Algae, plants and cyanobacteria perform a process called photosynthesis, in which carbon dioxide and water are converted into oxygen and energy-rich carbon compounds. The first step of this process involves an enzyme called photosystem II, which uses light energy to extract electrons from water to help capture the carbon dioxide.

If the photosystem absorbs too much light, compounds known as reactive oxygen species are produced in quantities that damage the photosystem and kill the cell. To ensure that the photosystem works efficiently and to protect it from damage, about half of the energy from the absorbed light is dissipated as heat, while the rest of the energy is stored in the products of photosynthesis.

The standard form of photosystem II uses the energy of visible light, but some cyanobacteria contain different types of photosystem II, which do the same chemical reactions using lower energy far-red light. One type of far-red photosystem II is found in Acaryochloris marina, a cyanobacterium living in stable levels of far-red light, shaded from visible light. The other type is found in a cyanobacterium called Chroococcidiopsis thermalis, which can switch between using its far-red photosystem II when shaded from visible light and using its standard photosystem II when exposed to it. Being able to work with less energy, the two types of far-red photosystem II appear to be more efficient than the standard one, but it has been unclear if there were any downsides to this trait.

Viola et al. compared the standard photosystem II with the far-red photosystem II types from C. thermalis and A. marina by measuring the efficiency of these enzymes, the quantity of reactive oxygen species produced, and the resulting light-induced damage. The experiments revealed that the far-red photosystem II of A. marina is highly efficient but produces elevated levels of reactive oxygen species if exposed to high light conditions. On the other hand, the far-red photosystem II of C. thermalis is less efficient in collecting and using far-red light, but is more robust, producing fewer reactive oxygen species.

Despite these tradeoffs, engineering crop plants or algae that could use far-red photosynthesis may help boost food and biomass production. A better understanding of the trade-offs between efficiency and resilience in the two types of far-red photosystem II could determine which features would be beneficial, and under what conditions. This work also improves our knowledge of how the standard photosystem II balances light absorption and damage limitation to work efficiently in a variable environment.

Photosystem II (PSII) is the water/plastoquinone photo-oxidoreductase, the key energy converting enzyme in oxygenic photosynthesis. The near-universal type of PSII, found in all photosynthetic eukaryotes and in most cyanobacteria, contains 35 chlorophylls a (Chl-a) and 2 pheophytins a (Phe). Four of the Chl molecules (P_D1, P_D2, Chl_D1, and Chl_D2) and both Phe molecules are located in the reaction centre (Diner and Rappaport, 2002). The remaining 31 Chl-a in the PSII core constitute a peripheral light-collecting antenna. When antenna chlorophylls are excited by absorbing a photon, they transfer the excitation energy to the primary electron donor, Chl_D1, the red-most chlorophyll in the reaction centre, although it’s been reported that charge separation from P_D1 can occur in a fraction of centres (Diner and Rappaport, 2002; Holzwarth et al., 2006; Romero et al., 2010; Cardona et al., 2012). The initial charge separation, forming the first radical pair Chl_D1⁺Phe^- (assuming Chl_D1 as primary donor), is quickly stabilized by the formation of the second radical pair, P_D1⁺Phe^-, and then by further electron transfer steps (Figure 1A) that lead to the reduction of plastoquinone and the oxidation of water.

Figure 1

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PSII activity is energy demanding. In Chl-a-PSII, the primary donor absorbs red photons at 680 nm, and this defines the energy available for photochemistry (1.82 eV) with a high quantum yield for the forward reactions. The energy stored in the products of the reaction (reduced plastoquinone and molecular oxygen) and in the trans-membrane electrochemical gradient is ~1 eV, while the remaining ~0.82 eV is released as heat helping to ensure a high quantum yield for the forward reaction and minimize damaging and wasteful side- and back-reactions. The 1.82 eV was suggested to be the minimum amount of energy required for an optimum balance of efficiency versus resilience to photodamage, and responsible for explaining the ‘red limit’ (~680 nm) for oxygenic photosynthesis (Nürnberg et al., 2018; Rutherford et al., 2012).

The first reported case in which the red limit is exceeded was the chlorophyll d (Chl-d)-containing cyanobacterium Acaryochloris marina (A. marina) (Miyashita et al., 1996). Chl-d-PSII contains 34 Chl-d and 1 Chl-a (proposed to be in the P_D1 position Renger and Schlodder, 2008) and uses less energy, with the proposed Chl-d primary donor in the Chl_D1 position absorbing far-red photons at ~720 nm (Schlodder et al., 2007), corresponding to an energy of ~1.72 eV (Figure 1B).

Recently, it was discovered that certain cyanobacteria use an even more red-shifted pigment, chlorophyll f (Chl-f), in combination with Chl-a (Chen et al., 2010; Gan et al., 2014). When grown in far-red light, these cyanobacteria replace their Chl-a-PSII with Chl-f-PSII, that has far-red specific variants of the core protein subunits (D1, D2, CP43, CP47, and PsbH) and contains ~90% of Chl-a and ~10% of Chl-f (Nürnberg et al., 2018; Gan et al., 2014). The Chl-f-PSII from Chroococcidiopsis thermalis PCC7203 (C. thermalis), which contains 30 Chl-a, 4 Chl-f, and 1 Chl-d, was shown to have a long wavelength primary donor (originally proposed to be either Chl-f or d, in the Chl_D1 position Nürnberg et al., 2018) absorbing far-red photons at ~720 nm (Figure 1C), the same wavelength as in A. marina (Nürnberg et al., 2018; Judd et al., 2020). A recent cryo-EM structure has also argued for Chl_D1 being the single Chl-d in the Chl-f-PSII of Synechococcus sp. PCC7335 (Gisriel et al., 2022). This suggests that this could be the case also in the Chl-f-PSII of C. thermalis, because of the conservation of the amino acids coordinating Chl_D1 in the far-red PSII of the two species. The facultative, long-wavelength species that use Chl-f are thus the second case of oxygenic photosynthesis functioning beyond the red-limit (Nürnberg et al., 2018), but the layout of their long wavelength pigments is quite different from that of the Chl-d-PSII.

Assuming that Chl-a-PSII already functions at an energy red limit (Rutherford et al., 2012), the diminished energy in Chl-d-PSII and Chl-f-PSII seems likely to increase the energetic constraints. Thus, if the far-red PSII variants store the same amount of energy in their products and electrochemical gradient, as seems likely, then it was suggested that they should have decreased photochemical efficiency and/or a loss of resilience to photodamage (Nürnberg et al., 2018; Cotton et al., 2015; Davis et al., 2016). These predicted energetic constraints are worth investigating to generate knowledge that could be beneficial for designing strategies aimed at engineering of far-red photosynthesis into other organisms of agricultural or technological interest (Chen and Blankenship, 2011).

Here we report a comparison of the enzyme turnover efficiency, forward reactions, and back-reactions in the three known types of PSII: Chl-a-PSII, and the two far-red types, the Chl-f-PSII from C. thermalis and the Chl-d-PSII from A. marina. To compare the enzymatic properties of the three types of PSII and minimize the effects of physiological differences between strains, isolated membranes rather than intact cells were used. The use of isolated membranes allows the minimization of potential effects due to: (i) the transmembrane electric field, which affects forward electron transfer (Diner and Joliot, 1976) and charge recombination (Joliot and Joliot, 1980), (ii) the uncontrolled redox state of the plastoquinone pool in whole cells, which can affect the Q_B/Q_B^- ratio present in dark-adapted PSII, (iii) differences in the size and composition of the phycobilisomes and in their association with PSII, and (iv) the presence of photoprotective mechanisms such as excitation energy quenching and scavengers of reactive oxygen species.

We investigated several functional properties of the two different types of far-red PSII, (i) the constitutive Chl-d-PSII of A. marina, and (ii) the facultative Chl-f-PSII of C. thermalis. We compared these properties with each other and with those of Chl-a-PSII, from either WL C. thermalis, Synechocystis or T. elongatus, looking for differences potentially related to the diminished energy available in the two long-wavelength PSII variants.

Back reactions and singlet oxygen production

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

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Figure 7—source data 1

D1 sequences used for multi-alignments.

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

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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 .

Effects of the pigment composition on the energetics of the far-red PSII

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

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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.

Table 4

Chl-a-PSII
State	E*	n	Pi
Bulk Chl-a/Phe-a	685	34	0.878
ChlD1680	685	1	0.026
F685	685	1	0.026
F695	695	1	0.071
Chl-d-PSII
State	E*	n	Pi
Chl-a/Phe-a	685	3	0.002
ChlD1720	725	1	0.029
Bulk Chl-d	725	33	0.969
Chl-f-PSII
State	E*	n	Pi
Bulk Chl-a/Phe-a	685	32	0.046
ChlD1721	726	1	0.075
F720/A715	720	1	0.043
F731/A726	731	1	0.117
F737/A732	737	1	0.2
F748/A743	748	1	0.52

1. The states are denoted as follows: Chl_D1 is the primary donor (Pi highlighted in bold), Bulk indicates the antenna pigments considered as isoenergetic, and F indicates the antenna pigments considered separately from the bulk, with the fluorescence emission wavelength indicated. In the case of the far-red pigments in Chl-f-PSII the peak absorptions (A) are also indicated, as taken from Judd et al., 2020.

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.

All data generated and analysed during this study have been included in the manuscript and supporting file and provided as source data files.

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Decision letter after peer review:

Thank you for submitting your article "Impact of energy limitations on function and resilience in long-wavelength Photosystem II" for consideration by eLife. Your article has been reviewed by 4 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Benoît Kornmann as the Senior Editor. The following individuals involved in the review of your submission have agreed to reveal their identity: Jingcheng Huang (Reviewer #1); Elisabet Romero (Reviewer #2).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

This manuscript describes the energetic mechanisms by which two quite different cyanobacteria use far-red light.

Four excellent and thoughtful reviews were obtained. They all expressed that the experiments and data analysis were carefully done and generally support their conclusions. However, some important issues were brought up that should be addressed.

Depending on the specific replies, it will most likely be possible to address these without further experimentation, but some additional work may be required to rule out some alternative explanations.

See in particular comments on the PQ pool and S-states status and misses, and distinguishing singlet oxygen produced by reaction centers and antenna complexes, by Reviewers 1 and 3. Addressing the other (quite constructive) comments should also help clarify and increase the impact of the text.

Reviewer #2 (Recommendations for the authors):

The authors present a comprehensive investigation of the photochemical efficiency and resilience to photodamage of two low-energy Chl-d and Chl-f PSII systems by applying a collection of carefully designed and executed spectroscopic and electrochemical methods. The data presentation in the figures and the explanations provided in the text are clear and concise, and the large number of experimental biological replicas measured together with the comparison with the relevant controls ensure the consistency of the data presented. The conclusions reached are fully supported by the data presented in the main text and further confirmed by the large dataset presented in the supplementary information. Excellent work, in my opinion.

Overall, I consider that this work is methodologically and scientifically rigorous, objectively presented, and of broad interest, and therefore it fulfills the high-quality standards of eLife. For all these reasons I recommend this manuscript for publication in eLife.

Reviewer #3 (Recommendations for the authors):

This is a generally solid manuscript and the contents will be of great value to the photosynthesis research community, as discussed in my public review. However, there are a few points which should be addressed:

1) Have all the authors reviewed this? Stefano Santabarbara's name appears to be misspelled.

2) Particularly in the Introduction, this could use a quick review for spelling and grammar.

3) The abbreviation for pheophytin is given as both "Phe" and "Pheo".

4) Line 102: "the locations of the 4 antenna in the peripheral antenna" – I believe what there are 4 of here would be Chl-f.

5) Lines 117-118: The decision to remove phycobilisomes is understandable and even necessary considering the techniques used, but undercuts the ecological/agricultural interpretations of the results to some extent. Modification of the antenna alongside FRL adaptation is normal, and the lack of this component removes the potential regulating factor of another energy funnel. It might be worth mentioning in the Discussion.

6) Throughout this work, charge recombination is treated as being entirely with the S2 state. Is there any evidence that this is the case? Providing initial S-state populations as obtained from the Kok model used to generate misses might help here. I don't consider this terribly necessary for the main text but it would probably be helpful SI; regardless, all recombination observed is probably with a mixed population of mostly S2 that would be difficult to further clarify but how much of this is actual S2 would be good to know.

7) Line 180-181: It's mentioned that there was not enough PSII in WL membranes from C. thermalis to use as a control for oximetry- this is a bit concerning considering there was apparently enough to generate singlet oxygen. Was using a higher density of membranes not an option? Or, for that matter, additional purification to cores as was done later?

8) What are the flash yields in Figure 3 normalized to? Is it possible to present this data in oxygen per chlorophyll or oxygen per PSII, since it appears the chlorophyll concentration would be known?

9) Line 211: What in C. thermalis membranes makes them scatter so much more light than other strains studied?

10) Line 214-215: For the sake of novice readers, it may be useful to clarify that PsbA3 has the highest sequence identity with Chl-f-PSII D1 out of the T. elongatus PsbA isoforms rather than out of known D1 sequences.

11) Line 218: Please define deltaI/I for first use, this is a fairly niche term.

12) Figure 4: Why is there no large non-oscillating component in T. elongatus absorption and did its absence have any impact on how data modeling was performed?

13) The subsection numbering skips from 3.2 to 3.5.

14) Figure 7 might be better off higher up – it's three pages below its first mention.

15) Line 586: What is the intensity and source of room light used?

16) Were any electron acceptors used in the flash oxygen evolution experiments? On one hand, this is directly discussed in the introduction as necessary to avoid unwanted regulation by the redox poise of the pool in whole cells; on the other, a bare platinum electrode is used.

17) Why was the Kok model described in reference 28 used? This is a fairly outdated model; its creator has developed more detailed ones in the meantime, as have several other groups. It might be worthwhile to reprocess with a newer method, especially when considering the issue of the initial S state distribution not being given.

18) Table S1: With no addition, the slow phase is provided with an amplitude but no time constant, which shouldn't be possible with the model provided- is that just y0 in place of the amplitude?

19) SI lines 163-170: this appears to say that first, Synechocystis membranes have faster fluorescence decay kinetics than intact cells, and later, that Synechocystis has faster decay kinetics in cells than in membranes. Please clarify.

20) Figure S6: The caption references dashed lines; there are no dashed lines in the figure.

Reviewer #4 (Recommendations for the authors):

Importantly, I encourage the authors to remove the ambiguity regarding which Chl type is found in the ChlD1 site of Chl-f-PSII. I find that the evidence for Chl d occupying this site in Synechococcus sp. PCC 7335 is overwhelming, and every aspect of that site is well conserved in C. thermalis and all other FaRLiP-capable cyanobacteria. If there is some argument against this sentiment, the authors could include it. For example, if the authors think the two organisms are different in which pigment is found in ChlD1, it could be mentioned.

Essential revisions:

This manuscript describes the energetic mechanisms by which two quite different cyanobacteria use far-red light.

Four excellent and thoughtful reviews were obtained. They all expressed that the experiments and data analysis were carefully done and generally support their conclusions. However, some important issues were brought up that should be addressed.

Depending on the specific replies, it will most likely be possible to address these without further experimentation, but some additional work may be required to rule out some alternative explanations.

See in particular comments on the PQ pool and S-states status and misses, and distinguishing singlet oxygen produced by reaction centers and antenna complexes, by Reviewers 1 and 3. Addressing the other (quite constructive) comments should also help clarify and increase the impact of the text.

We thank the reviewers and the editor for the thoughtful comments and suggestions. We have tried to address them all in the best possible way. The point-by-point answers are provided below, and the revised manuscript with changes highlighted in red is provided in the Related manuscript file.

Additionally, we have rearranged the format of the manuscript, to comply with eLife’s policy of incorporating as much information as possible within the main text of the article and to replace a single Supplementary Information file with either supplementary figures linked to main figures or appendices, where additional text is required. More specifically:

– Figure S1 (100 s fluorescence decay kinetics) is now provided as Figure 2—figure supplement 1, Tables S1 and S2 have been merged in Table 1 (all fluorescence decay kinetics fits)

– A new figure showing the non-normalised flash dependent O₂ evolution is now provided as Figure 3—figure supplement 1, new Table 2 contains the distribution of S-states calculated from the fits

– Figure S5 is now provided as Figure 5—figure supplement 1, Table S3 is now provided as Table 3 (plots and averages of TL biological replicates)

– Figure S6 and S8 have been merged into Figure 5—figure supplement 2 (Synechocystis TL and luminescence decay)

– Figure S14 is now included in the main text as Figure 7 (D1 multi-alignments)

– Table S5 is now provided as Table 4 (calculations of excited state distribution in the three types of PSII)

The rest of the Supplementary Information has been divided into 7 appendices:

– Appendix 1: comparison of fluorescence decay in Synechocystis and WL C. thermalis

– Appendix 2: period 4 oscillations of thermoluminescence

– Appendix 3: fits of period 4 oscillations of UV absorption

– Appendix 4: discussion of thermoluminescence intensity in present work compared to Nürnberg et al., 2018

– Appendix 5: analysis and interpretation of luminescence decay kinetics

– Appendix 6: all controls for singlet oxygen production

– Appendix 7: supplementary Materials and methods

– Additionally, all Source data files have been mentioned in the corresponding figure legends.

Reviewer #3 (Recommendations for the authors):

This is a generally solid manuscript and the contents will be of great value to the photosynthesis research community, as discussed in my public review. However, there are a few points which should be addressed:

1) Have all the authors reviewed this? Stefano Santabarbara's name appears to be misspelled.

Yes, sorry, all authors did review this article, including Stefano Santabarbara (who did so several times), but everyone (including SS) missed the typo in the name. This has been corrected.

2) Particularly in the Introduction, this could use a quick review for spelling and grammar.

We reviewed the text. In the absence of specific indications in eLife’s instructions for authors, British English spelling has been maintained.

3) The abbreviation for pheophytin is given as both "Phe" and "Pheo".

All corrected to Phe.

4) Line 102: "the locations of the 4 antenna in the peripheral antenna" – I believe what there are 4 of here would be Chl-f.

Yes, when the article was written this was deliberately ambiguous as Nurnberg et al., 2018 had identified the primary donor as being either Chl-d or Chl-f. As a result, the antenna could be 4 Chl-f or 3 Chl-f and 1 Chl-d. A cryo-EM structural model of Chl-f-PSII argued for Chl-d in the Chl_D1 position. While not definitive and obtained in another species, this is an argument in favour of Chl-d being the primary electron donor. Based on the suggestion by reviewer #4 we have opted to remove the ambiguity and this sentence is now modified:

“In (C) the single Chl-d is located in the Chl_D1 position, reflecting the assignment of the single Chl-d as the primary donor (13), leaving the remaining 4 Chl-f molecules as peripheral antenna”.

5) Lines 117-118: The decision to remove phycobilisomes is understandable and even necessary considering the techniques used, but undercuts the ecological/agricultural interpretations of the results to some extent. Modification of the antenna alongside FRL adaptation is normal, and the lack of this component removes the potential regulating factor of another energy funnel. It might be worth mentioning in the Discussion.

The reviewer is right, this has now been mentioned in the Discussion, L. 473–475:

“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 (11)”.

Additionally, the end of the Discussion (L. 491–493) now reads:

“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”.

6) Throughout this work, charge recombination is treated as being entirely with the S2 state. Is there any evidence that this is the case? Providing initial S-state populations as obtained from the Kok model used to generate misses might help here. I don't consider this terribly necessary for the main text but it would probably be helpful SI; regardless, all recombination observed is probably with a mixed population of mostly S2 that would be difficult to further clarify but how much of this is actual S2 would be good to know.

Based on the light pre-treatment followed by dark incubation that we used for all membrane samples, we expected a distribution of ~75% S₁ and ~25% S₀ (or TyrDS₁ which is equivalent in the context of the question) at the beginning of our measurements, as described in Appendix 3 (discussion on the fits of the UV absorption oscillations). Therefore, after one flash we would expect to form ~25% S₁Q_A^–, (which does not recombine) and ~75% S₂Q_A^–, that will recombine. For this reason, we refer to recombination as being with the S₂ state.

This is confirmed by the distribution of S-states obtained by fitting the flash dependent O₂ evolution data, that now have been added in Table 2. According to the fits, in the A. marina and Synechocystis samples there is an initial fraction of S₂ of about 10%, but this is not expected to greatly affect our conclusion that the charge recombination we measure is mostly S₂Q_A^–. It was shown that the other Sstate capable of charge recombination with Q_A^– (and Q_B^–) is S₃. For S₃Q_A^– recombination the kinetics are similar to those for S₂Q_A^– but are pH dependant and at lower pH become slightly less stable. The small fraction S₃Q_A^– predicted based on the S-state fitting are predicted to have a minor effect on the kinetics. The first sentence where recombination is mentioned in section 2.1 (L. 124–126), has been modified as “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 centers where forward electron transfer to Q_B/Q_B^– did not occur.”

7) Line 180-181: It's mentioned that there was not enough PSII in WL membranes from C. thermalis to use as a control for oximetry- this is a bit concerning considering there was apparently enough to generate singlet oxygen. Was using a higher density of membranes not an option? Or, for that matter, additional purification to cores as was done later?

We did measure the flash dependence of oxygen evolution in WL C. thermalis membranes, and we did observe oscillations with visible flashes (but not with far-red flashes, as expected), but the data were just not good enough to be able to perform any significant analysis, so we originally did not include it in the manuscript. The data have now been added in Figure 3—figure supplement 1, together with the non-normalised traces of all other samples, and the text has been modified accordingly.

Using higher density of membranes for the flash polarography would have not improved the situation, reducing the penetration of the light flashes in the sample, and, unfortunately, the case of WL C. thermalis we have never been able to isolate O₂-evolving cores, as stated in L. 194–195.

It is difficult to compare the sensitivity of the flash-dependent polarography and of the O₂ measurements performed with the Clark electrode, because the latter measure O₂ changes that accumulate over time and were performed using chlorophyll concentrations at which light penetration in the sample was not an issue. Additionally, we cannot exclude differences in the PSII content per chlorophyll between the samples used for the different experiments. We note that the physiology of C. thermalis cells has not been studied in detail yet; this means that it is difficult to control the levels of PSII accumulation, that we know depend on the growth stage (among other factors) in model species such as Synechocystis. This is for example visible in the inter-sample variability in the levels of O₂ evolution per chlorophyll measured with the Clark electrode, that have now been added in Appendix 6—table 1.

8) What are the flash yields in Figure 3 normalized to? Is it possible to present this data in oxygen per chlorophyll or oxygen per PSII, since it appears the chlorophyll concentration would be known?

The data were normalised to the oxygen yield of the last of the 40 flashes sequence, where no oscillation of oxygen release remained. This is now specified in the legend of Figure 3. The nonnormalised data (total yield for 10 µg chlorophyll) are now provided in Figure 3—figure supplement 1 (also for WL C. thermalis).

9) Line 211: What in C. thermalis membranes makes them scatter so much more light than other strains studied?

Based on our experience so far, we believe that this is because the cell disruption method we use generates bigger membrane fragments in the case of C. thermalis cells (both WL- and FR-grown) than in the case of other species. Indeed, C. thermalis membranes precipitate at lower centrifugation speed that other membrane samples and are also more difficult to solubilise. Why this is the case, we are not sure, but it could be because C. thermalis cells tend to form clusters encapsulated by a protective layer of excreted polymers, which makes them harder to break.

10) Line 214-215: For the sake of novice readers, it may be useful to clarify that PsbA3 has the highest sequence identity with Chl-f-PSII D1 out of the T. elongatus PsbA isoforms rather than out of known D1 sequences.

Sentence (L. 196–198) modified as

“Therefore, PSII cores from T. elongatus with the D1 isoform PsbA3 (33) 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, section 3.2)”.

11) Line 218: Please define deltaI/I for first use, this is a fairly niche term.

L. 200–202: “As expected, maximum absorption decrease (positive ΔI/I, as defined in Materials and Methods section 4.7) occurred on S₂ (flash 1,5,9 etc.) and maximum absorption increase (negative ΔI/I) on S₀ (flash 3,7,11 etc.)”.

In section 4.7, L. 569–570: “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”.

12) Figure 4: Why is there no large non-oscillating component in T. elongatus absorption and did its absence have any impact on how data modeling was performed?

The non-oscillating component was originally proposed by Lavergne to arise from non-active PSII centres, but we cannot exclude that other photo-induced phenomena could contribute to it in A. marina membranes and partially purified FR C. thermalis PSII. Since the T. elongatus sample is constituted by basically 100% active PSII, this non-oscillating component is smaller. The component is not expected to have any impact on the fit, since the first flash was excluded from the analysis as indicated in Appendix 3, that now reads:

“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 (28). 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”.

13) The subsection numbering skips from 3.2 to 3.5.

Corrected, thank you.

14) Figure 7 might be better off higher up – it's three pages below its first mention.

In the format required for the revised version of the manuscript figures are no longer embedded in the text.

15) Line 586: What is the intensity and source of room light used?

Room light was provided by a white fluorescent lamp (Philips Master TL5 High Efficiency), the intensity at which the samples are pre-illuminated was about 80 µmol photons m^–2 s^–1. This information has been added in the Materials and methods.

16) Were any electron acceptors used in the flash oxygen evolution experiments? On one hand, this is directly discussed in the introduction as necessary to avoid unwanted regulation by the redox poise of the pool in whole cells; on the other, a bare platinum electrode is used.

No, the flash oxygen evolution measurements were performed in absence of electron acceptors, and this could be one of the reasons for the slightly higher miss factor obtained in some samples, compared to those obtained with the UV measurements (that were done in presence of the electron acceptor PPBQ). This is mentioned in L. 185–187 (“Nevertheless, these differences, attributed to the combination of the absence of exogenous electron acceptors, and the relatively long and possibly not fully saturating flashes, were not significant.”) and L. 190–191 (“These measurements were done in the presence of the electron acceptor PPBQ and using single-turnover monochromatic saturating laser flashes”).

17) Why was the Kok model described in reference 28 used? This is a fairly outdated model; its creator has developed more detailed ones in the meantime, as have several other groups. It might be worthwhile to reprocess with a newer method, especially when considering the issue of the initial S state distribution not being given.

We are not sure about which more detailed model the reviewer is referring to for the fit of the period 4 oscillations at 291 nm. In contrast to the period 4 oscillation of O₂ evolution where only the S₃ to S₀ transition give a signal (the O₂ detected at the electrode), at 291 nm, each S-state transition is characterized by a different absorption change (3 in total, because the ΔI/I for the S₃ to S₀ transition is necessarily the negative sum of the 3 other transitions). This requires a specific treatment that was done only by Jérôme Lavergne, as far as we are aware. This model developed by Lavergne was more recently detailed in the supplementary material of https://doi.org/10.1074/jbc.M710583200.

The parameters to be minimized are the S₁/S₀ ratio (in fact, the apparent S₀, which includes the S₀Tyr_D^Δ and S₁Tyr_D states, as we discussed in the Supplementary Information, with the proportion of S₁ given in Figure S4), the miss parameter (in our case we do not take into account a double hit parameter since we are using a 6 ns laser flash illumination) and the 3 transition weights, T0(i), T1(i), T2(i), calculated from the proportion of each state taking into account the S₁/S₀ ratio and the miss parameter.

T0(i) = S0(i‐1) – S0(i)

T1(i) = T0(i) + S1(i‐1) – S1(i)

T2(i) = T1(i) + S2(i-1) – S2(i)

The ΔI/I upon each flash of the sequence were then calculated by using 3 arbitrary starting values for Δε0, Δε1 and Δε2, for the S₀ to S₁, S₁ to S₂ and S₂ to S₃ transitions, respectively, by using the straightforward formula:

Adjustment of (ΔI/I)(i) computed from the equation above to the experimental ΔI/I values was done by varying the S₁ and S₀ proportions, the 3 coefficients Δε0, Δε1 and Δε2 and the miss parameter.

More recent models include a miss parameter that can vary for each S-state transition. This results in a greater number of variables to minimize when compared to the number of experimental data points, with the risk that the procedure may result in a large variation in the value of some variables that is offset by too much variation in other variables.

For this reason, the fits were done using S-state independent misses, also in the case of the flash dependent O₂ evolution measurements (added in corresponding Materials and methods section).

18) Table S1: With no addition, the slow phase is provided with an amplitude but no time constant, which shouldn't be possible with the model provided- is that just y0 in place of the amplitude?

Yes, the reviewer is right, we did not describe it properly: we provide the cumulative amplitudes of the slowest exponential decay phase (A3) and of y₀, and for this reason we do not provide the time constant. The table (now Table 1) has been modified accordingly, and this has been specified in the table legend.

19) SI lines 163-170: this appears to say that first, Synechocystis membranes have faster fluorescence decay kinetics than intact cells, and later, that Synechocystis has faster decay kinetics in cells than in membranes. Please clarify.

There was a mistake in the first sentence, that is now corrected to

“The fluorescence decay kinetics measured here in Synechocystis membranes, as well as those measured in A. marina and C. thermalis membranes, are slower than those measured in Synechocystis intact cells in a previous work (44)” (beginning of Appendix 1). Overall, the Q_A^– decay kinetics are faster in cells than in isolated membranes for the reasons mentioned in Appendix 1 (corrected to remove the wrong reference to ref. 17): “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 (18). 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”.

20) Figure S6: The caption references dashed lines; there are no dashed lines in the figure.

The dashed lines were present in the original graph but were not preserved when transferring the graph from the Origin software to Word. The figure (now Figure 5—figure supplement 2A) has been modified to make them visible.

Author details

1. Stefania Viola

   Department of Life Sciences, Imperial College London, London, United Kingdom

   Contribution

   Conceptualization, Data curation, Formal analysis, Investigation, Writing – original draft, Writing – review and editing

   For correspondence

   s.viola@imperial.ac.uk

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-0773-8158

2. William Roseby

   Department of Life Sciences, Imperial College London, London, United Kingdom

   Contribution

   Investigation

   Competing interests

   No competing interests declared

3. Stefano Santabarbara

   Photosyntesis Research Unit, Consiglio Nazionale delle Ricerche, Milan, Italy

   Contribution

   Data curation, Formal analysis, Writing – review and editing

   Competing interests

   No competing interests declared

4. Dennis Nürnberg

   Physics Department, Freie Universität Berlin, Berlin, Germany

   Contribution

   Data curation, Formal analysis, Investigation

   Competing interests

   No competing interests declared

5. Ricardo Assunção

   Physics Department, Freie Universität Berlin, Berlin, Germany

   Contribution

   Data curation, Formal analysis, Investigation

   Competing interests

   No competing interests declared

6. Holger Dau

   Physics Department, Freie Universität Berlin, Berlin, Germany

   Contribution

   Resources, Formal analysis

   Competing interests

   No competing interests declared

7. Julien Sellés

   Institut de Biologie Physico-Chimique, UMR CNRS 7141 and Sorbonne Université, Paris, France

   Contribution

   Formal analysis, Investigation

   Competing interests

   No competing interests declared

8. Alain Boussac

   Institut de Biologie Intégrative de la Cellule, UMR9198, CEA Saclay, Gif-Sur-Yvette, France

   Contribution

   Formal analysis, Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

9. Andrea Fantuzzi

   Department of Life Sciences, Imperial College London, London, United Kingdom

   Contribution

   Conceptualization, Formal analysis, Funding acquisition, Project administration, Writing – review and editing

   Competing interests

   No competing interests declared

10. A William Rutherford

    Department of Life Sciences, Imperial College London, London, United Kingdom

    Contribution

    Conceptualization, Formal analysis, Funding acquisition, Writing – original draft, Project administration, Writing – review and editing

    For correspondence

    a.rutherford@imperial.ac.uk

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-3124-154X

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