The electron transfer properties of the three types of PSII were investigated by comparing the decay kinetics of the flash-induced fluorescence in membranes from A. marina, white-light (WL) grown C. thermalis and far-red-light (FR) grown C. thermalis. When forward electron transfer occurs (Figure 2A), the fluorescence decay comprises three phases (Crofts and Wraight, 1983; Vass et al., 1999): the fast phase (~0.5ms) is attributed to electron transfer from Q_A^- to Q_B or Q_B^- and the middle phase (~3ms) is generally attributed to Q_A^- oxidation limited by plastoquinone (PQ) entry to an initially empty Q_B site and/or by Q_BH₂ exiting the site prior to PQ entry (de Wijn and van Gorkom, 2001). These two phases had comparable time-constants in all samples (T₁=0.5–0.6 and T₂=3.5–5ms, Table 1). The fast electron transfer from Q_A^- to the non-heme iron possibly oxidized in a fraction of centres is too fast (t½~50 µs) to be detected here.

Figure 2 with 1 supplement see all

Download asset Open asset

Figure 2—source data 1

Fluorescence decay kinetics.

https://cdn.elifesciences.org/articles/79890/elife-79890-fig2-data1-v3.xlsx

Download elife-79890-fig2-data1-v3.xlsx

The slower decay phase is attributed to the charge recombination between Q_A^- and the Mn-cluster mostly in the S₂ state (see section 2.2) in centres where forward electron transfer to Q_B/Q_B^- did not occur. This phase was significantly slower in FR C. thermalis (T₃=14.3 ± 4.6 s) than in WL C. thermalis (T₃=5.6 ± 2.4 s) but had a similar amplitude in the two samples (Figure 2—figure supplement 1 and Table 1). In A. marina this phase had a bigger amplitude than in the two C. thermalis samples (Table 1), because it was superimposed to a non-decaying component of the fluorescence, that did not return to the original F₀ level even at 100 s after the flash (Figure 2—figure supplement 1). This non-decaying component, absent in the two C. thermalis samples, is attributed to centres without a functional Mn-cluster, in which P_D1⁺ is reduced by an electron donor that does not recombine in the minutes timescale (such as Mn²⁺, TyrD, or the ChlZ/Car side-path), with the consequence of stabilizing Q_A^- (Nixon et al., 1992; Debus et al., 2000). The fluorescence decay arising from the S₂Q_A^- recombination was slower in A. marina (T₃=10.8 ± 2.6 s) than in WL C. thermalis, but its overlap with the non-decaying component made the fit of its time-constant potentially less reliable.

Indeed, when the fluorescence decay due to charge recombination was measured in presence of the Q_B-site inhibitor DCMU (Figure 2B), the decay kinetics were bi-phasic in all samples, and no difference in the major S₂Q_A^- recombination phase (slow phase in Table 1, ~80% amplitude, T₃ ~6–7 s) was found between A. marina and WL C. thermalis. In contrast, the decay was significantly slower in FR C. thermalis, with the time-constant of the major S₂Q_A^- recombination phase (slow phase in Table 1, ~80% amplitude, T₃=10.4 ± 0.8 s) similar to that measured in the absence of DCMU. The shorter lifetime (~0.22–1 s) of the middle decay phase (amplitude 15–20%) was compatible with it originating from TyrZ^•(H⁺)Q_A^- recombination occurring either in centres lacking an intact Mn-cluster (Yerkes et al., 1983) or in intact centres before charge separation is fully stabilised, as proposed in Debus et al., 2000. The fluorescence decay in WL and FR C. thermalis also had an additional fast phase of small amplitude (5–6%), attributed to forward electron transfer in centres in which DCMU was not bound (Lavergne, 1983). Again, the A. marina traces included a non-decaying phase of fluorescence, attributed to centres lacking an intact Mn-cluster.

The fluorescence decay kinetics in membranes of Synechocystis sp. PCC6803 (Synechocystis), perhaps the best studied Chl-a containing cyanobacterium, were also measured as an additional control. The kinetics in Synechocystis membranes were comparable to those reported for WL C. thermalis (Appendix 1). The Synechocystis and A. marina fluorescence decay kinetics measured in membranes here are overall slower than those previously measured in cells (Cser et al., 2008). This difference is ascribed to pH and membrane potential effects, as discussed in Appendix 1, and illustrates the difficulty to use whole cells for such measurements.

To conclude, the forward electron transfer rates from Q_A^- to Q_B/Q_B^- are not significantly different in the three types of PSII. In contrast, the S₂Q_A^- recombination is slower in Chl-f-PSII of FR C. thermalis compared to Chl-a-PSII of WL C. thermalis and Chl-d-PSII of A. marina.

The efficiency of PSII water oxidation activity can be estimated by the flash-dependent progression through the S-states of the Mn-cluster. This can be measured by thermoluminescence (TL), which arises from radiative recombination of the S₂Q_B^- and S₃Q_B^- states (Rutherford et al., 1982). The TL measured in A. marina, WL C. thermalis, and FR C. thermalis membranes showed similar flash-dependencies in all three types of PSII (Appendix 2—figure 1), confirming and extending the earlier report (Nürnberg et al., 2018). Because the TL data presented some variability between biological replicates (Appendix 2), additional analyses were performed by polarography and absorption spectroscopy.

Figure 3 shows the flash-dependent oxygen evolution measured in A. marina, FR C. thermalis and Synechocystis membranes. The latter were used as a Chl-a-PSII control because the content of PSII in membranes of WL C. thermalis was too low to allow accurate O₂ polarography measurements (Figure 3—figure supplement 1D). As shown by fluorescence, no significant difference in forward electron transfer between the two types of Chl-a-PSII was observed (Appendix 1), and the use of Synechocystis membranes was therefore considered as a valid control.

Figure 3 with 1 supplement see all

Download asset Open asset

Figure 3—source data 1

Flash-dependent oxygen evolution.

https://cdn.elifesciences.org/articles/79890/elife-79890-fig3-data1-v3.xlsx

Download elife-79890-fig3-data1-v3.xlsx

The measurements were performed using white, red, and far-red flashes. As expected, in dark-adapted samples, with S₁ as the majority state (Table 2), the maximal O₂ evolution occurred on the 3rd flash with subsequent maxima at 4 flash intervals. These maxima reflect the occurrence of the S₃Y_Z^●/S₄ to S₀ transition in most centres as two water molecules are oxidized, resulting in the release of O₂. This oscillation pattern was the same in all samples and under all excitation conditions, except in Synechocystis membranes illuminated with far-red light, where the slow rise in O₂ evolution is due to the weak excitation of Chl-a-PSII by the short wavelength tail of the 730 nm flash.

The miss factor, indicating the fraction of PSII centres failing to progress through the S-states after a saturating flash excitation (Lavergne, 1991; Grabolle and Dau, 2007 and see Discussion ), was ≤20% in all the samples except in the Synechocystis sample illuminated with far-red flashes, where it was >80% (Figure 3D). For A. marina, the misses (13–17%) were very similar to those reported earlier (Shevela et al., 2006). The misses in FR C. thermalis and in Synechocystis when illuminated with the 613 nm LED were slightly higher (17%–20%). Nevertheless, these differences, attributed to the combination of the absence of exogenous electron acceptors, and the relatively long and possibly not fully saturating flashes (Figure 3—figure supplement 1), were not significant.

In order to confirm and expand the results obtained with polarography, we measured the S-state turnover as the flash-induced absorption changes at 291 nm (Figure 4), that reflect the redox state of the Mn ions in the oxygen evolving complex (Lavergne, 1991; Boussac et al., 2004). These measurements were done in the presence of the electron acceptor PPBQ and using single-turnover monochromatic saturating laser flashes. In the case of A. marina, the measurements could be done using membranes, but the membranes of WL and FR C. thermalis could not be used because of their high light-scattering properties in the UV part of the spectrum. In the case of the FR C. thermalis partially purified O₂ evolving Chl-f-PSII were made and used for the measurements, while difficulties were encountered in isolating O₂ evolving PSII from WL C. thermalis. Therefore, PSII cores from T. elongatus with the D1 isoform PsbA3 (Sugiura et al., 2008) were used as a Chl-a-PSII control. Among the three D1 present in T. elongatus, PsbA3 has the highest sequence identity with the D1 of Chl-f-PSII in FR C. thermalis (see Discussion).

Figure 4

Download asset Open asset

Figure 4—source data 1

Flash-dependent UV absorption.

https://cdn.elifesciences.org/articles/79890/elife-79890-fig4-data1-v3.xlsx

Download elife-79890-fig4-data1-v3.xlsx

The Chl-f-PSII was illuminated with flashes at wavelengths preferentially absorbed by Chl-a (680 nm) and by long-wavelength chlorophylls (720–750 nm) (Figure 4A). As expected, maximum absorption decrease (positive ΔI/I, as defined in Materials and methods, section UV transient absorption) occurred on S₂ (flash 1,5,9 etc.) and maximum absorption increase (negative ΔI/I) on S₀ (flash 3,7,11 etc.) (Lavergne, 1991). No differences could be observed in either the amplitude or the damping of the oscillations between the excitation wavelengths. When using sub-saturating flashes (~83% power), the damping of the oscillations was the same for all excitation wavelengths (Figure 4B), verifying that the illumination with 100% laser power was saturating at all the wavelengths. The equal amplitude of the oscillations obtained at all excitation wavelengths also indicates that the FR C. thermalis sample used does not contain any detectable Chl-a-PSII contamination. No differences in the oscillation patterns measured in FR C. thermalis Chl-f-PSII cores and in A. marina membranes, flashed at either 680 or 725 nm, were observed (Figure 4C). The PSII of T. elongatus showed a normal S-states progression when using 680 nm excitation, but no oscillation pattern when far-red flashes were used (Figure 4D). For all samples the calculated miss factor was ~10% (Appendix 3, discussion based on Styring and Rutherford, 1987; Velthuys and Visser, 1975; Vermaas et al., 1984; Sugiura et al., 2004).

In conclusion, the data reported here show that the overall efficiency of electron transfer from water to the PQ pool is comparable in all three types of PSII (independently of the Chl-a-PSII control used), as shown by the near-identical flash patterns of thermoluminescence (Appendix 2) and O₂ release (Figure 3), both measured without external electron acceptors. When the S-state turnover was measured by following the absorption of the Mn-cluster in the UV (Figure 4), the use of artificial electron acceptors and single-turnover saturating flashes allowed us to obtain better resolved flash patterns that were essentially indistinguishable in all three types of PSII and between excitation with visible or far-red light in the case of the Chl-d-PSII and Chl-f-PSII.

Charge recombination reactions were investigated by monitoring the thermoluminescence and luminescence emissions. The TL curves in Figure 5A and B show that both Chl-f-PSII and Chl-d-PSII are more luminescent than Chl-a-PSII, with Chl-f-PSII being the most luminescent. These differences, that are much larger than the variability between biological replicates (Figure 5—figure supplement 1C and D and Table 3), fit qualitatively with earlier reports (Nürnberg et al., 2018; Cser et al., 2008) (see Appendix 4 for more details). The high luminescence indicates that in the Chl-d-PSII and Chl-f-PSII there is an increase in radiative recombination, although the causes of this increase are likely to be different between the two photosystems, as detailed in the Discussion.

Figure 5 with 2 supplements see all

Download asset Open asset

Despite the large difference in TL intensity between the Chl-a-PSII and Chl-f-PSII, the peak temperatures corresponding to the S₂Q_B^- and S₂Q_A^- recombination were both similar in Chl-a-PSII and Chl-f-PSII. In Chl-d-PSII, the temperature of the S₂Q_B^- peak was only slightly lower, while the S₂Q_A^- peak was ~15 °C lower (Figure 5—figure supplement 1A and B and Table 3). Earlier TL reports comparing Chl-d-PSII in A marina cells with Chl-a-PSII in Synechocystis cells also showed that, while the peak position of S₂Q_B^- recombination was similar in the two samples, the S₂Q_A^- peak position was lower in A. marina (Cser et al., 2008), in agreement with the present results in membranes. The peak temperatures measured in cells were lower than those reported here, which can be explained by (i) the effect of the transmembrane electric field, as discussed for the fluorescence decay results, and (ii) by differences in the heating rates used (1 °C s^–1 here, 0.33 °C s^–1 in Cser et al., 2008). When performing the same measurements in Synechocystis membranes (Figure 5—figure supplement 2A), the S₂Q_B^- and S₂Q_A^- peak positions were comparable to those obtained in the two C. thermalis samples, confirming that the lower S₂Q_A^- peak temperature is a specific feature of Chl-d-PSII.

The S₂Q_A^- recombination in the presence of DCMU was also measured by luminescence decay kinetics at 10, 20, and 30°C, a range of temperatures that covers those of the S₂Q_A^- TL peaks of the three samples. Luminescence decay kinetics were recorded from 570ms for 300 seconds after the flash. In this time-range, the luminescence arises mainly from recombination via the back-reaction of S₂Q_A^- (Goltsev et al., 2009). The total S₂Q_A^- luminescence emission (Figure 5C) reflected the intensities of the TL peaks, as expected (Rutherford and Inoue, 1984), with the order of intensity as follows: Chl-f-PSII>Chl-d-PSII>Chl-a-PSII (although the variability between replicates made the difference between Chl-a-PSII and Chl-d-PSII less significant than that measured by TL). The total emissions did not vary significantly between 10°C and 30°C, although the decay kinetics were temperature-sensitive (Appendix 5—figure 1). The decay components identified by fitting the curves and their significance are discussed further in Appendix 5, based on Yerkes et al., 1983; Lavorel and Dennery, 1984; Tyystjarvi and Vass, 2007; Sugiura et al., 2014. The luminescence decay attributed to S₂Q_A^- recombination was bi-phasic (Appendix 5—table 1), with the kinetics of both phases being faster in Chl-d-PSII (~3 and~11 s) than in Chl-a-PSII (~4 and~25 s), but slower in Chl-f-PSII (~9 and~39 s). The average S₂Q_A^- luminescence decay lifetimes accelerated with increasing temperature in Chl-a-PSII and Chl-f-PSII but were always the fastest in Chl-d-PSII and the slowest in Chl-f-PSII (Figure 5D). The luminescence decay kinetics of the Chl-a-PSII in Synechocystis membranes were similar to those measured in WL C. thermalis (Figure 5—figure supplement 2B and C), suggesting, as seen with the TL data, that the differences in kinetics observed in the two types of far-red PSII are not due to differences between species.

In conclusion, both Chl-f-PSII and Chl-d-PSII show strongly enhanced luminescence, as previously reported (Nürnberg et al., 2018; Cser and Vass, 2007). However, the Chl-d-PSII differs from the Chl-a-PSII and Chl-f-PSII by having a lower S₂Q_A^- TL peak temperature and a faster S₂Q_A^- luminescence decay. This indicates that Chl-d-PSII has a smaller energy gap between Q_A^- and Phe compared to Chl-a-PSII and Chl-f-PSII, resulting in: (i) less heat required for the electron to be transferred energetically uphill from Q_A^- to Phe (manifest as lower TL peak temperature), and (ii) a bigger proportion of S₂Q_A^- recombination occurring via repopulation of P_D1⁺Phe^-, a route faster than direct P_D1⁺Q_A^- recombination (manifest as faster luminescence decay kinetics, see also Appendix 5). The lower TL temperature and faster luminescence decay for S₂Q_A^- recombination in Chl-d-PSII, but without a marked increase in its Q_A^- decay rate as monitored by fluorescence (Figure 2), could reflect differences in the competition between radiative and non-radiative recombination pathways in Chl-d-PSII compared to those in Chl-a-PSII and Chl-f-PSII. In contrast, in Chl-f-PSII the energy gap between Q_A^- and Phe does not appear to be greatly affected or could even be larger, as suggested by the slower S₂Q_A^- recombination measured by fluorescence (Figure 2) and luminescence (Figure 3) decay. The Q_B potentials appear to be largely unchanged, as manifested by the similar S₂Q_B^- stability in all three types of PSII, with the slightly lower S₂Q_B^- TL peak temperature in A. marina probably reflecting the decrease in the energy gap between Q_A^- and Phe.

The smaller energy gap between Q_A^- and Phe reported here in A marina is expected to result in enhanced singlet O₂ production and hence greater sensitivity to photodamage (Nürnberg et al., 2018; Cotton et al., 2015; Johnson et al., 1995; Vass and Cser, 2009). This was investigated by measuring the rates of ¹O₂ generation induced by saturating illumination in isolated membranes using histidine as a chemical trap (Figure 6A, representative traces in Figure 6—figure supplement 1A-C). ¹O₂ reacts with histidine to form the final oxygenated product, HisO₂, resulting in the consumption of O₂, as measured using the O₂ electrode. Without the histidine trap, most ¹O₂ is thought to be quenched by carotenoids (Telfer et al., 1994). When histidine was present in addition to DCMU, the Chl-d-PSII in A. marina membranes showed significant light-induced ¹O₂ formation. Under the same conditions, little ¹O₂ formation occurred in Chl-a-PSII or Chl-f-PSII in C. thermalis membranes. Similarly low levels of ¹O₂ were generated by Chl-a-PSII in Synechocystis membranes (Figure 6—figure supplement 1D). The His-dependent O₂ consumption in A. marina membranes showed the same light intensity dependence as O₂ evolution (Appendix 6—figure 1B), which suggests that ¹O₂ formation was related to Chl-d-PSII photochemistry. Sodium azide, a ¹O₂ quencher, suppressed the His-dependent oxygen consumption measured in A. marina in the presence of DCMU and when using the ¹O₂-generating dye Rose Bengal, confirming that it was due to the production of ¹O₂ (Figure 6—figure supplement 1E and F).

Figure 6 with 1 supplement see all

Download asset Open asset

Figure 6—source data 1

Singlet oxygen production membranes.

https://cdn.elifesciences.org/articles/79890/elife-79890-fig6-data1-v3.xlsx

Download elife-79890-fig6-data1-v3.xlsx

Figure 6—source data 2

Photoinhibition membranes.

https://cdn.elifesciences.org/articles/79890/elife-79890-fig6-data2-v3.xlsx

Download elife-79890-fig6-data2-v3.xlsx

The strikingly high amount of ¹O₂ generated by Chl-d-PSII prompted us to perform additional controls. (i) To test if the high ¹O₂ production was related to the intactness of the PSII donor side, Mn was removed from A. marina membranes by Tris-washing. This had little effect on the ¹O₂ formation with respect to the Mn-containing membranes (Appendix 6—figure 2), suggesting that the high ¹O₂ production in untreated A. marina membranes does not arise specifically from the fraction of centres lacking an intact Mn-cluster that are potentially responsible for the non-decaying fluorescence observed in Figure 2. (ii) The possibility that photosystem I (PSI) contributed to the light-induced O₂ consumption by reducing oxygen to O₂^•- in membranes was tested (Appendix 6—figure 3). In the presence of DCMU, PSI-driven O₂ reduction mediated by methyl viologen only took place when exogenous electron donors to PSI were provided. This indicates that there is no contribution from PSI-induced O₂ reduction in Figure 6A, where exogenous PSI donors are absent. (iii) The higher ¹O₂ production is also seen in A. marina cells compared to FR C. thermalis cells (Appendix 6—figure 4A), and thus is not an artefact associated with the isolation of membranes (e.g. damaged photosystems or free chlorophyll). WL C. thermalis cells also showed low levels of ¹O₂ production, similar to those measured in membranes (Appendix 6—figure 4B). The reliability of the His-trapping method to monitor ¹O₂ production in intact cyanobacterial cells has been previously demonstrated (Rehman et al., 2013).

Figure 6B shows the effect of 30 min of saturating illumination (red light) on the activity of the Chl-d-PSII, Chl-a-PSII and Chl-f-PSII. The results show that Chl-d-PSII is significantly more susceptible to light induced loss of activity compared to Chl-f-PSII, and to a lesser extent to Chl-a-PSII, and this can be correlated to the higher levels of ¹O₂ production in Chl-d-PSII.

The turnover of the water oxidation cycle is comparably efficient in all three types of PSII, as shown by their near-identical flash patterns in thermoluminescence (Appendix 2—figure 1), O₂ release (Figure 3), and UV spectroscopy (Figure 4). In PSII, a photochemical ‘miss factor’ can be calculated from the damping of the flash patterns of O₂ evolution. These misses, which are typically ~10% in Chl-a-PSII, are mainly ascribed to the µs to ms recombination of S₂TyrZ^•Q_A^- and S₃TyrZ^•Q_A^- states (Grabolle and Dau, 2007). Despite the diminished energy available, the miss factors in both types of far-red PSII were virtually unchanged compared to Chl-a-PSII, which also suggests that the misses have the same origin. If so, the energy gaps between TyrZ and P_D1, and thus their redox potentials, would be essentially unchanged. These conclusions agree with those in earlier work on Chl-d-PSII (Shevela et al., 2006) and on Chl-f-PSII (Nürnberg et al., 2018).

The similar flash-patterns also indicate that, after the primary charge separation, the electron transfer steps leading to water oxidation must have very similar efficiencies in all three types of PSII, that is, close to 90%, and that there are no major changes affecting the kinetics of forward electron transfer. In the case of Chl-f-PSII, this confirms earlier suggestions based on flash-dependent thermoluminescence measurements (Nürnberg et al., 2018). Indeed, electron transfer from Q_A^- to Q_B/Q_B^-, monitored by fluorescence, showed no significant differences in kinetics in the three types of PSII (Figure 2A).

The most striking difference between the three types of PSII is that the Chl-d-PSII of A. marina shows a decreased stability of S₂Q_A^-, indicated by the lower temperature of its TL peak and the correspondingly faster luminescent decay kinetics (Figure 5), and consequently a significant increase in ¹O₂ generation under high light (Figure 6A). This likely corresponds to the decrease in the energy gap between Phe and Q_A predicted to result from the ~100 meV lower energy available when using light at ~720 nm to do photochemistry (Nürnberg et al., 2018; Cotton et al., 2015). This is also supported by the estimates in the literature of the redox potential (E_m) values of Phe/Phe^- and Q_A/Q_A^- in Mn-containing Chl-d-PSII: compared to Chl-a-PSII, the estimated increase of ~125 mV in the E_m of Phe/Phe^- is accompanied by an estimated increase of only ~60 mV in the E_m of Q_A/Q_A^-, which implies that a normal energy gap between the excited state of the primary donor (Chl_D1^*) and the first and second radical pairs (Chl_D1⁺Phe^- and P_D1⁺Phe^-) is maintained, but the energy gap between P_D1⁺Phe- and P_D1⁺Q_A^- is significantly decreased (~325 meV vs ~385 meV) (Allakhverdiev et al., 2011). The changes in the D1 and D2 proteins of A. marina responsible for the changes in the E_m of Phe/Phe^- and Q_A/Q_A^- are currently unknown.

Our results indicate that in Chl-d-PSII, the decrease in the energy gap between Phe and Q_A favours charge recombination by the back-reaction route (via P_D1⁺Phe^-), forming the reaction centre chlorophyll triplet state (Rutherford et al., 1981), which acts as an efficient sensitizer for ¹O₂ formation (Johnson et al., 1995; Vass and Cser, 2009; Keren et al., 1995; Keren et al., 2000). Consequently, the Chl-d-PSII is more sensitive to high light (Figure 6B), reflecting the fact that this long-wavelength form of PSII has evolved in shaded epiphytic environments (Nürnberg et al., 2018; Miyashita et al., 1996; Cotton et al., 2015; Davis et al., 2016; Murakami et al., 2004; Miller et al., 2005; Kühl et al., 2005; Mohr et al., 2010). The increase in the proportion of recombination going via P_D1⁺Phe^- in Chl-d-PSII can also result in a higher repopulation of the excited state of the primary donor (Chl_D1^*), with a consequent increase in radiative decay (high luminescence).

In contrast to the Chl-d-PSII, the Chl-f-PSII shows no increased production of ¹O₂ and no increased sensitivity to high light compared to Chl-a-PSII, in the conditions tested here (Figure 6). The back-reactions appear to be little different from the Chl-a-PSII except for the more stable (more slowly recombining) S₂Q_A^-, as seen by fluorescence (Figure 2) and luminescence (Figure 5) decay. These properties may seem unexpected because this type of PSII has the same energy available for photochemistry as the Chl-d-PSII. In the Chl-d-PSII the lower energy of Chl_D1^* is matched by an increase in the E_m of Phe/Phe^-. In the Chl-f-PSII of C. thermalis and of the other Chl-f containing species, the E_m of Phe/Phe^- is also expected to be increased by the presence, in the far-red D1 isoform, of the strong H-bond from Glu130 (Figure 7), which is characteristic of high-light D1 variants in cyanobacteria (Sugiura et al., 2010). In Chl-a-PSII, this change has been reported to induce an increase in the E_m of Phe/Phe^- between ~15 and~30 mV (Sugiura et al., 2010; Merry et al., 1998): an increase of this size would only partially compensate for the ~100 meV decrease in the energy of Chl_D1^* in Chl-f-PSII, and this would result in a smaller energy gap between Chl_D1^* and the first and second radical pairs Chl_D1⁺Phe^- and P_D1⁺Phe^-. This would favour the repopulation of Chl_D1^* by back-reaction from P_D1⁺Phe^- (even if the repopulation of P_D1⁺Phe^- from the P_D1⁺Q_A^- state did not increase), resulting in the higher luminescence of Chl-f-PSII, as proposed earlier (Nürnberg et al., 2018). Increased decay of the P_D1⁺Q_A^- radical pair via the radiative route could in principle decrease the decay via the triplet route, but the overall small yield of luminescence means that this could be a minor effect.

Figure 7

Download asset Open asset

Figure 7—source data 1

D1 sequences used for multi-alignments.

https://cdn.elifesciences.org/articles/79890/elife-79890-fig7-data1-v3.txt

Download elife-79890-fig7-data1-v3.txt

Additionally, the longer lifetime of S₂Q_A^- recombination in Chl-f-PSII indicates that the E_m of Q_A/Q_A^- has increased to compensate the up-shift in the E_m of Phe/Phe^- and to maintain an energy gap between Phe and Q_A large enough to prevent an increase in P_D1⁺Phe^- repopulation and thus in reaction centre chlorophyll triplet formation. This situation occurs in the PsbA3-D1 high light variant of T. elongatus, although the protein changes responsible for the increase in the E_m of Q_A/Q_A^- are not known (Sugiura et al., 2010). A slower S₂Q_A^- recombination could also arise from an increase in the redox potential of P_D1/P_D1⁺ (Diner et al., 2001), but this would likely compromise forward electron transfer in Chl-f-PSII by decreasing the driving force for stabilization of Chl_D1⁺Phe^- by the formation of P_D1⁺Phe^-, if the redox potential of Chl_D1/Chl_D1⁺ was not increased accordingly. If this problem were avoided by an equivalent increase in the redox potential of Chl_D1/Chl_D1⁺, this would likely compromise charge separation, as a matching up-shift the Chl_D1⁺/Chl_D1^* couple would occur, thus decreasing the reductive power of Chl_D1^*. As a 720 nm pigment, the reducing power of Chl_D1^* seems likely to be already compromised in Chl-f-PSII. The results here (Figure 2A) show forward electron transfer and charge separation appear to be comparable to Chl-a PSII and so it is unlikely that redox tuning of P_D1 and Chl_D1 is responsible for the stabilisation of the S₂Q_A^- recombination .

In addition to changes in the redox potentials of Phe and Q_A, the size and pigment composition of the antennas of Chl-d-PSII and Chl-f-PSII could also contribute to the functional differences reported in the present work. These differences are summarized in Figure 8.

Figure 8

Download asset Open asset

In PSII, two factors will determine the yield of charge separation: (i) the relative population of the excited state of the primary donor, Chl_D1^*, which depends on the dynamics of excitation energy transfer between pigments, and (ii) the rate of population of the second radical pair, P_D1⁺Phe^-, that is more stable (less reversible) than the first radical pair, Chl_D1⁺Phe^-. This rate is determined by the rates of the primary charge separation (forming Chl_D1⁺Phe^-) and of its stabilization by secondary electron transfer (forming P_D1⁺Phe^-), and hence by the energetics of these electron transfer steps.

In the Chl-a-PSII core, the 37 chlorins absorb light between ~660 and~690 nm and are therefore almost isoenergetic to the Chl_D1 primary donor absorbing at 680 nm. Given the small energy differences, there is little driving force for downhill ‘funnelling’ of excitation energy to Chl_D1, making it a ‘shallow trap’. Different models have been proposed to explain the shallowness of the photochemical trap in Chl-a-PSII.

In the trap-limited model, the transfer of excitation between pigments is significantly faster than the electron transfer reactions leading to P_D1⁺Phe^- formation, and a near-complete equilibration of the excitation energy is established over all pigments, including Chl_D1, with a distribution that is determined by their individual site energies (van Mieghem et al., 1995; Schatz et al., 1987; Dau and Sauer, 1996). This leads to a low population of Chl_D1^* (Table 4), that is diminished as a function of the number of quasi-isoenergetic pigments with which it shares the excitation energy.

In the transfer-to-trap limited model, the small driving force for downhill ‘funnelling’ of excitation energy to Chl_D1 causes kinetic bottlenecks for excitation energy equilibration between the core antenna complexes CP43 and CP47 and for excitation energy transfer from these antennas to the reaction centre (Jennings et al., 2000; Pawlowicz et al., 2007; Raszewski and Renger, 2008; Kaucikas et al., 2016). In this scenario, there is not a full equilibration of the excitation energy over all pigments, but the relatively slow and reversible energy transfer from the core antennas to the reaction centre still leads to a relatively low population of Chl_D1^*.

Irrespectively of the differences in the details of the kinetic limitation to photochemical trapping between the two models, the common requirement for establishing a high quantum yield of charge separation is a sufficiently large overall energy gap (~160 meV, ) between Chl_D1* and P_D1⁺Phe^-, that comprises the primary charge separation (Chl_D1^* ↔ Chl_D1⁺Phe^-) and secondary electron transfer (Chl_D1⁺Phe^- ↔ P_D1⁺Phe^-), as shown in Figure 8. An energy gap of this magnitude is required to avoid rapid recombination to the excited state Chl_D1^*, thereby limiting the probability of its dissipation via non-photochemical relaxation to the ground state in the antenna (Raszewski and Renger, 2008; Schatz et al., 1988).

For Chl-d-PSII the antenna system is comparable to that in Chl-a-PSII: all 34 Chl-d molecules, including the primary donor Chl_D1 at ~720 nm, are close in wavelength and thus both systems are expected to have comparable Chl_D1^* population (Table 4), irrespective of the rate-limitation model assumed. Chl-a-PSII and Chl-d-PSII should therefore have the same energetic requirements to ensure a sufficiently high yield of charge separation. Given that the energy of Chl_D1^* is ~100 meV lower in Chl-d-PSII than in Chl-a-PSII, the energy level of the second and more stable radical pair, P_D1⁺Phe^-, needs to be decreased by ~100 meV in Chl-d-PSII relative to Chl-a-PSII. This corresponds to the published E_m of Phe/Phe^- (Allakhverdiev et al., 2011) and to the kinetic data (Figures 5 and 6), as detailed in the previous section.

In A. marina membranes, additional Chl-d containing antenna proteins, which form supercomplexes with PSII cores, have been reported to increase the Chl-d-PSII antenna size by almost 200% (Chen et al., 2005). This will likely result in an increased sharing of the excited state, leading to a diminished population of Chl_D1^*, and thus a bigger requirement for an energy drop between Chl_D1^* and P_D1⁺Phe^- to ensure efficient charge separation. At the same time, the larger near-isoenergetic antenna could also contribute to its higher luminescence, by increasing the probability of Chl_D1^* decay via radiative emission with respect to photochemical re-trapping (Rappaport and Lavergne, 2009). This is similar to what happens in plant PSII, where the yield of photochemical trapping of excitation energy is decreased by 10–15% by the association of the Light Harvesting Complex antennas (Engelmann et al., 2005).

The pigment layout of Chl-f-PSII is very different from that of Chl-a-PSII and Chl-d-PSII. The 30 Chl-a molecules are energetically separated from Chl_D1, absorbing at 720 nm, by >30 nm (>3 k_BT). This means the excitation energy resides predominantly on Chl_D1^* and on the other 4 far-red pigments. If the equilibration of the excitation energy between the 5 far-red pigments were significantly faster than charge separation, this pigment arrangement would result in a higher probability of populating Chl_D1^* in Chl-f-PSII than in Chl-a-PSII and Chl-d-PSII (Table 4). The higher Chl_D1^* population in Chl-f-PSII could ensure that sufficient yield of charge separation is achieved even when the E_m of Phe/Phe^- is increased by much less that the 100 meV needed to compensate for the nominally lower energy in Chl_D1^*.

However, thermal equilibration of the excitation energy over the entire antenna in Chl-f-PSII might not occur due to 3 of the 4 long-wavelength antenna chlorophylls absorbing at longer wavelength than Chl_D1. This type of antenna energetics could result in rapid excited state equilibration in each of the three main pigment-protein complexes (CP43, CP47 and reaction centre), due to rapid energy transfer from Chl-a to Chl-f/d (with visible light excitation) followed by slower transfer from the two postulated far-red antenna pools to Chl_D1, leading to a transfer-to-trap limited bottleneck. As a result, the kinetics of excitation energy transfer from the red and far-red antenna to the reaction centre could be more complex than in Chl-a-PSII and Chl-d-PSII, explaining the spread in charge separation kinetics that has been suggested based on ultrafast absorption data (Zamzam et al., 2020) and the slower excitation energy trapping kinetics measured by time-resolved fluorescence (Mascoli et al., 2020).

The driving force for charge separation is decreased in Chl-f-PSII also by the smaller energy gap between Chl_D1^* and P_D1⁺Phe^-, estimated to be ~80 meV in Chl-f-PSII compared to ~160 meV in Chl-a-PSII and Chl-d-PSII. This decrease in the energy gap between Chl_D1^* and P_D1⁺Phe^- is necessary in Chl-f-PSII to avoid the increased photosensitivity seen in Chl-d-PSII by maintaining a large energy gap between P_D1⁺Phe^- and P_D1⁺Q_A^- (~385 meV) (Figure 8). Nonetheless, the slower excitation energy transfer and the smaller energy gap between Chl_D1^* and P_D1⁺Phe^- could be partially compensated by the decreased dilution of the excitation energy on Chl_D1^* arising from the small number of long-wavelength antenna pigments, resulting in only a small loss of trapping efficiency (Mascoli et al., 2020) and a near-negligible effect on enzyme turnover efficiency (Figures 2—5).

This energetic balancing trick in Chl-f-PSII, which allows both reasonably high enzyme efficiency and high resilience to photodamage (by limiting recombination via the repopulation of P_D1⁺Phe^-) despite working with 100 meV less energy, comes with a very significant disadvantage: its absorption cross-section at long wavelength is ~7 times smaller than that of the Chl-a-PSII core antenna in visible light. In the case of Chl-f-PSII, evolution therefore seems to have prioritized the minimization of harmful charge recombination, by maintaining a big energy gap between Phe and Q_A, over light collection and photochemical quantum efficiency. This makes sense as this system has evolved as a facultative survival mechanism, that is not advantageous when visible light is available. It must be noted that in vivo the far-red antenna cross-section of Chl-f-PSII is increased by the presence of red-shifted phycobilisomes, that replace the visible light-absorbing phycobilisomes when the cells are adapted to far-red light (Gan et al., 2014).

In contrast, Chl-d-PSII seems to have maximized light collection at long wavelengths (with its full-size far-red antenna) and maximized the yield of charge separation (by maintaining the full Chl_D1^* to P_D1⁺Phe^- driving force). However, the energy shortfall at long wavelength is lost from the ‘energy headroom’ (mainly from the transmembrane energy gap between Phe and Q_A) that is proposed to minimize harmful charge recombination by buffering the effects of pulses of the trans-membrane electric field associated with fluctuations in light intensity (Davis et al., 2016; Davis et al., 2017). This seems to correspond well to the shaded and stable epiphytic niche that A. marina occupies (Nürnberg et al., 2018; Miyashita et al., 1996; Cotton et al., 2015; Davis et al., 2016; Murakami et al., 2004; Miller et al., 2005; Kühl et al., 2005; Mohr et al., 2010).

Chl-d-PSII and Chl-f-PSII have evolved different strategies to do oxygenic photosynthesis in far-red light and have been impacted differently by the decrease in energy available. Understanding how the redox tuning of the electron transfer cofactors and the layout of the far-red pigments determine the trade-off between efficiency and resilience in PSII is a necessary step to inform strategies aimed at using far-red photosynthesis for agricultural and biotechnological applications.

The present findings suggest the exchange of the full Chl-a manifold to long-wavelength chlorophylls, as seen in Chl-d-PSII (A. marina), should allow efficient oxygenic photosynthesis, but only under constant shading and stable light conditions: for example, for cultivation under LED light (vertical farming, etc). The more robust, facultative Chl-f PSII has an intrinsically low absorption cross-section in the far red; however, this could be beneficial in a shaded canopy, especially in combination with a suitably designed far-red external antenna.

Acaryochloris marina was grown in a modified liquid K-ESM medium containing 14 µM iron (Bailleul et al., 2008), at 30 °C under constant illumination with far-red light (750 nm, Epitex; L750-01AU) at ~30 μmol photons m^–2 s^–1. Chroococcidiopsis thermalis PCC7203 was grown in liquid BG11 medium (Stanier et al., 1979) at 30 °C, under two illumination conditions: white light at ~30 μmol photons m^–2 s^–1 (for WL C. thermalis samples) and far-red light (750 nm, Epitex; L750-01AU) at ~30 μmol photons m^–2 s^–1 (for FR C. thermalis samples). Synechocystis sp. PCC6803 was grown in liquid BG11 medium at 30 °C under constant illumination with white light at ~30 μmol photons m^–2 s^–1. The Thermosynechococcus elongatus ΔpsbA1, ΔpsbA2 deletion mutant (WT*3, Sugiura et al., 2008) was grown in liquid DNT medium at 45 °C.

Membranes were prepared as described in Appendix 7, frozen in liquid nitrogen and stored at –80 °C until use. Partially purified C. thermalis PSII cores retaining oxygen evolution activity were isolated by anion exchange chromatography using a 40 ml DEAE column. The column was equilibrated with 20 mM MES (2-(N-morpholino)ethanesulfonic acid)-NaOH pH 6.5, 5 mM CaCl₂, 5 mM MgCl₂ and 0.03% (w/v) β-DM (n-Dodecyl-β-D-maltoside) and elution was done using a linear gradient of MgSO₄ from 0 to 200 mM in 100 min (in the same buffer conditions as those used to equilibrate the column), with a flow rate of 4 ml min^–1. Fractions enriched in PSII were pooled, frozen in liquid nitrogen and stored at –80 °C. PSII-PsbA3 cores from T. elongatus WT*3 were purified as previously described (Sugiura et al., 2014).

Flash-induced chlorophyll fluorescence and its subsequent decay were measured with a fast double modulation fluorimeter (FL 3000, PSI, Czech Republic). Excitation was provided by a saturating 70 μs flash at 630 nm and the decay in Q_A^- concentration was probed in the 100 μs to 100 s time region using non-actinic measuring pulses following a logarithmic profile as described in Vass et al., 1999. The first measuring point was discarded during the data analysis because it contains a light artefact arising from the tail of the saturating flash used for excitation. Details on the analysis of the fluorescence curves (based on Crofts and Wraight, 1983; Vass et al., 1999; Cser et al., 2008) are provided in Appendix 7. Membrane samples were adjusted to a total chlorophyll concentration of 5 μg Chl ml^–1 in resuspension buffer, pre-illuminated with room light (provided by a white fluorescent lamp, ~80 μmol photons m^–2 s^–1) for 10 s and then kept in the dark on ice until used for measurements. Measurements were performed at 20 °C. Where indicated, 20 µM DCMU (3-(3,4-dichlorophenyl)–1,1-dimethylurea) was used.

Thermoluminescence curves and luminescence decay kinetics were measured with a laboratory-built apparatus, described in De Causmaecker, 2018. Membrane samples were diluted in resuspension buffer to a final concentration of 5 µg Chl ml^–1 in the case of A. marina and FR C. thermalis and of 10 µg ml^–1 in the case of WL C. thermalis and Synechocystis. The samples were pre-illuminated with room light (provided by a white fluorescent lamp,~80 μmol photons m^–2 s^–1) for 10 s and then kept in the dark on ice for at least one hour before the measurements. When used, 20 µM DCMU was added to the samples before the pre-illumination step. Excitation was provided by single turnover saturating laser flashes (Continuum Minilite II, frequency doubled to 532 nm, 5 ns FWHM). Details on the measurement conditions and on the analysis of the luminescence decay kinetics are provided in Appendix 7.

Oxygen evolution and consumption rates were measured with a Clark-type electrode (Oxygraph, Hansatech) at 25 °C. Membrane samples were adjusted to a total chlorophyll concentration of 5 μg Chl ml^–1. Illumination was provided by a white xenon lamp filtered with a heat filter plus red filter, emitting 600–700 nm light at 7100 µmol photons m^–2 s^–1 (Quantitherm light meter, Hansatech). When required, the light intensity was reduced by using neutral density filters (Thorlabs). For PSII maximal oxygen evolution rates, 1 mM DCBQ (2,5-Dichloro-1,4-benzoquinone) and 2 mM potassium ferricyanide were used as an electron acceptor system. Photoinhibitory illumination was performed at room temperature for 30 min with a laboratory-built red LED (660 nm, 2600 µmol photons m-² s^–1). For histidine-mediated chemical trapping of singlet oxygen, 20 µM DCMU, 5 mM L-Histidine and, where specified, 10 mM sodium azide (NaN₃) were used. PSI activity was measured as the rate of oxygen consumption in presence of 20 µM DCMU and 100 µM methyl viologen using 5 mM sodium ascorbate and 50 µM TMPD (N,N,N′,N′-tetramethyl-p-phenylenediamine) as electron donors. The dye Rose Bengal was used at a final concentration of 0.1 µM. After all necessary additions, samples were left to equilibrate with air in the measuring chamber for 1 minute, under stirring, to start with similar dissolved O₂ concentrations in all measurements. For each measurement, a dark baseline was recorded for 1 min before starting the illumination.

Time-resolved oxygen polarography was performed using a custom-made centrifugable static ring-disk electrode assembly of a bare platinum working electrode and silver-ring counter electrode, as previously described (Dilbeck et al., 2012). For each measurement, membranes equivalent to 10 µg of total chlorophyll were used. Three different light sources were used to induce the S-state transitions: a red LED (613 nm), a far-red LED (730 nm) and a Xenon flashlamp. Details on the experimental setup and on the lights used are provided in Appendix 7. Measurements were performed at 20 °C. For each measurement, a train of 40 flashes fired at 900ms time interval was given and the flash-induced oxygen-evolution patterns were taken from the maximal O₂ signal of each flash and fitted with an extended Kok model with flash-independent miss factor, as described in Nöring et al., 2008.

UV pump-probe absorption measurements were performed using a lab-built Optical Parametric Oscillator (OPO)-based spectrophotometer (Joliot et al., 1998) with a time resolution of 10 ns and a spectral resolution of 2 nm (see Appendix 7 for details on the setup). ΔI/I stands for differential absorption, a method that measures the changes in absorption depending on whether or not a sample is subjected to actinic light. Samples were diluted in resuspension buffer to a final concentration of 25 µg Chl ml^–1 for isolated C. thermalis and T. elongatus PSII cores and 40 µg Chl ml^–1 for A. marina membranes. Samples were pre-illuminated with room light (provided by a white fluorescent lamp,~80 μmol photons m^–2 s^–1) for 10 s and then kept in the dark on ice for at least one hour before the measurements. 100 µM PPBQ (Phenyl-p-benzoquinone) was added just before the measurements. The sample was refreshed between each train of flashes. For each measurement, a train of 20 flashes (6 ns FWHM) fired at 300ms time interval was given, and absorption changes measured at 100ms after each flash. The data were fitted according to Lavergne, 1991; Lavorel, 1976.

The fluorescence decay kinetics measured here in Synechocystis membranes (Appendix 1—figure 1), as well as those measured in A. marina and C. thermalis membranes (Figure 2), are slower than those measured in Synechocystis intact cells in a previous work (Cser and Vass, 2007). Additionally, a study of fluorescence decay times was previously reported comparing Q_A^- lifetimes in A. marina and Synechocystis but in cells rather than membranes. In A. marina cells, the forward (Q_A^- to Q_B) electron transfer rate was slower than in Synechocystis cells, while the S₂Q_A^- recombination rate A. marina cells was faster than in Synechocystis cells (Cser et al., 2008). In both organisms, the fluorescence decay kinetics were faster than the values measured here in membranes. The faster rates in cells compared to isolated membranes are intrinsic to the type of sample used. The transmembrane electric field, which is present in cells but not in isolated membranes, is known to accelerate Q_A^- decay in presence of DCMU (Joliot and Joliot, 1980). Additionally, the faster rates for Q_A^- to Q_B electron transfer in cells may be attributed to the Q_B site in living cells functioning optimally at higher pH rather than at the pH 6.5 used here to maintain PSII donor-side function.

Appendix 1—figure 1

Download asset Open asset

Appendix 1—figure 1—source data 1

Fluorescence decay kinetics Synechocystis.

https://cdn.elifesciences.org/articles/79890/elife-79890-app1-fig1-data1-v3.xlsx

Download elife-79890-app1-fig1-data1-v3.xlsx

Appendix 2—figure 1 shows the TL emission after a series of saturating flashes in A. marina (panels A, D and G), WL C. thermalis (panels B, E and H) and FR C. thermalis (panels C, F and I) membranes. Although no major differences in the flash patterns could be observed between the three samples, the flash dependence of the TL peak intensities (panels A, B and C) and their peak temperatures (panels D, E and F) showed variability between biological replicates. Representative TL glow curves obtained in one biological replicate for each sample after 1–6 flashes are shown in panels G, H and I. The differences in the flash patterns between replicates are easily explained by some variability in both the S₀/S₁ and Q_B/Q_B^- ratios present in the dark before the first flash, although the flash patterns suggest the presence of ~75% S₁/~25% S₀ and ~50% Q_B/~50% Q_B^-, based on Rutherford et al., 1981.

For WL C. thermalis, the smaller TL amplitude makes the peak temperature more difficult to estimate very precisely. For FR C. thermalis, a progressive broadening of the TL peak with increasing flash number made quantification less reliable, and for A. marina an increase in the baseline at high temperatures (also occurring to a smaller extent but still visible in WL C. thermalis) added to the difficulties in estimating the area of the TL peaks. For these reasons, the TL data are not precise enough to quantify potential differences in the S-state turnover efficiency in the different types of PSII, although they show that any such differences, if present, must be small (from the data in Appendix 2—figure 1A, B and C).

Appendix 2—figure 1

Download asset Open asset

Appendix 2—figure 1—source data 1

Flash dependent thermoluminescence.

https://cdn.elifesciences.org/articles/79890/elife-79890-app2-fig1-data1-v3.xlsx

Download elife-79890-app2-fig1-data1-v3.xlsx

Appendix 3—figure 1 shows the fit of the flash-induced absorption changes at 291 nm that reflect the progression through the S-states of the Mn-cluster (Lavorel, 1976; Boussac et al., 2004). The data are those reported in Figure 4: absorption changes measured in T. elongatus PsbA3-PSII cores with excitation at 680 nm, in A. marina membranes with excitation at 680 nm and in partially purified Chl-f-PSII cores from FR C. thermalis with excitation at 680 and 750 nm. The measurements were performed in presence of PPBQ, with intervals of 300ms between the flashes.

Appendix 3—figure 1

Download asset Open asset

The fit was done by taking the absorption changes corresponding to the S₀→S₁, S₁→S₂ and S₂→S₃ transitions determined in T. elongatus with the procedure established by Lavergne, 1991, and multiplying them by a factor γ which corresponds to the ratio in active PSII per chlorophyll of the given sample with respect to the T. elongatus sample. It is of note that the factor γ indicates the fraction of active PSII centres over the total PSII present only when comparing isolated cores, while in the data reported here, in which partially purified O₂-evolving Chl-f-PSII cores and A. marina membranes were used, it merely reflects the amounts of active PSII present in those samples for a given chlorophyll concentration. Since the measurements were done by using single-turnover excitation flashes (6 ns FWHM), the double-hit parameter β was considered to be zero. Using the formula developed by Lavorel, 1976, the miss parameter α and the proportion of the centres in S₁ state in the dark-adapted samples could be calculated. In these fits, the absorption changes on the first flash of the sequence was not taken into account because they may contain a non-oscillating component (Lavergne, 1991). This non-oscillating component is bigger in the A. marina and FR C. thermalis samples than in the T. elongatus sample, likely because of the lower content of active PSII per chlorophyll. The misses were comparable in all samples (~10%), and in the Chl-f-PSII cores from FR C. thermalis they did not significantly increase when using 750 nm excitation flashes.

The fits in Appendix 3—figure 1 indicate that the proportion of centres in S₁ in the dark-adapted samples was 75% in the A. marina and FR C. thermalis samples but 100% in the T. elongatus PsbA3-PSII cores. All samples were pre-illuminated in ambient light for ~10 s and then dark-adapted for >1 hr before the measurements and were therefore expected to be in 100% S₁ at the start of the flash sequence, with ~75% of the centres having TyrD^• and ~25% having TyrD (Styring and Rutherford, 1987). It has been shown that TyrD can reduce the S₂ and S₃ oxidation states of the Mn-cluster (Velthuys and Visser, 1975): in samples having starting populations of 75% TyrD^•S₁ and 25% TyrDS₁, some of the S₂ and S₃ states generated during the flash sequence will be re-reduced to S₁ and S₂, respectively, in centres where TyrD is present. This process will result in the apparent presence of 25% S₀ in the dark-adapted sample. This effect has been shown to depend on the spacing between excitation flashes (Vermaas et al., 1984): if the time between the flashes is not long enough to allow for TyrD donation, the flash pattern will reflect the initial presence of 100% S₁ (34). At room temperature, electron donation from TyrD is slower in T. elongatus PSII than in plant PSII (Sugiura et al., 2004): this could reflect the fact that T. elongatus is a thermophile and mesophilic cyanobacterial species such as A. marina and C. thermalis could be expected to have TyrD oxidation kinetics more similar to plants, thus explaining the difference in S₁ populations in our fits.

Figure 5—figure supplement 1 shows the plots of the peak amplitudes and temperature of the thermoluminescence arising from S₂Q_B^- and S₂Q_A^- in the three types of PSII. As mentioned in the main text, although our data fit qualitatively with earlier reports (Nürnberg et al., 2018; Cser et al., 2008), there is a degree of variability in both amplitude and temperature between biological replicates. Consequently, the average values reported in Table 3 present relatively high standard deviations. The variability in TL intensity between different membrane samples could depend on differences in the Q_B/Q_B^- ratios and distribution of S states present in the dark before applying the single-turnover flash (Rutherford et al., 1982).

These variabilities between biological replicates could also partially explain slight discrepancies between the data reported here and those in Nürnberg et al., 2018 regarding the ratio of luminescence intensity between the Chl-f-PSII and the Chl-a-PSII. In Nürnberg et al., 2018 the luminescence from both S₂Q_B^- and S₂Q_A^- were reported to be >25 times higher in FR C. thermalis membranes than in WL C. thermalis membranes, while the data reported here indicate that the luminescence of FR C. thermalis is between 5 and 16 times higher than in WL C. thermalis in the case of the S₂Q_B^- recombination, and between 3 and 15 times higher in the case of the S₂Q_A^- recombination. In the present work all measurements were performed at constant chlorophyll concentrations (5 µg Chl ml^–1 in the case of A. marina and FR C. thermalis, 10 µg ml^–1 in the case of WL C. thermalis), while in Nürnberg et al. the FR C. thermalis membranes were diluted to achieve a signal intensity comparable to that obtained in WL C. thermalis membranes. Although the dilution factor was included in the normalization on the O₂ evolution activities, these differences in the protocols used could contribute to the quantitative discrepancies, together with the biological variability, as at higher chlorophyll concentrations sample self-absorption can occur, thus skewing the measured TL intensity.

The S₂Q_A^- luminescence decay curves measured in A. marina, WL C. thermalis and FR C. thermalis at 10, 20, and 30°C (Appendix 5—figure 1) could be fitted with three exponential components (Appendix 5—table 1) and the differences in the kinetics between samples and between temperatures could be ascribed to differences in the amplitude and lifetimes of these components.

The luminescence decays at each temperature were similar in shape in Chl-a-PSII and Chl-f-PSII, while they were markedly different in Chl-d-PSII. Chl-a-PSII and Chl-f-PSII had a fast decay phase (T₁ ~0.5 and~1 s, respectively) absent in Chl-d-PSII. This phase, that has a bigger amplitude in Chl-a-PSII (~60%) than in Chl-f-PSII (~30%), is too fast to correspond to the S₂Q_A^- recombination and appears to match the rates of TyrZ^•(H⁺)Q_A^- recombination occurring either in centres lacking an intact Mn-cluster (Yerkes et al., 1983) or in intact centres before charge separation is fully stabilised, as proposed in Debus et al., 2000. The contribution of this fast component to the total luminescence emission was no more than 10% in the case of WL C. thermalis and 5% for FR C. thermalis. This decay phase was not detectable in the case of A. marina, suggesting that TyrZ^•(H⁺)Q_A^- recombination might be too fast in Chl-d-PSII to appear in our measurements. In A. marina an additional slower phase (~40 s) was present at 10 °C, but the very low amplitude made its contribution to the overall decay negligible.

The luminescence decay that we ascribe to the S₂Q_A^- back-reaction in the seconds to tens of seconds timescale, is comprised of two decay components, designated the middle and slow phases in Appendix 5—table 1. Both phases were faster in Chl-d-PSII (~3 and~11 s) than in Chl-a-PSII (~4 and~25 s), but slower in Chl-f-PSII (~9 and~39 s). In A. marina the lifetimes of these two decay components did not show a significant temperature dependence, resulting in only a minor acceleration of the overall luminescence decay of the Chl-d-PSII between 10°C and 30°C (Figure 5D). Indeed, the relative contribution of the two decay phases to the total luminescence changed little in function of temperature in this sample (Appendix 5—figure 1), with the changes being only at the level of the amplitude of the decay phases. The middle phase lifetimes did not show a significant temperature dependence in WL and FR C. thermalis either, but they were slower than in A. marina, especially in FR C. thermalis. The slow phase was also slower in the two C. thermalis samples and, additionally, its decay accelerated with increasing temperature, while its amplitude decreased. This resulted in its contribution to the overall luminescence decreasing between 10°C and 30°C (Appendix 5—figure 1) and the overall decay accelerating significantly (Figure 5D), especially in the FR C. thermalis.

It can be argued that the differences in kinetics between samples and their changes in function of temperature could represent changes in the relative contribution of different recombination pathways to the decay of S₂Q_A^-. It is not clear, though, whether each of the two decay components we identified represents a distinct recombination route or whether they derive from the combination of more complex kinetics. For instance, it has been suggested that the so-called ‘deactivation’ luminescence should follow a hyperbolic decay, rather than an exponential decay, due to the progressive decrease in the concentration of S₂Q_A^- resulting in a progressive slowing down of the rates of the various recombination routes (Lavorel and Dennery, 1984; Tyystjarvi and Vass, 2007). The data presented here could be satisfactorily fitted with exponentials but, given the considerations above and the uncertainty about how the evolution of luminescence reflects the actual concentrations of the charge separated states from which it originates, no assignment of the decay phase to specific recombination routes could be made.

Altogether, the data show that the luminescence kinetics in Chl-d-PSII are significantly different from those in Chl-a-PSII and Chl-f-PSII, pointing to a faster decay of the S₂Q_A^- charge separated state.

According to electron tunnelling calculations, the rate of P_D1⁺Q_A^- direct recombination to ground (10²–10³ s^–1) is much slower than P_D1⁺Phe^- recombination to ground (10⁶–10⁷ s^–1), although the limiting rate for S₂Q_A^- recombination via the repopulation of Phe^- is thought to be the migration of the electron hole from the Mn-cluster to TyrZ (~10³ s^–1) (Sugiura et al., 2014; Moser et al., 2005). Although the temperature dependence of the recombination routes is complex, an increase in temperature would have no effect on the rate of P_D1⁺Q_A^- direct recombination to ground, but would increase the rate of the backwards electron transfer from Q_A^- to Phe, as this is thermally activated, following the relationship

where k_rev and k_for are the rate constants of the backward and forward electron transfer, respectively, ΔG is the energy gap between the two cofactors, k_B is the Boltzmann constant and T is the temperature. Note that the rates of back-transfer of the positive charge from the Mn-cluster to P_D1 are also thermally activated and thus will accelerate with temperature and affect the rates of recombination from Phe^-. In this case, though, the smaller ΔGs involved should result in a less pronounced temperature dependence compared to the back electron transfer from Q_A^- to Phe (according to the equation above).

The acceleration of the luminescence decay kinetics with increasing temperature, observed for Chl-a-PSII and Chl-f-PSII, could reflect an increase in the contribution of S₂Q_A^- recombination route via repopulation of Phe^- in competition with the direct, non-radiative P_D1⁺Q_A^- recombination route. In Chl-d-PSII, the lower temperature of the S₂Q_A^- recombination thermoluminescence peak (Figure 5B and Figure 5—figure supplement 1) suggests a smaller ΔG between Q_A and Phe. This would result in a faster electron transfer from Q_A^- back to Phe, with this route already dominating the competition with the direct P_D1⁺Q_A^- recombination to ground, with a consequently high luminescence yield and a small temperature sensitivity of the decay rates.

Appendix 5—figure 1

Download asset Open asset

Appendix 5—figure 1—source data 1

Luminescence decay kinetics.

https://cdn.elifesciences.org/articles/79890/elife-79890-app5-fig1-data1-v3.xlsx

Download elife-79890-app5-fig1-data1-v3.xlsx

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

This work was supported by BBSRC grants BB/R001383/1, BB/V002015/1 and BB/R00921X. JS acknowledges funding from the Labex Dynamo (ANR-11-LABX-0011–01). AB was in part supported by the French Infrastructure for Integrated Structural Biology (FRISBI) ANR-10-INBS-05. SS acknowledges support from Fondazione Cariplo, project “Cyanobacterial Platform Optimised for Bioproductions” (ref. 2016–0667).

© 2022, Viola et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.