1. Plant Biology
  2. Structural Biology and Molecular Biophysics
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The structure of photosystem I from a high-light-tolerant cyanobacteria

  1. Zachary Dobson
  2. Safa Ahad
  3. Jackson Vanlandingham
  4. Hila Toporik
  5. Natalie Vaughn
  6. Michael Vaughn
  7. Dewight Williams
  8. Michael Reppert
  9. Petra Fromme
  10. Yuval Mazor  Is a corresponding author
  1. School of Molecular Sciences, Arizona State University, United States
  2. BiodesignCenter for Applied Structural Discovery, Arizona State University, United States
  3. Department of Chemistry, Purdue University, United States
  4. John M. Cowley Center for High Resolution Electron Microscopy, Arizona State University, United States
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Cite this article as: eLife 2021;10:e67518 doi: 10.7554/eLife.67518

Abstract

Photosynthetic organisms have adapted to survive a myriad of extreme environments from the earth’s deserts to its poles, yet the proteins that carry out the light reactions of photosynthesis are highly conserved from the cyanobacteria to modern day crops. To investigate adaptations of the photosynthetic machinery in cyanobacteria to excessive light stress, we isolated a new strain of cyanobacteria, Cyanobacterium aponinum 0216, from the extreme light environment of the Sonoran Desert. Here we report the biochemical characterization and the 2.7 Å resolution structure of trimeric photosystem I from this high-light-tolerant cyanobacterium. The structure shows a new conformation of the PsaL C-terminus that supports trimer formation of cyanobacterial photosystem I. The spectroscopic analysis of this photosystem I revealed a decrease in far-red absorption, which is attributed to a decrease in the number of long- wavelength chlorophylls. Using these findings, we constructed two chimeric PSIs in Synechocystis sp. PCC 6803 demonstrating how unique structural features in photosynthetic complexes can change spectroscopic properties, allowing organisms to thrive under different environmental stresses.

Introduction

Oxygenic photosynthesis evolved on earth about 2.5 billion years ago (Bekker, 2004). Plants, algae, and cyanobacteria carry out this process and are found in a wide variety of environments. Despite nearly 3 billion years of evolution, all oxygenic photosynthetic organisms use the same large pigment–protein complexes, known as photosystem I (PSI) and photosystem II (PSII), to convert solar energy to chemical energy (Fromme et al., 2003; Witt, 1996; Zouni et al., 2000). Both complexes use light to induce a charge separation event, then transport the high-energy electron, ultimately to be stored as a chemical bond (Fromme et al., 2003; Witt, 1996; Zouni et al., 2000). Although light is required for photosynthesis, an excess of light can be detrimental to photosynthetic organisms by damaging the photosynthetic proteins in a process called photoinhibition (Kok, 1956; Jones and Kok, 1966; Gururani et al., 2015). Therefore, the ability to adapt to different qualities and quantities of light is paramount for photosynthetic organisms to survive.

To prevent photoinhibition, photoprotective mechanisms such as changes in the photosystem content (Muramatsu and Hihara, 2012; Murakami and Fujita, 1991; Demmig-Adams and Adams, 1992; Miskiewicz et al., 2002), dissipation of excess energy as heat (Peltier and Schmidt, 1991; Levy et al., 1993; Jacob and Lawlor, 1993), and upregulation of antioxidant enzymes have been observed (Tsang et al., 1991; Camp, 1994; Zhang and Scheller, 2004; Ort and Baker, 2002; Horling, 2003). A response shown by the model cyanobacterium Synechocystis sp. PCC 6803 to high light is to decrease the PSI:PSII ratio, minimizing its light harvesting capacity and presumably protecting against the generation of reactive oxygen species around PSI (Hihara et al., 1998; Sonoike et al., 2001). This is in agreement with the response observed in vascular plants (Anderson, 1986). However, this response is not universal. In the terrestrial cyanobacteria, Synechococcus OS-B’, isolated from a microbial mat in a hot spring, the ratio of PSI:PSII increases upon high-light conditions (Kilian, 2007), demonstrating the importance of PSI in the high-light response of cyanobacteria.

A significant amount of work on photoinhibition has focused on PSII due to its rapid turnover in high light and the efficient repair mechanisms that evolved to cope with PSII-specific photodamage (Yao et al., 2012a; Yao et al., 2012b). PSI-specific damage, however, is irreversible and long lived due to a lack of repair mechanisms, requiring the biosynthesis of new PSI polypeptides (Li et al., 2004; Sonoike, 2010; Terashima et al., 1994; Sonoike et al., 1995; Lima-Melo et al., 2019). Fluctuating light and low temperatures have been attributed to PSI photoinhibition by causing an imbalance in the redox state of PSI donors and acceptors (Sonoike, 2010; Allahverdiyeva et al., 2015; Tiwari, 2016; Kudoh and Sonoike, 2002; Kono and Terashima, 2016). Recovery times in leaves for PSI-specific photodamage have been reported to be longer than a week, much longer than the 30 min half-life of the D1 protein of PSII (Zhang and Scheller, 2004; Yao et al., 2012b; Li et al., 2004; Kudoh and Sonoike, 2002; Zhou et al., 2004; Kanervo et al., 1993). PSI inhibition is therefore potentially more devastating than damage to PSII because it results in the over-reduction of the plastoquinone pool subsequently inhibiting PSII and thereby blocking the complete electron transfer chain (Sonoike, 2010). This has lead to the proposal that photoinhibition of PSII is a mechanism to protect PSI by reducing the amount of electrons sent to PSI (Barbato et al., 2020; Tikkanen et al., 2014). Furthermore, there are multiple mechanisms acting on both the lumen and stromal side to reduce PSI damage under these conditions (Sonoike, 2010; Tiwari, 2016; Barbato et al., 2020; Munekage, 2002; Munekage et al., 2008; Suorsa et al., 2013; Suorsa, 2015), suggesting PSI damage plays an important role in adaptation to stress. The first high-resolution structure of cyanobacterial PSI revealed a core antenna system comprised of 96 chlorophyll a (Chla) molecules, of which 6 are integral to the electron transport chain (ETC), while the remaining chlorophyll molecules function to harvest light and transfer excitation energy to the ETC (Fromme et al., 2006; Jordan et al., 2001). Still, the roles and properties of individual chlorophylls in the antenna remain largely unknown. While all chlorophyll molecules in cyanobacteria are chemically identical except one (one Chla of P700 is Chla’, the C13 epimer of Chla), the local environment of a few individual chlorophylls has been shown to extend their absorbance properties above 700 nm, giving rise to long-wavelength chlorophylls (LWC) (Fromme et al., 2006; Wientjes et al., 2012; Croce and van Amerongen, 2014).

Further studies have revealed that, in cyanobacteria, LWC are associated with PSI and the amount of these pigments vary between species, suggesting that the number of LWC is an important evolutionary adaptation (Shubin et al., 1991; Gobets et al., 2001). It has also been shown that LWC are strongly affected by their immediate chemical environment. The absorbance properties of specific chlorophylls are highly dependent on its coordinating residues as well as excitonic coupling between neighboring chlorophylls; and changing either can alter the spectroscopic properties of a chlorophyll. (Wientjes et al., 2011; Toporik, 2020; Khmelnitskiy et al., 2020). These characteristics have led to several suggestions to the physiological role of LWC such as directing energy to P700, extending light-harvesting capabilities into the far red, and photoprotective mechanisms (Valkunas et al., 1995; Trissl, 1993; Rivadossi et al., 1999; Schlodder et al., 2005; Herascu et al., 2016).

While the function of PSI is conserved in all photosynthetic organisms, the oligomeric state and subunit composition of PSI varies. Higher plants exclusively utilize monomeric PSI, whereas cyanobacteria utilize monomeric, trimeric, and tetrameric oligomers (Li et al., 2014; Li et al., 2019; Zheng et al., 2019). It has been suggested that trimerization is a way of modulating light harvesting in changing light conditions (Chitnis and Chitnis, 1993; Sener et al., 2004). The PsaL subunit has been shown to be vital for these larger oligomeric complexes to form (Li et al., 2019; Zheng et al., 2019; Chitnis and Chitnis, 1993; Malavath et al., 2018). Adding a single histidine to the C-terminus of PsaL was shown to completely disrupt trimerization in Synechocystis (Malavath et al., 2018; Netzer-El et al., 2018). The first crystal structures of PSI from Thermosynechococcus elongatus and Synechocystis sp. PCC. 6,803 (Synechocystis) showed that the C-terminus of PsaL coordinates a calcium ion together with the PsaL subunit of the adjacent monomer, which was suggested to stabilize trimer formation and emphasized the importance of the C-terminus of PsaL in oligomerization (Jordan et al., 2001; Mazor et al., 2013).

In order to understand how the light harvesting machinery has evolved to adapt to high-light conditions, we isolated a cyanobacterium from the Sonoran Desert, an environment with light intensities regularly exceeding 1600 µmol photon m–2s–1, in order to characterize its photosynthetic machinery. Genomic sequencing revealed that this cyanobacterium is a new strain of Cyanobacterium aponinum. Other strains of C. aponinum have been shown to grow in both freshwater and seawater, as well as extreme environments with temperatures reaching 45°C (Moro et al., 2007; Winckelmann et al., 2015). The ability of C. aponinum to survive in these vastly different environments make it a promising candidate for biofuel production (Ertugrul and Dönmez, 2011). Here we report the structure and the spectroscopic characterization of the trimeric PSI complex isolated from the high-light tolerant C. aponinum. We designed two chimeric PSIs in Synechocystis to test the functionality of structural variations between C. aponinum and Synechocystis. Our results demonstrate how the structure of PSI modulates its spectroscopic properties and elucidate the mechanisms controlling PSI oligomerization in cyanobacteria, bringing the ability to design large photosynthetic complexes with desired optical properties a step closer.

Results

C. aponinum 0216 is a high-light-tolerant cyanobacteria

To study a photosynthetic organism that exhibits the ability to grow in high-light environments, samples were taken from a biofilm growing on a south facing concrete wall of a freshwater reservoir in Tempe, AZ, that had a constant drip of fresh water and exposed to over 300 days of sunlight per year. Samples were taken in February, which has an average temperature of 18.7°C according to the National Climatic Data Center (NOAA) for this area (LOCAL CLIMATOLOGICAL DATA, 1946). Samples were cultivated in BG-11 growth media and exposed to light intensities exceeding 3000 µmol photon m–2s–1 of warm white light at 30°C for a week to select for organisms able to survive high-light conditions. One photoautotroph was able to survive these light intensities and was subsequently isolated through continuous streaking on BG-11 media agar plates supplemented with iron (Figure 1). Genomic DNA was extracted, and the 16 S rRNA was amplified to identify the organism (Nubel et al., 1997). The 16 S rRNA was compared to other cyanobacterial 16 s rRNA libraries revealing a close relationship to C. aponinum strains (Figure 1B).

Isolation of a high-light-tolerant cyanobacteria.

(A) Cross-sectional negative stained image of C. aponinum fixed in acrylic medium. (B) Phylogenetic analysis based on C. aponinum 16 S rRNA. Evolutionary analyses were conducted in MEGA7. (C) Serial dilutions of Synechocystis and C. aponinum on BG11 plates. Cells were serially diluted in ¼ steps and incubated at 30°C for 5 days (light intensities > 370 µmol photons m–2s–1) and 10 days (light intensity = 50 µmol photons m–2s–1) (D) in vivo absorption spectra (normalized to the max wavelength of the Qy transition) of C. aponinum cells grown in low light (45 µmol photons m–2s–1) and high light (450 µmol photons m–2s–1). (E) 77 K fluorescence spectra (normalized to the max emission wavelength) of whole cells excited at 440 nm.

The ability of C. aponinum to grow in high light was compared directly to Synechocystis across several light intensities (Figure 1C). Cells were serially diluted and exposed to a range of light intensities to determine the viability for growth. C. aponinum grew in much higher light intensities than Synechocystis, with the ability to survive in conditions as high as 1850 µmol photons m–2s–1. Under these conditions, Synechocystis cannot grow regardless of the density of the cells on the culture plate, thus emphasizing the innate ability of C. aponinum to grow under high-light conditions.

The response of C. aponinum to high-light conditions was measured by comparing C. aponinum grown under high- (450 µmol photons m–2s–1) and low-light (45 µmol photons m–2s–1) conditions. Absorption spectra revealed that cells grown in high light show an increase in absorption between 400 and 550 indicating a higher carotenoid content relative to chlorophyll, which is a known response to high light in photosynthetic organisms (Sozer, 2010; Figure 1D). Additionally, low-temperature fluorescence measurements (77 K) indicated that the F722:F685 ratio, a proxy for the distribution of excitation energy between PSI:PSII in vivo (Murakami, 1997), increases from 2.00 in low-light cells to 3.14 in cells grown in high light (Figure 1E). This increase under high light led us to investigate PSI and determine if it is involved in the high-light tolerance observed in C. aponinum.

C. aponinum PSI contains less LWC and has a modified PsaL

To explore possible adaptations of PSI to high light, the complex was isolated using anion exchange chromatography of solubilized thylakoid membranes. The sucrose density gradient shows the chlorophyll-containing species (Figure 2A). Comparing this sample to a known PSI sample from Synechocystis, SDS–PAGE shows similar bands for PSI subunits in Figure 2B, with notable shifts in the PsaD, PsaF, PsaL, and PsaC subunits.

Figure 2 with 1 supplement see all
Isolation and characterization of trimeric PSI.

(A) Ten percent to 30% sucrose gradient of solubilized C. aponinum membranes following an anion exchange chromatography. (B) SDS–PAGE of the main sucrose gradient band (Trimer) compared to PSI isolated from Synechocystis. A notable difference between the PsaL bands is clearly observable around 13 kD. (C) Absorption spectra of the purified trimer of C. aponinum (green) and Synechocystis (black) normalized to the area between 550 and 775 nm (D) Difference of the C. aponinum – Synechocystis absorbance spectra, shown are averages of three biological replicas, blue edges indicate± SD. (E) The negative peak at 701 nm of the absorbance difference spectrum (dashed black line) is fitted to a sum of three gaussian components colored blue, purple, and red with the sum as a solid black line and the residual of the fit (F) 77 K fluorescence of C. aponinum (green) and Synechocystis (black) using an excitation wavelength of 440 nm. Samples were normalized to their max peak.

To investigate the different migration of PSI subunits between C. aponinum and Synechocystis, C. aponinum genomic DNA was isolated and sequenced (NCBI:txid2676140). PSI genes were located, annotated, and compared to Synechocystis (Figure 2—figure supplement 1). The sequence of the PsaL gene revealed that the difference in migration is likely due to two substantial differences compared to Synechocystis: (1) a six amino acid insert located on the stromal side of the membrane between two transmembrane helixes and (2) a markedly different C-terminus (Figure 2—figure supplement 1) containing an extension of four amino acids. In addition, a seven amino acid insertion is seen in the PsaB gene of C. aponinum compared to Synechocystis (Figure 2—figure supplement 1). Genes for the remaining subunits (PsaD, PsaF, and PsaC) did not reveal differences that would correspond to these shifts (Figure 2—figure supplement 1). There were however sequence variations between C. aponinum and Synechocystis, which could cause gel shifting, a common occurrence when analyzing membrane proteins via SDS–PAGE (Rath et al., 2009).

Absorption spectra from PSI purified from C. aponinum and Synechocystis were normalized to the area between 550 nm to 775 nm to account for the individual contribution of each chlorophyll (Figure 2C). The difference spectrum of the absorption, C. aponinum–Synechocystis, shows a strong negative peak at 701 nm revealing that C. aponinum PSI contains less LWC than Synechocystis (Figure 2D). Based on the Gaussian deconvolution of this peak three components were identified at 699, 704, and 711 nm, the latter two in agreement with the site energy of previously reported LWC in Synechocystis that would be altered in C. aponinum (Toporik, 2020; Khmelnitskiy et al., 2020; Figure 2E).

Surprisingly, the 77 K emission spectra shows that the emission peak of C. aponinum PSI displays a 2 nm red shift compared to Synechocystis PSI (Figure 2F). We attribute this shift to enhanced emission from a red state common to both complexes, as we only detected loss of LWC in C. aponinum compared to Synechocystis. To identify which chlorophylls could be responsible for the different spectroscopic properties, we determine the structure of the PSI trimer from C. aponinum using Cryo-EM.

Structure of trimeric PSI from C. aponinum

Using single-particle cryo-EM, the structure of trimeric C. aponinum PSI seen in Figure 3A was determined to 2.7 Å resolution as assessed by the gold-standard Fourier shell correlation (FSC) criteria (Henderson, 2011) when imposing C3 symmetry (Table 1 and Figure 3—figure supplements 1 and 2). Local resolution calculations show that regions of the interior of the protein are resolved to the Nyquist limit of 2.1 Å, while surface- and membrane-exposed regions are resolved to 2.5 Å resolution (Figure 3—figure supplement 2). The complex contains 33 protein subunits, 288 chlorophylls, 72 carotenoids, 6 phylloquinones, and 9 iron–sulfur clusters as shown in Figure 3A. Interestingly, no density was observed for the Ca2+ ion in the PsaL subunit that was proposed to stabilize trimerization in previously solved trimeric PSI structures (Figure 3E,F; Jordan et al., 2001; Mazor et al., 2013).

Figure 3 with 6 supplements see all
The structure of trimeric PSI from C. aponinum.

(A) C. aponinum trimeric PSI (B) chlorophyll B40 shifts its position due to the insertion seen in the PsaB subunit in C. aponinum (green) compared to Synechocystis (black). (C) The PsaL subunits of C. aponinum (green) and Synechocystis (black) showing the difference of the overall structure of the PsaL C-terminus. (D) The C-terminus of the PsaL subunit of C. aponinum (green) and Synechocystis (black) displaying the coordination to the Ca2+ in the adjacent monomer in Synechocystis, but is absent in C. aponinum and the Red_d mutant of Synechocystis. (E) C. aponinum and its electron density map compared to (F) Synechocystis (PDBID 5OY0, shown with 2Fo-Fc map) clearly depicting no density for the Ca2+ ion in the map for C. aponinum.

Table 1
Cryo-EM data collection, refinement, and validation statistics.
PSI complex(EMD-21320, PDB-6VPV)
Data collection and processing
Calibrated pixel size (Å)Detector, physical pixel size (µm)1.05K2 summit, 5
Voltage (kV)300
Total electron dose (e2)61
Defocus range (μm)–1.5 to – 3.0
Super pixel size (Å)0.525
Symmetry imposedC3
Initial particle images (no.)256,410
Final particle images (no.)73,984
Map resolution (Å)2.7
FSC threshold0.143
Map resolution range (Å)2.1–4.1
Refinement
Initial model used (PDB code)5OY0
Model resolution (Å)2.7
FSC threshold0.143
Model resolution range (Å)2.1–4.1
Map sharpening B factor (Å2)–72.48
Model composition
Nonhydrogen atoms71,814
Protein residues6,743
Ligands384
B factors (Å2)
Protein50.00/137.33/86.61
Ligand27.10/131.48/54.64
R.m.s. deviations
Bond lengths (Å)0.005
Bond angles (°)0.894
Validation
MolProbity score1.82
Clashscore10.36
Poor rotamers (%)0.0
Ramachandran plot
Favored (%)95.89
Allowed (%)4.11
Disallowed (%)0

Comparison of the PSI chlorophyll arrangement between C. aponinum and Synechocystis reveals a high degree of conservation, except for chlorophyll B40 (Figure 3B). This chlorophyll is located next to a seven amino acid insertion seen in the sequence alignment of PsaB (Figure 3—figure supplement 3) that creates a loop sterically forcing chlorophyll B40 to shift its orientation in C. aponinum. As a result, the coupling between chlorophyll B40 and B19 changes. The significance of this conformation was determined using a combination of mutagenesis and modeling (see below). Shorter insertions at the same PsaB location are also observed in PSI sequences from other photoautotrophs, including T. elongatus and Pisum sativum, marking this PsaB loop as a unique, variable region, in the core PSI (shown in Figure 3—figure supplement 3). The structure of T. elongatus PSI shows the corresponding B40 chlorophyll is not present due to an additional subunit, PsaX, which results in a different conformation of this loop relative to C. aponinum (Jordan et al., 2001), sterically blocking the binding site of chlorophyll B40. In the structure of P. sativum PSI, chlorophyll B40 is present; however, it exhibits a similar shift compared to C. aponinum because of a two amino acids insertion (Figure 3—figure supplement 3).

In addition to the rearrangement around chlorophyll B40, an additional chlorophyll molecule was modeled on the stromal side of the PsaK subunit in the C. aponinum structure. This chlorophyll was not modeled in early cyanobacteria PSI structures; however, it has been recently resolved in PSI structures from Synechocystis and Synechococcus sp. PCC 7942 (henceforth Synechococcus) (Toporik, 2020; Cao et al., 2020).

The PsaL subunit in C. aponinum reveals two drastically different features compared to Synechocystis: First, a large loop on the stromal side of the membrane (Figure 3—figure supplement 4) and second, a C-terminus that does not bridge adjacent monomers through the coordination of a Ca2+ ion (Figure 3C,D; Chitnis and Chitnis, 1993; Malavath et al., 2018). The loop on the stromal side in C. aponinum lays relatively flat along the membrane plane and only differs slightly in conformation compared to plant PSI due to the PsaH subunit, which is missing in cyanobacteria. In the plant PSI structure this loop is raised to accommodate the binding of PsaH shown in Figure 3—figure supplement 4.

Previously solved PSI structures from cyanobacteria reveal that the C-terminus of PsaL from one monomer coordinates a Ca2+ ion together with a negatively charged residue from the adjacent PsaL subunit (Jordan et al., 2001; Mazor et al., 2013). However, in the structure from C. aponinum this interaction does not occur, as shown in Figure 3C. Comparing the structures of Synechocystis and C. aponinum revealed that at the position of aspartate 73, one of the Ca2+ coordinating residues in Synechocystis, a leucine is present in C. aponinum (Figure 3D). Unlike aspartate, leucine is an uncharged species which prevents the coordination of this Ca2+ ion. Furthermore, the C-terminus in C. aponinum is longer than Synechocystis (Figure 2—figure supplement 1 and Figure 3—figure supplement 4) and would sterically prevent oligomerization if adopting the same orientation as Synechocystis.

A similar orientation of PsaL was shown in the structure of the PSI-IsiA antenna super-complex from Synechococcus. In this structure, there is also no Ca2+ ion modeled and an asparagine is in the coordination position (Cao et al., 2020). To understand the frequency of cyanobacterial PSI that coordinate a calcium ion, a protein alignment of 680 cyanobacteria PsaL sequences was constructed and sorted by their position on an evolutionary tree. This alignment revealed that the position of the calcium coordinating residue is variable in cyanobacteria. Early structures suggest a negatively charged residue is crucial for the calcium coordination (Jordan et al., 2001; Malavath et al., 2018); however, negatively charged residues were present in only 48% of sequences in this alignment (Figure 3—figure supplement 5). The next most prevalent residue in this position is asparagine, occurring in 30% of the sequences. The recent PSI-IsiA structure from Cao et al., 2020 contains an asparagine at this position, demonstrating that asparagine does not coordinate an calcium ion. Additionally, in eukaryotic organisms, which contain monomeric PSI, the residue at this position is as asparagine in 81% of sequences (Figure 3—figure supplement 6). Therefore, other interactions must be present for trimerization to occur, as less than 50% of cyanobacteria contain a residue capable of coordinating Ca2+ at this position.

Structural changes lead to spectral shifts around the Qy transition

To explore the functional significance of the structural differences highlighted in Figure 3 between C. aponinum and Synechocystis (henceforth WT Synechocystis), two mutant strains of Synechocystis were constructed. One, Red_c, contains the sequence of the PsaB loop from C. aponinum (Figure 3B, Figure 3—figure supplement 3) and the other, Red_d, contains a point mutation at the calcium coordinating aspartic acid in PsaL to a leucine as observed in C. aponinum (Figure 3D–F).

Trimeric PSI was purified from both Red_c and Red_d (Figure 4A). Subsequence SDS–PAGE shows that these samples have the same subunit composition seen in the WT Synechocystis PSI (Figure 4B). Absorbance spectra were normalized to the area between 550 nm and 775 nm (Figure 4C), and the spectrum of WT Synechocystis PSI was subtracted from both Red_c and Red_d (Figure 4D).

Figure 4 with 1 supplement see all
Spectroscopic analysis of Red_c.

(A) Ten percent to 30% sucrose gradient of solubilized membranes from WT Synechocystis, Red_c, and Red_d after purification by anion exchange. (B) SDS–PAGE of the main sucrose gradient bands in comparison with PSI isolated from WT Synechocystis. (C) Absorption spectra of the purified trimer of Red_c mutant (blue), Red_d (red), and WT Synechocystis (black) normalized to the area between 550 and 775 nm (D) Difference spectra of the Red_c – WT Synechocystis (blue) and the Red_d – WT Synechocystis absorbance spectra (red). (E) Room temperature emission using an excitation wavelength of 440 nm. Samples were normalized to their max peak. (F) 77 K fluorescence of C. aponinum (green), WT Synechocystis (black), Red_c (blue), and Red_d (red) using an excitation wavelength of 440 nm. Samples were normalized to their max peak.

The difference spectrum of Red_c – WT Synechocystis around the chlorophyll Qy transitions revealed a positive peak with a maximum at 669 nm and a negative peak with a minimum at 685 nm, but relatively little change in wavelengths above 700 nm. We attribute these differences to the altered orientation of chlorophyll B40 in Red_c (see below). The difference spectrum of Red_d – WT Synechocystis (Figure 4D) showed that removal of the Ca2+ clearly affect the absorption of PSI in the far-red region of the Qy transition, evident by the negative peak centered at 704 nm. In contrast to affecting individual chlorophyll coordination or orientation (by point mutations or loop insertion) removing the positive charge of the Ca2+ ion is expected to have a more global effect, possibly influencing several chlorophylls in different ways (depending on or their orientation and distance from the Ca2+ ion). We attribute the additional features in the Red_d difference spectra to interactions between the Ca2+ and neighboring chlorophylls which are not part of the LWC in PSI. The identity of the LWC affected by the removal of Ca2+ ion is discussed further below.

Further confirmation that LWC absorption is modified by Ca2+ binding in PSI is seen in room temperature emission from PSI. While the emission from trimers isolated from Red_c strains is similar to the wild-type trimer at the red regions of the spectra, the emission from PSI isolated from Red_d strains is different and its intensity at the red region of the spectra is significantly lower than emission from wild-type trimers (Figure 4E). Surprisingly, despite removing LWC in the Red_d strain, we did not resolve differences in the 77 K emission peak between WT Synechocystis and Red_d (Figure 4F). These remaining differences show that additional mutations are needed to fully account for the differences between the two strains (see Discussion).

It was previously shown that disruption (deletion or C terminal extension) of the PsaL subunit prevents trimerization in PSI (Chitnis and Chitnis, 1993; Malavath et al., 2018). A PSI trimer was readily isolated from Red_d, showing that Ca2+ binding is not essential for trimer formation. However, we investigated the distribution of the trimer configuration in native membranes by solubilizing and running a sucrose gradient without additional purification steps (Figure 4—figure supplement 1). These experiments showed a higher monomer to trimer ratio in the Red_d strain compared to WT Synechocystis, suggesting that although Ca2+ is not required for trimerization, Ca2+ coordination does play a role in stabilizing the trimeric organization in WT Synechocystis.

Modified excitonic interactions explains the observed spectral differences

The shape of the difference spectra between Red_c and WT Synechocystis (Figure 4D) is easily understood as a consequence of oscillator strength redistribution amongst a coupled cluster of chlorophyll pigments, an effect often observed in photosynthetic hole burning spectra (Reppert et al., 2010; Reppert et al., 2009; Reppert et al., 2008). Briefly, the shift in conformation of chlorophyll B40 is expected to modify both its transition dipole orientation and excitonic coupling interactions with its neighboring pigments, particularly chlorophylls B18 and B19. These altered interactions affect both the transition energies and oscillator strengths of the B18/B19/B40-cluster exciton states, presumably (based on the experimental absorption difference spectrum) shifting absorption intensity from a low-energy exciton near 686 nm to a higher-energy band near 670 nm (Reppert et al., 2010; Reppert et al., 2009; Reppert et al., 2008).

To test this explanation, we calculated electronic transition dipoles and coupling elements amongst chlorophylls B18, B19, and B40 using the transition electrostatic potential (TrESP) method (Madjet et al., 2006). For the TrESP calculation, we used the gas-phase transition charges previously calculated (Madjet et al., 2006) and rescaled here to ensure that each pigment carried a Qy dipole moment strength of 4.3 Debye (Reppert et al., 2010; Reppert et al., 2008). Calculated coupling values are displayed in Table 2 and confirm that excitonic interactions are significantly modified between the two structures. In the table, entries above the diagonal correspond to coupling elements in WT Synechocystis, and values below the diagonal correspond to the C. aponinum structure, presumed to be similar to the Red_c mutant; all values are in units of cm–1. The largest difference is the B19/B40 coupling element which decreases in magnitude from –106 cm–1 in the WT structure to only –70 cm–1 in the Red_c mutant. In addition, as illustrated in Figure 5A,B, the transition dipole moment for pigment B40 is rotated (predominantly out of plane) by approximately 40° in the C. aponinum structure relative to WT Synechocystis.

Table 2
Site energies and calculated coupling values amongst chlorophylls B18, B19, and B40 for WT Synechocystis and C. aponinum structures.
B18B19B40
B1814,600–71–20
B19–5914,950–106
B40–10–7014,950
Calculated transition dipole vectors for chlorophylls B18, B19, and B40.

Viewed from above (A) or beside (B) the plane of pigment B19. The structures are aligned relative to the main ring atoms of chlorophylls B18 and B19. Red atoms/dipoles refer to WT Synechocystis, while blue atoms/dipoles refer to the Red_c mutant. (C) Simulated (red curve) Red_c – WT Synechocystis absorption difference spectra compared with the corresponding experimental spectrum (black curve).

To evaluate the impact of these changes on the Qy absorption spectrum, we performed excitonic structure calculations using the TrESP coupling and dipole parameters for each structure using the PigmentHunter app at nanoHUB.org (Safa et al., 2021). Since no quantitative method exists for assigning pigment site energies based on the structural data, we chose the average site energy values for each pigment to achieve reasonable agreement with the experimental absorption difference spectrum (Red_c – WT Synechocystis). The same site energy values (reported on the diagonal of Table 2) were used for both complexes; only the dipole moments and coupling values were altered according to the TrESP values calculated from the C. aponinum and WT Synechocystis structures. Each Qy absorption spectrum was calculated as an average over 1,000,000 iterations with site energies for each pigment sampled randomly from a 300 cm–1 (full width at half-maximum) Gaussian around the respective average value. The final spectrum was convolved with a 10 cm–1 Gaussian (full width at half-maximum) for visualization. As seen in Figure 5C, the calculated spectrum is in excellent qualitative agreement with the experimental data. (The lack of absorption intensity near and above 650 nm in the calculated spectrum is due to the absence of coupling to vibrational modes in our model.) Since the site energies are chosen in these calculations to match experimental data, these results do not, of course, imply that pigment site energies are identical in the WT Synechocystis and Red_c complexes. They do, however, demonstrate that the modified coupling values and dipole orientations reflected in the structural data are sufficient to explain the observed spectroscopic changes.

Theoretical modeling of the Red_d mutant is more difficult than Red_c for two reasons. First, Ca2+ removal may induce changes in protein structure (as evidenced by the modified monomer/trimer ratio noted above), which could modify the site energies and couplings of multiple pigments. Second, Ca2+ removal almost certainly produces significant site-energy shifts for at least the five pigments closest to the Ca2+ binding site (B6, B7, A31, A32, and L3); Chl site energy prediction is, in general, very challenging due to the large number of factors that contribute to environment-induced frequency shifts (e.g., local electrostatics, pigment deformation, electron induction effects, etc.) (Müh and Zouni, 2020; Renger and Müh, 2013; Curutchet and Mennucci, 2017; Lahav et al., 2021). For these reasons, we leave detailed modeling of the Red_d mutant for a future study.

Comparing the local environment of chlorophylls between C. aponinum and WT Synechocystis

Previous theoretical calculations have suggested that the protein electrostatic environments are significant factors in shifting the absorption wavelengths of chlorophylls in addition to strong coupling to neighboring chlorophylls (Saito et al., 2020; Adolphs and Renger, 2006; Fiedor et al., 2008). This led us to examine the structural features that could result in different local electrostatic environments of key chlorophylls between C. aponinum and WT Synechocystis. To find these features, all chlorophyll rings within 4 Å of an amino acid difference between C. aponinum and WT Synechocystis were selected. This selection was then compared to predicted LWC sites resulting in the identification of chlorophylls A32 and B7 (Figure 6A; Jordan et al., 2001). Interestingly, the WT Synechocystis PSI structure reveals a vastly different environment around chlorophyll B7 compared to C. aponinum. In WT Synechocystis, two indole groups of tryptophan residues (I-W20 and L-W65) are located 3.1 Å and 3.9 Å from chlorophyll B7, both along the Qy dipole axis (Figure 6B). However, the structure of C. aponinum reveals a leucine and a phenylalanine (PsaI-L27 and PsaL-F64), respectively, at these positions (Figure 6B). Because the removal of the Ca2+ ion does not fully explain the spectral differences between C. aponinum and WT Synechocystis above 700 nm, the absence of the indole electrons could be responsible for the remaining difference in LWC content between C. aponinum and WT Synechocystis.

Figure 6 with 4 supplements see all
Local protein environment of predicted LWC.

(A) The location of chlorophyll B7 and A32 within each monomer, C. aponinum (green), and WT Synechocystis (gray, LWC in red). (B) The surrounding environment for chlorophyll B7 and A32 with C. aponinum (green) and WT Synechocystis (gray, LWC in red).

Additionally, due to its proximity to the B7-A32 dimer, we think chlorophyll A31 could also affect the LWC in WT Synechocystis. PsaA-F446 in WT Synechocystis is changed to a tryptophan in C. aponinum (PsaA-W455) providing an indole group 3.2 Å from A31 along the extended Qy dipole in C. aponinum. Similarly to B7, the presence of the indole group would allow for electron donation to the chlorophyll, altering the electrostatic environment and potentially shifting this chlorophyll absorbance.

Discussion

The unique environment that C. aponinum was isolated from is exposed to high light and a constant supply of moisture. In deserts where water is scarce, cyanobacteria are mostly active after rainstorms, when there is cloud cover, or around dawn when dew is present, both low-light environments (Harel et al., 2004; Potts, 1994). Under drought conditions, cyanobacteria have adapted to decrease PSI and PSII activity, protecting cells from desiccation (Satoh, 2002; Lawlor, 2002; Potts and Friedmann, 1981; Chaves, 1991). The photoinactivation caused by drought may have prevented any evolutionary pressure toward tolerating high light (Harel et al., 2004; Potts, 1994). This makes C. aponinum a unique candidate to study adaptations in cyanobacteria to survive in high-light conditions.

C. aponinum was isolated from environmental samples of a microbial mat and grown under ~3200 µmol photons m–2s (Bekker, 2004) in the lab, greatly exceeding the maximum amount of solar irradiance it would naturally experience. Strains of C. aponinum were already known to grow in high temperatures, and now it is shown that they grow in high light, demonstrating the innate ability of this organism to survive in extreme environments. When exposed to high light, C. aponinum increases its carotenoid content as well as changing the distribution of excitation energy between PSI and PSII to favor PSI, suggesting that PSI is important for growth under high light.

Purified PSI from C. aponinum is found as a trimer and its subunit composition is identical to WT Synechocystis. Protein sequence alignments identified insertions in PsaB and PsaL, as well as a drastically different C-terminus of PsaL. Absorbance measurements show C. aponinum PSI absorbs less in the far-red than WT Synechocystis PSI, indicating a lower content of LWC in C. aponinum. Gaussian deconvolutions indicate that these LWC contribute to absorbance bands at 704 nm and 711 nm. Surprisingly, the 77 K emission peak is shifted 2 nm red compared to WT Synechocystis. This is a seemingly contradictory result, as C. aponinum contains less LWC, which are the main emitters of the 77 K emission. We suggest that the elimination of some LWC in C. aponinum cause a greater emission from a different either C706 or C714 (Toporik, 2020; Khmelnitskiy et al., 2020; Riley et al., 2007) LWC with a lower energy. This is consistent with the findings that there is less red absorption in C. aponinum together with the red shifted emission at 77 K.

To understand the structural features that could explain these spectroscopic differences, the structure of PSI was determined by cryo-EM. The structure was similar to trimeric PSI structures from cyanobacteria with a few differences. One of which being an insertion in PsaB manifests as a loop on the stromal side of the protein near three chlorophyll molecules, chlorophylls B18/B19/B40.

The Insertion in PsaB causes chlorophyll B40 to alter its orientation in comparison to WT Synechocystis (Figure 3B). To test whether the orientation of this chlorophyll is responsible for differences in the spectroscopic properties, we constructed a chimeric PSI, Red_c, with this insertion in WT Synechocystis. Red_c demonstrates that insertion of the loop results in a blue shift within the Qy transition attributed to the change in chlorophyll B40 position. Although the difference spectrum shows a minimum absorbance of 685 nm (Figure 4D), occupying an energetic position lower than most of the antenna chlorophylls, chlorophyll B40 is not a LWC (in WT Synechocystis) according to the accepted definition.

Mutating the Ca2+ coordinating residue in PsaL to mimic the C. aponinum sequence resulted in a decrease in absorption at 704 nm, indicating that the Ca2+ ion tunes LWC in WT Synechocystis and its absence in C. aponinum probably contributes to the observed loss of LWC absorption. A similar effect of Ca2+ binding was shown previously in the light harvesting complex-1 of purple bacteria (Kimura et al., 2008; Swainsbury et al., 2017). The Ca2+ ion in PsaL of WT Synechocystis is located near known LWCs, chlorophylls B7 and A32 (Khmelnitskiy et al., 2020; Jordan et al., 2001), and our results support its contribution to the low-energy states of this chlorophyll pair (Figure 4D,E). We do not resolve any changes in the 77 K emission between WT Synechocystis and Red_c or Red_d, this is in agreement with previous results showing that red shifting the B7-A32 dimer contributes very little to the maximum emission from the PSI trimer in WT Synechocystis, even at 4 K (Khmelnitskiy et al., 2020). This suggests that the terminal emitter for PSI is a different LWC.

The Red_d mutation does not account for all the changes in LWC absorption seen between C. aponinum and WT Synechocystis. We suggest that the indole groups of tryptophan residues proximal to chlorophylls B7–A32–A31 also contribute to the spectral differences observed between PSI from C. aponinum and WT Synechocystis. The lone pair from the tryptophan indole group could affect the HOMO/LUMO energy levels of these chlorophylls, thus changing the site energies (Saito et al., 2020). In the case of chlorophylls B7–A32–A31 in C. aponinum, the combination of losing PsaI-W20 and PsaL-W65 to uncharged species and gaining an indole lone pair in PsaA-W455 compared to WT Synechocystis would change the local environment along the transition dipole of this chlorophyll aggregate (Figure 6; Saito et al., 2020). Interestingly, similar amino acid variations are also seen in PSI structures isolated from plants, green algae, red algae and diatoms, presumably altering the local environment in a similar way throughout the evolution of photosynthesis in eukaryotic organisms (Figure 6—figure supplement 1 and Figure 6—figure supplements 3 and 4). These differences around LWC chlorophylls probably account for additional differences in the absorption spectra between C. aponinum and WT Synechocystis. To confirm this and to quantify the effect of each tryptophan residue on these chlorophylls, additional studies are needed.

The orientation of the PsaL C-terminus in C. aponinum differs from other structures of trimeric PSI from cyanobacteria (but is similar to Synechococcus; Cao et al., 2020). As previously reported, the deletion of PsaL or the addition of a terminal histidine to the C-terminus PsaL in WT Synechocystis leads to the complete dissociation of trimeric PSI, underlining the importance and sensitivity of PsaL to trimerization (Malavath et al., 2018; Netzer-El et al., 2018). In C. aponinum, the C-terminus is shifted away from the adjacent monomer and there is no Ca2+ ion observed in the density of the structure (Figure 4). High resolution structures of PSI showed that the C-terminus of PsaL interacts with an adjacent monomer through a Ca2+ ion coordinated by an aspartic acid (Netzer-El et al., 2018; Mazor et al., 2015). We now show that Ca2+ contributes but is not essential for trimer assembly. Unexpectedly, we also show that Ca2+ binding also tunes and red-shifts LWC in PSI. This agrees with the fact that in monomeric PSI from eukaryotes, a Ca2+ ion is not seen (Mazor et al., 2015), together with the lower LWC content of eukaryotic core complexes (Croce et al., 1998). In addition, upon monomerization, the cyanobacterial PSI loses LWC absorption, as monomerization is also accompanied by the loss of Ca2+, our findings suggest that Ca2+ loss contributes to these differences (Gobets et al., 2001).

How prevalent is Ca2+ binding in PSI? Comparing prokaryotic and eukaryotic PsaL shows differences in conservation in Ca2+ coordinating residue in cyanobacteria. This residue exhibits high conservation as an asparagine across eukaryotes (Figure 3—figure supplement 6), while an alignment of 680 cyanobacterial PsaL sequences revealed that despite the high conservation around the Ca2+ coordinating residue in prokaryotes, this specific residue is variable in cyanobacteria (Figure 3—figure supplement 5). Residues containing a negative charge are seen at this location in ~50% of species, including WT Synechocystis and T. elongatus, which likely corresponds to coordination of Ca2+. In Synechococcus however, the coordinating residue is an asparagine. The recently solved PSI structure from Synechococcus lacks a Ca2+ ion, giving further support to the importance of this residue in coordinating a calcium ion (Cao et al., 2020). While the presence of Ca2+ aids in the stabilization of trimerization for some species, the structure of trimeric PSI from C. aponinum and Synechococcus shows it is not required. It is interesting to note that the majority of differences in sequence between C. aponinum and WT Synechocystis occur in the intermonomer space. These differences probably stabilize the monomer–monomer interaction in C. aponinum despite the absence of the PsaL Ca2+ ion.

The mechanism underlying the contribution of PSI to photoprotection in C. aponinum is not presently clear; however, PSI is potentially the most potent quencher of excitation energy in the cell. The optical properties of PSI from C. aponinum are similar to the eukaryotic core PSI family, with a low number of LWC. The contribution of PSI to cellular resistance to high light can stem from its interaction with other components of the light reactions, such as PSII, PBS, or the stress-induced antenna, IsiA, as has been suggested in the past (Tiwari, 2016). The role played by LWC in these interactions is not clear to a large degree because specific mutations or precise information on the architecture of LWC in PSI is scarce. Previously, it was hypothesized that LWCs can serve several functions within the core PSI antenna. Light harvesting in the far-red region is one clear suggestion, but it was also shown that low energy states associated with LWC are quenched by an oxidized P700+ and that this varies between cyanobacterial species (Schlodder et al., 2005; Herascu et al., 2016; Schlodder et al., 2011). This shows that energy transfer in the core PSI antenna varies depending on the oxidation state of P700 and this can be important during high-light conditions when P700 is in its oxidized state. A specific LWC in PSI, in close proximity to the ETC (Schlodder et al., 2005; Herascu et al., 2016; Schlodder et al., 2011), is responsible for this effect and can play a role in photoprotection. In this scenario, the optical adaptations that are seen in C. aponinum facilitate energy transfer through this LWC, this is consistence with our 77 K emission results, which show red shifted emission from C. aponinum compared to WT Synechocystis, probably due to the contribution from this site. Overall, it is necessary to develop an understanding of individual LWC as specific sites (these sites probably include more than one chlorophyll molecule) that carry out specific functions. An essential step on this route is identifying specific LWC and factors that tune them in PSI, as the present study does.

The environment that C. aponinum was isolated from provides ideal conditions for using P700 as a photoprotective mechanism. PSI-specific photodamage is known to be induced by fluctuating light and cold temperatures (Sonoike, 2010; Terashima et al., 1994; Sonoike et al., 1997); however, the environment C. aponinum was isolated from rarely experiences these conditions. Constant light and temperature may allow P700 to withstand more irradiance without causing photoinhibition.

PSI displays high conservation in sequence and structure across domains despite the vastly different environments occupied by photosynthetic organisms. The results of this work show that small structural variations can have large effects, highlighting the sensitivity of pigments to the local electronic environment, and potentially giving rise to physiological advantages (Wientjes et al., 2012; Wientjes et al., 2011). The structural effects on spectroscopic properties of PSI observed in this work lay the foundations for intelligently designing photosynthetic organisms with absorption spectra tuned to specific light environments. The structure and biochemical characterization of trimeric PSI from C. aponinum advances the fundamental understanding of the photosynthetic machinery in organisms that can survive extreme light conditions and reinforces the need to study extremophiles and their adaptations to fully understand the photosynthetic process.

Materials and methods

Selection conditions

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Crude biofilm samples were placed in 50 ml of BG-11 media in Erlenmeyer flasks and exposed to >3000 µmol photons m–2s (Bekker, 2004). Samples were not stirred. Once noticeable growth had occurred, about 2 weeks, samples were agitated by aggressively swirling growth flasks. One milliliter was then plated onto a BG-11 agar plate and allowed to grow. Single colonies were picked and continually streaked to achieve a pure culture of the photosynthetic organism.

16S rRNA sequencing

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A sterile culture of C. aponinum was grown and harvested mid log phase growth. Genomic DNA was extracted by a modified phenol chloroform extraction as previously described (Adolphs and Renger, 2006). Using primers designed by Li et al., 2014, the 16 S rRNA gene was amplified and sent for sanger sequencing.

Genomic DNA sequencing

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Illumina compatible Genomic DNA libraries were generated on the Apollo 384 liquid handler using KAPA Biosystem’s LTP library preparation kit (KK8232). DNA was sheared to approximately 600 bp fragments using a Covaris M220 ultrasonicator, end repaired, and A-tailed as described in the Kapa protocol. Illumina-compatible adapters with unique indexes (IDT #00989130v2) were ligated on each sample individually. The adapter ligated molecules were cleaned using Kapa pure beads (Kapa Biosciences, KK8002) and amplified with Kapa’s HIFI enzyme (KK2502). Each library was then analyzed for fragment size on an Agilent’s Tapestation and quantified by qPCR (KAPA Library Quantification Kit, KK4835) on Thermo Fisher Scientific’s Quantstudio 5. Libraries were then multiplexed and sequenced on 2 × 250 flow cell on the MiSeq platform (Illumina) at the ASU’s Genomics Core facility.

Genomic analysis

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The raw Illumina MiSeq 2 × 250 bp reads (14,485,288 pairs of reads) were quality checked using FastQC v0.10.1, followed by adapter trimming and quality clipping by Trimmomatic 0.35. Any reads shorter than 150 bp were dropped. Any reads with start, end, or the average quality within 4 bp window falling below quality scores 18 were trimmed. A clean 12,273,912 read pairs survived for further insert size estimation. Kmer analysis was ran by Jellyfish 2.2.4 over both entire 14,485,288 read pairs and clean 12,273,912 read pairs for genome size estimation. Cleans reads were aligned to C. aponinum PCC 10605 (cyanobacteria) reference genome (https://www.ncbi.nlm.nih.gov/assembly/GCF_000317675.1/) by bwa mem 0.7.15 for insert size estimation. Spades 3.7.1 with mismatch corrector mode was applied for whole-genome assembly with kmer size 21,33,55,77,99,127. Best whole genome assembly with kmer size 127 was evaluated by comparing to C. aponinum PCC 10605 reference genome by Quast 4.5 (http://bioinf.spbau.ru/quast). When sorting contigs from largest to smallest, first 80 contigs with minimum length 1000 bp were extracted. CAR, a novel reference-based contig assembly and scaffolding tool (http://genome.cs.nthu.edu.tw/CAR/), was applied on the 80 contigs for scaffolds. In order to improve assembly, SSPACE was applied to scaffold pre-assembled contigs using NGS paired-read data. Eighty contigs were kept in final 55 scaffolds. Quast was used to evaluate assembly with 55 scaffolds. BUSCO, a tool for assessing genome assembly and annotation completeness with benchmarking universal single copy orthologs (http://busco.ezlab.org/), indicated 97.3% genome completeness with 812 complete BUSCOs out of 834 total BUSCOs defined in cyanobacteria database. Total of 989 genes were predicted de novo by Glimmer (Gene Locator and Interpolated Markov Model ER, https://ccb.jhu.edu/software/glimmer/). Additional genome annotation was performed by protein homology-based tblastn (blast +2.3.0) approach using protein sequences of C. aponinum PCC 10605 reference genome. Three thousaand three hundred and thirty-three hits were identified. Genomic data was deposited to NCBI (NCBI:txid2676140).

Culture conditions

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C. aponinum used for the structural studies was cultured in BG11 medium supplemented with 6 µg/ml ferric ammonium citrate under continuous white light (∼40 µmol photons m–2s–1) in 30°C.

Growth tests

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C. aponinum and Synechocystis sp. PCC 6803 cells were cultured in BG11 liquid medium supplemented with 6 µg/ml ferric ammonium citrate under continuous white light (∼40 µE) in 30°C. The optical density was adjusted to 5 at 730 nm for Synechocystis and three for C. aponinum. Each culture was diluted ×5, ×25, ×125, ×625, ×3125, ×15,625 and grew on BG11 plates in different light intensities to determine cell viability.

Thylakoid preparation

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Cells were harvested during log phase growth by centrifugation at 12,000 rpm for 3 min at room temperature. Cells were washed in STN1 buffer (30 mM Tricine–NaOH pH 8, 15 mM NaCl, 0.4 M sucrose) and pelleted again to remove any excess growth media, then lysed in a cooled Constant Cells Disruptive Systems French press for three cycles at 30,000 psi. The lysate was cleared of cell debris by centrifuging 12,000 rpm for 5 min in a F20−12 × 50 LEX rotor (Thermo Scientific). Membranes present in supernatant were then pelleted by ultracentrifugation (Ti70 rotor) for 2 hr at 45,000 rpm and 4°C. Membranes were then resuspended in STN1 with 150 mM NaCl and allowed to incubate on ice for 15 min before ultracentrifugation (Ti70 rotor) for 2 hr at 45,000 rpm and 4°C. The pellet was then resuspended in 15 ml of STN1 and stored at –80°C.

PSI purification n-dodecyl β-maltoside (DDM, Glycon) was added to the membrane stock to achieve a ratio of 15:1 DDM-to-chlorophyll ratio, and the samples were manually mixed a few times then allowed to incubate for 30 min on ice. Membranes were centrifuged for 30 min at 45,000 rpm at 4°C to remove any insoluble material. The supernatant was then loaded onto a diethylaminoethyl column (toyopearl DEAE 650 C) and eluted with a linear NaCl gradient (15–350 mM) in 30 mM Tricine–NaOH pH 8, 0.2% DDM. The dark green peak corresponding to the PSI trimer was collected and precipitated with 8% PEG3350 (Hampton Research) and 150 mM NaCl. This was then centrifuged at 5000 rpm for 5 min at 4°C and the supernatant discarded. The pellet was resuspended in 30 mM Tricine–NaOH pH 8, 15 mM NaCl, 0.1% DDM and loaded onto a 10–30% sucrose gradient (30 mM Tricine–NaOH pH 8, 75 mM NaCl, 0.05% DDM). This was centrifuged (Beckman SW40 rotor) for 16 hr at 36,000 rpm. The dark green band was collected and used for subsequent experiments.

Absorption and fluorescence spectroscopy

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Absorption spectra were recorded on a Cary 4,000 UV–Vis spectrophotometer (Agilent Technologies). Fluorescence spectra were recorded on a Fluoromax-4 spectrofluorometer (HORIBA Jobin-Yvon). The slit width was set to 5 nm on both the entrance and exit monochromators for room temperature measurements. For 77 K measurements, slit width of 5 nm and 3 nm were used for the entrance and exit monochromators, respectively. Samples were diluted to an optical density of 1 and 0.1 at 680 nm for absorption and fluorescence measurements respectively, using buffer containing 30 mM Tricine–NaOH pH 8, 15 mM NaCl, and 0.05% β-DDM. The resulting spectra were normalized to the area of the chlorophyll Q bands between 550 and 775 nm. Whole-cell measurements were performed using an integrating diffuse reflectance sphere (DRA 900) to correct for scattering by the cells. For 77 K fluorescence measurements, samples were adjusted to an OD680 of 0.1 in a buffer of 50% glycerol 30 mM tricine pH 8.0, 15 mM NaCl, and 0.02% β-DDM. An Oxford instruments Cryostat was used to cool the sample to 77 K (cells were plunged into liquid nitrogen and measured immersed in liquid nitrogen). Figures were prepared using OriginPro (OriginLab).

Sample preparation for single particle cryo-EM

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The PSI band from the sucrose gradient was collected, NaCl was added to a final concentration of 150 mM and the complex was precipitated using 9% PEG3350. After centrifugation (5000 rpm, 5 min in an Eppendorf tabletop), the green precipitate was resuspended in buffer (30 mM Tricine–NaOH pH 8, 150 mM NaCl, and 0.02% DDM), and any undissolved material was removed by repeating the centrifugation step (14,000 rpm, 5 min). The chlorophyll concentration in the soluble material was adjusted to 1.2 mg/ml using the above buffer. Three microliters of the PSI complex was added to holey carbon grids (C-flat 1.2/1.3 Cu 400 mesh grids [Protochips, Raleigh, NC]) after soaking the grids in buffer. The sample was vitrified by flash plunging the grid into liquid ethane using manual plunger with blotting time of 6 s. The grids were stored in liquid nitrogen until data collection.

Data acquisition

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The cryo-EM specimens were imaged on a Titan Krios transmission electron microscope (Thermo Fisher - FEI, Hillsboro, OR). The electron images were recorded using a K2 Summit direct electron detect camera (Gatan, Pleasanton, CA) in super-resolution counting mode. Image collection was automated with SerialEM (Fiedor et al., 2008) utilizing scripting of stage shifts between hole exposures. The defocus was set to vary between 0.8 and 2.6 μm, corresponding to a super-resolution pixel size of 0.525 Å at the specimen level. The counting rate was adjusted to 7.614 e2 s. Total exposure time was 8 s accumulating to a dose of 61 e2.

Data processing

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A flow chart describing data handling is shown in Figure 3—figure supplement 1. MotionCor2 (Harel et al., 2004) was used to register the translation of each sub-frame, and the generated averages were Fourier-cropped to 1.5 times and dose-weighted (Potts, 1994). CTF parameters for each movie were determined using CTFFIND4 (Satoh, 2002). Relion was then used for the subsequent data processing (Lawlor, 2002). A set of manually picked particles (~1000) from an early data set was subjected to a few rounds of unsupervised 2D classification and then used to generate an initial 3D volume. This volume was then used on a later data set as the template for the automated particle picking procedure as implemented in Relion which yielded 256,410 particles. This particle set was subjected to several rounds of unsupervised 2D classification (Relion) resulting in a set of 132,677 particles. This particle set was then subjected to a focused 3D classification on the PsaL subunit. This procedure yielded eight classes with one dominate class. This class was selected yielding 73,984 particles. 3D refinement (C3 symmetry) using this set yielded a volume at a resolution of 3.71 Å. CTF refinement was used (Potts and Friedmann, 1981), followed by 3D refinement (C3 symmetry), yielding the final resolution of 3.79 Å. Particle polishing was implemented followed by 3D refinement (C3 symmetry) yielding a resolution of 3.75 Å. This was followed by three cycles of CTF refinement and 3D refinement (C3 symmetry), yielding a resolution of 2.88 Å. The detergent signal was subtracted from the PSI trimer, followed by a C3 expansion of the particles, yielding a total particle count of 221,952. This particle set was used for a 3D refinement (C1 symmetry) resulting in a map of 3.0 Å. Multibody refinement was used to generate the final map, of 2.74 Å according to the gold standard FSC criteria (Chitnis and Chitnis, 1993). The final map was sharpened using the postprocessing procedure in Relion, and the b-factor used for sharpening was –72.48. Local resolution was estimated using ResMap.

Model building and refinement

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The initial PSI model was taken from the 2.5 Å x-ray structure of the trimeric PSI from Synechocystis (PDBID: 5OY0) (Toporik, 2020). The model was docked into the map using PHENIX (Chaves, 1991). The final model was refined against the cryo-EM density map using phenix.real_space_refine (Riley et al., 2007; Kimura et al., 2008). Final model statistics are shown in Table 1, and side chain resolvability was calculated using MapQ (Swainsbury et al., 2017; Supplementary file 1a). PyMOL (Jordan et al., 2001) and UCSF Chimera (Mazor et al., 2015) were used to generate all images.

Strain construction and growth

Red_c construction

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A plasmid (p60) containing the entire PsaAB operon marked with Kanamycin resistance gene was previously constructed (Tikkanen et al., 2014). The Red_c mutation was constructed into p60 by adding the loop sequence observed in C. aponinum using the p60_red_c_Forward_insert and P60_red_c_Reverse_insert primers and p60 as a template. The two fragments were assembled using the NEBuilder HiFi DNA Assembly Master Mix. All plasmids were sequenced before being used to transform Synechocystis sp. PCC6803 according to standard protocols. Complete and correct replacement of PsaB was verified by PCR and sequencing.

Red_d construction

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A plasmid (PsaLPsaI) containing both the PsaL and PsaI genes marked with a chloramphenicol resistance gene 0.1 kb upstream to PsaI was constructed from four PCR amplified fragments (Tikkanen et al., 2014). PsaL and PsaI fragments, with up an 0.8 kb upstream fragment, were amplified using the PsaL_R and PsaL_F primer pairs from the Synechocystis sp. PCC6803 genome. The chloramphenicol resistance gene was amplified from previously constructed plasmids in our lab, using Cm_F and Cm_R primer pairs. A 0.8 kb downstream fragment from PsaI was amplified using Down_F and Down_R primer pairs. A pJET backbone was amplified using the primer pairs Backbone_F and Backbone_R. The four fragments were assembled using the NEBuilder HiFi DNA Assembly Master Mix. The Red_d mutation was constructed into PsaLPsaI by creating a point mutation observed in C. aponinum using the Ca_D2L_F/PsaL_R and Ca_D2L_R/PsaL_F primer pairs and PsaLPsaI as a template. The two fragments were assembled using the NEBuilder HiFi DNA Assembly Master Mix. All plasmids were sequenced before being used to transform Synechocystis sp. PCC6803 according to standard protocols. Complete and correct replacement of all aspects was verified by PCR and sequencing. All primer sequences are listed in Supplementary file 1b.

Excitonic structure calculations

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Electronic transition dipoles and coupling elements amongst Chls B18, B19, and B40 for WT Synechocystis were calculated using the TrESP method (Moro et al., 2007). The PDB: 6UZV model of trimeric PSI was used as WT Synechocystis, and the C. aponinum PSI structure (PDB:6VPV) was used to approximate the Red_c structure. For the TrESP calculation, we used the gas-phase transition charges previously calculated (Moro et al., 2007) and rescaled here to ensure that each pigment carried a Qy dipole moment strength of 4.3 Debye (Malavath et al., 2018; Mazor et al., 2013). Excitonic structure calculations using the TrESP coupling and dipole parameters for each structure using the PigmentHunter app at nanoHUB.org (Winckelmann et al., 2015). Site energy values for each pigment were varied to achieve reasonable agreement with the experimental absorption difference spectrum (Red_c - WT Synechocystis). The same site energy values (reported on the diagonal of Table 1) were used for both complexes; only the dipole moments and coupling values were altered according to the TrESP values calculated from the C. aponinum and WT Synechocystis structures. Each Qy absorption spectrum was calculated as described previously (Mazor et al., 2013) as an average over 1,000,000 iterations with site energies for each pigment sampled randomly from a 300 cm–1 (full width at half-maximum) Gaussian around the respective average value. The final spectrum was convolved with a 10 cm–1 Gaussian (full width at half-maximum) for visualization; no vibrational or phonon-sideband structure was included.

Data availability

The final model (PDBID 6VPV) and map (EMD-21320) were deposited in the Protein Databank and Electron Microscopy Database, respectively. C. aponinum genomic DNA was deposited in NCBI genebank under NCBI:txid2676140.

The following data sets were generated
    1. Dobson Z
    2. Vaughn N
    3. Vaughn M
    4. Fromme P
    5. Mazor Y
    (2019) NCBI BioProject
    ID PRJNA580528. Cyanobacterium aponinum 0216, whole genome shotgun sequencing project.
    1. Dobson Z
    2. Vaughn N
    3. Vaughn M
    4. Fromme P
    5. Mazor Y
    (2021) RCSB Protein Data Bank
    ID 6VPV. Trimeric Photosystem I from the High-Light Tolerant Cyanobacteria Cyanobacterium Aponinum.
    1. Dobson Z
    2. Vaughn N
    3. Vaughn M
    4. Fromme P
    5. Mazor Y
    (2021) EMD
    ID 21320. Trimeric Photosystem I from the High-Light Tolerant Cyanobacteria Cyanobacterium Aponinum.

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Decision letter

  1. David M Kramer
    Reviewing Editor; Michigan State University, United States
  2. Olga Boudker
    Senior Editor; Weill Cornell Medicine, United States
  3. Geoffry A Davis
    Reviewer; Michigan State University, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

The work presents high resolution structure and functional measurements of a photosystem I complex from a high light tolerant species of cyanobacteria. The results give important clues about the adaptations of photosynthetic systems that allow them to function under such harsh environmental conditions. The revised version of the paper clears up most of the questions presented by the reviewers and adds data pointing to a possible new insights on the function of long wavelength chlorophylls in photosystem I light harvesting complexes.

Decision letter after peer review:

Thank you for submitting your article "The Structure of Photosystem I from a High-Light Tolerant Cyanobacteria" for consideration by eLife. Your article has been reviewed by 4 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Olga Boudker as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Geoffry A. Davis (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.

All of the reviewers felt that the structural work was both interesting and well done. Also, there was considerable enthusiasm for the approach of comparing structures from diverse species that might reveal interesting adaptations to different environments.

On the other hand, all of the reviewers had substantial questions about how the identified structural variations relate to the biophysical/spectroscopic measurements or whether these differences were relevant to any physiological changes.

It should be possible to address the spectroscopy issues by reanalyzing the data along the lines of that proposed by reviewer 4.

It will probably be more difficult to deal with the issue of physiological relevance. As stated by several of the reviewers, given the many changes that are likely to have occurred between the two species, it is difficult to know if the specific structural variations identified in the paper are important, or indeed, if they have any physiological consequences by themselves. It may be, as pointed out by one reviewer, that the big story is not that there are vast differences, but that there are so few differences.

It was also pointed out that there is an inherent difficulty with comparing light acclimation between two species, especially those that are quite distantly related and grow under very different light regimes.

From these comments, it would seem that directly addressing the question of physiological relevance will require some additional data, though it may be possible to approach to achieve this by extending the bioinformatics approaches rather than additional experimentation.

A second approach would be to better focus the discussion to aspects of the structure that are more directly supported by the data itself, and discuss the potential implications more hypothetically. It would be great if this could be achieved while retaining the intended high impact.

Reviewer #1:

The paper can be improved by: (1) better describing the fitting methods and their bases; (2) describing alternative models; (3) providing more evidence and arguments that the structural differences that we observed are relevant to the selection pressure or physiological differences required for growth in the different light regimes; and (4) acknowledging and discussing that the light environments are relative, e.g. that using Synechococcus, which is generally found growing at very low light, may not be an adequate (sole) comparison for a high light species.

1) "…had a constant drip of fresh water and exposed to extreme light conditions." How is 1300 uE an extreme light? Are all terrestrial vascular plants now extremophiles?

2) In this regard, choosing Synechocystis as a comparison is a bit of a "straw man" and it would be more appropriate to compare to a wider range of species/strains. In other words, wouldn't it be more accurate to say that Synechocystis is adapted to LOW light, rather than C. aponiunum adapted to high? If this is true, then the story may be about selection pressure for low light rather than photoprotectiom.

3) The work shows that C. aponium can grow at higher light than Synechocystis. However, it was not possible to attribute this difference to PSI properties. This would be a pretty high bar to set, of course, but can the authors make the case that the differences they do observe are relevant?

4) The experiments exposed plates to 3000 µmol photons m-2 s-1, which is higher than that these cells experience in the real world. What is the argument that 3000 uE is physiologically relevant? Is it not the case that this selects for something not relevant to nature?

5) The 77K emission spectrum is not a measure of PSI:PSII content, but on the distribution of excitation energy. Further, the interpretation of these spectra is dependent on the specific excitation wavelength. For instance, if cyan light will excite the PBS, which can dump energy differently into PSI and PSII, compared to, e.g. blue light. The emission spectrum is also impacted by cell density and other factors. None of these issues is acknowledged in the text, nor is that excitation wavelength described in the Methods, as is clear from the scant description given in the text: "An Oxford instruments Cryostat was used to cool 26 the sample to 77 K (cells were plunged into liquid nitrogen and measured immersed in liquid nitrogen)." which does not give any indication of the instrument, excitation, cell concentration, etc., though Figure 2 mentions that particular data was collected with excitation at 440 nm. What is the resolution of the spectrometer?

6) Figure 3. The spectra are interpreted as apparent shifts based on subtracting pairs of spectra normalized to their peaks. This seems to be based on a subjective interpretation and other interpretations are possible. Changing the normalization point could result in an entirely different interpretation. For instance, normalizing to a point on the red side of the peak could give the appearance of an increase in absorbance or emission on the blue edge of the spectrum (rather than a shift). The deconvolution of the absorbance spectra using three Gaussian components as in Figure 2D is likewise based on an assumption or interpretation that there are only these components.

7) The text needs to be more circumspect and/or directly address this as a possibility, since such alternative normalization could indicate a quite different story. For example, the text reads: "The difference spectrum of C. aponinumSynechocystis shows a strong negative peak at 701 nm revealing that C. aponinum PSI contains less LWC 150 than Synechocystis (Figure 2D)." It suggests this, but there are other interpretations. This issue is more evident further in the manuscript, in Figure 4, a different method of normalization was used, as stated in the following: "The absorbance spectra were normalized to the area between 650 nm to 775 nm and the spectrum of Wild Type Synechocystis was 238 subtracted from Red_c (Figure 4C and 4D)." As argued above, changing the normalization method can give rise to different interpretations, so some real justification must be presented for using a particular approach, and an even stronger argument needs to be made when changing the normalization between samples. None is given.

8) Figure 4 D compares isolated complexes from two different species and the differences spectra can be interpreted as a series of narrowing and shifting of spectra, but all of which are likely to be dependent on the method of normalization, which is not adequately described. Moever, the method used to make the differences spectra were not consistently applied, e.g. "[t]he absorbance spectra were normalized to the area between 650 nm to 775 nm and the spectrum of Wild Type Synechocystis was subtracted from Red_c (Figure 4C and 4D)." What were the criteria for using the different procedures and how was bias avoided?

9) The shifts in absorbance and fluorescence emission are quite small, especially realizing that the bands tend to sharpen at 77K. At RT, there may be essentially no functionally-relevant differences in the spectra. How can this be addressed?

10) Can the text be expanded to further discuss the very large apparent difference in PsaF?

11) Cyanobacterium aponinum (C. 78 aponinum). should be both Ital. Is this an official name?

Reviewer #2:

The manuscript by Dobson et al., describes photosystem I (PSI) of a high-light tolerant cyanobacterium, Cyanobacterium aponinum 0216, isolated from an environmental sample at both the sequence and structural level and attempts to test how differences in the C. aponinum (PSI) sequences may result in the structural and spectroscopic differences characterized. The authors solved the Cryo-EM structure of trimeric PSI of C. aponinum and noted two major differences with the model cyanobacterium Synechocystis PCC 6803 trimeric PSI, namely insertions in the PsaB and PsaL sequences, potentially resulting in a shifted position of chlorophyll B40 and loss of calcium ion coordination by PsaL. With the construction of a Synechocystis mutant containing the C. aponinum PsaB sequence, the authors attempted to clarify if the structural changes around chlorophyll B40 could explain the loss of long wavelength chlorophylls and absorbance differences seen in C. aponinum. The work carried out by the authors is useful in the understanding of structural and sequence variability in PSI for understanding chlorophyll energy tuning as well as the variability in amino acid sequences throughout cyanobacterial PSI subunits and how they may be adapted for specific light conditions.

The spectral characterization of C. aponinum PSI relative to Synechocystis provides the crux of the rationale for the work done in the manuscript, however the discussion and interpretation by the authors is limited to primarily the baseline differences between the spectra. As C. aponinum is presented as a cyanobacterium isolated via propagation of environmental samples at 3,500 umol photons m-2s-1, discussion of the physiological significance of the PSI differences is minimal.

The C. aponinum PSI Cryo-EM is well discussed and presented by the authors to highlight the structural features that may be involved in shifted the absorbance spectrum and the loss of far-red wavelength chlorophylls. Calculations of the excitonic interactions based on the solved structure and the shifted position of chlorophyll B40 (relative to the Synechocystis position) highlighted the utility of solving the PSI structure rather than relying on sequence differences modeled onto reference structures.

Similarly beneficial to the objective of trying to understand how C. aponinum differences impact the function of PSI is the introduction of C. aponium PsaB sequence into Synechocystis and analysis of the resulting spectrum and far-red chlorophyll presence. As the authors found a large shift in the position of chlorophyll B40 in C. aponinum PSI, potentially caused by amino acid changes in the PsaB gene, the introduction and testing in Synechocystis is a reasonable first method to understand the impact. The discussion of this chimeric PSI (Red_c) should be addressed better to make clear what is being referenced, the Red_c mutant versus C. aponinum itself, as discussion of the differences seems to misstate at times what is actually being compared.

While the manuscript presents data in a clear, concise format, there are some outstanding issues that should be addressed to improve the understanding of the results and significance of the findings.

The selection strategy was using 3,000 umol photons m-2s-1 light to select for high light tolerant strains. As the sample was from the Sonoran desert, it would be useful to clarify if/what temperature was used for the selection as well, as the growth temperature of 30C is not particularly high for many cyanobacteria.

The figures detailing the characterization of PSI between different samples could benefit from a clearer delineation between the samples shown. For most samples, the spectra are nearly identical, and the differences, or the fact that two samples are shown is nearly impossible to determine (figures 2C, 4C).

Similarly, it would be useful to discuss the other differences in PSI subunit composition. From the analysis of isolated PSI (figure 2), the authors discuss the migration differences between PsaL. However, there are clear differences between the migration of other subunits as well. Whether the primary sequences are markedly different should be mentioned. The authors also state that the expected number of bands is observed for C. aponinum (line 137). It is unclear from the figure if that is the case, as five distinct bands are visible in the Synechocystis sample but only four in C. aponinum. If PsaF migrates almost together with PsaD that should be clarified.

Lines 176-177 should be clarify which chlorophylls are discussed relative to the changes in B40 coupling.

The description of the chimeric Synechocystis mutant needs considerable clarification. Figure 4B: what is Blue6803? The legend for Figure 2G does not seem to be correct. It should also be clarified that the amino acids present in C. aponinum are not mutations (ex: line 253, 399), they are the wild-type sequences and are different relative to Synechocystis. The discussion within the section of the excitonic coupling calculations should also be addressed to clarify the distinction between the C. aponinum structure and the Synechocycsist Red_C mutant.

As the PsaB changes around chlorophyll B40 did not explain either the absorbance spectrum changes or the long-wavelength chlorophyll differences, the authors also discuss amino acid differences in the vicinity between the two species. It would be beneficial to do a similar comparison between these positions and amino acid composition between difference cyanobacterial species as was done for the calcium binding residue position. This may also provide more information to discuss these changes rather than a preliminary list of the differences between the two structures.

It is interesting that this mutant did not reproduce either the C. aponinum absorbance spectrum or the loss of far-red wavelength chlorophylls, again highlighting the utility of doing both the sequence and structure comparisons. While it would increase the impact of this work to also include mutation comparisons introducing the C. aponinum PsaL into Synechocystis, as well as the double C. aponinum PsaB/PsaL chimera, that work may not be feasible under current circumstances. However, because the Red_c mutant did not explain the differences between absorbance spectra, a more detailed discussion of how the various changes may be involved in those changes should be carried out.

Based on the presentation in the manuscript, it is unclear why full genome sequencing was performed, as PSI genes could have presumably be sequenced via PCR methods. The authors do themselves a disservice at having done this analysis and essentially not addressing the results. It would be interesting to know if there are any genes within the genome beside those highlighted for PSI that may be of interest in understanding the high-light capacity of this cyanobacterium.

The methods section should be tidied up. Many explanations are unnecessary and likely could be better described via referencing previous works.

In general the manuscript requires further proof-reading throughout to rectify typos, italicization issues, unit labels, etc.

Reviewer #3:

In this work, Dobson Z et al., target the PSI complex of a high-light adaptive cyanobacterium C. aponinum 0216. The cyanobacterial PSI complexes carry a number of long wavelength chlorophylls (LWCs) in their core antennae. The LWCs were suggested to increase the absorption cross section of PSI, and to serve in photoprotective function. In this work, they suggest that the C. aponinum PSI contains less LWC compared with Synechocystis PSI, based on their spectroscopic results. They further determine the 2.7 Å resolution cryo-EM structure of PSI complex from the high-light tolerant strain to investigate if it possesses some special structural features. The structure shows that one chlorophyll, B40, shifts its orientation which is likely induced by an inserted loop in PsaB of C. aponinum PSI. Therefore, they assume that the altered loop region and chlorophyll B40 are responsible for the spectral property of less LWC in C. aponinum PSI. To confirm this assumption, they construct a mutant form of Synechocystis strain containing the sequence of the PsaB loop from C. aponinum. However, spectra of the mutant PSI indicate that chlorophyll B40 is not responsible for the difference of LWC. Then they analyze the local environment of chlorophylls in C. aponinum and Synechocystis PSI, and suggest that the local environment of chlorophylls B7-A32-A31 accounts for some of the differences in the absorption spectra.

Overall, the structural data in this work is solid. Generating a mutant strain possessing the different peptide sequence to induce different chlorophyll arrangement based on the structural data is useful in analyzing the spectral property and potential function of individual chlorophyll molecules. It was widely applied in the previous studies to generate a mutant strain possessing different pigment arrangement and to analyze the spectral difference of photosynthetic complexes from the mutant strain, now with the high-resolution structural information, the interpretation of the biochemical and spectral data are more reliable, as the structure provides details regarding the position, orientation and environment of each chlorophyll molecules.

In this work, although the red LWCs or the factors affecting the spectral features of this high-light tolerant cyanobacterial PSI are not identified, they do have some suggestions/ideas worthy of further exploration based on their present results. Therefore the findings about C. aponinum PSI are beneficial to the photosynthesis research field, including the artificial photosynthesis research. The conclusions and suggestions put forward by the authors are largely supported by their data, but some aspects of data analysis need to be clarified and amended.

The interpretation of spectral analysis of C. aponinum PSI is questionable. They conclude that C. aponinum PSI contains less LWC than Synechocystis PSI based on the differences of absorption spectra between the two PSIs shown in Figures 2C and 2D. However, the difference is very small, with the amplitude of δ-absorbance lower than 0.04, which seems within the error range, while the absorbance of the PSI complex is around 1.5. On the contrary, the 2-nm red-shift of 77K fluorescence emission of C. aponinum PSI is quite evident (Figure 2F). Therefore it is possible that C. aponinum PSI contains more, but not less, LWC.

They construct a mutant form of Synechocystis strain containing the sequence of the PsaB loop from C. aponinum, and find that the 77K fluorescence emission of the mutant PSI is the same with Synechocystis PSI. Thus conclude that chlorophyll B40 is not responsible for the different absorption of LWC. Although they confirm the complete and correct replacement of PsaB in the mutant strain, they do not validate that B40 orientates the same with that in C. aponinum PSI. Since the loop regions are usually highly flexible and may adopt quite different conformation even with the same primary sequence, it is possible that B40 is indeed responsible for the different absorption of LWC in C. aponinum, but B40 in the mutant does not change its orientation, i.e. adopts the same orientation as that in Synechocystis PSI.

C. aponinum is a high-light tolerant cyanobacterium, and it does not grow well under low light conditions as shown in Figure 1C. However, in this study, they culture the cells at low light (∼40 μE), which may lead to some changes of the photosynthetic complexes, therefore the structural features of PSI important for C. aponinum to survive in the high light environment cannot be identified.This reviewer understands that the difference of absorption spectra between PSI from C. aponinum and Synechocystis is very small, as only a few chlorophylls are changed. However, if they would like to use the spectral data, they need to prove that the differences of absorption spectra are indeed due to the different spectral property of PSI, but not the standard error. The spectra should be repeated at least three times for each PSI complexes and the differences of the repeated spectra should be shown in the paper.

It's better to determine the structure of the PSI complex from the constructed mutant to show the orientation of chlorophyll B40. If it is the same as that in C. aponinum PSI, this result will further strengthen their conclusion. If it is not, then chlorophyll B40 should be further investigated.

It will be more convincing if they purify and determine the structure of the PSI complex from C. aponinum cultured under high-light conditions.

Figure 4 please explain Blue6803 and Red_c.

Line 276, B39/B40 -> B19/B40

Line 363, The orientation of the C-terminus -> The orientation of the C-terminus of PsaL

Reviewer #4:

In this work the authors present the structure and the biochemical and spectroscopic properties of the photosystem I complex of a high light tolerant cyanobacterium, C. aponimum. They find that this complex has slightly different spectroscopic features compared to PSI from the model cyanobacterium Synechocystis, which grows in low/medium light, and try to relate them to the differences in light growth conditions. Next they proceed to mutate Synechocystis PSI introducing some of the structural features present in C. aponimum PSI. By comparing the spectroscopic properties of the two complexes, they could identify the absorption property of one specific chlorophyll out of the 96 present in the complex.

The work is interesting and nicely combines structural and spectroscopic data. The conclusions on the second part of the work are sound and show how the protein environment can tune the spectroscopic properties of the pigments. On the other hand the link between the observed PSI features and the high light tolerance of this organism is less convincing.

General comments:

The authors discuss the PSI/PSII ratio of the cells and relate it to the light stress experienced by the two organisms. There are two relevant points

1. The correlation between PSI/PSII ratio and light intensity is less straight forward than indicated in the manuscript. This ratio depends on the species and in plants does not change in different light conditions.

2. The authors estimate the PSI/PSII ratio using the relative intensity of the 680- and 720-nm peaks of the 77-K fluorescence spectra. This quantification is not reliable because it depends on the excited-state lifetimes of PSI and PSII complexes, which might be different in the two organisms. The PSI/PSII ratio should be determined biochemically (i.e. by looking at the protein content) and/or via physiological measurements (electrochromic shift, P700 oxidation rates, etc.). On the other hand, this information is not essential for this manuscript, since the discussion about changes in PSI/PSII ratio is not needed (see the previous point) and PSI is the main focus of the manuscript.

The authors noted – correctly – that the PSI of the high-light growing C. aponimum has a lower content of red Chls in comparison to PSI of Synechocystis. However, the correlation of the red Chl content to light stress conditions remains unclear, and the related discussion (page 21) is not very convincing. In general, the authors seem to favor the conclusion that red Chls are detrimental for PSI in high light, which would explain the reduced red Chl content of C. aponimum PSI. For instance, the authors hypothesize that C. aponimum PSI might carry less red Chls to reduce its trapping time and maximize photoprotection. This claim is not supported by data, since the authors do not measure the trapping time of the two PSI. Moreover, it is known from the analysis of other organisms that the differences in PSI trapping times with different red Chl content are typically small, and that the PSI of cyanobacteria always behaves as a very efficient trap (lifetime < 50 ps) and is therefore well protected independent of the red Chl content. The other hypothesis regarding the IsiA ring (i.e. that the IsiA ring is mainly there to drag excitations away from the red Chls) is also not sound.

The last section in the results is not as strong as the rest of the work and could be removed/reduced. Indeed, the differences in amino acids (mostly aromatic) around Chls B7, A31, A32 do not seem enough to justify the presence/absence of a red Chl cluster. Indeed, red Chls in PSI are usually ascribed to charge-transfer states, which are expected to be influenced more by changes in surrounding charged/polar residues rather than aromatic ones. Also, the mentioned calcium ion seems to be too distant from the Chl cluster to really play an effect in this sense. Finally, the hypothesis of electron donation to a Chl by an aromatic amino acid having a large influence on the spectral properties (discussion, page 20) is not sound from a chemical perspective.

Detailed points:

Lines 34-39. PSI is certainly central in photosynthesis, but I do not see how the authors can conclude it from the fact that the PSI/PSII ratio changes in high light (and please check the recent literature).

Line 55. The discovering of the two photosystems by using different excitations was due to the spectroscopic work of Lou Duysens ( Duysens LNM, Amesz J, Kamp BM (1961) Two photochemical systems in photosynthesis. Nature 190:510-511)

Line 118. A higher carotenoid content relative to chlorophylls is observed in all organisms when grown in high light compared to low light.

Line 120. The ratio between the fluorescence peaks at 77K cannot be used to measure PSI/PSII. The fluorescence signal is much more complex. It can at best give indication that there are changes, but those need to be validated with other techniques.

Line 121-123. The rational for the investigation of PSI does not appear very convincing.

Lines 136-136, Figure 2B. The protein pattern differs between the two PSI. It makes sense that some of the proteins are slightly different, but the authors should not conclude that they could see "the expected bands". The change in mobility of PsaL is explained, but what about PsaF and PsaC?

Line 177. Which two Chls?

Line 233, it should be a different supplementary figure.

Line 277. The decrease is only in the absolute value, because the numbers are negative.

Figure 1. For the dilution experiment, please specify for how long cells were incubated at the given light intensity before taking the picture (C).

Figure 2. the differences in absorption and fluorescence between the PSI of the two strains are very small. Are they reproducible? The normalization is tricky because the authors do not know if the number of pigments is the same in the two PSI.

A number of the references seems to be outdated/incorrect. It is absolutely true that pioneering work of a high quality was done in the 80' and 90', but the introduction of new techniques in the last 30 years has led to the revision of some of these early reports. I would suggest the authors to consult some more recent literature.

Finally, the manuscript needs careful editing.

https://doi.org/10.7554/eLife.67518.sa1

Author response

Reviewer #1:

1) "…had a constant drip of fresh water and exposed to extreme light conditions." How is 1300 uE an extreme light? Are all terrestrial vascular plants now extremophiles?

This raises a good point and we have revised the wording of the text accordingly. That being said, the Sonoran Desert experiences one of the highest light conditions on the planet, experiencing over 300 days of sunlight per year. We theorize that an organism isolated from this type of desert environment could have adapted unique properties. Additionally, the south facing wall (with no shade) provides an optimal environment for studying an organism adapted to experiencing high light. In addition, an even higher light intensity was used during the selection process that ultimately revealed the ability of C. aponinum to grow in conditions up to 3000 µmol photons m-2s-1 to explore the limits of light adaptation.

In our opinion comparing these light intensities to the 200 µmol photons m-2s-1 used to culture other desert cyanobacteria in (Alwathnani, A. and Jeffrey, R. Cyanobacteria in Soils from a Mojave Desert Ecosystem. 5, 71–89 (2021)) and the 500 µmol photons m-2s-1 used for 20 minutes to induce high light inducible proteins in Synechocystis sp. PCC 6803 (Komenda, J. and Sobotka, R. Biochimica et Biophysica Acta Cyanobacterial high-light-inducible proteins — Protectors of chlorophyll – protein synthesis and assembly. BBA – Bioenerg. 1857, 288–295 (2016)) warrants the labeling of C. aponinum as an extremophile.

2) In this regard, choosing Synechocystis as a comparison is a bit of a "straw man" and it would be more appropriate to compare to a wider range of species/strains. In other words, wouldn't it be more accurate to say that Synechocystis is adapted to LOW light, rather than C. aponiunum adapted to high? If this is true, then the story may be about selection pressure for low light rather than photoprotectiom.

Our main motivation for using Synechocystis is that it is used in laboratories worldwide as the model cyanobacterium leading to its PSI being well characterized. Synechocystis can also be genetically modified relatively easily, so we are able to modify its PSI to test how structural variations effect spectroscopic properties. This allows us to validate the significance of the structural differences we observe in C. aponinum. Having a well characterized “baseline” to compare is paramount in order to reach any conclusions on the PSI core antenna. Additionally, Synechocystis is relatively close to C. aponinum on an evolutionary tree. While it may be possible to argue that Synechocystis is adapted to low light; “high-light” treatment is usually no more than 500 µmol photons m-2s-1 for short durations of time (see Muramatsu, M. and Hihara, Y. Acclimation to high-light conditions in cyanobacteria: From gene expression to physiological responses. J. Plant Res. 125, 11–39 (2012)).

3) The work shows that C. aponium can grow at higher light than Synechocystis. However, it was not possible to attribute this difference to PSI properties. This would be a pretty high bar to set, of course, but can the authors make the case that the differences they do observe are relevant?

Thank you for this comment. This is an important question. Under high light conditions, PSII is known to become damaged. In C. aponinum we show that there is an increased dependence on PSI for cells grown under high light. Even with the contribution of PBS’s, this still means that the balance between the two available photochemical traps changes and PSI is now more dominant. It is true that we cannot yet fully explain how PSI acts to confer high light resistance, but we can show that it is an important conduit of excitation energy at high light. In the revised version of this manuscript, we demonstrate that the red states of PSI are affected by the presence of a ca2+ ion located on the luminal side of PSI in the PsaL subunit. This explains the spectroscopic differences between the oligomeric states of PSI, a subject which has been studied for some time, as well as some of the properties of the core PSI complex in the green lineage. We think that identifying precise ways to manipulate the properties of PSI is integral to being able to address its contribution to high light stress and identify the underlying mechanism.

4) The experiments exposed plates to 3000 µmol photons m-2 s-1, which is higher than that these cells experience in the real world. What is the argument that 3000 uE is physiologically relevant? Is it not the case that this selects for something not relevant to nature?

3000mE was used during the selection on liquid cultures and not on plates. By exposing cells to extreme environments, we can explore the limits of light adaptation and thereby expose unique properties. This has been done in many genetic screens and has led to the elucidation of highly relevant cellular mechanisms. One example that comes to mind is DNA repair, where cells are often exposed to high levels of UV radiation or DNA damaging agents, still, the repair pathways that were discovered proved to be functional under everyday growth. While 3000mE is not something any cyanobacteria would experience in nature, the fact that C. aponinum could survive those conditions was the first indication that photosynthetic organisms can survive high light conditions. C. aponinum’s tolerance to high light conditions was also demonstrated on agar plates in direct comparison to Synechocystis (Figure 1).

5) The 77K emission spectrum is not a measure of PSI:PSII content, but on the distribution of excitation energy. Further, the interpretation of these spectra is dependent on the specific excitation wavelength. For instance, if cyan light will excite the PBS, which can dump energy differently into PSI and PSII, compared to, e.g. blue light. The emission spectrum is also impacted by cell density and other factors. None of these issues is acknowledged in the text, nor is that excitation wavelength described in the Methods, as is clear from the scant description given in the text: "An Oxford instruments Cryostat was used to cool 26 the sample to 77 K (cells were plunged into liquid nitrogen and measured immersed in liquid nitrogen)." which does not give any indication of the instrument, excitation, cell concentration, etc., though Figure 2 mentions that particular data was collected with excitation at 440 nm. What is the resolution of the spectrometer?

Thank you for this comment. We altered the description in the Results section and improved the description in the method section. We refer to the 77K measurements as a way to approximate the reliance on PSI:PSII, and that is now better reflected in the text. We excited our sample at 440 nm which preferentially excite Chls. While some backflow to PBS may occur, the physiological relevant parameter is the PSI/PSII utilization which ultimately shows more reliance on (or energy distribution to) PSI after growing in high light. We used a Horiba Fluoromax-4 for these measurements with a stated resolution of +/- 0.5 nm. We have indicated our slit settings on both monochromators in the method section for both RT and 77 K measurements.

6) Figure 3. The spectra are interpreted as apparent shifts based on subtracting pairs of spectra normalized to their peaks. This seems to be based on a subjective interpretation and other interpretations are possible. Changing the normalization point could result in an entirely different interpretation. For instance, normalizing to a point on the red side of the peak could give the appearance of an increase in absorbance or emission on the blue edge of the spectrum (rather than a shift). The deconvolution of the absorbance spectra using three Gaussian components as in Figure 2D is likewise based on an assumption or interpretation that there are only these components.

We now normalized the absorbance spectra to the area between 550-775 nm because the structures of each PSI reveal the same number of chlorophylls, making this method a reasonable approximation (this is different from the previous manuscript in which we used the 650-775 nm interval for integration). Normalizing the area between 550-775 normalized the amount of light being absorbed exclusively by chlorophylls. The presence of long wavelength features in the difference spectra persisted using several normalization methods and integration intervals. When computing the deconvolution of the peak, we used the least number of components that adequately fit the data, resulting in 3 total peaks. It’s entirely possible (even probable) that these peaks contain contributions from multiple Chls, we are not assigning these gaussians to specific chlorophylls in our work and our statements that these gaussians represent the contribution of LWC still holds. Methods with higher optical resolution are required to provide finer details on the electronic structure of LWC. Our choice of deconvolution and normalization represents, in our view, the most conservative treatment of our data.

7) The text needs to be more circumspect and/or directly address this as a possibility, since such alternative normalization could indicate a quite different story. For example, the text reads: "The difference spectrum of C. aponinum – Synechocystis shows a strong negative peak at 701 nm revealing that C. aponinum PSI contains less LWC 150 than Synechocystis (Figure 2D)." It suggests this, but there are other interpretations. This issue is more evident further in the manuscript, in Figure 4, a different method of normalization was used, as stated in the following: "The absorbance spectra were normalized to the area between 650 nm to 775 nm and the spectrum of Wild Type Synechocystis was 238 subtracted from Red_c (Figure 4C and 4D)." As argued above, changing the normalization method can give rise to different interpretations, so some real justification must be presented for using a particular approach, and an even stronger argument needs to be made when changing the normalization between samples. None is given.

The indication of different normalization methods were the results of typos that we apologies for, all the previous absorbance data was normalized using the same method to the area between 650 to 775 nm. We completely agree that the same normalization method should be applied to all relevant data and that any changes should be well reasoned. In this version of the manuscript all the presented absorbance spectra were normalized to the area between 550-775 nm to completely account for both Qy and Qx transitions contributions, so comparisons between species are consistent and there are no changes in normalization methods between absorbance samples as we have clarified in the text. Room temperature and 77K emission spectra were normalized to the max wavelength for comparison of peaks ratios and shapes.

8) Figure 4 D compares isolated complexes from two different species and the differences spectra can be interpreted as a series of narrowing and shifting of spectra, but all of which are likely to be dependent on the method of normalization, which is not adequately described. Moever, the method used to make the differences spectra were not consistently applied, e.g. "[t]he absorbance spectra were normalized to the area between 650 nm to 775 nm and the spectrum of Wild Type Synechocystis was subtracted from Red_c (Figure 4C and 4D)." What were the criteria for using the different procedures and how was bias avoided?

This was addressed by our answer above.

9) The shifts in absorbance and fluorescence emission are quite small, especially realizing that the bands tend to sharpen at 77K. At RT, there may be essentially no functionally-relevant differences in the spectra. How can this be addressed?

This is a good point and was addressed in the new version of the manuscript. The addition of a new mutant (Red_d) to the updated manuscript shows a difference in LWC at room temperature in both absorption and emission measurements, but not 77K, and therefore we included the room temperature emission of all species/mutants to this version. These differences could be physiological significance, by possibly affecting the rate of chl triplet accumulation or any other damage inducing mechanism known to occur in PSI. Currently, C. aponinum, is not genetically amenable and we cannot test these mutations in this organism. On the other hand, transferring them to Synechocystis unavoidably leaves additional differences (in PSI and other cellular systems) between these two organisms unaccounted for. Generating a detailed understanding on the mechanisms that tune energy transfer and damage avoidance in PSI can ultimately provide the answer to this question, but the complexity of this system currently prevents a simple explanation, so more work should be done in the future to fully unravel the mechanisms.

10) Can the text be expanded to further discuss the very large apparent difference in PsaF?

To explain this we have run, and included, a new gel that better separate the PsaF and PsaD bands (Figure 2B) and expanded an explanation in the manuscript. In the new gel, size differences can still be seen between some subunits, notably, PsaC, PsaD, and PsaF. We examined the DNA sequences for these genes carefully and found no evidence for additional sequences that can be translated at both the N and C termini. Based on the protein sequence data, the mass of PsaC for C. aponinum and Synechocystis is 8.82 kD and 8.83 kD, respectively; the mass of PsaD for C. aponinum and Synechocystis is 16.18 kD and 15.65 kD, respectively; and the mass of PsaF for C. aponinum and Synechocystis is 18.69 kD and 18.25 kD. These masses do not explain the difference in migration for either species, and the bands we observe on the gel do not correspond precisely to these molecular weights. However, due to the variations of sequences between species, some changes in detergent binding are likely different between these samples. This is known to cause gel shifting, and is common when analyzing membrane proteins via SDS-PAGE and can also account for soluble proteins like PsaC and PsaD. This is well described in (Rath, A., Glibowicka, M., Nadeau, V. G., Chen, G. and Deber, C. M. Detergent binding explains anomalous SDS-PAGE migration of membrane proteins. 106, 1760–1765 (2009)).

11) Cyanobacterium aponinum (C. 78 aponinum). should be both Ital. Is this an official name?

This has been corrected so it is in italics, and it is the official name as deposited in the NCBI database (NCBI:txid2676140). The updated manuscript reflects this in the Results section and the methods section, as well as provided in the data availability statement.

Reviewer #2:

While the manuscript presents data in a clear, concise format, there are some outstanding issues that should be addressed to improve the understanding of the results and significance of the findings.

The selection strategy was using 3,000 umol photons m-2s-1 light to select for high light tolerant strains. As the sample was from the Sonoran desert, it would be useful to clarify if/what temperature was used for the selection as well, as the growth temperature of 30C is not particularly high for many cyanobacteria.

Environmental samples were collected in February 2016. According to the National centers for Environmental Information, the average temperature in February for Tempe, AZ where C. aponinum was isolated from is 18.7 C. This information was added to the text to expand on the description of isolation. We used 30 C during selection because that it is used in ours (and others) lab for cyanobacteria cultivation. Over the course of the last five years, we have grown C. aponinum at 22 C as well and found that it grows very well at this temperature which is closer to the average environmental value at the time of isolation. Other members of the aponinum family were isolated from hot springs and this family of cyanobacteria appear to grow in many different environments.

The figures detailing the characterization of PSI between different samples could benefit from a clearer delineation between the samples shown. For most samples, the spectra are nearly identical, and the differences, or the fact that two samples are shown is nearly impossible to determine (figures 2C, 4C).

This stems from the large number of chlorophylls present in PSI, which means single chlorophyll differences make about 1% of the total Qy intensity. We’ve done our best to visually improve the graphs but this is still a challenge due to the magnitude of the differences. We’ve included difference spectra when appropriate but think that showing the complete spectra is important to accurately report on the extent of the differences and not only their shapes. In emission curves we have included enlarged sections to improve the visibility of the differences.

Similarly, it would be useful to discuss the other differences in PSI subunit composition. From the analysis of isolated PSI (figure 2), the authors discuss the migration differences between PsaL. However, there are clear differences between the migration of other subunits as well. Whether the primary sequences are markedly different should be mentioned. The authors also state that the expected number of bands is observed for C. aponinum (line 137). It is unclear from the figure if that is the case, as five distinct bands are visible in the Synechocystis sample but only four in C. aponinum. If PsaF migrates almost together with PsaD that should be clarified.

To explain this we have run, and included, a new gel that better separate the PsaF and PsaD bands (Figure 2) and expanded an explanation in the manuscript. See our response to reviewer 1, point 10 above.

“Comparing this sample to a known PSI sample from Synechocystis, SDS-PAGE shows similar bands for PSI subunits in Figure 2B, with notable shifts in the PsaD, PsaF, PsaL and PsaC subunits.

To investigate the different migration of PSI subunits between C. aponinum and Synechocystis, C. aponinum genomic DNA was isolated and sequenced (NCBI:txid2676140). PSI genes were located and annotated, then compared to Synechocystis (Figure 2 —figure supplement 1). The gene for PsaL revealed the difference in migration of is likely due to two substantial differences compared to Synechocystis: (1) a 6 amino acid insert located on the stromal side of the membrane between two transmembrane helixes and (2) a markedly different C-terminus (Figure 2 —figure supplement 1) containing an extension of four amino acids. In addition, a 7 amino acid insertion is seen in the PsaB gene of C. aponinum compared to Synechocystis (Figure 2 —figure supplement 1). Genes for the remaining subunits (PsaD, PsaF, and PsaC) did not reveal size differences that would correspond to these shifts (Figure 2 —figure supplement 1). There were however sequence variations between C. aponinum and Synechocystis which likely cause gel shifting, a common occurrence when analyzing membrane proteins via SDS-PAGE60.”

Lines 176-177 should be clarify which chlorophylls are discussed relative to the changes in B40 coupling.

We have changed this sentence to use chlorophyll numbers in the manuscript text.

The description of the chimeric Synechocystis mutant needs considerable clarification. Figure 4B: what is Blue6803? The legend for Figure 2G does not seem to be correct. It should also be clarified that the amino acids present in C. aponinum are not mutations (ex: line 253, 399), they are the wild-type sequences and are different relative to Synechocystis. The discussion within the section of the excitonic coupling calculations should also be addressed to clarify the distinction between the C. aponinum structure and the Synechocycsist Red_C mutant.

Thank you for pointing these out. Blue6803 was regrettably a typo and has been fixed, it is now referred to as Red_c. We do not have a figure 2G, however we did correct a mistake on the legend of figure 4G. We have clarified that the sequences in C. aponinum are natural variations and refer to them as variations/differences in the text. To clarify which models were used in calculations we added the following statement to our method section under “Excitonic Structure Calculations:” – “The PDB: 6UZV model of trimeric PSI was used as WT Synechocystis and the C. aponinum PSI structure (PDB:6VPV) was used to approximate the Red_c structure.”

As the PsaB changes around chlorophyll B40 did not explain either the absorbance spectrum changes or the long-wavelength chlorophyll differences, the authors also discuss amino acid differences in the vicinity between the two species. It would be beneficial to do a similar comparison between these positions and amino acid composition between difference cyanobacterial species as was done for the calcium binding residue position. This may also provide more information to discuss these changes rather than a preliminary list of the differences between the two structures.

We thank the reviewer for this comment. The new results on the Red_d mutation now explains some of the differences in the LWC absorption. Additionally, Figure 6 —figure supplement 1 compare the structure of Synechocystis, C. aponinum, and Pisum Sativum (Pea plant), a known eukaryotic PSI structure. Plants have less LWC in its core PSI (Croce, Roberta, Giuseppe Zucchelli, Flavio M. Garlaschi, and Robert C. Jennings. 1998. “A Thermal Broadening Study of the Antenna Chlorophylls in PSI- 200, LHCI, and PSI Core.” Biochemistry.). Further, we have added multiple sequence alignment statistics of the residues shown in Figure 6, to the supplemental figures, investigating the identity of these residues across bacteria and eukaryotic databases (Figure 6 —figure supplements 2-4), demonstrating that these specific residues vary between cyanobacteria and eukaryotic organisms, and that C. aponinum contains residues that are more prevalent in eukaryotic organisms than in cyanobacteria. It’s important to note that we don’t want to discount the possibility that water molecules (which we cannot resolve at the current resolution) could be causing the shifts in the spectral characteristics, however the amino acids nearby could also be the underlying reason for a different organization of water molecules.

It is interesting that this mutant did not reproduce either the C. aponinum absorbance spectrum or the loss of far-red wavelength chlorophylls, again highlighting the utility of doing both the sequence and structure comparisons. While it would increase the impact of this work to also include mutation comparisons introducing the C. aponinum PsaL into Synechocystis, as well as the double C. aponinum PsaB/PsaL chimera, that work may not be feasible under current circumstances. However, because the Red_c mutant did not explain the differences between absorbance spectra, a more detailed discussion of how the various changes may be involved in those changes should be carried out.

We included a new mutant, Red_d, in this manuscript to explain the consequence of calcium ion binding which partially answers this remark. We agree that generating additional combinations of site mutations is important, however more information is needed before we approach this large task, and we think this should be addressed in a separate work.

Based on the presentation in the manuscript, it is unclear why full genome sequencing was performed, as PSI genes could have presumably be sequenced via PCR methods. The authors do themselves a disservice at having done this analysis and essentially not addressing the results. It would be interesting to know if there are any genes within the genome beside those highlighted for PSI that may be of interest in understanding the high-light capacity of this cyanobacterium.

Because the 16S based tree suggested this is a new species, we sequenced the entire genome to confirm this, and this data is now publicly available for the community to study (NCBI:txid2676140). To enumerate and validate all the sequence changes that contribute to high light resistance requires a separate experimental design, beyond the scope of the current manuscript in our opinion.

The methods section should be tidied up. Many explanations are unnecessary and likely could be better described via referencing previous works.

We have removed some sections (such as phenol chloroform DNA extraction) from the methods section, corrected additional typos and added a unified primers table.

In general the manuscript requires further proof-reading throughout to rectify typos, italicization issues, unit labels, etc.

We have carefully proofread the entire manuscript and made many corrections.

Reviewer #3:

The interpretation of spectral analysis of C. aponinum PSI is questionable. They conclude that C. aponinum PSI contains less LWC than Synechocystis PSI based on the differences of absorption spectra between the two PSIs shown in Figures 2C and 2D. However, the difference is very small, with the amplitude of δ-absorbance lower than 0.04, which seems within the error range, while the absorbance of the PSI complex is around 1.5. On the contrary, the 2-nm red-shift of 77K fluorescence emission of C. aponinum PSI is quite evident (Figure 2F). Therefore it is possible that C. aponinum PSI contains more, but not less, LWC.

A 0.04 difference is well above the baseline variation of our spectrophotometer (which is app. 0.0005 OD over the measured wavelength range). To better demonstrate the robustness of this measurement, PSI was isolated from both C. aponinum and Synechocystis 3 separate times and absorption spectra were taken on 3 separate days and compared to each other. These spectra were then normalized according to our methods section, and the difference spectra of these measurements are shown in Figure 2D, including the standard deviation of the measurements along the entire wavelength range, showing that around the Qy transition (which includes the LWCs) the variation is negligible compared to the size of the difference signal. 77K emission is not a good indication for the total number of LWC, as it is affected by transfer processes that still occur at 77K, therefore this measurement is a better indication of the terminal emitter in PSI. Our hypothesis is that eliminating some of the LWC in C. aponinum causes increased energy transfer to a different LWC, which now has a stronger contribution to 77 K emission. This is consistent with the findings that C. aponinum contains less LWC but displays a red shifted emission peak at 77K. The principles of this hypothesis are that the contribution of a specific LWC to the 77K emission depends not only on its transition energy but also on its connectivity. This has been shown by many studies, but our recent work on site specific mutations in Synechocystis is relevant for this particular case (Khmelnitskiy, A., Toporik, H., Mazor, Y. and Jankowiak, R. On the Red Antenna States of Photosystem I Mutants from Cyanobacteria Synechocystis PCC 6803. J. Phys. Chem. B (2020). doi:10.1021/acs.jpcb.0c05201).

They construct a mutant form of Synechocystis strain containing the sequence of the PsaB loop from C. aponinum, and find that the 77K fluorescence emission of the mutant PSI is the same with Synechocystis PSI. Thus conclude that chlorophyll B40 is not responsible for the different absorption of LWC. Although they confirm the complete and correct replacement of PsaB in the mutant strain, they do not validate that B40 orientates the same with that in C. aponinum PSI. Since the loop regions are usually highly flexible and may adopt quite different conformation even with the same primary sequence, it is possible that B40 is indeed responsible for the different absorption of LWC in C. aponinum, but B40 in the mutant does not change its orientation, i.e. adopts the same orientation as that in Synechocystis PSI.

The general statement that loop regions are more flexible than regions with secondary structure is of course correct, but we do not think it applies in this particular case. The first indication for this is the fact that the B40 loop is well resolved in our cryo-EM structure, this means that it adopts a relatively stable conformation in C. aponinum. This can be explained by the additional interactions between this loop and the chlorophylls it binds, together with the stacking interactions between the chlorophylls themselves. We have shown previously that chlorophyll binding loops can be transferred between different PSIs, and that the conformation of the loop remain essentially identical to the original structure even in the absence of adjacent subunits (Toporik, H. et al. The structure of a red-shifted photosystem I reveals a red site in the core antenna. Nat. Commun. (2020). doi:10.1038/s41467-020-18884-w). Further support for our assertion that the B40 loop adopts a similar configuration comes from the high sequence similarity of sequences juxtaposed to this loop, making the immediate environments between the two different PSI’s highly similar (see Figure 3 —figure supplement 3).

C. aponinum is a high-light tolerant cyanobacterium, and it does not grow well under low light conditions as shown in Figure 1C. However, in this study, they culture the cells at low light (∼40 μE), which may lead to some changes of the photosynthetic complexes, therefore the structural features of PSI important for C. aponinum to survive in the high light environment cannot be identified.

We would like to point out that C. aponinum grows fine under low light conditions. In the growth study in Figure 1 (and described in the methods and in the updated manuscript’s figure legend), C. aponinum was plated at an OD730 of 3 and Synechocystis was plated at an OD730 of 5, and both were diluted in ¼ steps. This was done to best depict the difference in viability between the two cyanobacteria in increasing light. While it would be ideal to grow the cyanobacteria at higher light, technical issues arise, mainly that proteins isolated from stressed cells are not ideal for structural analysis. We also prepared PSI from highlight grown cells and from low light grown cells and did not observe any differences in absorbance or protein content (SDS PAGE), we conclude that PSI itself does not change in response to highlight.

This reviewer understands that the difference of absorption spectra between PSI from C. aponinum and Synechocystis is very small, as only a few chlorophylls are changed. However, if they would like to use the spectral data, they need to prove that the differences of absorption spectra are indeed due to the different spectral property of PSI, but not the standard error. The spectra should be repeated at least three times for each PSI complexes and the differences of the repeated spectra should be shown in the paper.

It's better to determine the structure of the PSI complex from the constructed mutant to show the orientation of chlorophyll B40. If it is the same as that in C. aponinum PSI, this result will further strengthen their conclusion. If it is not, then chlorophyll B40 should be further investigated.

It will be more convincing if they purify and determine the structure of the PSI complex from C. aponinum cultured under high-light conditions.

Figure 4 please explain Blue6803 and Red_c.

Line 276, B39/B40 -> B19/B40

Line 363, The orientation of the C-terminus -> The orientation of the C-terminus of PsaL

Reviewer #4:

The authors discuss the PSI/PSII ratio of the cells and relate it to the light stress experienced by the two organisms. There are two relevant points

1. The correlation between PSI/PSII ratio and light intensity is less straight forward than indicated in the manuscript. This ratio depends on the species and in plants does not change in different light conditions.

2. The authors estimate the PSI/PSII ratio using the relative intensity of the 680- and 720-nm peaks of the 77-K fluorescence spectra. This quantification is not reliable because it depends on the excited-state lifetimes of PSI and PSII complexes, which might be different in the two organisms. The PSI/PSII ratio should be determined biochemically (i.e. by looking at the protein content) and/or via physiological measurements (electrochromic shift, P700 oxidation rates, etc.). On the other hand, this information is not essential for this manuscript, since the discussion about changes in PSI/PSII ratio is not needed (see the previous point) and PSI is the main focus of the manuscript.

Reviewer 4 correctly points out that PSI/PSII ratio is complex and adds that these values are a measure of the distribution of excitation energy between photosystems. The distribution of excitation energy between the photosystems is probably a more relevant parameter then the PSI/PSII protein ratio. We have changed the text to better reflect that the 77K measurements is a way to approximate the distribution of excitation energy between PSI and PSII. This demonstrates that under high light, C. aponinum directs more excitation energy to PSI, relative to PSII, and this is the opposite of what was reported for Synechocystis.

The authors noted – correctly – that the PSI of the high-light growing C. aponimum has a lower content of red Chls in comparison to PSI of Synechocystis. However, the correlation of the red Chl content to light stress conditions remains unclear, and the related discussion (page 21) is not very convincing. In general, the authors seem to favor the conclusion that red Chls are detrimental for PSI in high light, which would explain the reduced red Chl content of C. aponimum PSI. For instance, the authors hypothesize that C. aponimum PSI might carry less red Chls to reduce its trapping time and maximize photoprotection. This claim is not supported by data, since the authors do not measure the trapping time of the two PSI. Moreover, it is known from the analysis of other organisms that the differences in PSI trapping times with different red Chl content are typically small, and that the PSI of cyanobacteria always behaves as a very efficient trap (lifetime < 50 ps) and is therefore well protected independent of the red Chl content. The other hypothesis regarding the IsiA ring (i.e. that the IsiA ring is mainly there to drag excitations away from the red Chls) is also not sound.

In the context of the PSI core antenna, we do not think there is a general all-inclusive idea explaining the physiological role of red chlorophylls. We favor the position that specific LWC (including groups of coupled chlorophylls) contribute to specific functions. A prerequisite for this approach is identifying and being able to manipulate LWC in vivo and in vitro and this work provides an important step towards this goal. We have included a new section in the discussion expanding on this point. We do think that specific red sites are important for interactions with antenna systems (PBS and IsiA) and this is clearly relevant to high light growth. At the same time, we can only propose a general mechanism to account for the contribution of PSI to high light growth in C. aponinum.

The last section in the results is not as strong as the rest of the work and could be removed/reduced. Indeed, the differences in amino acids (mostly aromatic) around Chls B7, A31, A32 do not seem enough to justify the presence/absence of a red Chl cluster. Indeed, red Chls in PSI are usually ascribed to charge-transfer states, which are expected to be influenced more by changes in surrounding charged/polar residues rather than aromatic ones. Also, the mentioned calcium ion seems to be too distant from the Chl cluster to really play an effect in this sense. Finally, the hypothesis of electron donation to a Chl by an aromatic amino acid having a large influence on the spectral properties (discussion, page 20) is not sound from a chemical perspective.

The inclusion of Red_d, which removes the ca2+ ion in question, in this manuscript demonstrates its effect on LWC of Synechocystis.

With regards to the distribution of aromatic amino acids, we compared the environment of A31-B7 cluster between Synechocystis, C. aponinum, diatom, green algae, red algae, and plant PSI, the amino acids discussed are identical between c. aponinum and eukaryotic structures and different in Synechocystis. Together with the fact that the plant (and green algae) core PSI was shown to contain less LWC than the cyanobacterial PSI (Synechocystis and T. elongatus) this supports our suggestion that these changes affect LWC is reasonable (see new Figure 6 —figure supplements 2-4). Published structures show that the electrons on the tryptophan indole are 3.1 Å from the chlorophyll ring in Synechocystis, which is closer than the ring-ring distance known to effect chlorophyll absorbance. We think this is within the range required to influence the absorbance of a chlorophyll molecule.

Detailed points:

Lines 34-39. PSI is certainly central in photosynthesis, but I do not see how the authors can conclude it from the fact that the PSI/PSII ratio changes in high light (and please check the recent literature).

We’ve significantly revised the text to address this issue.

Line 55. The discovering of the two photosystems by using different excitations was due to the spectroscopic work of Lou Duysens ( Duysens LNM, Amesz J, Kamp BM (1961) Two photochemical systems in photosynthesis. Nature 190:510-511)

We included this citation in the current version.

Line 118. A higher carotenoid content relative to chlorophylls is observed in all organisms when grown in high light compared to low light.

We agree and we included a statement regarding this in our text.

Line 120. The ratio between the fluorescence peaks at 77K cannot be used to measure PSI/PSII. The fluorescence signal is much more complex. It can at best give indication that there are changes, but those need to be validated with other techniques.

We have changed the text to better reflect that the 77K measurements is a way to approximate the reliance on PSI:PSII.

Line 121-123. The rational for the investigation of PSI does not appear very convincing.

Our rational is mainly derived from our 77K whole cells measurements. It is very reasonable that other mechanism contributes to the ability of c. aponinum to grow at high light.

Lines 136-136, Figure 2B. The protein pattern differs between the two PSI. It makes sense that some of the proteins are slightly different, but the authors should not conclude that they could see "the expected bands". The change in mobility of PsaL is explained, but what about PsaF and PsaC?

To explain this we have run, and included, a new gel that better separate the PsaF and PsaD bands (Figure 2) and expanded an explanation in the manuscript. See our response to reviewer 1, point 10 above.

Line 177. Which two Chls?

Line 233, it should be a different supplementary figure.

This was fixed

Line 277. The decrease is only in the absolute value, because the numbers are negative.

We have changed the text to “decreases in magnitude”.

Figure 1. For the dilution experiment, please specify for how long cells were incubated at the given light intensity before taking the picture (C).

This was added to the description:

“(C) Serial dilutions of Synechocystis and C. aponinum on BG11 plates. Cells were serially diluted in ¼ steps and incubated at 30°C for 5 days (light intensities > 370 µmol photons m-2s-1) and 10 days (light intensity = 50 µmol photons m-2s-1)”

Figure 2. the differences in absorption and fluorescence between the PSI of the two strains are very small. Are they reproducible? The normalization is tricky because the authors do not know if the number of pigments is the same in the two PSI.

This is very reproducible, we modified figure 2D to include the results +- SD. From the structure work, we know that both species of PSI contain the same number of chlorophylls.

A number of the references seems to be outdated/incorrect. It is absolutely true that pioneering work of a high quality was done in the 80' and 90', but the introduction of new techniques in the last 30 years has led to the revision of some of these early reports. I would suggest the authors to consult some more recent literature.

We added citations throughout the manuscript and changed a paragraph in the introduction (lines 42-61) to better acknowledge recent work on PSI photoinhibition:

“… A significant amount of work on photoinhibition has focused on PSII due to its rapid turnover in high-light and the efficient repair mechanisms that evolved to cope with PSII specific photodamage24,25. PSI specific damage, however, is irreversible and long lived due to a lack of repair mechanisms, requiring the biosynthesis of new PSI polypeptides26–30. Fluctuating light and low temperatures have been attributed to PSI photoinhibition by causing an imbalance in the redox state of PSI donors and acceptors27,31–34. …”

Finally, the manuscript needs careful editing.

We carefully edited the manuscript.

https://doi.org/10.7554/eLife.67518.sa2

Article and author information

Author details

  1. Zachary Dobson

    1. School of Molecular Sciences, Arizona State University, Tempe, United States
    2. BiodesignCenter for Applied Structural Discovery, Arizona State University, Tempe, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Software, Validation, Visualization, Writing - original draft, Writing - review and editing
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4951-1701
  2. Safa Ahad

    Department of Chemistry, Purdue University, West Lafayette, United States
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Software, Writing - original draft, Writing - review and editing
    Competing interests
    none
  3. Jackson Vanlandingham

    1. School of Molecular Sciences, Arizona State University, Tempe, United States
    2. BiodesignCenter for Applied Structural Discovery, Arizona State University, Tempe, United States
    Contribution
    Investigation
    Competing interests
    none
  4. Hila Toporik

    1. School of Molecular Sciences, Arizona State University, Tempe, United States
    2. BiodesignCenter for Applied Structural Discovery, Arizona State University, Tempe, United States
    Contribution
    Formal analysis, Investigation, Methodology, Software, Supervision, Visualization, Writing - review and editing
    Competing interests
    none
  5. Natalie Vaughn

    1. School of Molecular Sciences, Arizona State University, Tempe, United States
    2. BiodesignCenter for Applied Structural Discovery, Arizona State University, Tempe, United States
    Contribution
    Conceptualization
    Competing interests
    none
  6. Michael Vaughn

    1. School of Molecular Sciences, Arizona State University, Tempe, United States
    2. BiodesignCenter for Applied Structural Discovery, Arizona State University, Tempe, United States
    Contribution
    Conceptualization, Methodology
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9357-094X
  7. Dewight Williams

    John M. Cowley Center for High Resolution Electron Microscopy, Arizona State University, Tempe, United States
    Contribution
    Investigation, Methodology
    Competing interests
    none
  8. Michael Reppert

    Department of Chemistry, Purdue University, West Lafayette, United States
    Contribution
    Conceptualization, Methodology, Resources, Software, Supervision, Visualization, Writing - original draft, Writing - review and editing
    Competing interests
    none
  9. Petra Fromme

    1. School of Molecular Sciences, Arizona State University, Tempe, United States
    2. BiodesignCenter for Applied Structural Discovery, Arizona State University, Tempe, United States
    Contribution
    Conceptualization, Funding acquisition, Methodology, Resources, Supervision, Writing - original draft, Writing - review and editing
    Competing interests
    none
  10. Yuval Mazor

    1. School of Molecular Sciences, Arizona State University, Tempe, United States
    2. BiodesignCenter for Applied Structural Discovery, Arizona State University, Tempe, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Software, Supervision, Validation, Visualization, Writing - original draft, Writing - review and editing
    For correspondence
    yuval.mazor@asu.edu
    Competing interests
    none
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5072-0928

Funding

National Institute of Food and Agriculture (2020-67034-31742)

  • Zachary Dobson

Biodesign, Center of Applied Structural Discovery

  • Zachary Dobson

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

Senior Editor

  1. Olga Boudker, Weill Cornell Medicine, United States

Reviewing Editor

  1. David M Kramer, Michigan State University, United States

Reviewer

  1. Geoffry A Davis, Michigan State University, United States

Publication history

  1. Received: February 13, 2021
  2. Accepted: August 25, 2021
  3. Accepted Manuscript published: August 26, 2021 (version 1)
  4. Version of Record published: September 9, 2021 (version 2)

Copyright

© 2021, Dobson 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.

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Further reading

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    Phycobilisome (PBS) is the main light-harvesting antenna in cyanobacteria and red algae. How PBS transfers the light energy to photosystem II (PSII) remains to be elucidated. Here we report the in situ structure of the PBS–PSII supercomplex from Porphyridium purpureum UTEX 2757 using cryo-electron tomography and subtomogram averaging. Our work reveals the organized network of hemiellipsoidal PBS with PSII on the thylakoid membrane in the native cellular environment. In the PBS–PSII supercomplex, each PBS interacts with six PSII monomers, of which four directly bind to the PBS, and two bind indirectly. Additional three ‘connector’ proteins also contribute to the connections between PBS and PSIIs. Two PsbO subunits from adjacent PSII dimers bind with each other, which may promote stabilization of the PBS–PSII supercomplex. By analyzing the interaction interface between PBS and PSII, we reveal that αLCM and ApcD connect with CP43 of PSII monomer and that αLCM also interacts with CP47' of the neighboring PSII monomer, suggesting the multiple light energy delivery pathways. The in situ structures illustrate the coupling pattern of PBS and PSII and the arrangement of the PBS–PSII supercomplex on the thylakoid, providing the near-native 3D structural information of the various energy transfer from PBS to PSII.

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    Research Article

    The chloroplast proteome contains thousands of different proteins that are encoded by the nuclear genome. These proteins are imported into the chloroplast via the action of the TOC translocase and associated downstream systems. Our recent work has revealed that the stability of the TOC complex is dynamically regulated by the ubiquitin-dependent chloroplast-associated protein degradation (CHLORAD) pathway. Here, we demonstrate that the TOC complex is also regulated by the SUMO system. Arabidopsis mutants representing almost the entire SUMO conjugation pathway can partially suppress the phenotype of ppi1, a pale-yellow mutant lacking the Toc33 protein. This suppression is linked to increased abundance of TOC proteins and improvements in chloroplast development. Moreover, data from molecular and biochemical experiments support a model in which the SUMO system directly regulates TOC protein stability. Thus, we have identified a regulatory link between the SUMO system and the chloroplast protein import machinery.