Plants acclimate to Photosystem I photoinhibition by readjusting the photosynthetic machinery

Abstract Photosynthetic light reactions require strict regulation under dynamic environmental conditions. Still, depending on environmental constraints, photoinhibition of Photosystem (PSII) or PSI occurs frequently. Repair of photodamaged PSI, in sharp contrast to that of PSII, is extremely slow and leads to a functional imbalance between the photosystems. Slow PSI recovery prompted us to take advantage of the PSI‐specific photoinhibition treatment and investigate whether the imbalance between functional PSII and PSI leads to acclimation of photosynthesis to PSI‐limited conditions, either by short‐term or long‐term acclimation mechanisms as tested immediately after the photoinhibition treatment or after 24 h recovery in growth conditions, respectively. Short‐term acclimation mechanisms were induced directly upon inhibition, including thylakoid protein phosphorylation that redirects excitation energy to PSI as well as changes in the feedback regulation of photosynthesis, which relaxed photosynthetic control and excitation energy quenching. Longer‐term acclimation comprised reprogramming of the stromal redox system and an increase in ATP synthase and Cytochrome b6f abundance. Acclimation to PSI‐limited conditions restored the CO2 assimilation capacity of plants without major PSI repair. Response to PSI inhibition demonstrates that plants efficiently acclimate to changes occurring in the photosynthetic apparatus, which is likely a crucial component in plant acclimation to adverse environmental conditions.


| INTRODUCTION
Photosynthesis utilizes light to assimilate CO 2 into organic compounds.
Photons are transduced to chemical energy in Photosystem II (PSII) and Photosystem I (PSI) reaction centres, which eject electrons to the linear electron transfer chain (LET) generating NADPH. Concomitant proton pumping to the thylakoid lumen establishes a proton motive force (pmf) that is utilized by ATP synthase to produce ATP from Pi and ADP. This energy transduction requires the concerted function of the four membrane-embedded protein complexes, PSII, PSI, cytochrome b 6 f complex (Cyt b 6 f), and ATP synthase. Photosystems and Cyt b 6 f are connected by mobile electron carriers, membrane soluble plastoquinone (PQ), and lumenal plastocyanin (PC), whereas two soluble proteins, ferredoxin (Fd) and ferredoxin NADP + oxidoreductase (FNR) on the stromal side of PSI finalize the reduction of NADP + to NADPH. Most of the ATP and reducing power generated in LET are used to reduce CO 2 to triose phosphates and to regenerate ribulose 1,5-bisphosphate in the Calvin-Benson-Bassham (CBB) cycle. Other major sinks for reducing power in chloroplasts include photorespiration, antioxidant system, nitrogen assimilation (NA) and, to a smaller extent, also sulfur assimilation and lipid biosynthesis, all depending on environmental conditions (Asada, 1999;Foyer et al., 2009;Walker et al., 2020). In addition to these stromal reaction pathways, a part of the reducing power is exported from chloroplast through the malate valve (Selinski & Scheibe, 2018) to be utilized in other cellular compartments (Shameer et al., 2019). The distribution of reducing power between these different sinks needs to be regulated according to the metabolic state of the cell, predominantly occurring via the thioredoxin (Trx) system (Geigenberger et al., 2017;Schürmann & Buchanan, 2008).
Prompt and accurate regulation of photosynthetic light reactions is crucial, as the imbalance between PSII and PSI, as well as that between the light reactions and stromal sinks, leads to the formation of harmful reactive oxygen species (ROS) (Asada, 2006). Despite multilayered antioxidant systems scavenging ROS in chloroplasts, both PSII and PSI are prone to oxidative damage under harsh environmental conditions. The control of ROS production in LET is important not only to avoid uncontrolled oxidative damage but also to allow site-specific ROS production in LET as an important secondary messenger for regulation of gene expression and promotion of plant long-term acclimation to changed environmental conditions (Chan et al., 2016;de Souza et al., 2017;Farmer & Mueller, 2013;Fitzpatrick et al., 2022).
Photodamage and inhibition of PSI under conditions challenging the capacity of the regulatory and scavenging systems have been known to exist already for decades, particularly in chilling sensitive plants (Havaux & Davaud, 1994;Terashima et al., 1994). Nevertheless, the demonstration of PSI photoinhibition in Arabidopsis thaliana proton gradient regulation 5 (PGR5) mutant in natural fluctuating light conditions (Suorsa et al., 2012) boosted PSI photoinhibition research. In sharp contrast to the fast repair mechanisms of PSII upon photoinhibition (Aro et al., 1993;Nishiyama et al., 2011), PSI repair is known to be extremely slow (Kudoh & Sonoike, 2002) and to require the synthesis and assembly of the entire PSI complex. Depending on environmental conditions, the complete repair takes from days to weeks and during this time the photosynthetic light reactions can become limited by PSI (Lima-Melo et al., 2019;Zhang & Scheller, 2004;Zhang et al., 2011;Zivcak et al., 2015). Thus, it is likely that plants mitigate the consequences of PSI inhibition by acclimation to the PSI complex deficiency.
To disclose putative mechanisms that drive the acclimation of plants to PSI deficiency, without being hampered by concomitant low-temperature stress, we subjected wild-type Arabidopsis plants to moderate and severe PSI photoinhibition, using specific light treatments at normal growth temperature (Tikkanen & Grebe, 2018) to induce 60% and 85% PSI photoinhibition. The five sets of plants, (i) untreated, (ii) 60% PSI photoinhibited, (iii) 85% PSI photoinhibited, as well as (iv) 60% PSI photoinhibited and subsequently 'recovered' for 24 h in growth conditions, and (v) 85% PSI photoinhibited and subsequently 'recovered' for 24 h in growth conditions, were subjected to analyses of their major photosynthetic complexes, the function and regulation of photosynthetic light reactions, ATP synthase and carbon assimilation. These measurements were further complemented by analyses of the redox regulation of photosynthetic enzymes, as well as the phosphorylation and supercomplex formation of thylakoid proteins and complexes in different light conditions. The experimental setup is outlined in Figure 1. Whole plants were treated with a specific fluctuating light regime to induce PSI photoinhibition (Tikkanen & Grebe, 2018

| Biophysical analyses
CO 2 assimilation, chlorophyll a fluorescence, P700, Fd and PC redox states were recorded concurrently with GSF-3000 infra-red gas analyser which was connected with a 3010-Dual gas exchange cuvette to Dual-KLAS-NIR (Heinz Walz GmbH). The used light intensities are described in Table 2. Actinic light was supplemented with 10% blue light to ensure proper stomatal regulation. Measurements were done after a minimum of 30 min dark acclimation. CO 2 assimilation was recorded every 10 s during the light curve. The gas flow rate was set to 400 µmol s −1 , the cuvette temperature was kept at a constant 25°C and the concentration of CO 2 and H 2 O were set to 400 and 18 000 p.p.m., respectively. Assimilation was calculated ACCLIMATION TO PSI PHOTOINHIBITION F I G U R E 1 Experimental setup to investigate the recovery of Arabidopsis from Photosystem I (PSI) photoinhibition. For biophysical analyses, plants were PSI photoinhibited for 4 h (moderate inhibition) or 8 h (severe inhibition) after which half of the plants were returned to growth conditions to recover for 24 h. Control plants were taken directly from growth conditions. Five differently treated plant groups were dark acclimated for at least 30 min before light curves were recorded to investigate the function and regulation of photosynthetic light reactions, ATPase and CO 2 assimilation. For biochemical analyses, another set of dark acclimated plants was divided into four groups, which were treated for 1 h in darkness (0 µmol photons m −2 s −1 ), low light (35 µmol photons m −2 s −1 ), growth light (165 µmol photons m −2 s −1 ), or high light (635 µmol photons m −2 s −1 ). After these light treatments, leaf samples were collected for thylakoid isolation and redox labelling of proteins, to study the regulation of photosynthesis at the protein level. according to (von Caemmerer & Farquhar, 1981). Saturating pulse of 4000 µmol photons m −2 s −1 was given every minute to determine the fluorescence and absorbance parameters described below.
Chlorophyll a fluorescence was detected with pulse-modulated 540 nm measuring light. P700, Fd and PC redox states were determined by deconvolution of pulse-modulated dual-wavelength 785-840, 810-870, 870-970 and 795-970 nm signals. Deconvolution was performed using differential model plots measured from control plants (Klughammer & Schreiber, 2016). PSI inhibition prevents the formation of P700 + , which means that only the functional reaction centres contribute to the P700 signal in PSI photoinhibited plants. Therefore, we calculated the yields of PSI using the average of control P M measured with the NIR MAX script (Klughammer & Schreiber, 2016) for all treatments, following the approach used previously (Zivcak et al., 2015).
This approach assigns the damaged PSI reaction centres as acceptor side limited. The photochemical quantum yield of PSI (Φ I ), donor side limitation (Φ ND ) and acceptor side limitation (Φ NA ) were calculated according to (Klughammer & Schreiber, 1994, 2008b were calculated according to Genty (Genty et al., 1989;Klughammer & Schreiber, 2008a). Fd results were not analysed further since the signal is partly originating from PSI iron-sulfur clusters (Klughammer & Schreiber, 2016), which get damaged in the photoinhibition treatment.
The distribution of reducing power (electrons) to carbon assimilation was calculated according to equation 1.
Distribution of electrons to CO assimilation = 4 × Assimilation ETRII = 4 × Assimilation Φ × 0.5 × 0.84 × PPDF 2 II Equation 1: Distribution of reducing power to CO 2 assimilation Electrochromic shift (ECS) was recorded with Dual-PAM-100 equipped with P515/535 module (Heinz Walz GmbH). The used light intensities are described in Table 3. The electrochromic shift was determined by the difference between 515 and 550 nm signals (Schreiber & Klughammer, 2008). Dark interval of 250 ms was applied every minute to quantify the thylakoid proton conductivity (g H+ ) and the pmf (ECS t ). The g H+ parameter was calculated as the inverse of the time constant of the first-order exponential fit to the decay of the ECS signal during the dark interval (Kanazawa & Kramer, 2002). ECS t was calculated as a difference between the ECS in the light before the dark interval and the ECS dark baseline calculated from the first-order exponential fit. ECS t was normalized to the chlorophyll content of leaf discs (10 mm in diameter) cut from leaves after the measurements. Chlorophyll content and chlorophyll a/b ratio were determined from dimethylformamide (DMF) leaf extracts (Inskeep & Bloom, 1985).

| Biochemical analyses
Western blot analysis was performed as described in Rantala et al. (2020), with primary antibodies raised against the following proteins: PsaB, AtpF, PetA (Agrisera product numbers: AS10 695, AS10 1604 and AS20 4377, respectively) and PsbA (Kettunen et al., 1996). Infrared dye-labelled secondary antibody (IRDye ® 800CW Goat anti-Rabbit IgG Secondary Antibody [1:20 000 in 1% milk/TTBS, Li-Cor]) was used in protein immune detection with Odyssey CLx imager (Li-Cor). The antibody signal was quantified with the Image Studio programme (Li-Cor). The relative amounts of proteins were interpolated from the linear regression of signals of the control dilution series.
Thylakoid proteins were separated with sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gel containing 12% acrylamide and 6 M urea. ProQ and Sypro Ruby staining of separated thylakoidal proteins were performed according to manufacturer's instructions (Invitrogen). Gels were imaged with Perkin Elmer Geliance 1000 using Cy3 filter for ProQ and UV-filter for Sypro-dyed gels. 77 K fluorescence measurements were performed with a chlorophyll concentration of 10 µg/ml. Thylakoids were excited by 480 nm light and fluorescence was detected with Ocean Optics S2000 spectrophotometer. Spectra were normalized to 685 nm peak and the ratio between 685 and 735 nm peaks was calculated to illustrate the distribution of excitation energy between PSII and PSI.
Blue native gel electrophoresis was performed as described in (Järvi et al., 2011). Isolated thylakoids were suspended into ice-cold was added to the supernatant. Solubilized thylakoid protein complexes were separated with 3%-12% acrylamide gradient gels.
Protein redox labelling followed the protocol (Peled-Zehavi et al., 2010). Leaves frozen in liquid nitrogen were homogenized in 10% trichloroacetic acid and the proteins were precipitated with centrifugation. Precipitated proteins were first labelled with 50 mM N-ethyl maleimide to block the protein thiols followed by the reduction of residual disulfide bridges in proteins with dithiothreitol and subsequent labelling of exposed thiols with 10 mM pegylated maleimide (5 kDa) as described in Nikkanen et al. (2016). For estimation of protein content in the samples, the labelled proteins were separated with sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gel containing 12% acrylamide and 6 M urea and the gel was stained with Sypro Ruby protein stain and imaged with Perkin Elmer Geliance 1000.
The protein content of the samples was normalized to the amount of rubisco small subunit. An equal amount of proteins in labelled samples was separated with SDS-PAGE gel with 5%-15% acrylamide gradient with 6 M urea and electroblotted to polyvinylidene difluoride membrane (Millipore). Fructose 1,6-bisphosphatase (FBPase) was identified with a specific antibody (a kind gift from Dr M. Sahrawy), using an enhanced chemiluminescence detection kit (GE Healthcare) and Perkin Elmer Geliance 1000 imager.

| Photoinhibition treatment is specific for PSI
We first tested the specificity and effectivity of the PSI photoinhibi- recovery to 45% inhibition was recorded after 24 h in growth conditions. Severe PSI photoinhibition treatment of leaves resulted in around 85% inhibition, with a recovery to 60% inhibition during 24 h in growth conditions. Partial recovery of P M * indicated that plants can repair some of the damaged iron-sulfur clusters of PSI via a still unidentified mechanism as discussed before (Tiwari et al., 2016). PSII maximal quantum efficiency (F V /F M ) was only slightly affected by the moderate or severe PSI photoinhibition treatment and was partially restored during 1 day in growth conditions ( Figure 2b). Leaf chlorophyll content and chlorophyll a/b ratio were not substantially changed by the inhibition treatment (Figure 2c,d).

| Consequences of PSI photoinhibition to carbon assimilation and functionality of light reactions
Effects of PSI photoinhibition on CO 2 assimilation and functional characteristics of photosynthetic light reactions were analysed from intact leaves with infra-red gas analyser GSF-3000. The gas analyser was connected to P700, PC, Fd and chlorophyll a fluorescence measuring system Dual-KLAS-NIR with a Dual-PAM-100 gasexchange cuvette for parallel measurement of all parameters.
Moderate and severe PSI photoinhibition altered differently the CO 2 assimilation rate as a response to applied light intensity 3.3 | 24 h recovery from PSI photoinhibition treatment alters the distribution of electrons to carbon assimilation under growth light illumination CO 2 assimilation appeared to recover faster from PSI photoinhibition than the function of PSII (Figures 2e and 3a). Therefore, we estimated the relative proportion of electrons allocated to CO 2 assimilation  inhibited plants (Figure 6a), whereas the amount of the PSII subunit PsbA was not affected by the photoinhibition treatment ( Figure 6b).
The effect of PSI photoinhibition on ATP synthase subunit AtpF ( Figure 6c) and Cyt b 6 f complex subunit PetA (Figure 6d) was more notable. AtpF amount increased during the inhibition treatment and remained at an elevated level during the subsequent 24 h recovery. A similar trend was observed for PetA, but the changes were smaller than those for AtpF.
As the amounts of ATP synthase and Cyt b 6 f complex subunits (AtpF and PetA) were affected in PSI photoinhibited plants, we next studied the composition of thylakoid protein complexes with blue native gel electrophoresis (BN-PAGE), which maintains the photosynthetic protein complexes intact. As expected, the amount of Cyt b 6 f was increased in PSI photoinhibited plants (Figure 6e). ATP synthase, unfortunately, co-migrates with the PSII dimer (slightly lower) and therefore could not be assessed from the BN-gels.
In addition to changes in the amounts of major complexes, PSI photoinhibition treatment altered the composition of PSII-LHCII supercomplexes and the PSI-inhibited samples showed reduced amounts of large supercomplexes (C 2 S 2 M 2 and C 2 S 2 M) (Figure 6e).
This was accompanied by an increase in the free moderately bound M-LHCII, which comprises LHCII trimers associated with tightly bound minor light-harvesting antennae proteins, Lhcb4 and Lhcb6.
Conversely, the amount of loosely bound L-LHCII was not noticeably affected by the PSI photoinhibition treatment.
3.6 | Plasticity and dynamics of thylakoid protein complexes upon PSI photoinhibition As shown above, PSI photoinhibition altered the formation of PSII-LHCII supercomplexes (Figure 6e), which prompted us to check whether this change in the structural organization of light-harvesting is reflected at the functional level. To this end, we isolated thylakoids from PSI photoinhibited and control plants illuminated for 1 h in the four different light intensities, which were also applied in functional studies of intact leaves (Figure 1). The distribution of excitation energy between PSII and PSI was addressed by recording the low-temperature fluorescence emission spectra from highly diluted suspensions of isolated thylakoids.
PSI inhibition did not alter the peak positions of the 77 K fluorescence spectra, which implied that the light-harvesting antennae remained connected to the reaction centres (Supporting Information: Figure 4).
However, the ratio of fluorescence emitted from PSII (peaking at 685 and 695 nm) to that emitted from PSI (peaking at 735 nm) was affected by PSI photoinhibition, but the difference was highly dependent on preillumination light intensity (Figure 7a). Although the effect was minor in darkness, the low light and growth light illumination for 1 h enhanced the excitation of PSI, with no restoration during the 24 h recovery period.
After 1 h preillumination of plants in high light, the situation was different and the excitation energy distribution between PSII and PSI was closer to that in control plants.
To get further insights into the redox conditions that drive the dynamics of excitation energy distribution between PSII and PSI in the thylakoid membrane, we next analysed how the phosphorylation levels

| DISCUSSION
The photosynthetic apparatus is harnessed with mechanisms that continuously control the excitation energy distribution to and between the two photosystems to keep ETC optimally oxidized (Bassi & Dall'Osto, 2021;Tikkanen & Aro, 2014). ETC oxidation, on the other hand, is highly dependent on the capacity of the CBB cycle, which functions as the main electron sink from ETC. CBB cycle is likewise impacted by different biotic and abiotic factors, which heavily modify CO 2 availability and the rate of chemical reactions. After the light treatments leaf samples were immediately frozen in liquid nitrogen, followed by isolation of proteins. Protein thiols were blocked with N-ethyl-maleimide followed by the reduction of residual protein disulfides with dithiothreitol (DTT) and labelling of exposed thiols with pegylated maleimide that increases the apparent molecular mass of the protein by 5 kDa per thiol group. Labelled proteins were separated with SDS-PAGE and transferred to a polyvinylidene difluoride (PVDF) membrane. Amounts of differentially labelled proteins were determined with specific antibody. [Color figure can be viewed at wileyonlinelibrary.com] Keeping the ETC optimally oxidized is important, as any condition that results in the accumulation of excess electrons in ETC, such as an abrupt increase in light intensity, easily leads to PSI photoinhibition . The sensitivity of PSI to photoinhibition under high or fluctuating light is dependent on plant species (Huang et al., 2018;Terashima et al., 2021;Yamori et al., 2016). Moreover mutants, with problems in regulation of electron transfer, particularly the PGR5 deficient mutants, are especially prone to PSI photoinhibition Suorsa et al., 2012;Yamori et al., 2016). PSI shows extremely slow recovery capacity from photoinhibition, both when induced by illumination of plants at low temperature (Kudoh & Sonoike, 2002;Zhang & Scheller, 2004)

| Restoration of the CO 2 assimilation capacity after PSI photoinhibition is not dependent on PSI recovery
It was of primary importance to understand how the reduced number of functional PSI centres (Figure 2a) impacts the CO 2 assimilation rate in different light intensities and whether the subsequent acclimation of plants to the PSI-limited state during the 24 h recovery period changes the relationship between the abundance of functional PSI and CO 2 assimilation rate (Figure 2a,e). Our results demonstrate that the decrease in the abundance of functional PSI, recorded immediately after the PSI photoinhibition treatment, slowed down the CO 2 assimilation rate only when measurements were conducted under low and growth light intensities (Figure 2e). On the contrary, under higher light intensity the difference in CO 2 assimilation rate between the PSI photoinhibited and control plants strongly diminished ( Figure 2e). These results imply that the plants make use of only a small fraction of their entire PSI pool to reach a maximal CO 2 assimilation rate in high light conditions. This might allow plants to lower the PSI to PSII ratio as generally seen during long-term high light acclimation (Bailey et al., 2001;Flannery et al., 2021). Nonetheless, during the 24 h recovery period after PSI photoinhibition treatment, the CO 2 assimilation rates which were recorded under low and growth light illumination increased from that recorded immediately after the photoinhibition treatment (Figure 2e). These results demonstrate that the 24 h recovery period after the PSI inhibition treatment, while not long enough for major PSI repair, nevertheless provides the means for the photosynthetic machinery to partially restore the CO 2 assimilation rate also under low and moderate light intensities. In addition to these effects, it is likely that the excitation energy distribution to photoinhibited PSI centres decreases and conversely the energy transfer to functional PSI centres increases.
Although we could not obtain experimental evidence of this process, it is something that must occur at the latest when the inhibited PSI centres are being degraded. Thus, this process could partially explain the observed increase in CO 2 assimilation rate in light-limited conditions upon recovery for 24 h in growth conditions (Figure 2e).

| PSI photoinhibition limits LET
PSI photoinhibition has two main consequences on light reactions.
Firstly, the damaged and therefore permanently acceptor side limited PSI centres still receive excitation energy, which hinders the availability of light for functional photosystems (Zivcak et al., 2015).
This aggravates the light limitation of photosynthesis, especially at low light intensity (Figures 3b and 2e). Secondly, severe damage of PSI, in connection with only minor inhibition of PSII, results in a high reduction state of the PC pool ( Figure 3d) and, more importantly, of the PQ pool, which leads to acceptor side limitation at PSII, seen as low qP (Supporting Information: Figure  catalysed by the STN7 and STN8 kinases, respectively (Bellafiore et al., 2005;Bonardi et al., 2005), was substantially enhanced in PSI photoinhibited leaves (Figure 7b). Both kinases are redox-regulated and principally activated by the reduction of the PQ pool. STN7 kinase is additionally regulated by the stromal Trx system (Rintamäki et al., 1997(Rintamäki et al., , 2000, being inhibited by reduced Trx in conditions of excess light (Ancín et al., 2019). Nevertheless, the stromal redox network of PSI photoinhibited plants was maintained fairly oxidized ( Figure 7d), contrary to that of the PQ pool, thereby favouring high STN7 kinase activity and LHCII phosphorylation in the thylakoid membrane throughout the different light conditions, also under high light illumination (Figure 7b). The high level of LHCII phosphorylation leads to formation of the PSI-LHCI-LHCII complex (Figure 7c), which allows PSI to receive more excitation energy from the LHCII-lake (Benson et al., 2015;Grieco et al., 2015;Schiphorst et al., 2021). PSII core phosphorylation, on the other hand, destabilizes the larger C 2 S 2 M 2 and C 2 S 2 M PSII supercomplexes (Figure 6e), lowering the excitation energy partitioning from LHCII-lake to PSII reaction centres (Dietzel et al., 2011). The reducing capacity of the stromal Trx system was restored to control levels during the 24 h recovery period after PSI photoinhibition (Figure 7d), although the amount of functional PSI did not recover completely (Figure 2a). Restoration of Trx system activity was reflected in an increase in the relative amount of reduced FBPase in light (Figure 7d), which can be seen as lower Φ NA F (Figure 3c).
Restoration of the Trx system was also seen in the activity of STN7 kinase, demonstrated by a lowered phosphorylation level of LHCII in high light-illuminated leaves (Figure 7b), which in turn was reflected in excitation energy distribution ( Figure 7a). This data implies that the capacity of light reactions to reduce these stromal components increased during the 24 h recovery period.
Directly after PSI photoinhibition, the CO 2 assimilation and photochemical yield of PSII (Φ II ) decreased in synchrony (Figures 2e   and 3a). However, the situation was changed after the 24 h recovery period and, especially under growth light illumination, the CO 2 assimilation recovered more than the photochemical yield of PSII (Φ II ).
Such a large discrepancy between the photochemical yield of PSII (Φ II ) and the CO 2 assimilation rate can only be explained by the CBB cycle being favoured over other electron sinks downstream from ETC ( Figure 4). Under growth light illumination, NA and malate valve comprise other probable sinks for reducing power. However, Fd has a lower affinity for nitrite reductase than for FNR, which implies that nitrite reduction functions efficiently only when Fd supply exceeds its consumption in NADPH formation (Baysdorfer & Robinson, 1985;Rachmilevitch et al., 2004). In addition, the chloroplastic malate dehydrogenase, functioning in the malate valve, is efficiently activated F I G U R E 8 Scheme of Photosystem I (PSI) photoinhibition and subsequent induction of regulatory mechanisms postulated to allow PSI-deficient plants to restore CO 2 assimilation. (a) Balanced function of photosynthetic light reactions and stromal metabolism under steadystate environmental conditions. Photosynthetic control, and the phosphorylation of light-harvesting complex II (LHCII) and PSII core proteins, are modestly activated and reducing power from light reactions is fluently allocated to CO 2 assimilation and other stromal sinks. (b) PSI photoinhibition limits linear electron transfer (LET). Damaged PSI reaction centres quench the excitation energy, and the plastoquinone (PQ) pool gets reduced limiting PSII activity. (c) PSI photoinhibition exerts direct effects on regulation mechanisms of photosynthesis. Firstly, PSI deficiency restricts electron flow to stromal acceptors and key reducing enzymes activating the Calvin-Benson-Bassham (CBB) cycle and other stromal components. Secondly, relaxation of the proton gradient (pmf) diminishes photosynthetic control at cytochrome b 6 f complex (Cyt b 6 f), facilitating electron flow to the plastocyanin (PC) pool. Thirdly, the reduction of the PQ pool further enhances the activity of Stn7 and Stn8 kinases (phosphorylate the LHCII and PSII core proteins, respectively) leading to the reorganization of the light-harvesting antenna system to favour excitation of PSI. Notably, the canonical inhibition of LHCII phosphorylation in high light is prevented due to the lack of reduced Trx in the stroma. only when excess NADPH accumulates (Selinski & Scheibe, 2018).
Therefore, it is conceivable that the CBB cycle is favoured over nitrite reduction and/or malate valve in these conditions since the sink capacity of the CBB cycle is increased.
4.4 | PSI inhibition-related increase in ATP synthase and Cyt b 6 f amounts modulates the regulation of light reactions As a response to PSI photoinhibition, an increase in the abundance of ATP synthase and Cyt b 6 f took place and the elevated levels were maintained during the subsequent 24 h recovery period (Figures 6c-e). The factors controlling the stoichiometric biosynthesis of Cyt b 6 f and ATP synthase are poorly understood (Schöttler et al., 2015), yet it is clear that Cyt b 6 f, ATP synthase and the enzymes of the CBB cycle are tightly coregulated to maintain the balance between light reactions and stromal metabolism (Schöttler & Tóth, 2014;Vanlerberghe et al., 2019;Yamori et al., 2010). It is therefore likely that PSI photoinhibition-induced imbalance in ETC initiates a redox cascade to upregulate the synthesis of these protein complexes. This is in line with previous observations showing that chronic high reduction state of the PQ pool in stn7 mutant and PSIinhibited pgr5 mutant induces an accumulation of ATP synthase (Suorsa et al., 2012;Tikkanen et al., 2006).
The elevated abundance of ATP synthase (Figure 6c) was reflected as an increase in thylakoid proton conductivity (g H+ ) (Figure 5b), although the contribution by several ion channels and transporters (Armbruster et al., 2017;Spetea et al., 2017) cannot be neglected. Nevertheless, the higher g H+ lowers light-induced pmf (ECS t ) and thus decreases the photosynthetic control and non-photochemical quenching of excitation energy (Figures 5a and 3a,d). Consequently, the electron flow through Cyt b 6 f is enhanced not only by the increase in the abundance of the complex but also by a decrease in photosynthetic control. These modulations in Cyt b 6 f increase the reduction state of PC ( Figure 3d) and decrease PSI donor side limitation (Figure 3b), which collectively allow the remaining undamaged PSI centres to function more efficiently ( Figure 3c). It is evident that the increase in Cyt b 6 f and ATP synthase improves the capacity of the photosynthetic apparatus when the abundance of functional PSI is strongly reduced.
Noteworthy, even though the abundance of ATP synthase was maintained at a high level also during the 24 h recovery period ( Figure 6c), g H+ returned close to that of control plants (Figure 5b). This provides evidence that upon the recovery period plants restore the capability to tune the activity of the ATP synthase. The Trx system contributes to the regulation of proton conductivity, which declines if the thiol-redox state of chloroplast rises (Nikkanen et al., 2018). The lack of PGR5 protein is known to abolish the Trx-dependent control of proton conductivity, suggesting that this phenomenon is mediated by PGR5 (Avenson et al. 2005;Nikkanen et al., 2018). Thus, the re-establishment of the thiol-redox state during the recovery period ( Figure 7d) likely contributes to the restoration of control-type proton conductivity in PSI photoinhibited plants, restoring the photosynthetic control and nonphotochemical quenching (Figure 3a,d). Our results highlight that the regulatory plasticity of the photosynthetic machinery provides the capacity to acclimate not only to changes in light conditions but also provides the ability to acclimate to any factor disturbing the homoeostasis of the electron transfer chain. We conclude that the light acclimation is composed not only of the responses to changes in light conditions but also of the mechanism mitigating the consequences of photoinhibition.