would lead to acidification of the thylakoid lumen. Moreover, greater partitioning to the ΔpH component of pmf at the expense of the membrane potential component, Δψ, as reported previously as a response to Pi deprivation (Carstensen et al. 2018) would add to acidification of the lumen. This acidification would then trigger photoprotective nonphotochemical quenching (NPQ) processes and reduce photosynthetic efficiency (ΦPSII) (Demmig Adams 1992; Kanazawa and Kramer 2002; Li et al. 2004).

Indeed, the observed changes in NPQt and ΦPSII under growth conditions were consistent with a Pi concentration-dependent acidification of the lumen (Fig. 7). However, proton conductivity of the ATP synthase (gH+) did not differ significantly between mutants and wild-type plants under growth conditions (Supplementary Fig. S10), suggesting an alternate mechanism for lumen acidification. It is unlikely that stromal Pi influences the generation of protons via water splitting in the lumen based on subcellular locations, and there is also no evidence to suggest an effect of stromal Pi on proton translocating components of the electron transport chain since LEF was unaffected under growth conditions, although LEF was reduced at higher light intensities (Supplementary Fig. S11A). It is therefore tempting to speculate that stromal Pi influences thylakoid transporters that participate in pH homeostasis. One intriguing candidate is K+ EXCHANGE ANTIPORTER3 (KEA3), which catalyzes export of H+ from the lumen in exchange for stromal K+ (Spetea et al. 2017; Correa Galvis et al. 2020). If low stromal Pi levels directly or indirectly restrict K+/H+ antiport, then this would promote lumen acidification while other passive ion channels could then allow dissipation of membrane potential to maintain pmf. Such compensatory effects for pmf may become limited under some conditions, as seen with high light intensities (Fig. 8C) but be sufficient under growth conditions to mask the relatively small increases in pmf expected from the thermodynamic effect of reduced Pi concentration on ATP synthesis (ΔGATP).

A kinetic effect of reduced stromal Pi concentration on ATP synthase activity was observed only when plants were transitioned from dark to light (Fig. 9) and even then, the effect was transient with readjustment to wild-type levels within minutes. One possible explanation for these results is that the ATP synthase is regulated at the level of its affinity for Pi to sustain ATP production. That is, the rate of ATP synthesis could be relatively constant provided that the Km for Pi is responsive to stromal Pi concentration. Such an acclimatory response may, like the observed changes in NPQt and ΦPSII, be modulated by acidification of the lumen. Indeed, experiments conducted with isolated thylakoids suggest that the affinity of ATP synthase for Pi rises with thylakoid acidification (Pänke and Rumberg 1996). It is also possible that stromal Pi levels influence the fraction of the ATP synthase pool that is active (Vanlerberghe et al. 2019), but a change in the abundance of active ATP synthase would change the basal proton conductivity of the thylakoid, altering the relationship between light driven proton fluxes and pmf. Nevertheless, changes in the abundance and affinity of the ATP synthase are not mutually exclusive. Although it has been shown previously that ATP synthase activity can be reduced in response to Pi starvation (Karlsson et al. 2015; Carstensen et al. 2018), it is unclear if the reported responses reflect an extreme limitation that is beyond the acclimatory capacity of the ATP synthase. Additional studies are needed to resolve this intriguing question.

Materials and methods

Transgenic plant lines and growth conditions

Arabidopsis (A. thaliana) plants expressing stroma-targeted versions of the cpFLIPPi-5.3m Pi sensor (Mukherjee et al. 2015), the Pi-insensitive control sensor, cpFLIPPi-Null, (Banerjee et al. 2016), and single fluorescent protein controls contained an N-terminal 237-bp fragment encoding the 79-amino acid RbcS chloroplast transit peptide (Lee et al. 2006). All transgenic lines contain the suppressor of gene silencing3–13 (sgs3-13) mutation to ensure stable fluorescence. The Pi transporter double mutants were isolated from genetic crosses. Transgenic plants expressing the Pi sensor or controls were selected on 0.5× Murashige and Skoog (MS) (Murashige and Skoog 1962) agar plates with 10 μm phosphinothricin, and subcellular localizations were verified by confocal microscopy (experimental setup detailed in the next section). For plant growth, seeds were plated on 0.5× MS medium (Murashige and Skoog 1962) containing 250 μm Pi and 0.25% (w/v) sucrose. Seedlings were grown for 1 wk with 120 μmol m−2 s−1 light intensity, 16-h light/8-h dark photocycle, and then transferred to soil and grown for an additional 2 wk before imaging.

Stromal Pi imaging by confocal microscopy

The third or fourth true leaf was detached, treated with perfluorodecalin (PFD), and placed under a small glass brick either on the adaxial or abaxial surface depending on the tissues being images. PFD increases leaf tissue image resolution by eliminating air spaces, which enhances signal to noise ratio (Littlejohn et al. 2010, 2014). Mesophyll and epidermal cells were imaged at 6 different time points during the 16-h photoperiod using an inverted Olympus IX81 microscope equipped with a Yokogawa CSU-X1 Spinning Disk confocal unit, an iXon3 897 EMCCD camera (Andor Technology, Concord, MA, USA), and a 40× (numerical aperture 1.3) oil immersion objective. A 445 nm excitation laser and 483/32 nm and 542/27 nm emission filters were used to detect sensor eCFP and FRET-derived cpVenus, respectively. A 515 nm laser was used for direct excitation of cpVenus. Raw FRET-derived cpVenus emission was corrected for donor spectral bleed-through and acceptor cross-excitation values using location-matched controls to yield sensitized FRET emission. At least 50 to 60 chloroplasts from 6 or more independent plants were imaged for each data point.

Chlorophyll fluorescence-based photosynthetic parameters and ECS measurements

Photosynthetic parameters were measured using a MultiSpeQ instrument with PhotosynQ v2. Platform. The photosynthesis RIDES protocol was used to measure quantum efficiency of photosystem II (ΦPSII) and nonphotochemical quenching (NPQ) estimated using the NPQt method and thus designated NPQt (Tietz et al. 2017). ECSt, representing the magnitude of the pmf formed in the light, was used to estimate conductivity of ATP synthase (gH+) and light driven proton fluxes (vH+) using DIRK analysis (Sacksteder and Kramer 2000; Kanazawa and Kramer 2002). The measuring beam provided in the MultispeQ is a broadband emission peaking at 525 nm coupled with a BG-18 filter for signal detection (Kuhlgert et al. 2016). MultispeQ was used to measure relative chlorophyll contents. Data cleaning was performed using code from a GitHub repository (https://github.com/protonzilla/Light-Potentials-in-Field) as described (Kanazawa et al. 2021).

PHT2;1 and TPT complementation

The PHT2;1-mCherry construct contained 1,776 bp PHT2;1 promoter sequence plus its full coding region. The mCherry coding sequence was cloned in-frame at the 3′ end of PHT2;1. The TPT-mCherry construct was assembled similarly but included 1,402 bp TPT promoter plus full coding region. These constructs were introduced into the pht2;1 and tpt mutants, respectively, by Agrobacterium-mediated transformation. Transformed plants were selected for hygromycin resistance and then screened for mCherry fluorescence. T2 progeny were used for complementation assays and for analysis of transporter protein abundance.

Phosphomolybdate assay for total Pi

Third and fourth leaves of 3-wk-old plants were harvested from each genotype and Pi contents of leaves were determined using a phosphomolybdate colorimetric assay as previously described (Ames 1966; Versaw and Harrison 2002). All assays were conducted with 3 independent tissue samples.

Voxel counting to estimate stromal and cytosolic volumes

Leaves expressing cytosolic Pi sensor were treated with PFD. For cytosol volume estimation, a 515 nm laser was used for direct excitation of cpVenus with a 542/27 nm emission filter. For chloroplast stroma volume estimation, chlorophyll auto-fluorescence was detected using the 448 nm laser excitation and 633 LP emission filter. Z-stacks were acquired using a 40× silicone oil objective (RI = 1.404, which matches closely to that of cell cytosol). Step size was set at 0.5 μm to yield voxels of a defined size of 0.06 μm3. Z-stacks were thresholded to represent features of interest (chloroplast/cytosol). The resulting binary image stack was used for further analysis. The Voxel Counter ImageJ plugin was used to count the total number of thresholded voxels (in μm3) in each cell and this was converted to picoliters (1,000 μm3 = 1 pL).

Imaging with variable exposure

Leaves of 3-wk-old wild-type plants expressing the plastid-targeted cpFLIPPi-5.3m sensor were imaged for FRET and CFP emissions using the 445 nm excitation wavelength at the following exposure times: 30 ms, 50 ms, 100 ms, 200 ms, 300 ms, 400 ms, 500 ms, 700 ms, 800 ms, 1,000 ms, 1,200 ms, 1,500 ms, and 2,000ms. Imaging was carried out sequentially with 1 min dark treatment before each subsequent exposure. Imaging regions-of-interest (ROIs) were obtained from adaxial epidermal and mesophyll cells.

Statistical analyses

Student's t-test was applied to perform pairwise comparisons of relative Pi-dependent FRET ratios between genotypes and time points in the photoperiod. ANOVA was utilized to examine the genotypic differences in FRET emission ratios across leaf tissues and the photoperiod.

Accession numbers

Accession At5g23570 (SGS3) can be found under EMBL/GenBank data libraries. Mutants used in this study include sgs3-13 (SALK_039005), tpt-3 (SALK_028503), and pht2;1-1 (CS19883) and were obtained from the Arabidopsis Biological Resource Center. The pht4;4-1 (ET4970) mutant was obtained from the Cold Spring Harbor Laboratory.

Supplementary Material

kiae241_Supplementary_Data

Acknowledgments

We thank Dr. Abira Sahu (MSU-DOE Plant Research Laboratory) and Dr. Swayoma Banerjee (Intel Corporation) for their participation in assembling materials and for their insight.

Author contributions

A.S.R. and W.K.V. designed the research; A.S.R. performed the research; A.S.R., D.M.K., and W.K.V. analyzed the data; and A.S.R. and W.K.V. wrote the article.

Supplementary data

The following materials are available in the online version of this article.

Supplementary Figure S1. Ratiometric analysis of a chloroplast stroma-localized Pi sensor in adaxial mesophyll cells.

Supplementary Figure S2. Effect of pht4;4 mutation on stromal Pi in double mutants of pht2;1 pht4;4, and tpt pht4;4.

Supplementary Figure S3. No significant spatial or temporal variation in FRET ratio for the stroma-targeted control sensor.

Supplementary Figure S4. Mesophyll tissue-specific contributions of TPT and PHT2;1 toward stromal Pi concentration.

Supplementary Figure S5. Abundance of TPT and PHT2;1 proteins does not differ between tissues nor over time in the photoperiod.

Supplementary Figure S6. Change in PHT2;1 protein abundance during light/dark transitions.

Supplementary Figure S7. Voxel counting estimates that chloroplast stroma occupies larger volume than cytosol in the leaf mesophyll cells.

Supplementary Figure S8. Fv/Fm measured prior to onset of photosynthesis.

Supplementary Figure S9. Relative chlorophyll between genotypes.

Supplementary Figure S10. No significant differences between mutants and wild-type plants for either gH+ or pmf (ECSt × 103).

Supplementary Figure S11. Cyclic electron flow is not increased in chloroplastic Pi transporter mutants.

Funding

Financial support was provided by the U.S. Department of Energy, Office of Science, Bioimaging Technology Program (DE–SC0014037) and Basic Energy Sciences, (DE-FG02-04ER15559). A.S.R. was supported in part by funds from the Hagler Institute for Advanced Study, Texas A&M University.

Data availability

The data underlying this article are available in the article and its online supplementary material.

Dive Curated Terms

The following phenotypic, genotypic, and functional terms are of significance to the work described in this paper:

GPT Gramene: AT2G41490

GPT Araport: AT2G41490

SGS3 Gramene: AT5G23570

SGS3 Araport: AT5G23570

PHT2;1 Gramene: AT3G26570

PHT2;1 Araport: AT3G26570

TPT Gramene: At5g46110

TPT Araport: At5g46110

PHT4;4 Gramene: At5g20380

PHT4;4 Araport: At5g20380
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