Abstract
Elysia chlorotica is a kleptoplastic sea slug that preys on Vaucheria litorea, stealing its plastids which then continue to photosynthesize for months inside the animal cells. We investigated the native properties of V. litorea plastids to understand how they withstand the rigors of photosynthesis in isolation. Transcription of specific genes in laboratory-isolated V. litorea plastids was monitored up to seven days. The involvement of plastid-encoded FtsH, a key plastid maintenance protease, in recovery from photoinhibition in V. litorea was estimated in cycloheximide-treated cells. In vitro comparison of V. litorea and spinach thylakoids was applied to investigate ROS formation in V. litorea. Isolating V. litorea plastids triggered upregulation of ftsH and translation elongation factor EF-Tu (tufA). Upregulation of FtsH was also evident in cycloheximide-treated cells during recovery from photoinhibition. Charge recombination in PSII of V. litorea was found to be fine-tuned to produce only small quantities of singlet oxygen (1O2). Our results support the view that the genetic characteristics of the plastids themselves are crucial in creating a photosynthetic sea slug. The plastid’s autonomous repair machinery is likely enhanced by low 1O2 production and by upregulation of FtsH in the plastids.
Highlight Isolated Vaucheria litorea plastids exhibit upregulation of tufA and ftsH, key plastid maintenance genes, and produce only little singlet oxygen. These factors likely contribute to plastid longevity in kleptoplastic slugs.
Introduction
Functional kleptoplasty in photosynthetic sea slugs depends on two major components: the first is a slug capable of stealing plastids and retaining them functional within its cells, the second a plastid with a specific genetic repertoire (de Vries et al., 2015). All species that are able to do this belong to the Sacoglossan clade (Rumpho et al., 2011; de Vries et al., 2014). These slugs are categorized based on their plastid retention times, i.e. no retention, short-term retention (hours to ~10 days) and long-term retention species (≥10 days to several months) (Händeler et al., 2009). The record holding slug Elysia chlorotica can retain plastids for roughly a year (Green et al., 2000). The mechanisms utilized by the slugs to selectively sequester plastids from their prey algae remain uncertain, although recent studies have shown that in E. chlorotica it is an active process reminiscent of that observed for symbiotic algae and corals (Chan et al., 2018). The slugs possibly rely on scavenger receptors and thrombospondin type-1 repeat proteins for plastid recognition (Clavijo et al., 2020).
The sacoglossan’s ability to sequester plastids tends to distract attention from the unique features of the sequestered organelle, forming the second component of a photosynthetic slug system. Long-term retention sea slugs are only able to maintain functional plastids from a restricted list of siphonaceous algae and usually from only one species. Some sacoglossa have a wide selection of prey algae, but long-term retention of plastids is still limited to specific algal sources (Christa et al., 2013; de Vries et al., 2013). The native robustness of some plastid types was noticed decades ago, and early on suggested to contribute to their functionality inside animals (Giles and Sarafis, 1972; Trench et al., 1973 a, b). Studies focusing on the specific properties of the algal plastids, however, are scarce. Reduction of the plastid genome (plastome) during evolution has stripped the organelle of many genes required for self-maintenance (Martin, 2003), but genomic analysis of algal plastomes suggests that three genes (tufA, ftsH and psbA) could be among those critical for plastid maintenance inside a slug cell (de Vries et al., 2013). Out of the three, psbA remains in all plastomes, including those of higher plants, whereas tufA and ftsH are encoded by most algal plastid genomes (Baldauf and Palmer, 1990; Oudot-Le Secq et al., 2007; de Vries et al., 2013). It has been suggested that the plastid-encoded translation elongation factor EF-Tu (tufA) helps maintain translation, specifically of the thylakoid maintenance protease FtsH (ftsH) involved in the repair cycle of Photosystem II (PSII) (de Vries et al., 2013). FtsH degrades the D1 protein (psbA) of damaged PSII before the insertion of de novo synthesized D1 into PSII (Mulo et al., 2012; Järvi et al., 2015). Without continuous replacement of the D1 protein, light-induced damage to PSII would rapidly curtail photosynthesis (Tyystjärvi and Aro, 1996).
Unlike all other known plastid sources of long-term retention slugs, Vaucheria litorea (Fig. 1), the sole prey of E. chlorotica, is not a green but a yellow-green alga, with plastids derived from red algal lineage through secondary endosymbiosis (Cruz et al., 2013) (Fig. 1B). The plastome of V. litorea possesses the three important genes (de Vries et al., 2013). Furthermore, the plastid-encoded FtsH of V. litorea has been shown to carry the critical metalloprotease domain that is not present in ftsH of other prey algae of long-term retention slugs (Christa et al., 2018). Here, we show that isolated plastids of V. litorea (Fig. 1C) maintain highly specific transcription of their genes, and exhibit adequate genetic autonomy in their capability to recover from light induced damage of PSII, i.e. photoinhibition. We also estimated reactive oxygen species (ROS) production in the thylakoid membranes of V. litorea. While our results highlight the importance of terminal electron acceptors downstream of Photosystem I (PSI) in limiting ROS production, we show that PSII of V. litorea is fine-tuned to decrease the yield of the highly reactive singlet oxygen (1O2). The consequences of our findings to light-induced damage and longevity of the plastids inside photosynthetic sea slugs are discussed in detail.
Materials and Methods
Organisms and culture conditions
Spinach, Spinacia oleracea L. Matador (Nelson Garden, Tingsryd, Sweden), and V. litorea C. Agardh 1823 (SCCAP K-0379) were grown in SGC 120 growth chambers (Weiss Technik UK, Loughborough, United Kingdom) in 8/12 h and 12/12 h light/dark cycles, respectively. Growth light (Master TL-D 36W/840; Philips, Amsterdam, The Netherlands) photosynthetic photon flux density (PPFD) was set to 40 μmol m−2s−1 for both species. Temperature was maintained at 22 °C for spinach and 17 °C for V. litorea. Spinach plants used in the experiments were approximately 2 months old. V. litorea was grown in 500 ml flasks in f/2 culture medium (modified from Guillard and Ryther, 1962) made in 1% (m/v) artificial sea water (Sea Salt Classic, Tropic Marin, Wartenberg, Germany). V. litorea cultures were routinely refreshed by separating 1-4 g of inoculate into new flasks, and cultures used in the experiments were 1-2 weeks old. Nuclei of V. litorea were stained for microscopy with Hoechst 33342 (Thermo Scientific, Waltham, MA, USA) using standard protocols. In vivo transmission electron microscope (TEM) images were taken after freeze-etch fixation. The sea slug Elysia timida and its prey alga Acetabularia acetabulum were routinely maintained as described earlier (Schmitt et al., 2014; Havurinne and Tyystjärvi, 2020).
Gene expression of isolated V. litorea plastids
Plastid isolation from V. litorea was performed based on Green et al. (2005). Briefly, filaments were cut to small pieces, resuspended in 40 ml of isolation buffer (see Table 1) and homogenized with ULTRA-TURRAX® (IKA, Staufen, Germany) using four short bursts at 8000 rpm. The homogenate was filtered twice through a layer of Miracloth (Calbiochem, Darmstadt, Germany), centrifuged (1900 x g, 5 min) and the pellet was resuspended in 1 ml of isolation buffer. Percoll solution containing 0.25 M sucrose was diluted to a 75 and 30% solution with 1xTE buffer containing 0.25 M sucrose. The sample was layered between the two dilutions and the assemblage was centrifuged (3500 x g, 20 min) in a swing-out rotor with no deceleration. Intact plastids were collected from the interphase and washed twice by centrifugation (2200 x g, 3 min) with isolation buffer lacking BSA. All steps were carried out at 4 °C in the dark. TEM imaging of the plastids was done after fixing the samples using glutaraldehyde and cryo-fixation followed by freeze substitution.
Plastids were kept in isolation buffer for seven days in routine culturing conditions. RNA was isolated at different time points using Spectrum™ Plant Total RNA Kit (Sigma-Aldrich, St. Louis, MO, USA). Aliquots with 50 ng RNA were subjected to DNAse treatment (Thermo Scientific), and treated aliquots amounting to 10 ng RNA were used for cDNA synthesis (iScript™ cDNA Synthesis Kit, BioRad, Hercules, CA, USA). Quantitative real-time PCR was carried out using a StepOnePlus (Applied Biosystems, Foster City, CA, USA) and reagents from BioRad. The primers used in the qPCR were designed using Primer3 (http://frodo.wi.mit.edu/primer3); the primer sequences are listed in Supplementary Table S1 at JXB online. Every reaction was done with technical triplicates and results were analyzed using the ΔΔCt method (Pfaffl, 2001), in which the qPCR data were double normalized to rbcL and time point 0 (immediately after plastid isolation).
In vivo photoinhibition
The capacity to recover from photoinhibition was tested in spinach leaves and V. litorea cells in the presence of cycloheximide (CHI), a cytosolic translation inhibitor. Spinach leaf petioles were submerged in water containing 1 mM CHI and incubated for 24 h in the dark. The incubation was identical for V. litorea cells, except that the cells were fully submerged in f/2 medium supplemented with 1 mM CHI. Control samples were treated identically without CHI. The samples were then exposed to white light (PPFD 2000 μmol m−2 s−1) for 60 min and subsequently put to dark and thereafter low light (PPFD 10 μmol m−2 s−1) to recover for 250 min. Temperature was maintained at growth temperatures of both species using a combination of a thermostated surface and fans. The petioles of spinach leaves were submerged in water (-/+ CHI) during the experiments. Cell clusters of V. litorea were placed on top of the thermostated surface on a paper towel moistened thoroughly with f/2 medium (-/+ CHI). PSII activity was estimated by measuring the ratio of variable to maximum fluorescence (FV/FM) (Genty et al., 1989) with PAM-2000 (Walz, Effeltrich, Germany) fluorometer. During the light treatments, FV/FM was measured from samples that were dark acclimated for <5 min, except for the final time point, where the samples were dark acclimated for 20 min. The light source used for all high-light treatments discussed in this study was an Artificial Sunlight Module (SLHolland, Breda, The Netherlands).
Membrane proteins were isolated at timepoints indicated in the figures. The same area where FV/FM was measured (approximately 1 cm2) was cut out of the leaves/algal clusters and placed in a 1 ml Dounce tissue grinder (DWK Life Sciences, Millville, NJ, USA) filled with 0.5 ml of osmotic shock buffer (Table 1) and ground thoroughly. The homogenate was filtered through one layer of Miracloth and centrifuged (5000 x g, 5 min). The pellet containing the membrane protein fraction was resuspended in 50 μl of thylakoid storage buffer. The samples were stored at −80 °C until use. Membrane protein samples containing 1 μg total Chl were solubilized and separated by electrophoresis on a 10 % SDS-polyacrylamide gel using Next Gel solutions and buffers (VWR, Radnor, PA, USA). Proteins were transferred to Immobilon-P PVDF membranes (MilliporeSigma, Burlington, MA, USA). FtsH was immunodetected using antibodies raised against Arabidopsis thaliana FtsH5, reactive with highly homologous proteins FtsH1 and FtsH5, or FtsH2, reactive with FtsH2 and FtsH8 (Agrisera, Vännäs, Sweden). Western blots were imaged using goat anti-rabbit IgG (H+L) alkaline phosphatase conjugate (Life Technologies, Carlsbad, CA, USA) and CDP-star Chemiluminescence Reagent (Perkin-Elmer, Waltham, MA, USA). Protein bands were quantified with Fiji (Schindelin et al., 2012).
Experiments with E. timida were performed on freshly fed individuals. Slugs were kept in the dark overnight both in the absence and presence of 10 mg/ml lincomycin in 3.7 % artificial sea water and then exposed to high light (PPFD 2000 μmol m−2 s−1) in wells of a 24 well-plate filled with artificial sea water for 40 min. Temperature was maintained at 23 °C throughout the treatment. The slugs were then put to recover overnight in low light (PPFD <20 μmol m−2 s−1) in their growth conditions. FV/FM was measured with PAM-2000 after a minimum 20 min dark period as described earlier (Havurinne and Tyystjärvi, 2020).
Isolation of functional thylakoids for in vitro experiments
Functional thylakoids were isolated as described earlier (Hakala et al., 2005) after 24h dark incubation. One spinach leaf per isolation was ground in a mortar in thylakoid isolation buffer (Table 1). The homogenate was filtered through a layer of Miracloth and pelleted by centrifugation (5000 x g, 5 min). The pellet was resuspended in osmotic shock buffer, centrifuged (5000 x g, 5 min) and the resulting pellet was resuspended in thylakoid storage buffer. Chl concentration was determined spectrophotometrically in 90 % acetone using extinction coefficients for Chls a and b (Jeffrey and Humphrey, 1975). Thylakoid isolation from V. litorea was performed using the same procedure, by grinding 2-5 g of fresh cell mass per isolation. The cell mass was briefly dried between paper towels before grinding. Chl concentration from V. litorea thylakoids was determined in 90% acetone using coefficients for Chls a and c1 + c2 (Jeffrey and Humphrey, 1975). Protein concentrations of the thylakoid suspensions were determined with DC™ Protein Assay (Bio-Rad, Hercules, CA, USA). Thylakoids used in functional experiments were kept on ice in the dark and always used within a few hours of isolation.
Photosystem stoichiometry
Photosystem stoichiometry was measured from thylakoid membranes with an EPR spectroscope Miniscope MS5000 (Magnettech GmbH, Berlin, Germany) as described earlier (Tiwari et al., 2016; Nikkanen et al., 2019). EPR spectra originating from oxidized tyrosine-D residue of PSII (TyrD+) and reaction center Chl of PSI (P700+) of concentrated thylakoid samples (2000 μg Chl ml−1 in storage buffer) were measured in a magnetic field ranging from 328.96 to 343.96 mT during illumination (PPFD 4000 μmol m−2 s−1) (Lightningcure LC8; Hamamatsu Photonics, Hamamatsu City, Japan) and after a subsequent 5 min dark period in the absence and presence of 50 μM 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU). The dark stable TyrD+ EPR signal (PSII signal), measured after the post illumination period in the absence of DCMU, and the P700+(PSI signal), measured during illumination in the presence of DCMU, were double integrated to determine photosystem stoichiometry.
In vitro photoinhibition
For in vitro photoinhibition experiments, thylakoids were diluted to a total Chl concentration of 100 μg ml−1 in photoinhibition buffer (Table 1), and 1 ml sample was loaded into a glass beaker submerged in a water bath kept at 22 °C. The samples were exposed to white light (PPFD 1000 μmol m−2 s−1) and mixed with a magnet during the 60 min treatments. Aliquots were taken at set intervals to determine PSI or PSII activities using a Clark-type oxygen electrode (Hansatech Instruments, King’s Lynn, England). The sample concentration in the activity measurements was 20 μg total Chl ml−1 in 0.5 ml of PSI or PSII measuring buffer (Table 1). PSI activity was measured as oxygen consumption, whereas PSII activity was measured as oxygen evolution. Both activities were measured at 22 °C in strong light (PPFD 3200 μmol m−2 s−1) from a slide projector. The rate constant of PSII photoinhibition (kPI) was obtained by fitting the loss of oxygen evolution to a first-order reaction equation with Sigmaplot 13.0 (Systat Software, San Jose, CA, USA), followed by dark correction, i.e. subtraction of the dark inactivation rate constant from the initial kPI.
Lipid peroxidation was measured by detecting malondialdehyde (MDA) formation (Heath and Packer, 1968). A thylakoid suspension aliquot of 0.4 ml was mixed with 1 ml of 20 % trichloroacetic acid containing 0.5 % thiobarbituric acid, incubated at 80 °C for 30 min and cooled down on ice for 5 min. Excess precipitate was pelleted by centrifugation (13500 x g, 5 min), and the difference in absorbance between 532 and 600 nm (Abs532-600) was measured as an indicator of the relative amount of MDA in the samples. Protein oxidation was determined by detecting protein carbonylation with Oxyblot™ Protein Oxidation Detection Kit (MilliporeSigma, Burlington, MA, USA). Thylakoid aliquot amounting to a protein content of 45 μg was taken at set time points and 10 mM dithiothreitol was used to prevent further protein carbonylation. The samples were prepared according to the manufacturer’s instructions and proteins were separated in 10 % Next Gel SDS-PAGE (VWR). Carbonylated proteins were detected with Immobilon Western Chemiluminescent HRP Substrate (MilliporeSigma).
The maximum oxidation of P700 (PM) was estimated in an additional experiment. Thylakoids equivalent to 25 ug Chl in 50 μl of photoinhibition buffer were pipetted on a Whatman filter paper (grade 597; Cytiva, Marlborough, MA, USA). The filter was placed inside the lid of a plastic Petri dish, and the bottom of the Petri dish was placed on top of the lid. Photoinhibition buffer was added to the sample from the small openings on the sides of the assemblage. The thylakoids were then illuminated with high light (PPFD 1000 μmol m−2 s−1) and the temperature was maintained at 22 °C using a thermostated surface. FV/FM and PM were measured using a 700 ms high-light pulse (PPFD 10000 μmol m−2 s−1) with Dual-PAM 100 (Walz) (Schreiber, 1986; Schreiber and Klughammer, 2008) at set intervals. The high-light treated samples were dark acclimated for <5 min prior to the measurements.
1O2 measurements
1O2 was measured from thylakoids diluted to 100 μg total Chl ml−1 in 0.3 ml of photosystem measuring buffer, using the histidine method described earlier (Telfer et al., 1994; Rehman et al., 2013). Continuously stirred thylakoid samples were exposed to high light (PPFD 3200 μmol m−2 s−1) from a slide projector at 22 °C in the presence and absence of 20 mM histidine. Oxygen consumption was measured for 60 s using an oxygen electrode (Hansatech), and the difference in the oxygen consumption rates in the presence and absence of histidine was taken as an indicator of 1O2 production. PSII electron transfer activity (H2O to DCBQ) in the same conditions was 124.7 (SE±15.4) and 128.4 (SE±10.7) μmol O2 mg Chl−1 h−1 in spinach and V. litorea samples, respectively, containing 20 μg Chl ml−1.
PSII charge recombination measurements
Flash-induced oxygen evolution was recorded at room temperature using a Joliot-type bare platinum oxygen electrode (PSI, Brno, Czech Republic) (Joliot and Joliot, 1968) from thylakoids diluted in photosystem measuring buffer to 50 μg Chl ml−1 and supplemented with 50 mM KCl, essentially as described in Antal et al. (2009). 200 μl of sample was pipetted on the electrode and kept in the dark for 10 min before the measurements. The samples were then exposed to a flash train consisting of 15 single-turnover flashes (4 ns/pulse) at one second intervals, provided by a 532 nm Nd:YAG laser (Minilite, Continuum, San Jose, CA, USA). Charge recombination within PSII was probed by exposing the samples to a preflash and different dark times between the preflash and the flash train used for recording the oxygen traces.
The decay of Chl a fluorescence yield after a 30 μs single turnover flash (maximum PPFD 100 000 μmol m−2 s−1) were measured at room temperature from 1 ml samples of thylakoids using FL200/PS fluorometer (PSI). Measurement length was 120 s and 8 datapoints/decade were recorded (2 in the presence of DCMU). The first datapoint was recorded 150 μs after the flash. Single turnover flash and measuring beam voltages were set to 100 % and 60 % of the maximum, respectively. The samples were diluted in photosystem measuring buffer to a total Chl concentration of 20 μg ml−1. A set of samples was poisoned with 20 μM DCMU to block electron transfer at the reducing side of PSII.
Thermoluminescence was measured from thylakoids using a custom setup (Tyystjärvi et al., 2009). Thylakoids were diluted to a total Chl concentration of 100 μg ml−1 in photosystem measuring buffer (Table 1) in the presence and absence of 20 μM DCMU, and a volume of 100 μl was pipetted on a filter paper disk that was placed inside the cuvette of the measuring apparatus. The samples were dark acclimated for 5 min before the onset of cooling to −20 °C by a Peltier element (TB-127-1,0-0,8; Kryotherm, Carson City, NV, USA). The samples were then exposed to a flash (E = 1 J) from a FX-200 Xenon lamp (EGandG, Gaithersburg, MD, USA) and heated at a rate of 0.47 °C s−1 up to 60 °C while simultaneously recording luminescence emission.
In vivo P700 redox kinetics
Redox kinetics of P700 were measured as described by Shimakava et al., (2019) using Dual-PAM 100 (Walz). Spinach plants and V. litorea cells were kept in darkness for at least 2 h before the measurements. Anaerobic conditions were obtained using a custom cuvette described in Havurinne and Tyystjärvi (2020). For spinach leaf cutouts, the cuvette was flushed with nitrogen. A combination of glucose oxidase (8 units/ml), glucose (6 mM) and catalase (800 units/ml) in f/2 culture medium was used to create anaerobic conditions for V. litorea cells. All samples were treated with 15 s of far red light (PFD 120 μmol m−2 s−1) and a subsequent darkness lasting 25 s prior to firing a high-light pulse (780 ms, PPFD 10 000 μmol m-2 s-1).
Results
Isolated V. litorea plastids maintain regulated gene expression
Laboratory-isolated V. litorea plastids exhibited differentially regulated gene expression even after seven days in isolation (Fig. 2). The orientations of selected genes in V. litorea plastome are shown in Fig. 2A. PSII core subunit genes psbA, psbB, psbC and psbD were downregulated after day 3 of the isolation period, while psbB and psbD, encoding CP47 and D2 proteins of PSII, reached a stationary level after five days, and the transcription of the genes encoding PSII proteins CP43 (psbC) and D1 (psbA) were among those downregulated most significantly (Fig. 2B). The main protein of PSII targeted for degradation after photoinhibition is D1, whereas release of CP43 from the PSII core has been suggested to precede D1 degradation in higher plants (Aro et al., 2005). One gene, psbH, encoding a small PSII subunit involved in proper PSII assembly in cyanobacteria (Komenda et al., 2005), exhibited stationary transcript levels throughout the isolation period, similar to the gene encoding PSI reaction center subunit PsaA. Transcription of ftsH and tufA, encoding the maintenance protease FtsH and the translation elongation factor EF-Tu, followed an upward trajectory throughout the experiment (Fig. 2B). We also tested the genetic autonomy of plastids sequestered by E. timida that feeds on A. acetabulum. Subjecting the slugs to high light for 40 min resulted in a drastic decrease in PSII photochemistry (FV/FM), but the kleptoplasts inside the slugs were capable of restoring PSII activity back to 78 % of the initial level during a 20 h recovery period. Subjecting the slugs to lincomycin, a plastid specific translation inhibitor (Mulo et al., 2003), however, almost completely prevented the recovery (Fig. 2C).
FtsH translation is enhanced in functionally isolated plastids of V. litorea during recovery from photoinhibition
Treating spinach leaves with CHI, a cytosolic translation inhibitor, resulted in faster loss of PSII activity in high light (Fig. 3A). Also PSII repair was impaired by CHI in spinach. V. litorea showed almost no effect of CHI during the same photoinhibition and recovery treatment (Fig. 3B). Using two different FtsH antibodies (FtsH 1+5 and FtsH 2+8), we tested the possible involvement of plastid-encoded FtsH of V. litorea in the unaffected PSII photochemistry in CHI treated samples. There were no differences in the relative protein levels of FtsH between control and CHI treated spinach during the experiment (Fig. 3C). Genes for FtsH reside in the nucleus in spinach, and our results suggest that the CHI treatment did not inhibit cytosolic translation in the leaves entirely, although de novo synthesis of proteins could not be tested by radiolabeling experiments. In V. litorea, CHI treatment increased FtsH levels towards the end of the experiment (Fig. 3D). This suggests that not only is expression of plastome genes active in functionally isolated plastids of V. litorea, but the translation of specific genes such as ftsH can be upregulated when the plastids are deprived from normal cytosolic governance.
Thylakoids of V. litorea exhibit moderate photoinhibition of PSII and elevated ROS damage, but produce little 1O2
Basic photosynthetic parameters of isolated thylakoids from spinach and V. litorea are shown in Table 2. Photoinhibition of PSII during a 60 min high-light treatment of isolated thylakoids proceeded according to first-order reaction kinetics (Tyystjärvi and Aro, 1996) in both species (Fig. 4A). However, spinach thylakoids were more susceptible to damage, as indicated by the larger rate constant of dark-corrected PSII photoinhibition (kPI) (Table 2). General oxidative stress assays of lipids and proteins of the thylakoid membranes exposed to high light showed more ROS damage in V. litorea than in spinach thylakoids during the treatment (Fig. 4B,C). Measurements of 1O2 production, the main ROS produced by PSII (Krieger-Liszkay, 2005; Pospíšil, 2012), from isolated thylakoids showed that the rate of 1O2 production in V. litorea is only half of that witnessed for spinach (Fig. 5A). This suggests that the main ROS, causing the in vitro oxidative damage to lipids and proteins (Fig. 4B,C) in V. litorea, are partially reduced oxygen species produced by PSI.
V. litorea produces only little 1O2, likely due to slow PSII charge recombination
We probed charge recombination reactions within PSII using three different methods to investigate the role of PSII in the low 1O2 yield in V. litorea thylakoids (Fig. 5A). First, we measured flash-induced oxygen evolution from isolated thylakoids of spinach and V. litorea. After 10 min dark acclimation, thylakoids from both species exhibited a typical pattern of oxygen evolution, i.e. the third flash caused the highest oxygen yield due to the predominance of the dark-stable S1 state of the oxygen evolving complex (OEC), after which the oxygen yield oscillated with a period of four until dampening due to misses and charge recombination reactions (Fig. 5B, top curves). A single turnover pre-flash treatment makes S2 the predominant state. A 10 s dark period after the pre-flash treatment was not long enough to cause noticeable changes in the S-state distribution in either species, as can be seen from the middle curves of Fig. 5B, where the second flash of the flash train causes the highest yield of oxygen. In spinach, 100 s darkness after the pre-flash treatment resulted in nearly complete restoration of the original S-states, whereas in V. litorea the second flash still yielded a considerable amount of oxygen (Fig. 5B, bottom curves). This is likely due to slow charge recombination between QB− and the S2 state of the OEC in V. litorea (Pham et al., 2019). The modeled percentage S-state distributions of OEC from spinach and V. litorea after different dark times between the pre-flash and the flash train are shown in Supplementary table S2.
Next, we measured the decay of Chl a fluorescence yield after a single turnover flash from thylakoids in the absence and presence of the PSII electron transfer inhibitor DCMU. Fluorescence decay in the absence of DCMU reflects QA− reoxidation mainly by electron donation to QB and QB−. In the presence of DCMU, fluorescence decay is indicative of QA− reoxidation through various charge recombination reactions (Mamedov et al., 2000), some of which generate the harmful triplet P680 Chl through the intermediate P680+Pheo− radical pair (Sane et al., 2012). The decay of fluorescence yield was slower in V. litorea thylakoids than in spinach both in the absence and presence of DCMU (Fig. 5C). In the absence of DCMU, the slower kinetics in V. litorea shows that electron transfer from QA− to QB is not as favorable as in spinach. The slow decay of fluorescence in the presence of DCMU indicates slow S2QA− charge recombination.
Thermoluminescence Q and B bands from thylakoids in the presence and absence of DCMU, respectively, were also measured. For a description on the interpretation of thermoluminescence data, see Tyystjärvi and Vass, (2004) and Sane et al., (2012). Briefly, the thylakoid samples were dark acclimated for 5 min, cooled down to −20 °C, flashed with a single turnover Xenon flash and then heated with a constant rate. The luminescence emitted by the samples at different temperatures is proportional to the rate of the luminescence-producing charge recombination reactions between the S-states of the OEC and downstream electron acceptors, more specifically S2/QA− (Q band) and S2,3/QB− (B band). The Q and B band emission peaks in spinach were at 15 and 28 °C, whereas in V. litorea they were at 14 and 24 °C (Fig. 5D). The lower peak temperatures in V. litorea would actually suggest that both QA− and QB− are less stable at room temperature in V. litorea than in spinach. However, the multiple pathways of recombination (Rappaport and Lavergne, 2009) obviously allow the luminescence-producing minor pathway to suggest destabilization of QA− in V. litorea (Fig. 5D) even if the total recombination reaction is slower in V. litorea than in spinach (Fig. 5B,C and Supplementary table S2). The thermoluminescence signal intensity was lower in V. litorea than in spinach, suggesting that the luminescence-producing reaction has a low yield in V. litorea. The narrow energy gap between QA and QB in V. litorea favors the probability of an electron residing with QA. Furthermore, a small QA−QB energy gap also increases the probability that S3QB− or S2QB− recombine directly and non-radiatively without producing triplet P680 and subsequently 1O2 (Ivanov et al., 2003; Sane et al., 2003; Ivanov et al., 2008; Sane et al., 2012).
In vitro high-light treatment lowers electron donation to methyl viologen and maximal oxidation of P700 in V. litorea
When PSI activity was estimated as electron transfer from DCPIP to methyl viologen (oxygen consumption), spinach PSI remained undamaged during in vitro high-light treatment, while V. litorea seemed highly susceptible to photoinhibition of PSI (Fig. 6A,B). We repeated the photoinhibition experiment, but this time PSII and PSI activities were monitored with Chl fluorescence and P700 absorption changes. Again, thylakoid membranes of spinach were more sensitive to photoinhibition of PSII during the high-light treatment than V. litorea (Fig. 6C,D). However, this time PSI functionality of both species decreased similarly when estimated as the maximum oxidation of P700 (PM). The decrease in PM was strong during the first 15 (V. litorea) or 30 min (spinach) of the light treatment, whereafter PM remained at a somewhat stationary level (Fig. 6C,D). The decrease in PM depended on electron transfer from PSII, as PM did not decrease in high light in spinach thylakoids in the presence of DCMU (Supplementary Fig. S4).
In both spinach and V. litorea, redox kinetics of P700, measured in aerobic conditions from thylakoids (Fig. 6E,F) were similar as their respective in vivo kinetics (Fig. 6E,F insets), i.e. P700 in V. litorea remained more oxidized during a light pulse than in spinach. Isolating thylakoids from V. litorea did, however, cause a decrease in P700 oxidation capacity. Unlike in spinach, P700 remains oxidized during a high-light pulse in intact V. litorea cells if oxygen is present, indicating that alternative electron sinks, such as flavodiiron proteins, function as efficient PSI electron acceptors V. litorea (Fig. 6E,F insets), probably protecting PSI against formation of ROS (Allahverdiyeva et al., 2015; Ilík et al., 2017; Shimakawa et al., 2019). In both species, P700 redox kinetics changed in the same way during the course of the high-light treatment of isolated thylakoids. The tendency of both species to maintain P700 oxidized throughout the high-light pulse in measurements done after 15 min treatment in high light is possibly due to decreasing electron donation caused by photoinhibition of PSII. At 45 min timepoint the damage to PSI is more severe, as indicated by a clear slowing down of P700 oxidation, which could be associated with problems in electron donation to downstream electron acceptors of PSI, such as ferredoxin (Fig. 6E,F).
Discussion
Upregulation of FtsH at the center of V. litorea plastid longevity
Previous studies have shown that the kleptoplasts stemming from V. litorea carry out de novo protein translation and are generally quite robust inside E. chlorotica (Green et al., 2000; Rumpho et al., 2001; Green et al., 2005). Our transcriptomic analysis of V. litorea plastids demonstrates active and regulated transcription of the plastome throughout the seven days of isolation we tested (Fig. 2), deepening our knowledge about the factors underpinning their native robustness. Considering gene orientation of the up- and downregulated genes suggests that e.g. ftsH and psbB, neighboring genes sharing the same orientation, do not constitute an operon (Fig. 2A).
Our results highlight the upregulation of ftsH and tufA during a period of several days after isolation of V. litorea plastids. Active transcription of these genes also occurs in the plastids of E. timida after a month of starvation (de Vries et al., 2013). FtsH protease is critical for the PSII repair cycle, where it is responsible for degradation of the D1 protein after pulling it out of the PSII reaction center. Recent findings in cyanobacteria, green algae and higher plants imply that FtsH is also important for quality control of a multitude of thylakoid membrane proteins and thylakoid membrane biogenesis (reviewed by Kato and Sakamoto, 2018). These findings may suggest that already the removal of the D1 protein from damaged PSII serves to protect from further photodamage and the production of ROS. The results of our photoinhibition experiments on the long-term retention slug E. timida may serve as a model of photoinhibition in other slugs, as they indicate that the kleptoplasts of E. timida possess a genetic toolkit capable of maintaining a PSII repair cycle (Fig. 2C).
We showed that the capacity of V. litorea plastids to recover from photoinhibition of PSII in the presence of CHI is nearly unaffected (Fig. 3B). While our CHI experiments on spinach need further exploration in terms of CHI effects, studies on the green alga Chlamydomonas reinhardtii (that also lacks ftsH in its plastome) have shown severe defects in PSII repair both during high-light and subsequent recovery when exposed to CHI (Fig. 3A, Wang et al., 2017). C. reinhardtii mutant lines have also been used to show that abundant FtsH offers protection from photoinhibition of PSII and enhances the recovery process (Wang et al., 2017). In C. reinhardtii, the FtsH hetero-oligomers responsible for D1 degradation are comprised of FtsH1 (A-type) and FtsH2 (B-type) (Malnoë et al., 2014). We probed the relative FtsH protein levels of V. litorea during the photoinhibition experiment using antibodies raised against A. thaliana A-(FtsH 1+5) and B-type FtsH (FtsH 2+8) in the absence and presence of CHI (Fig. 3D). At the end of the recovery period the CHI treated cells showed elevated levels of FtsH according to both tested antibodies. The elevated FtsH abundance did not enhance the recovery from photoinhibition of PSII in our experimental setup (Fig. 3B), but our results do point to a tendency of both, truly isolated (Fig. 2) and functionally isolated (Fig. 3) V. litorea plastids, to upregulate FtsH.
Low 1O2 yield does not prevent photoinhibition of PSII, but can help maintain efficient repair processes in V. litorea
A green alga that is nearly immune to photoinhibition of PSII, Chlorella ohadii, has been isolated from the desert crusts of Israel (Treves et al., 2013; 2016). Its resilience against photoinhibition of PSII has largely been attributed to very narrow energetic gap between QA and QB, favoring non-radiative charge recombination pathways within PSII that do not lead to 1O2 production (Treves et al., 2016). While V. litorea does not have as small energetic gap between QA and QB as C. ohadii (temperature difference of V. litorea Q- and B-band thermoluminescence peaks was 10 °C, whereas in C. ohadii it is only 2-4 °C), PSII charge recombination reactions of V. litorea appear to be very slow compared to those of spinach (Fig. 5B-D). Furthermore, the low 1O2 yield in V. litorea (Fig. 5A) suggests that the charge recombination reactions favor the direct non-radiative pathway. The low 1O2 yield in V. litorea likely factors into the lower dark-corrected rate constant of PSII photoinhibition in comparison to that of spinach thylakoids (Table 2) (Vass, 2011). All of our experiments, however, show that V. litorea does experience quite regular levels of PSII photoinhibition. This could indicate that the most important effect of the low 1O2 yield is protection of the autonomous maintenance machinery of the plastids, as 1O2 has been shown to be specifically harmful for the PSII repair cycle (Nishiyama et al., 2004).
V. litorea thylakoids are highly vulnerable to ROS in the absence of regular stromal electron sinks
Despite the lower rate constant of PSII photoinhibition (Table 2) and 1O2 yield (Fig. 5A), V. litorea thylakoids exhibited drastic oxidative damage to lipids and proteins under high light (Fig. 4C,D). Isolated thylakoids are stripped of the main electron sink of PSI, the Calvin-Benson-Bassham cycle, and comparing P700 redox kinetics of V. litorea cells and isolated thylakoids (Fig. 6F and inset) reveals that they are also, at least partially, devoid of a Mehler-like reaction that safely reduces oxygen to water (Allahverdiyeva et al., 2013). This suggests that catalysts of oxygen reduction in V. litorea are likely soluble and therefore lost during the isolation procedure. Angiosperm plants like spinach do not rely on a Mehler-like reaction and are susceptible to photoinhibition of PSI in fluctuating light (Shimakawa et al., 2019). The PSI photoprotection by Mehler-like reaction has been assigned to enhanced electron sink capacity that lowers the probability of one-electron reduction of oxygen to superoxide by PSI. In comparison to spinach, this would make intact plastids of V. litorea less reliant on other ROS detoxification components that detoxify superoxide and hydrogen peroxide in the water-water cycle (Asada, 1999). Conversely, loss of the Mehler-like reaction during thylakoid isolation would leave the thylakoids highly conducive for ROS production by PSI and very susceptible to oxidative damage of the entire photosynthetic machinery. This is likely behind the finding that V. litorea thylakoids lose the ability to reduce methyl viologen in a high-light treatment that does not affect spinach thylakoids (Fig. 6A,B). When damage to PSI was estimated as a decrease in PM, spinach and V. litorea thylakoids showed very similar responses to high light, with both species exhibiting a decrease in PSI activity until electron donation from PSII was diminished due to photoinhibition of PSII (Fig. 6C,D), as suggested earlier (Sonoike, 1995; 1996). This, in addition to the highly similar changes in the redox kinetics of P700 during the photoinhibition treatment (Fig. 6E,F) between the two species, would suggest that the decrease in oxygen consumption in V. litorea thylakoids is caused by a further, more severe damage to PSI than the process causing the decrease in PM. The nature of this reaction is not known but it may be caused by production of ROS due to continuing electron flow through PSI in thylakoids of V. litorea exhibiting a low rate constant of PSII photoinhibition (Table 2) and normally relying on stromal electron acceptors for protection of PSI.
PSI of V. litorea is not particularly prone to photoinhibition, but our results do confirm that the electron sinks of photosynthesis must be functional in order to avoid large scale oxidative damage. This is especially relevant for animals that host a foreign organelle where uncontrolled ROS production is detrimental (de Vries et al. 2015). Our recent results on the LTR slug E. timida show that oxygen functions as an alternative electron sink in the slug plastids (Havurinne and Tyystjärvi, 2020), but whether the record-holding E. chlorotica utilizes the oxygen dependent electron sinks provided by V. litorea (Fig. 6F inset) remains to be tested. As for the main electron sink of photosynthesis, the carbon fixation rates of the plastids inside E. chlorotica are comparable to the rates measured from V. litorea cells after incorporation (Rumpho et al., 2001), suggesting that carbon fixation is not a problem in E. chlorotica.
Conclusion
Plastids of V. litorea are genetically more autonomous than those of embryophytes, containing genes that help to maintain plastid functionality. Isolating the plastids triggers upregulation of the translation elongation factor EF-Tu and the central maintenance protease FtsH – a phenomenon that may be important for plastid longevity in the foreign cytosol of a sea slug. Low 1O2 yield protects the functionality of the plastid-encoded maintenance machinery and may slow down photoinhibition of PSII. Interruption of oxygen dependent alternative electron sinks upstream of PSI leads to large scale oxidative damage in V. litorea, suggesting that carbon fixation, the main electron sink of photosynthesis, needs to remain in near perfect working order to avoid destruction of the plastids. Our results support decades old data (Trench et al., 1973 a, b) suggesting that the native stability and associated peculiar functionality of the plastids themselves hold the key to long-term kleptoplast longevity in sacoglossans. Nature has evolved an elaborate suite of photoprotective mechanisms and the unique animal-kleptoplast association allows to explore them and even identify new ones.
Abbreviations
- 1O2
- singlet oxygen
- CHI
- cycloheximide
- DCBQ
- 2,6-dichloro-1,4-benzoquinone
- DCMU
- 3-(3,4-dichlorophenyl)-1,1-dimethylurea
- DCPIP
- 2,6-dichlorophenolindophenol
- FV/FM
- maximum quantum yield of PSII photochemistry
- kPI
- rate constant of PSII photoinhibition
- MDA
- malondialdehyde
- OEC
- oxygen evolving complex of PSII
- P680
- reaction center Chl of PSII
- P700
- reaction center Chl of PSI
- PM
- maximum oxidation of P700
- PPFD
- photosynthetic photon flux density
- PSI
- Photosystem I
- PSII
- Photosystem II
- ROS
- reactive oxygen species
- TEM
- transmission electron microscope
- TyrD+
- oxidized tyrosine-D residue of PSII
Supplementary data
Supplementary data are available at JXB online.
Table S1. List of primers used in qPCR experiment.
Table S2. Modeled S-state distribution of the OEC in spinach and V. litorea.
Fig. S1. EPR spectra from spinach and V. litorea thylakoids.
Fig. S2. Dark control treatments of in vitro PSII photoinhibition in spinach and V. litorea.
Fig. S3. Dark control treatments of in vitro PSI and PSII photoinhibition in spinach and V. litorea.
Fig. S4. In vitro PSI and PSII photoinhibition in DCMU treated spinach thylakoids.
Author Contributions
VH, SBG and ET planned the experiments. VH did all photosynthesis and 1O2 measurements and wrote the paper with comments from all authors; MH, supervised by SBG, did the gene expression measurements and TEM imaging; MA measured lipid peroxidation, protein oxidation and EPR spectra; SK developed the cuvette system for P700+ measurements; ET supervised the work.
Data Availability
The data that support the findings of this study are openly available in Mendeley Data at http://doi.org/10.17632/535dcxjt2d.1.
Acknowledgements
This work was supported by Academy of Finland (grant 333421, ET). VH was supported by Finnish Cultural Foundation, Finnish Academy of Science and Letters (Vilho, Yrjö and Kalle Väisälä fund), Turku University Foundation and University of Turku Graduate School. Sofia Vesterkvist is thanked for help with E. timida photoinhibition measurements. SBG would like to thank the German research council for funding through the CRC 1208-267205415 – and the SPP2237, and the group of U.-G. Maier (Marburg) for help with electron microscopy.
Footnotes
vetahav{at}utu.fi, maria.handrich{at}hhu.de, mikkoantinluoma{at}gmail.com, sergey.khorobrykh{at}utu.fi, gould{at}hhu.de, esatyy{at}utu.fi