Time-resolved comparative molecular evolution of oxygenic photosynthesis

Abbreviations: MRCA, most recent common ancestor; LUCA, the last universal common ancestor or the most recent common ancestor of life; MSV, Margulisbacteria, Sericytochromatia and Vampirovibrionia; PSII, photosystem II; PSI, photosystem I; CBP, chlorophyll-binding protein; RC, reaction centre; ROS, reactive oxygen species; GOE, Great Oxidation Event; dD0, duplication leading to the origin of D1 and D2 subunits of photosystem II; dCP, duplication leading to the origin of CP43 and CP47 subunits of photosystem II; dAB, duplication leading to the origin of Alpha and Beta subunits of ATP synthase; ΔT, the span of time between dD0, dCP, dAB, or the LUCA, and the MRCA of Cyanobacteria; Ma, Mega-annum, million years; Ga, Giga-annum, billion years; ν, Rate of amino acid substitutions per site; δ Ga⁻¹, Amino acid substitutions per site per billion years

Keywords: Origin of photosynthesis, Origin of life, Cyanobacteria, Photosystem, Reaction centre, Water oxidation

1.1. Evolution of Cyanobacteria

The origin of oxygenic photosynthesis is considered a turning point in the history of life, marking the transition from the ancient world of anaerobes into a productive aerobic world that permitted the emergence of complex life [1]. Oxygenic photosynthesis starts with photosystem II (PSII), the water-oxidizing and O₂-evolving enzyme of Cyanobacteria and photosynthetic eukaryotes. PSII is a highly conserved, multicomponent, membrane protein complex, which was inherited by the most recent common ancestor (MRCA) of Cyanobacteria in a form that is structurally and functionally similar to that found in all extant species [2]. Thus, the origin of oxygenic photosynthesis antedates the MRCA of Cyanobacteria by an undetermined amount of time.

Cyanobacteria's closest living relatives are the clades known as Vampirovibrionia (formerly Melainabacteria) [3,4], followed by Sericytochromatia [5] and Margulisbacteria [6]. Described Margulisbacteria include intracellular symbionts of placozoans [7] or are found as part of a chain of complex symbiotic interactions in the gut of termites [8]. Vampirovibrionia also include gut symbionts as well as specialist predators of green algae [9]. Less is known of the Sericytochromatia [4]. Currently, no photosynthetic strains have been described in these groups of uncultured bacteria and this has led to the hypothesis that oxygenic photosynthesis arose during the time spanning the divergence of Vampirovibrionia and the MRCA of Cyanobacteria, starting from an entirely non-photosynthetic ancestral state [5,10]. Recent molecular clock studies have suggested that the span of time between the divergence of Cyanobacteria and Vampirovibrionia could be of several hundred million years [11,12]. However, it is still unclear from molecular clock analyses and the microfossil record when exactly the MRCA of Cyanobacteria occurred [13].

1.2. Evolution of photosystem II

The heart of PSII is made up of a heterodimeric reaction centre (RC) core coupled to a core antenna. The two subunits of the RC core of PSII are known as D1 and D2, and these are associated respectively with the antenna subunits known as CP43 and CP47. D1 and CP43 make up one monomeric half of the RC, and D2 and CP47, the other half. Water oxidation is catalysed by a Mn₄CaO₅ cluster coordinated by ligands from both D1 and CP43 [14,15]. The cluster is functionally coupled to a redox active tyrosine-histidine pair (Y_Z-H190) also located in D1, which relays electrons from Mn to the oxidized chlorophyll pigments of the RC during charge separation [16]. In a cycle of four consecutive light-driven charge separation events, O₂ is released in the decomposition of two water molecules.

Photosystems evolved first as homodimers [17,18]: therefore, the core and the antenna of PSII originated from ancestral gene duplication events that antedated the MRCA of Cyanobacteria. In this way, CP43/D1 retain sequence and structural identity with CP47/D2. The conserved structural and functional traits between CP43/D1 and CP47/D2 suggest that the ancestral PSII homodimer—prior to the duplication events—was not only structurally similar to heterodimeric PSII, but also that it could have split water and evolved protective mechanisms against the formation of reactive oxygen species (ROS) [[18], [19], [20]].

In a previous study, we attempted to measure the span of time between the duplication that led to D1 and D2 (dD0) and a point that approximated the MRCA of Cyanobacteria: a period of time that we called ΔT [19]. We observed that the magnitude of ΔT can be very large, well over a billion years. Such a large ΔT suggested that the origin of a water-oxidizing PSII substantially antedated the MRCA of Cyanobacteria and highlighted that the evolution of the process, prior to this ancestor, was likely marked by loss of photosynthesis and extinction events. This implies that despite their specialized heterotrophic lifestyles, an ancestral photosynthetic state at the divergence of Margulisbacteria, Sericytochromatia, and Vampirovibrionia (MSV) cannot be ruled out. However, our study neither provided an absolute age for the MRCA of Cyanobacteria nor the duplication event itself, as we simulated a comprehensive range of scenarios. Instead, we showed that even when the time-span of ΔT is over a billion years, the rate of protein evolution at the duplication point (dD0) needed to be over 40 times greater than any rate ever observed for D1 and D2, which decreased exponentially during the Archean and stabilized at current rates during the Proterozoic. Thus, the smaller the value of ΔT, the faster the rate at dD0, with the rate increasing following a power law function and reaching unrealistic values even when ΔT is still in the order of several hundred million years [19]. It was unclear then if such patterns of molecular evolution were unique to the core subunits of PSII or whether other systems have experienced similar evolutionary trajectories.

Here, to help in understanding the evolution of oxygenic photosynthesis, Cyanobacteria and MSV as a function of time, we compared the age of duplication leading to the RC antenna subunits, CP43 and CP47, with several well-defined ancient and more recent events. These include the duplication of the core catalytic subunits of ATP synthase, a very ancient event generally accepted to have occurred before the last universal common ancestor (LUCA) [[21], [22], [23], [24], [25]]; and the evolution of RNA polymerase catalytic subunit β (RpoB) and ribosomal proteins, which are universally conserved and widely accepted to have originated before the LUCA [[26], [27], [28], [29]]. We further constrain our analysis using in silico ancestral sequence reconstruction of PSII and through strict structural and functional rationales. We show that the core subunits of PSII show patterns of molecular evolution that are usually associated with some of the oldest transitions in the evolution of life. We also show that all events leading from a primordial homodimeric photosystem to Cyanobacteria's heterodimeric PSII can be logically reconstructed.

2.1. Phylogenetic overview

The phylogenies of CP43 and CP47 show that there is a much greater diversity of CP43 and CP43-derived subunits than CP47 (Fig. 1). This difference is the result of a greater number of gene duplication in CP43 than CP47 and mirrors the evolution of D1 compared with D2 [2], in which D1 has undergone more duplication events than D2. CP43 can be divided into two major groups: those that are assembled into PSII and can bind the Mn₄CaO₅ cluster, and those which have evolved to be used only as light harvesting complexes [30,31], known as chlorophyll binding proteins (CBP). The CBP are characterized by the loss of the extrinsic loop between the 5th and 6th transmembrane helices, where the ligands to the cluster are located (Fig. 1b). This large extrinsic loop is found in both CP43 and CP47 and interacts directly with the electron donor side of PSII, within D1 and D2 respectively. The unrooted tree of CP43 is consistent with CBP having a single origin likely occurring before the MRCA of Cyanobacteria (Supplementary Fig. S1) but have undergone an extensive duplication-driven diversification process. It also mirrors the evolution of D1 in that duplications appear to have occurred before the MRCA of Cyanobacteria [2]. The earliest of these D1 duplications also led to variants that lack the capacity to bind the Mn₄CaO₅ cluster [2,32], but are likely used in other supporting functions. Most notably, during chlorophyll f synthesis [33] in the far-red light acclimation response (FaRLiP) [34].

CP43 and CP47 are also distantly related to the antenna domain of cyanobacterial PSI and other type I RCs of phototrophic Chlorobia, Acidobacteriota and Heliobacteria (Supplementary Fig. S2). We found that the level of sequence identity between any two type I RC proteins is always greater than between a type I RC protein and CP43/CP47 (Supplementary Table S1). Therefore, the distance between CP43/CP47 and other type I antenna domains is the largest distance in the molecular evolution of RC proteins after that between type I core and type II RC core domains. The phylogenetic relationships between type I and type II RCs have been reviewed in detail before [20,35,36].

The phylogenetic distribution of Alpha and Beta subunits of the F-type ATP synthase showed that all Cyanobacteria have an F-type ATP synthase, and a fewer number of strains have an additional Na⁺-translocating ATPase (N-ATPase) of the bacterial F-type, as had been reported before [37]. We found that MSV have a standard F-type ATP synthase (Supplementary Fig. S3), but some N-ATPase Alpha and Beta sequences were also found in Vampirovibrionia and Sericytochromatia datasets, but not in Margulisbacteria. In this study we focused on the standard F-type ATP synthase of Cyanobacteria for further analysis.

We also constructed a phylogeny of bacterial RNA polymerase subunit β (RpoB) for divergence time estimation. This focused on Cyanobacteria and MSV, as well as phyla with known phototrophic representatives and included Thermotogae and Aquificae as potential outgroups (Supplementary Fig. S4). The tree was largely consistent with previous observed relationships between the selected groups [38], within Cyanobacteria and MSV, and within other phototrophs and their non-phototrophic relatives [39,40]. The only exception was Aquificae, which branched as a sister clade to Acidobacteriota, a feature that had been reported before for RpoB [28], and likely represents an ancient horizontal gene transfer event.

2.2. Rates of evolution

2.3. Species divergence

To understand the evolution of MSV relative to Cyanobacteria we wished to apply a molecular clock to a system where the calculated rates could be compared to observed rates, as determined by distances between species of known divergence times or at similar taxonomic ranks. We found RpoB to be suitable for this because it has been inherited vertically with few instances of horizontal gene transfer and had enough signal to resolve known phylogenetic relationships between and within clades. We implemented a set of 12 calibrations across bacteria, including two calibrations on Margulisbacteria and two in Vampirovibrionia with the aim of covering both slower and faster evolving lineages. The following results are based on an autocorrelated log normal molecular clock using CAT+Γ, a root constrained with a broad interval ranging from between 4.52 and 3.41 Ga, and as described in Materials and Methods and Supplementary Text S3 (Fig. 5). We found this to perform well and provided results comparable to other independent studies that did not combine a full set of MSV sequences and other clades with phototrophs in a single tree (Table 2). Nonetheless, a pipeline of sensitivity experiments tested the dependency of these results on models and prior assumptions: these are shown and described in Supplementary Figs. S6 and S7.

The root of the tree (divergence of Thermotogae) was timed at 3.64 Ga (95% CI: 3.42–4.11 Ga) and the divergence of Cyanobacteria at 2.74 Ga (95% CI: 2.46–3.12 Ga). Thus, the span of time between the mean age of the root of the bacterial tree and the mean age of the MRCA of Cyanobacteria was calculated to be 0.89 Ga and that between Vampirovibrionia and the latter was found to be 0.44 Ga (Figs. 4f and 5). This result is consistent with previous studies using different rationales, datasets and calibrations [11,12].

We also noted an exponential decrease in the rates of evolution of RpoB through the Archean, which stabilized at current levels in the Proterozoic (Fig. 4f). The rate at the root node was calculated to be 2.37 ± 0.45 δ Ga⁻¹ and the average rate of evolution of RpoB during the Proterozoic was found to be 0.19 ± 0.06 δ Ga⁻¹. The average rate of cyanobacterial RpoB was 0.14 ± 0.04 δ Ga⁻¹; for Margulisbacteria was 0.44 ± 0.17 δ Ga⁻¹, and for Vampirovibrionia 0.19 ± 0.05 δ Ga⁻¹: about 3.1× and 1.3× the mean cyanobacterial rate, respectively. These rates agree reasonably well with the observed distances (Supplementary Table S2), further indicating that the calibrations used in these clades performed well. Nevertheless, we suspect that the values for MSV represent underestimations of the true rates of evolution (slower than they should be), as some of the clades that include symbionts still appear much older than anticipated from their hosts. This is probably due to the relaxed nature of molecular clocks, which tends to smooth rates out [50]. For example, Gastranaerophilales were timed to have originated long before the emergence of animals (95% CI: 1.91–2.74 Ga). This dichotomy in the rates was also detected for Rickettsidae (0.40 ± 0.11 δ Ga⁻¹) made up of symbionts and parasites, when compared to Caulobacteridae (0.17 ± 0.05 δ Ga⁻¹) of the Alphaproteobacteria, which included free-living and phototrophic strains. We also noted the same potential underestimation for many strains of Rickettsidae, which appeared to be much older than their symbiotic associations would suggest, despite the provided constraints.

A more complex, but commonly used model like CAT+GTR+Γ implementing a birth-death prior with ‘soft bounds’ on the calibrations, resulted in rates that were smoothed out, and which translated into spread-out divergence times with Margulisbacteria and Vampirovibrionia evolving at 1.9× and 0.7× times the cyanobacterial rates, respectively (Supplementary Fig. S7, model n to p). These rate effects are thus translated into a very late Mesoproterozoic age for Cyanobacteria, and a relatively older divergence time for Vampirovibrionia: results that replicate those presented in ref. [46].

To investigate the effect of the age of the root on the divergence time of MSV and Cyanobacteria we also varied the root prior from 3.2 to 4.4 Ga (Supplementary Fig. S6). We noted that regardless of the time of origin of Bacteria (approximated by the divergence of Thermotogae in our analysis), a substantially faster rate is required during the earliest diversification events, decreasing through the Archean and stabilizing in the Proterozoic. This matches well the patterns of evolution of ATP synthase and PSII core subunits as shown in the previous section.

We then compared the above RpoB molecular clock result with a different clock that included a set of 112 diverse sequences from Archaea, in addition to the sequences from Thermotogae, MSV, and Cyanobacteria, but removing all other bacterial phyla (Fig. 6a). We found that the calculated average rate of evolution of bacterial RpoB during the Proterozoic was slower (0.09 ± 0.03 δ Ga⁻¹) than in the absence of archaeal sequences (0.19 ± 0.06 δ Ga⁻¹), resulting in overall older mean ages (see Fig. 4g). However, the rate at the Bacteria/Archaea divergence point was 6.87 ± 1.17 δ Ga⁻¹, similar to the rate for dAB and dCP, requiring therefore an exponential decrease in the rates similar to that observed for ATP synthase and the PSII core subunits. Notably, this rate and exponential decrease is associated with a large span of time between the mean estimated ages of the LUCA and the MRCA of Cyanobacteria, calculated in this tree to be 1.21 Ga. The span between Vampirovibrionia and Cyanobacteria was found to be 0.46 Ga (Fig. 6a).

Fig. 5 also highlights that the none of the most recent common ancestors of the groups containing anoxygenic phototrophs nor their divergence from their closest non-phototrophic relatives appears to be older than one of the oldest and best accepted geochemical pieces of evidence for photosynthesis at 3.41 Ga [51] (Table 2). For example, the MRCA of Heliobacteria or of phototrophic Chlorobia, containing homodimeric type I RCs, are likely to have existed only after the GOE, even allowing for large uncertainties in the calculations. These clades have traditionally been described as harbouring primitive forms of photosynthesis.

Finally, we performed a similar molecular clock analysis on a subset of concatenated ribosomal proteins published by Hug et al. [39] that included Archaea, Thermotogae, Margulisbacteria, Vampirovibrionia and Cyanobacteria. Even though this dataset generated younger ages compared to RpoB (see Table 2 and Fig. 6), a similar exponential decrease in the rates was observed with a rate at the Bacteria/Archaea divergence point measured at 4.62 ± 0.71 δ Ga⁻¹ and an average rate of bacterial ribosomal protein evolution of 0.30 ± 0.07 δ Ga⁻¹ (Fig. 4h). These rates were associated with a span of time between the mean estimated ages of the LUCA and the MRCA of Cyanobacteria of 1.45 Ga; and between Vampirovibrionia and Cyanobacteria 0.54 Ga (Fig. 6b).

2.4. Structural analysis

A fundamental premise of our investigation is that water oxidation started before the duplication of D1 and D2, and CP43 and CP47. The rationale behind this premise has been laid out before [18,52], and more extensively recently [19]. If this rationale is correct, it raises the question of how the CP47/D2 side of the RC lost its capacity to carry out water-splitting catalysis.

3.1. Origin of oxygenic photosynthesis and rates of evolution

This work was not explicitly aimed at providing an exact time of origin for the LUCA, the MRCA of Cyanobacteria, or the origin of photosynthesis. The use of different datasets, calibration strategies, chosen clock parameters, and researchers' preferences can result in widely varying calculated rates, which can translate into disparate estimated ages, see for example [46,57]. Instead, rather than arguing in favour or against a particular age or methodology, our objective was to contrast the evolution of oxygenic photosynthesis with the evolution of enzymes whose origin is less controversial. The duplication of ATP synthase's ancestral catalytic subunit, and the archaeal/bacterial divergence of RNA polymerase and the ribosome, are some of the oldest evolutionary events known in biology [[21], [22], [23], [24], [25], [26], [27], [28], [29]]. When taken on their own, their phylogenies are often interpreted as evidence of the earliest origin. We have shown here and in our previous work [19] that PSII has patterns of molecular evolution that closely parallel those of these most ancient systems. These patterns do not emerge from the application of any particular molecular clock approach or model of rate change, but from the inherently long phylogenetic distance that separates Alpha and Beta, archaeal and bacterial RNA polymerase and ribosomes, or the core subunits of PSII, in combination with the slow rates of evolution that these enzymes have featured through their multibillion-year diversification process. These results have profound implications because it cannot now be taken for granted that there was ever a long period of time between the origin of life and the origin of anoxygenic photosynthesis, followed by another long period of time between the origin of anoxygenic photosynthesis and the origin of oxygenic photosynthesis.

The exponential decrease in the rates of evolution observed in the studied systems, even when the span of time between the ancestral duplications (or the LUCA) and the MRCA of Cyanobacteria was considered to be well over a billion years, is consistent with our assessment that a large ΔT for the evolution of the core subunits of ATP synthase and PSII cannot be explained entirely by duplication-driven effects. Instead, we speculate that these effects are the result of the faster rates of evolution that are expected to have occurred during the earliest history of life [[58], [59], [60], [61], [62], [63]]. It is worth highlighting that ATP synthase, RNA polymerase, the ribosome, and the photosystems, are all complex molecular machines, with crucial functions and under strict regulation. These features largely explain their slow rates of evolution and the high degree of sequence (structural and functional) conservation through geological time. A similar level of complexity, however, can be traced back to the last common ancestor of each one of these molecular systems regardless of how ancient they truly are. Now, given that the rates of evolution of the core of PSII have remained slower than those of ATP synthase for billions of years, it could imply that the duplications of the core of PSII occurred before the duplication leading to Alpha and Beta. In consequence, to argue that oxygenic photosynthesis is a late evolutionary innovation relative to the origin of life, one must first demonstrate that the rates of protein evolution of PSII and ATP synthase catalytic core subunits, RNA polymerase, and the ribosome, have not remained proportional to each other during the Archean. That is to suggest that PSII—exclusively—experienced unprecedentedly faster rates of evolution during the heterodimerization process, and then, the rates stopped abruptly with the MRCA of Cyanobacteria. One could potentially argue that the origin of water oxidation itself could account for unprecedentedly faster rates. However, a decrease in ΔT of about 60% (from 1.1 to 0.46 Ga, for example) would result in a 30-fold additional increase in the rates at the earliest stages of diversification, resulting in photosystems that would be evolving at rates that surpass those of the fastest evolving peptide toxins (Fig. 4d and Supplementary Text S1). Furthermore, it should be noted that a ΔT of over a billion years already includes a period of extremely rapid evolution at the origin of all these ancient systems.

3.2. Diversification of bacteria

It has been postulated before that the diversification of the major phyla of Bacteria occurred very rapidly, starting at about 3.4 Ga and peaking at about 3.2 Ga, in what has been dubbed the Archean Genetic Expansion [64], but see also [46,65]. A recent study suggested that the major groups of Bacteria and Archaea diversified rapidly after the LUCA and hypothesized that the long distance between archaeal and bacterial ribosomal proteins could be attributed to fast rates of ribosome evolution during their divergence [66], but see also [67,68]. Our analyses suggest that the more explosive the diversification of prokaryotes, the greater the chance that photosynthetic water-splitting is an ancestral trait of life. Yet, if phototrophic communities—whether oxygenic or not—already existed 3.2 to 3.4 Ga ago, as it is supported by the geochemical record [51,69], then the earliest bacteria were likely photosynthetic. This is consistent with the evolution of RC proteins, which suggests that the structural and functional specialization that led to the two photosystem types, antedated the diversification processes leading to the known phyla containing photosynthetic bacteria [35]. It also explains why none of the groups of extant photosynthetic bacteria appear to be older than the earliest geochemical evidence for photosynthesis or hardly older than the GOE. The presented data is also consistent with recent studies of the oxygenation of the planet, which suggests that even if photosynthetic O₂ evolution started as early as the oldest rocks, the properties of early Earth biogeochemistry would have maintained very low concentrations of O₂ over the Archean [[70], [71], [72]].

3.3. Photosystem structural constraints

We have shown that the phylogenetic distance between CP43/CP47 and other type I RC proteins is the largest distance after that between type I and type II RC proteins (Supplementary Fig. S2). The antenna domain extends to the 8th helix, both in type I RCs and in PSII, with the latter retaining one antenna chlorophyll in the equivalent 8th helix (marked Z in Fig. 8) and substantial sequence identity around this chlorophyll's binding site, as shown before [35,73]. These conserved features indicate that at no time during the evolution of PSII was it devoid of core antenna proteins. In other words, CP43/CP47 are a descendant of the ancestral core antenna of type II photosystems. Both structural and phylogenetic evidence is in agreement and indicates that the ancestral type II RC, at the dawn of photosynthesis, was architecturally like water-splitting PSII.

Sequence reconstruction of the ancestral subunit to D1 and D2 is consistent with the existence of a highly oxidizing homodimeric photosystem, though transient and short-lived [19], that could split water in either side of the RC [52]. The structural comparisons between CP43/D1 and CP47/D2 suggest a mechanism for heterodimerization and loss of catalysis on one side that accounts for all ligands. These include direct amino acid substitutions, the loop swap within the extrinsic region of the ancestral protein to CP47, and the gene overlap between psbC (CP43) and psbD (D2). It would be difficult to reconcile these unique structural and genetic features with a scenario in which PSII evolved water oxidation at a late stage, or starting as a purple anoxygenic type II RC, once the heterodimerization process was well underway or completed, or in the absence of core antenna domains, given that these interact directly with the electron donor side of PSII, in a manner strikingly similar to that of the Ca-site in the homodimeric type I RC of the Firmicutes [20]. A conserved Ca-binding site has also been recently confirmed for the RC of the Chlorobia [74], as predicted by Cardona and Rutherford [20]. What is more, these structural and functional features, together with the ASR analysis (Supplementary Fig. S8 and S9) suggest that the atypical forms of D1, lacking ligands to the Mn₄CaO₅ cluster, such as the chlorophyll f synthase [33,75], or the “rogue” or “sentinel” D1 [32,76], diverged from forms of D1 that were able to support water oxidation, but that could antedate the MRCA of Cyanobacteria [2].

We have calculated that (oxygenic) PSII has experienced the slowest rates of evolution of all photosystems, with the core of the anoxygenic type II RC evolving on average about five times faster than the core of PSII [19]. That faster rate has led to conspicuous structural changes of the anoxygenic RC relative to PSII and type I photosystems. The outcome of these differences in the rates are not only visually apparent (Fig. 8), but also the anoxygenic type II RC among all photosystems has experienced the greatest erosion of sequence and structural symmetry and even complete loss of structural domains [20,77]. These differences in the rates and associated structural differences appear inconsistent with a scenario in which PSII experienced unprecedentedly fast rates of evolution. Counterintuitively, because PSII is the slowest evolving of all photosystems and among the slowest evolving enzymes known; and because that has been the case for a couple of billions of years, if not much longer, it follows then that PSII is the most likely RC to have changed the least since the origin of photosynthesis.

3.4. Origin of homodimeric water-splitting photosystems

Compelling scenarios that discuss the origin of water oxidation propose Mn-oxidizing photosynthesis as a key transitional stage [18,[78], [79], [80], [81], [82]]. Few of these explicitly state whether this transition should have occurred before or after the duplications that led to CP43/D1 and CP47/D2. Cardona et al. [2] correlated the phylogenetic branching pattern of the D1 tree to transitional stages from a Mn^II-oxidizing photosystem towards the emergence of the full Mn₄CaO₅ cluster. This correlation was later revisited based on the extended analysis presented in ref. [19], in line with previous suggestions [52], and it is indeed not now supported by ASR analysis as further indicated in this work.

In another scenario, Fischer et al. [78] hypothesized that a Mn^II-oxidizing photosystem occurred at the homodimeric stage and considered that the D1 and D2 duplication enabled water oxidation. Fischer et al. [78] also speculated using geochemical evidence from Mn deposits in the ~2.41 Ga Koegas Subgroup of the Kaapvaal Craton of South Africa [83], that the duplication occurred just before the GOE [78]. The data we present here can be consistent with a Mn-oxidizing transitional stage, but does not support an early Paleoproterozoic ancestral photosystem core duplication event. An alternative speculative scenario that could reconcile the Koegas Subgroup deposits with phylogenetics and other geochemical data [84], is that Mn oxidation in this locality was catalysed by a PSII using an atypical D1 form similar or ancestral to the rogue D1 or the ChlF proteins as recently suggested by Chernev et al. [82], but derived from water-splitting variants. It is also plausible that this occurred as a transitional stage towards the emergence of heterotrophy in cyanobacterial ancestors—even perhaps in any of the direct ancestors of MSV—under the selection pressure of other competing lineages of photosynthetic Cyanobacteria, linked to ecological changes occurring around the GOE [85,86].

The data presented here and in [19] suggest that water oxidation antedated the core duplications. We have taken this evidence before at face value to envision a water-splitting homodimeric photosystem that could perform catalysis on both branches of the RC. Such a system, however short-lived or inefficient, raises questions about the actual mechanism of catalysis [52]. For example, Fischer et al. considered the idea of a homodimeric water-splitting photosystem “highly unlikely” [87], because it would “require the coordination of eight electron transfers”. This thinking implicitly assumes that water oxidation can only occur through a mechanism that is identical to that of PSII, extrapolated to both sides of the RC, at the same time. This is but one of several scenarios that are consistent with water oxidation prior to core duplication. For example, partial oxidation of water to hydrogen peroxide is a plausible evolutionary transition before the full catalytic cycle of PSII was completed. Hydrogen peroxide release from partially oxidized water can occur in the lower oxidized states of the catalytic cycle in centres that have been perturbed by depletion of Ca²⁺ or Cl⁻ [88,89], removal of the extrinsic polypeptides [90], or by heating a sample [91]. Alternatively, a homodimeric system does not necessarily imply entirely homodimeric function. Like PSII, oxidation of Mn on one side of the RC could create asymmetric electrostatic effects on the photochemical pigments, P [92], and the quinone acceptor [93] (Fig. 8). In addition, the rate of Y_Z oxidation by P⁺ accelerates by two orders of magnitude in the presence of an assembled cluster [94]. This acquired asymmetry in a homodimeric photosystem upon Mn oxidation could shift redox equilibria increasing the probability of further oxidation events on one side relative to the other. It is conceivable that a catalytic site could turn over before the other is fully assembled, with the unassembled site assuming a supporting role similar to Y_D on D2 [95,96].

3.5. Duplications

Before natural selection or drift can drive the heterodimerization of the photosystem, a gene duplication must occur first. The attention can thus be shifted away from asking if the duplication occurred before or after the origin of water oxidation, to asking why the RC gene duplicated to begin with. This is crucial because within the described diversity of photosynthetic organisms, all known RC gene duplications have occurred exclusively in the context of oxygenic photosynthesis. In particular, but not exclusively, duplications of the subunits that allow catalysis, D1 and CP43. In other words, and to the best of our knowledge, there are no known gene duplication events of PufL, PufM, PshA or any version of PscA described for any known anoxygenic phototroph (Fig. 9). Therefore, based on available observations, the only driving force known to lead to RC gene duplications is water oxidation to oxygen, and the subsequent need for sustained repair and protection against ROS-mediated damage. Duplicated subunits can then drift away from their main functions and become the canvas of novel adaptations in a process that has never stopped [2]. This is spectacularly exemplified in the evolution of FaRLiP [97], as chlorophyll f synthesis was likely enabled by a drifting D1 variant accumulating just two amino acid substitutions at the interface with CP43 [98].

Another conspicuous duplication is that leading to the heterodimeric core of PSI, which has been discussed to have occurred as an adaptation to oxygenic photosynthesis [[99], [100], [101], [102]]. In fact, Orf et al. [103] recently suggested that PSI started to accumulate adaptations to avoid singlet oxygen formation even before the duplication of the core. Furthermore, we have recently shown that the duplication leading to L and M, likely post-dated the evolution of oxygenic photosynthesis [19]. We have also discussed how the heterodimerization of the anoxygenic type II RC core could have occurred under pressure to minimize ROS-mediated damage [19].

The only other discussed, yet more ambiguous duplication is that which gave rise to type I and type II RC proteins, unanimously and implicitly assumed to have occurred before the origin of water oxidation [18,[104], [105], [106], [107]]. While we find compelling some of the rationales supporting a duplication at the origin of type I and II RCs, Cardona [77] raised issues with the implicit assumption because, like Mn-oxidizing photosynthesis without water oxidation, it invokes a transitional form of photosynthesis for which no evidence has been found within the known diversity of anoxygenic phototrophs. In this case, the emergence of anoxygenic photosynthesis using both RC types in series. Both phylogenetic and structural evidence converge towards a scenario in which photosynthetic water-splitting started at an early stage during the evolution of life and long before the rise of what we understand today as Cyanobacteria. We have shown evidence suggesting that the ancestral type II RC was similar to PSII, which is consistent with the hypothesis that the type I and type II split, likely enabled by a duplication event, indeed occurred in the context of the establishment of oxygenic photosynthesis [73,77], as illustrated in Fig. 9. We argue that this perspective offers more explanatory power than conventional scenarios. Not only it explains the structural and functional characteristics of PSII, but it also explains the unique coordination sphere of the Mn₄CaO₅ cluster in a manner consistent with phylogenetic relationships. Distinctly, this perspective does not require the involvement of speculative forms of photosynthesis. More importantly yet, it also provides a mechanism for the generation of genetic diversity via serial gene duplications from where photosystem innovation can occur, and, for which ample evidence exists within the genomes of most Cyanobacteria today.

We think it is plausible that there was never a discrete origin of photosynthesis, but that the process may trace back to abiogenic photochemical reactions, some of which may have resulted in the oxidation of water, and at the interface of nascent membranes, membrane proteins, photoactive tetrapyrroles and other inorganic cofactors: much in the same way that ribosomes may have originated at the interface of nascent genetics and protein synthesis [108]. A photosynthetic origin of life is not a new idea [[109], [110], [111]] and abiotic photosynthesis-like chemistry has been recently proposed to have occurred at Gale Crater on Mars [112], and to occur even on Earth [113].

4.1. Sequence alignments and phylogenetic analysis

The first dataset was retrieved on the 31st of October 2017 to initiate this project. It included a total of 1389 type I RC, CP43 and CP47 protein sequences from the NCBI refseq database using BLAST. This dataset did not contain CBP proteins. A second dataset was retrieved on the 10th of October 2018 that included 392 CP43, 465 CBP proteins, and 375 CP47 subunits. 40 CP43 and 40 CP47 sequences from a diverse set of photosynthetic eukaryotes were selected manually and included. In addition, a selection of cyanobacterial and plastid CP43, CP47, AtpA (Alpha), AtpB (Beta), and FtsH were manually selected for an analysis of the rates of evolution as described below and in Supplementary Text S2.

A dataset of Alpha and Beta subunits belonging to cyanobacterial F-type ATP synthase were retrieved from the NCBI refseq on the 31st of August 2019. 507 and 529 cyanobacterial Alpha and Beta sequences were obtained respectively. We retrieved Alpha and Beta homologous for Margulisbacteria (19 and 18 sequences respectively), Sericytochromatia (4 and 2), and Vampirovibrionia (66 and 49) using AnnoTree [114] and searching with the KEGG codes K02111 (F-type Alpha) and K02112 (F-type Beta). A total of 111 AtpA (subunit A) and 176 AtpB (subunit B) from archaeal V-type ATP synthase were retrieved using the sequences from Methanocaldococcus jannaschii as BLAST queries.

On the same date (31st of August 2019), a dataset of 366 bacterial RNA polymerase subunit β (RpoB) were collected from the NCBI refseq. We focused on Cyanobacteria (65 sequences) and other phyla known to contain phototrophic representatives, as well as for their potential for allocating calibration points as described below. These included sequences from Firmicutes (32), Chloroflexota (32), Proteobacteria (102), Acidobacteriota (34), Chlorobia (26), Gemmatimonadetes (3), Aquifecae (12) and Thermotogae (24). Sequences for Candidatus Eremiobacterota (2) recently reported to include phototrophic representatives [115], Margulisbacteria (13), Sericytochromatia (2), and Vampirovibrionia (11) were obtained from AnnoTree using KEGG code K03043 as query. In addition, three extra margulisbacterial RpoB sequences were collected: from Termititenax persephonae and Termititenax aidoneus retrieved from the refseq database, and from margulisbacterium Candidatus Ruthmannia eludens obtained from https://www.ebi.ac.uk/ena/data/view/PRJEB30343 [7].

A second dataset of RpoB subunits containing only sequences from Cyanobacteria and MSV, in addition to sequences from Archaea was also used. To retrieve the archaeal sequences the subunit B′ (RpoB1) of Methanocaldococcus jannaschii was used as a BLAST query. A total 213 sequences across the entire diversity of Archaea were obtained. Two version of these were observed, those with split B (RpoB1 and RpoB2) and those with full-length B subunits: from the latter, 112 full-length B subunits were selected for molecular clock analysis. Finally, we also prepared a dataset of concatenated ribosomal proteins containing 2596 aligned sites. This dataset was obtained from an independent study [39] and we selected a total of 157 sequences: 56 cyanobacterial, 14 sequences from Vampirovibrionia, 9 from Margulisbacteria, 9 from Thermotogae and 78 from Archaea.

Sequences were aligned with Clustal Omega using 5 combined guided trees and Hidden Markov Model Iterations [116]. Given that RpoB sequences are known to contain many clade specific indels [28], we further processed this particular dataset using Gblocks [117] to remove these indels, allowing smaller final blocks, gap positions within the final blocks, and less strict flanking positions. This procedure left a total of 903 well-aligned sites for the dataset with only bacterial sequences and 788 for the dataset with archaeal sequences. The dataset of concatenated ribosomal proteins was made available by the authors aligned: after curating for unused taxa, remaining gap-only sites were removed without further processing.

Maximum Likelihood phylogenetic analysis was performed with the PhyML online service using the Smart Model Selection mode implementing the Bayesian Information Criterion for parameter selection [118,119]. Tree searching operations were calculated with the Nearest Neighbour Interchange model. Support was computed using the average likelihood ratio test [120]. Trees were visualized with Dendroscope V3.5.9 [121].

4.2. Distance estimation

Distance estimation was performed as a straightforward and intuitive approach to detect possible variations in the rate of sequence change. Distance trees were plotted using BioNJ [122] as implemented in Seaview V4.7 [123] using observed distances and 100 bootstrap replicates. Within-group mean distances were calculated for RpoB, Alpha, Beta, and the ribosomal dataset using three different substitution models as implemented in the package MEGA-X [124]: no. of differences, a Poisson model, and a JTT model. A gamma distribution to model rates among sites was used with a parameter of 1.00 and 500 bootstrap replicates.

To compare the level of sequence identity between RC proteins, two datasets of 10 random amino acid sequences were generated using the Sequence Manipulation Suit [125]. The datasets contained sequences of 350 and 750 residues. These were independently aligned as described above, resulting in 45 pairwise sequence identity comparisons for each dataset. These random sequence datasets were used as a rough minimum threshold of identity. Alignments of RC proteins were generated using three representative sequences spanning known diversity. Cyanobacterial CP43, CP47, standard D1 and D2 sequences were from Gloeobacter violaceus, Stanieria cyanosphaera, and Nostoc sp. PCC 7120; Heliobacterial PshA from Heliobacterium modesticaldum, Heliobacillus mobilis, and Heliorestis convoluta; PscA from Chlorobium tepidum, Prosthecochloris aestuarii, Chlorobium sp. GBchlB, Chloracidobacterium thermophilum and Chloracidobacterium sp. CP2-5A; Proteobacterial L and M from Methylobacterium sp. 88A, Roseivivax halotolerans, and Blastochloris sulfoviridis; and L and M from Chloroflexus sp. Y-400-fl, Roseiflexus castenholzii, and Oscillochloris trichoides.

4.3. Rates and molecular clocks

To measure the rates of amino acid substitution of PSII, ATP synthase and FtsH subunits under varying evolutionary scenarios we followed a methodology modified from Cardona et al. [19] and as detailed in Supplementary Text S2 Quantification of rates of evolution, Rationale and Method. For this experiment, all used rates of amino acid substitutions and divergence times were calculated with Phylobayes 3.3. We applied a log normal autocorrelated model under the CAT+Γ non-parametric model of amino acid substitution and a uniform distribution of equilibrium frequencies. We preferred a CAT model instead of a CAT+GTR as the latter only outperforms the former on sequence alignments of over 1000 sites and it is also much less computationally expensive [126]. Four discrete categories for the gamma distribution were used and four chains were executed in parallel through all experiments carried out in this study. The rates of evolution were retrieved from the output files of Phylobayes and expressed as amino acid substitutions per site per Ga. These rates are calculated by the software as described by the developers elsewhere [127,128].

To have a closer understanding of the spans of time between Cyanobacteria and MSV and their associated rates of evolution under different evolutionary scenarios, we applied a molecular clock to the phylogeny of RpoB sequences described above. We implemented 12 calibrations and several root priors, which are described and illustrated in Supplementary Text S3 Calibrating the RpoB and ribosomal proteins phylogenies. Molecular clocks were calculated as described above. We also compared a range of models and parameters including an uncorrelated gamma model of rates of amino acid substitution, +GTR, and the application of birth-death priors combined with soft bounds. When soft bounds were used a 2.5% tail probability was allowed to fall outside the minimum and maximum boundary, or 5% in the case of a single boundary. The full set of comparisons is presented in Supplementary Figs. S6 and S7.

4.4. Ancestral sequence reconstruction

Ancestral sequence reconstruction (ASR) of D1 and D2 amino acid sequences was carried out with a dataset collected on the 17th of November 2017. Duplicates and partial sequences were removed, leaving 755 D1 and 248 D2 sequences. CD-HIT [129] was used to remove sequences with greater than 92% sequence identity to create a representative sample. The L and M RC subunits from 5 strains of Proteobacteria were used as outgroup. The final alignment did not include the atypical variant D1 sequence from Gloeobacter kilaueensis (NCBI accession AGY58976.1) as this showed an unstable phylogenetic position in this dataset. Maximum likelihood trees used as input for ASR were computed with PhyML using Smart Model Selection [119]. The LG substitution model with observed amino acid frequencies and four gamma rate categories exhibited the best log likelihood (LG+Γ+F) (Supplementary Table S7). We used the top four models for tree reconstruction (LG+Γ+F, LG+Γ+I+F, LG+Γ and LG+Γ+I) in addition to another tree computed using the WAG substitution model with observed amino acid frequencies and four gamma rate categories (WAG+Γ+F). These trees were used as input trees to calculate maximum likelihood ancestral states at each site for the node corresponding to the homodimeric reaction protein, D0. Three ASR programs were used for the reconstructions: FastML [130], Lazarus [131] (a set of Python scripts which wraps PAML [132]) and MEGA-7 [133]. The substitution model used by all three programs corresponded to the substitution model used by PhyML to generate the specific input tree. Branch lengths of the trees were fixed as these were already estimated for the input trees. In FastML, a maximum likelihood method of indel reconstruction using a probability cut-off of 0.7 was used. In MEGA-7, all sites were used for analysis with no branch swap filter. In Lazarus, the ‘—gapcorrect’ option was used to parsimoniously place indels.

4.5. Structural analysis

The following crystal structures were used in this work: the crystal structure of PSII from Thermosynechococcus vulcanus and Cyanidium caldarium, PDB ID: 3wu2 and 4yuu respectively [14,134]; the anoxygenic type II RC of Thermochromatium tepidum, PDB ID: 5y5s [135]; the homodimeric type I RC from Heliobacterium modesticaldum, 8v5k [136]; PSI from Thermosynechococcus elongatus, PDB ID: 1jb0 [137]; the cryo-EM IsiA structure from Synechocystis sp. PCC 6803 [138]. Structures were visualized using Pymol™ V. 1.8.2.2 (Schrodinger, LLC) and structural overlaps were carried out with the CEAlign plugin [139].

Supplementary material: sequence alignments and trees

Supplementary figures and text

Supplementary tables

• 1.Sánchez-Baracaldo P., Cardona T. On the origin of oxygenic photosynthesis and Cyanobacteria. New Phytol. 2020;225:1440–1446. doi: 10.1111/nph.16249. [DOI] [PubMed] [Google Scholar]
• 2.Cardona T., Murray J.W., Rutherford A.W. Origin and evolution of water oxidation before the last common ancestor of the Cyanobacteria. Mol. Biol. Evol. 2015;32:1310–1328. doi: 10.1093/molbev/msv024. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 3.Di Rienzi S.C., Sharon I., Wrighton K.C., Koren O., Hug L.A., Thomas B.C., Goodrich J.K., Bell J.T., Spector T.D., Banfield J.F., Ley R.E. The human gut and groundwater harbor non-photosynthetic bacteria belonging to a new candidate phylum sibling to Cyanobacteria. Elife. 2013;2 doi: 10.7554/eLife.01102. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 4.Soo R.M., Hemp J., Hugenholtz P. Evolution of photosynthesis and aerobic respiration in the cyanobacteria. Free Radic. Biol. Med. 2019;140:200–205. doi: 10.1016/j.freeradbiomed.2019.03.029. [DOI] [PubMed] [Google Scholar]
• 5.Soo R.M., Hemp J., Parks D.H., Fischer W.W., Hugenholtz P. On the origins of oxygenic photosynthesis and aerobic respiration in Cyanobacteria. Science. 2017;355:1436–1440. doi: 10.1126/science.aal3794. [DOI] [PubMed] [Google Scholar]
• 6.Anantharaman K., Brown C.T., Hug L.A., Sharon I., Castelle C.J., Probst A.J., Thomas B.C., Singh A., Wilkins M.J., Karaoz U., Brodie E.L., Williams K.H., Hubbard S.S., Banfield J.F. Thousands of microbial genomes shed light on interconnected biogeochemical processes in an aquifer system. Nat. Commun. 2016;7:13219. doi: 10.1038/ncomms13219. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 7.Gruber-Vodicka H.R., Leisch N., Kleiner M., Hinzke T., Liebeke M., McFall-Ngai M., Hadfield M.G., Dubilier N. Two intracellular and cell type-specific bacterial symbionts in the placozoan Trichoplax H2. Nat. Microbiol. 2019;4:1465–1474. doi: 10.1038/s41564-019-0475-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 8.Utami Y.D., Kuwahara H., Igai K., Murakami T., Sugaya K., Morikawa T., Nagura Y., Yuki M., Deevong P., Inoue T., Kihara K., Lo N., Yamada A., Ohkuma M., Hongoh Y. Genome analyses of uncultured TG2/ZB3 bacteria in ‘Margulisbacteria’ specifically attached to ectosymbiotic spirochetes of protists in the termite gut. ISME J. 2019;13:455–467. doi: 10.1038/s41396-018-0297-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 9.Soo R.M., Woodcroft B.J., Parks D.H., Tyson G.W., Hugenholtz P. Back from the dead; the curious tale of the predatory cyanobacterium Vampirovibrio chlorellavorus. PeerJ. 2015;3 doi: 10.7717/peerj.968. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 10.Carnevali P.B.M., Schulz F., Castelle C.J., Kantor R.S., Shih P.M., Sharon I., Santini J.M., Olm M.R., Amano Y., Thomas B.C., Anantharaman K., Burstein D., Becraft E.D., Stepanauskas R., Woyke T., Banfield J.F. Hydrogen-based metabolism as an ancestral trait in lineages sibling to the Cyanobacteria. Nat. Commun. 2019;10:463. doi: 10.1038/s41467-018-08246-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 11.Shih P.M., Hemp J., Ward L.M., Matzke N.J., Fischer W.W. Crown group Oxyphotobacteria postdate the rise of oxygen. Geobiology. 2017;15:19–29. doi: 10.1111/gbi.12200. [DOI] [PubMed] [Google Scholar]
• 12.Magnabosco C., Moore K.R., Wolfe J.M., Fournier G.P. Dating phototrophic microbial lineages with reticulate gene histories. Geobiology. 2018;16:179–189. doi: 10.1111/gbi.12273. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 13.Schirrmeister B.E., Sánchez-Baracaldo P., Wacey D. Cyanobacterial evolution during the Precambrian. Int. J. Astrobiol. 2016;15:187–204. doi: 10.1017/S1473550415000579. [DOI] [Google Scholar]
• 14.Umena, Y., K. Kawakami, J.R. Shen, and N. Kamiya, Crystal structure of oxygen-evolving Photosystem II at a resolution of 1.9 Å. Nature, 2011. 473: 55–60. DOI: 10.1038/nature09913. [DOI] [PubMed]
• 15.Ferreira K.N., Iverson T.M., Maghlaoui K., Barber J., Iwata S. Architecture of the photosynthetic oxygen-evolving center. Science. 2004;303:1831–1838. doi: 10.1126/science.1093087. [DOI] [PubMed] [Google Scholar]
• 16.Cardona T., Sedoud A., Cox N., Rutherford A.W. Charge separation in Photosystem II: a comparative and evolutionary overview. Biochim. Biophys. Acta. 2012;1817:26–43. doi: 10.1016/j.bbabio.2011.07.012. [DOI] [PubMed] [Google Scholar]
• 17.Rutherford, A.W., T. Mattioli, and W. Nitschke, The FeS-type photosystems and the evolution of photosynthetic reaction centers, in Origin and evolution of biological energy conversion, H. Baltscheffsky, Editor. 1996, VCH: New York, N. Y. 177-203.
• 18.Rutherford, A.W. and W. Nitschke, Photosystem II and the quinone–iron-containing reaction centers, in Origin and evolution of biological energy conversion, H. Baltscheffsky, Editor. 1996, VCH: New York, N. Y. 143–175.
• 19.Cardona T., Sánchez-Baracaldo P., Rutherford A.W., Larkum A.W.D. Early Archean origin of Photosystem II. Geobiology. 2019;17:127–150. doi: 10.1111/gbi.12322. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 20.Cardona T., Rutherford A.W. Evolution of photochemical reaction centres: more twists? Trends Plant Sci. 2019;24:1008–1021. doi: 10.1016/j.tplants.2019.06.016. [DOI] [PubMed] [Google Scholar]
• 21.Mulkidjanian A.Y., Makarova K.S., Galperin M.Y., Koonin E.V. Inventing the dynamo machine: the evolution of the F-type and V-type ATPases. Nat. Rev. Microbiol. 2007;5:892–899. doi: 10.1038/nrmicro1767. [DOI] [PubMed] [Google Scholar]
• 22.Gogarten J.P., Kibak H., Dittrich P., Taiz L., Bowman E.J., Bowman B.J., Manolson M.F., Poole R.J., Date T., Oshima T., Konishi J., Denda K., Yoshida M. Evolution of the vacuolar H+-ATPase: implications for the origin of eukaryotes. Proc. Natl. Acad. Sci. U. S. A. 1989;86:6661–6665. doi: 10.1073/pnas.86.17.6661. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 23.Lane N., Allen J.F., Martin W. How did LUCA make a living? Chemiosmosis in the origin of life. Bioessays. 2010;32:271–280. doi: 10.1002/bies.200900131. [DOI] [PubMed] [Google Scholar]
• 24.Ouzounis C.A., Kunin V., Darzentas N., Goldovsky L. A minimal estimate for the gene content of the last universal common ancestor: exobiology from a terrestrial perspective. Res. Microbiol. 2006;157:57–68. doi: 10.1016/j.resmic.2005.06.015. [DOI] [PubMed] [Google Scholar]
• 25.Muller V., Gruber G. ATP synthases: structure, function and evolution of unique energy converters. Cell. Mol. Life Sci. 2003;60:474–494. doi: 10.1007/s000180300040. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 26.Werner F., Grohmann D. Evolution of multisubunit RNA polymerases in the three domains of life. Nat. Rev. Microbiol. 2011;9:85–98. doi: 10.1038/nrmicro2507. [DOI] [PubMed] [Google Scholar]
• 27.Kyrpides N., Overbeek R., Ouzounis C. Universal protein families and the functional content of the Last Universal Common Ancestor. J. Mol. Evol. 1999;49:413–423. doi: 10.1007/Pl00006564. [DOI] [PubMed] [Google Scholar]
• 28.Lane W.J., Darst S.A. Molecular evolution of multisubunit RNA polymerases: sequence analysis. J. Mol. Biol. 2010;395:671–685. doi: 10.1016/j.jmb.2009.10.062. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 29.Harris J.K., Kelley S.T., Spiegelman G.B., Pace N.R. The genetic core of the universal ancestor. Genome Res. 2003;13:407–412. doi: 10.1101/gr.652803. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 30.Chen M., Zhang Y., Blankenship R.E. Nomenclature for membrane-bound light-harvesting complexes of cyanobacteria. Photosynth. Res. 2008;95:147–154. doi: 10.1007/s11120-007-9255-0. [DOI] [PubMed] [Google Scholar]
• 31.Chen M., Hiller R.G., Howe C.J., Larkum A.W.D. Unique origin and lateral transfer of prokaryotic chlorophyll-b and chlorophyll-d light-harvesting systems. Mol. Biol. Evol. 2005;22:21–28. doi: 10.1093/molbev/msh250. [DOI] [PubMed] [Google Scholar]
• 32.Murray J.W. Sequence variation at the oxygen-evolving centre of Photosystem II: a new class of ‘rogue’ cyanobacterial D1 proteins. Photosynth. Res. 2012;110:177–184. doi: 10.1007/s11120-011-9714-5. [DOI] [PubMed] [Google Scholar]
• 33.Ho, M.Y., G. Shen, D.P. Canniffe, C. Zhao, and D.A. Bryant, Light-dependent chlorophyll f synthase is a highly divergent paralog of PsbA of Photosystem II. Science, 2016. 353: aaf9178. DOI: 10.1126/science.aaf9178. [DOI] [PubMed]
• 34.Gan, F., G. Shen, and D.A. Bryant, Occurrence of far-red light photoacclimation (FaRLiP) in diverse cyanobacteria. Life (Basel), 2014. 5: 4–24. DOI: 10.3390/life5010004. [DOI] [PMC free article] [PubMed]
• 35.Cardona T. A fresh look at the evolution and diversification of photochemical reaction centers. Photosynth. Res. 2015;126:111–134. doi: 10.1007/s11120-014-0065-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 36.Cardona T. Reconstructing the origin of oxygenic photosynthesis: do assembly and photoactivation recapitulate evolution? Front. Plant Sci. 2016;7:257. doi: 10.3389/fpls.2016.00257. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 37.Dibrova D.V., Galperin M.Y., Mulkidjanian A.Y. Characterization of the N-ATPase, a distinct, laterally transferred Na+-translocating form of the bacterial F-type membrane ATPase. Bioinformatics. 2010;26:1473–1476. doi: 10.1093/bioinformatics/btq234. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 38.Battistuzzi F.U., Feijao A., Hedges S.B. A genomic timescale of prokaryote evolution: insights into the origin of methanogenesis, phototrophy, and the colonization of land. BMC Evol. Biol. 2004;4:44. doi: 10.1186/1471-2148-4-44. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 39.Hug L.A., Baker B.J., Anantharaman K., Brown C.T., Probst A.J., Castelle C.J., Butterfield C.N., Hernsdorf A.W., Amano Y., Ise K., Suzuki Y., Dudek N., Relman D.A., Finstad K.M., Amundson R., Thomas B.C., Banfield J.F. A new view of the tree of life. Nat. Microbiol. 2016;1:16048. doi: 10.1038/nmicrobiol.2016.48. [DOI] [PubMed] [Google Scholar]
• 40.Parks D.H., Rinke C., Chuvochina M., Chaumeil P.A., Woodcroft B.J., Evans P.N., Hugenholtz P., Tyson G.W. Recovery of nearly 8,000 metagenome-assembled genomes substantially expands the tree of life. Nat. Microbiol. 2017;2:1533–1542. doi: 10.1038/s41564-017-0012-7. [DOI] [PubMed] [Google Scholar]
• 41.Soo R.M., Skennerton C.T., Sekiguchi Y., Imelfort M., Paech S.J., Dennis P.G., Steen J.A., Parks D.H., Tyson G.W., Hugenholtz P. An expanded genomic representation of the phylum Cyanobacteria. Genome Biol. Evol. 2014;6:1031–1045. doi: 10.1093/Gbe/Evu073. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 42.López-García, P. and D. Moreira, Physical connections: prokaryotes parasitizing their kin. Environ. Microbiol. Rep., 2020. DOI: 10.1111/1758-2229.12910. [DOI] [PubMed]
• 43.Castelle C.J., Brown C.T., Anantharaman K., Probst A.J., Huang R.H., Banfield J.F. Biosynthetic capacity, metabolic variety and unusual biology in the CPR and DPANN radiations. Nat. Rev. Microbiol. 2018;16:629–645. doi: 10.1038/s41579-018-0076-2. [DOI] [PubMed] [Google Scholar]
• 44.Coleman, G.A., A.A. Davín, T. Mahendrarajah, A. Spang, P. Hugenholtz, G.J. Szöllősi, and T.A. Williams, A rooted phylogeny resolves early bacterial evolution. bioRxiv, 2020: 2020.07.15.205187. DOI: 10.1101/2020.07.15.205187. [DOI] [PubMed]
• 45.Meheust R., Burstein D., Castelle C.J., Banfield J.F. The distinction of CPR bacteria from other bacteria based on protein family content. Nat. Commun. 2019;10:4173. doi: 10.1038/s41467-019-12171-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 46.Betts H.C., Puttick M.N., Clark J.W., Williams T.A., Donoghue P.C.J., Pisani D. Integrated genomic and fossil evidence illuminates life's early evolution and eukaryote origin. Nat. Ecol. Evol. 2018;2:1556–1562. doi: 10.1038/s41559-018-0644-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 47.Shao S., Cardona T., Nixon P.J. Early emergence of the FtsH proteases involved in photosystem II repair. Photosynthetica. 2018;56:163–177. doi: 10.1007/s11099-018-0769-9. [DOI] [Google Scholar]
• 48.Innan H., Kondrashov F. The evolution of gene duplications: classifying and distinguishing between models. Nat. Rev. Genet. 2010;11:97–108. doi: 10.1038/nrg2689. [DOI] [PubMed] [Google Scholar]
• 49.Lynch M., Conery J.S. The evolutionary fate and consequences of duplicate genes. Science. 2000;290:1151–1155. doi: 10.1126/science.290.5494.1151. [DOI] [PubMed] [Google Scholar]
• 50.Ho S.Y.W., Duchene S. Molecular-clock methods for estimating evolutionary rates and timescales. Mol. Ecol. 2014;23:5947–5965. doi: 10.1111/mec.12953. [DOI] [PubMed] [Google Scholar]
• 51.Tice M.M., Lowe D.R. Photosynthetic microbial mats in the 3,416-Myr-old ocean. Nature. 2004;431:549–552. doi: 10.1038/nature02888. [DOI] [PubMed] [Google Scholar]
• 52.Rutherford A.W., Faller P. Photosystem II: evolutionary perspectives. Philos. Trans. Royal Soc. B. 2003;358:245–253. doi: 10.1098/rstb.2002.1186. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 53.Nixon P.J., Diner B.A. Aspartate 170 of the Photosystem II reaction center polypeptide D1 is involved in the assembly of the oxygen-evolving manganese cluster. Biochemistry. 1992;31:942–948. doi: 10.1021/Bi00118a041. [DOI] [PubMed] [Google Scholar]
• 54.Adachi Y., Kuroda H., Yukawa Y., Sugiura M. Translation of partially overlapping psbD-psbC mRNAs in chloroplasts: the role of 5′-processing and translational coupling. Nucleic Acids Res. 2012;40:3152–3158. doi: 10.1093/nar/gkr1185. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 55.Carpenter, S.D., J. Charite, B. Eggers, and W.F. Vermaas, The psbC start codon in Synechocystis sp. PCC 6803. FEBS Lett., 1990. 260: 135–7. DOI: 10.1016/0014-5793(90)80085-w. [DOI] [PubMed]
• 56.Chisholm D., Williams J.G. Nucleotide sequence of psbC, the gene encoding the CP-43 chlorophyll a-binding protein of Photosystem II, in the cyanobacterium Synechocystis 6803. Plant Mol. Biol. 1988;10:293–301. doi: 10.1007/BF00029879. [DOI] [PubMed] [Google Scholar]
• 57.Garcia-Pichel F., Lombard J., Soule T., Dunaj S., Wu S.H., Wojciechowski M.F. Timing the evolutionary advent of cyanobacteria and the later great oxidation event using gene phylogenies of a sunscreen. Mbio. 2019;10 doi: 10.1128/mBio.00561-19. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 58.Cockell C.S. Ultraviolet radiation and the photobiology of earth’s early oceans. Orig. Life Evol. Biospheres. 2000;30:467–499. doi: 10.1023/a:1006765405786. [DOI] [PubMed] [Google Scholar]
• 59.Lewis C.A., Crayle J., Zhou S.T., Swanstrom R., Wolfenden R. Cytosine deamination and the precipitous decline of spontaneous mutation during Earth’s history. Proc. Natl. Acad. Sci. U. S. A. 2016;113:8194–8199. doi: 10.1073/pnas.1607580113. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 60.Levy M., Miller S.L. The stability of the RNA bases: implications for the origin of life. Proc. Natl. Acad. Sci. U. S. A. 1998;95:7933–7938. doi: 10.1073/pnas.95.14.7933. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 61.Eisen, J.A. and P.C. Hanawalt, A phylogenomic study of DNA repair genes, proteins, and processes. Mutat. Res./DNA Repair, 1999. 435: 171–213. DOI: 10.1016/s0921-8777(99)00050-6. [DOI] [PMC free article] [PubMed]
• 62.Koonin, E.V. and M.Y. Galperin, The major transitions in evolution: A comparative genomic perspective in Sequence — Evolution — Function: Computational Approaches in Comparative Genomics. 2003, Springer-Science+Business Media, B.V>. 252–292. DOI: 10.1007/978-1-4757-3783-7. [DOI]
• 63.Woese C. The universal ancestor. Proc. Natl. Acad. Sci. U. S. A. 1998;95:6854–6859. doi: 10.1073/pnas.95.12.6854. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 64.David L.A., Alm E.J. Rapid evolutionary innovation during an Archaean genetic expansion. Nature. 2011;469:93–96. doi: 10.1038/Nature09649. [DOI] [PubMed] [Google Scholar]
• 65.Marin J., Battistuzzi F.U., Brown A.C., Hedges S.B. The timetree of prokaryotes: new insights into their evolution and speciation. Mol. Biol. Evol. 2017;34:437–446. doi: 10.1093/molbev/msw245. [DOI] [PubMed] [Google Scholar]
• 66.Zhu, Q., U. Mai, W. Pfeiffer, S. Janssen, F. Asnicar, J.G. Sanders, P. Belda-Ferre, G.A. Al-Ghalith, E. Kopylova, D. McDonald, T. Kosciolek, J.B. Yin, S. Huang, N. Salam, J.-Y. Jiao, Z. Wu, Z.Z. Xu, K. Cantrell, Y. Yang, E. Sayyari, M. Rabiee, J.T. Morton, S. Podell, D. Knights, W.-J. Li, C. Huttenhower, N. Segata, L. Smarr, S. Mirarab, and R. Knight, Phylogenomics of 10,575 genomes reveals evolutionary proximity between domains Bacteria and Archaea. Nat. Commun., 2019. 10: 5477. DOI: 10.1038/s41467-019-13443-4. [DOI] [PMC free article] [PubMed]
• 67.Berkemer S.J., McGlynn S.E. A New Analysis of Archaea–Bacteria Domain Separation: Variable Phylogenetic Distance and the Tempo of Early Evolution. Mol. Biol. Evol. 2020 doi: 10.1093/molbev/msaa089. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 68.Moody, E.R.R., T.A. Mahendrarajah, N. Dombrowski, J.W. Clark, C. Petitjean, P. Offre, G.J. Szollosi, A. Spang, and T.A. Williams, Universal markers support a long inter-domain branch between Archaea and Bacteria. bioRxiv, 2021: 2021.01.19.427276. DOI: 10.1101/2021.01.19.427276. [DOI]
• 69.Homann, M., C. Heubeck, A. Airo, and M.M. Tice, Morphological adaptations of 3.22 Ga-old tufted microbial mats to Archean coastal habitats (Moodies Group, Barberton Greenstone Belt, South Africa). Precambrian Res., 2015. 266: 47–64. DOI: 10.1016/j.precamres.2015.04.018. [DOI]
• 70.Alcott L.J., Mills B.J.W., Poulton S.W. Stepwise Earth oxygenation is an inherent property of global biogeochemical cycling. Science. 2019;366:1333–1337. doi: 10.1126/science.aax6459. [DOI] [PubMed] [Google Scholar]
• 71.Kadoya S., Catling D.C., Nicklas R.W., Puchtel I.S., Anbar A.D. Mantle data imply a decline of oxidizable volcanic gases could have triggered the great oxidation. Nat. Commun. 2020;11:2774. doi: 10.1038/s41467-020-16493-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 72.Reinhard C.T., Planavsky N.J. Biogeochemical controls on the redox evolution of Earth’s oceans and atmosphere. Elements. 2020;16:191–196. doi: 10.2138/gselements.16.3.191. [DOI] [Google Scholar]
• 73.Cardona T. Photosystem II is a chimera of reaction centers. J. Mol. Evol. 2017;84:149–151. doi: 10.1007/s00239-017-9784-x. [DOI] [PubMed] [Google Scholar]
• 74.Chen, J.-H., H. Wu, C. Xu, X.-C. Liu, Z. Huang, S. Chang, W. Wang, G. Han, T. Kuang, J.-R. Shen, and X. Zhang, Architecture of the photosynthetic complex from a green sulfur bacterium. Science, 2020. 370: eabb6350. DOI: 10.1126/science.abb6350. [DOI] [PubMed]
• 75.Trinugroho J.P., Beckova M., Shao S., Yu J., Zhao Z., Murray J.W., Sobotka R., Komenda J., Nixon P.J. Chlorophyll f synthesis by a super-rogue photosystem II complex. Nat Plants. 2020;6:238–244. doi: 10.1038/s41477-020-0616-4. [DOI] [PubMed] [Google Scholar]
• 76.Wegener K.M., Nagarajan A., Pakrasi H.B. An atypical psbA gene encodes a sentinel D1 protein to form a physiologically relevant inactive Photosystem II complex in Cyanobacteria. J. Biol. Chem. 2015;290:3764–3774. doi: 10.1074/jbc.M114.604124. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 77.Cardona T. Thinking twice about the evolution of photosynthesis. Open Biol. 2019;9:180246. doi: 10.1098/rsob.180246. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 78.Fischer W.W., Hemp J., Johnson J.E. Manganese and the evolution of photosynthesis. Orig. Life Evol. Biospheres. 2015;45:351–357. doi: 10.1007/s11084-015-9442-5. [DOI] [PubMed] [Google Scholar]
• 79.Allen J.F., Martin W. Evolutionary biology - out of thin air. Nature. 2007;445:610–612. doi: 10.1038/445610a. [DOI] [PubMed] [Google Scholar]
• 80.Allen J.P., Olson T.L., Oyala P., Lee W.J., Tufts A.A., Williams J.C. Light-driven oxygen production from superoxide by Mn-binding bacterial reaction centers. Proc. Natl. Acad. Sci. U. S. A. 2012;109:2314–2318. doi: 10.1073/pnas.1115364109. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 81.Williamson A., Conlan B., Hillier W., Wydrzynski T. The evolution of Photosystem II: insights into the past and future. Photosynth. Res. 2011;107:71–86. doi: 10.1007/s11120-010-9559-3. [DOI] [PubMed] [Google Scholar]
• 82.Chernev P., Fischer S., Hoffmann J., Oliver N., Assunção R., Yu B., Burnap R.L., Zaharieva I., Nürnberg D.J., Haumann M., Dau H. Light-driven formation of manganese oxide by today’s photosystem II supports evolutionarily ancient manganese-oxidizing photosynthesis. Nat. Commun. 2020;11:6110. doi: 10.1038/s41467-020-19852-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 83.Johnson J.E., Webb S.M., Thomas K., Ono S., Kirschvink J.L., Fischer W.W. Manganese-oxidizing photosynthesis before the rise of cyanobacteria. Proc. Natl. Acad. Sci. U. S. A. 2013;110:11238–11243. doi: 10.1073/pnas.1305530110. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 84.Planavsky N.J., Asael D., Hofmann A., Reinhard C.T., Lalonde S.V., Knudsen A., Wang X., Ossa Ossa F., Pecoits E., Smith A.J.B., Beukes N.J., Bekker A., Johnson T.M., Konhauser K.O., Lyons T.W., Rouxel O.J. Evidence for oxygenic photosynthesis half a billion years before the great oxidation event. Nat. Geosci. 2014;7:283–286. doi: 10.1038/ngeo2122. [DOI] [Google Scholar]
• 85.Hodgskiss M.S.W., Crockford P.W., Peng Y.B., Wing B.A., Horner T.J. A productivity collapse to end Earth’s great oxidation. Proc. Natl. Acad. Sci. U. S. A. 2019;116:17207–17212. doi: 10.1073/pnas.1900325116. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 86.Bekker A., Holland H.D. Oxygen overshoot and recovery during the early Paleoproterozoic. Earth Planet. Sci. Lett. 2012;317:295–304. doi: 10.1016/j.epsl.2011.12.012. [DOI] [Google Scholar]
• 87.Fischer W.W., Hemp J., Johnson J.E. Evolution of oxygenic photosynthesis. Annu. Rev. Earth Planet. Sci. 2016;44:647–683. doi: 10.1146/annurev-earth-060313-054810. [DOI] [Google Scholar]
• 88.Fine P.L., Frasch W.D. The oxygen-evolving complex requires chloride to prevent hydrogen peroxide formation. Biochemistry. 1992;31:12204–12210. doi: 10.1021/bi00163a033. [DOI] [PubMed] [Google Scholar]
• 89.Arato A., Bondarava N., Krieger-Liszkay A. Production of reactive oxygen species in chloride- and calcium-depleted photosystem II and their involvement in photoinhibition. Bba-Bioenergetics. 2004;1608:171–180. doi: 10.1016/j.bbabio.2003.12.003. [DOI] [PubMed] [Google Scholar]
• 90.Hillier W., Wydrzynski T. Increases in peroxide formation by the Photosystem II oxygen evolving reactions upon removal of the extrinsic 16, 22 and 33 kDa proteins are reversed by CaCl2 addition. Photosynth. Res. 1993;38:417–423. doi: 10.1007/BF00046769. [DOI] [PubMed] [Google Scholar]
• 91.Thompson L.K., Blaylock R., Sturtevant J.M., Brudvig G.W. Molecular basis of the heat denaturation of photosystem II. Biochemistry. 1989;28:6686–6695. doi: 10.1021/bi00442a023. [DOI] [PubMed] [Google Scholar]
• 92.Ishikita H., Saenger W., Biesiadka J., Loll B., Knapp E.W. How photosynthetic reaction centers control oxidation power in chlorophyll pairs P680, P700, and P870. Proc. Natl. Acad. Sci. U. S. A. 2006;103:9855–9860. doi: 10.1073/pnas.0601446103. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 93.Johnson G.N., Rutherford A.W., Krieger A. A change in the midpoint potential of the quinone Q(a) in Photosystem II associated with photoactivation of oxygen evolution. Biochim. Biophys. Acta. 1995;1229:202–207. doi: 10.1016/0005-2728(95)00003-2. [DOI] [Google Scholar]
• 94.Schlodder E., Meyer B. pH dependence of oxygen evolution and reduction kinetics of photooxidized chlorophyll aII (P-680) in Photosystem II particles from Synechococcus sp. Biochim. Biophys. Acta. 1987;890:23–31. doi: 10.1016/0005-2728(87)90064-8. [DOI] [Google Scholar]
• 95.Rutherford A.W., Boussac A., Faller P. The stable tyrosyl radical in photosystem II: why D? Biochim. Biophys. Acta. 2004;1655:222–230. doi: 10.1016/j.bbabio.2003.10.016. [DOI] [PubMed] [Google Scholar]
• 96.Styring S., Sjoholm J., Mamedov F. Two tyrosines that changed the world: interfacing the oxidizing power of photochemistry to water splitting in photosystem II. Bba-Bioenergetics. 2012;1817:76–87. doi: 10.1016/j.bbabio.2011.03.016. [DOI] [PubMed] [Google Scholar]
• 97.Gan F., Zhang S., Rockwell N.C., Martin S.S., Lagarias J.C., Bryant D.A. Extensive remodeling of a cyanobacterial photosynthetic apparatus in far-red light. Science. 2014;345:1312–1317. doi: 10.1126/science.1256963. [DOI] [PubMed] [Google Scholar]
• 98.Trinugroho, J.P., M. Beckova, S.X. Shao, J.F. Yu, Z.Y. Zhao, J.W. Murray, R. Sobotka, J. Komenda, and P.J. Nixon, Chlorophyll f synthesis by a super-rogue photosystem II complex. Nature Plants, 2020. 6: 238−+. DOI: 10.1038/s41477-020-0616-4. [DOI] [PubMed]
• 99.Rutherford A.W., Osyczka A., Rappaport F. Back-reactions, short-circuits, leaks and other energy wasteful reactions in biological electron transfer: redox tuning to survive life in O2. FEBS Lett. 2012;586:603–616. doi: 10.1016/j.febslet.2011.12.039. [DOI] [PubMed] [Google Scholar]
• 100.Ben-Shem A., Frolow F., Nelson N. Evolution of photosystem I - from symmetry through pseudosymmetry to asymmetry. FEBS Lett. 2004;564:274–280. doi: 10.1016/S0014-5793(04)00360-6. [DOI] [PubMed] [Google Scholar]
• 101.Jagannathan, B., G.Z. Shen, and J.H. Golbeck, The evolution of Type I reaction centers: the response to oxygenic photosynthesis, in Functional genomics and evolution of photosynthetic systems, R.L. Burnap and W. Vermaas, Editors. 2012, Springer Science: Dordrecht. 285-316.
• 102.Cardona T. Early Archean origin of heterodimeric Photosystem I. Heliyon. 2018;4 doi: 10.1016/j.heliyon.2018.e00548. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 103.Orf, G.S., C. Gisriel, and K.E. Redding, Evolution of photosynthetic reaction centers: insights from the structure of the heliobacterial reaction center. Photosynth. Res., 2018. DOI: 10.1007/s11120-018-0503-2. [DOI] [PubMed]
• 104.Allen J.F. A redox switch hypothesis for the origin of two light reactions in photosynthesis. FEBS Lett. 2005;579:963–968. doi: 10.1016/j.febslet.2005.01.015. [DOI] [PubMed] [Google Scholar]
• 105.Sousa F.L., Shavit-Grievink L., Allen J.F., Martin W.F. Chlorophyll biosynthesis gene evolution indicates photosystem gene duplication, not photosystem merger, at the origin of oxygenic photosynthesis. Genome Biol. Evol. 2013;5:200–216. doi: 10.1093/Gbe/Evs127. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 106.Olson J.M., Pierson B.K. Origin and evolution of photosynthetic reaction centers. Orig. Life Evol. Biospheres. 1987;17:419–430. doi: 10.1007/Bf02386479. [DOI] [Google Scholar]
• 107.Mulkidjanian A.Y., Koonin E.V., Makarova K.S., Mekhedov S.L., Sorokin A., Wolf Y.I., Dufresne A., Partensky F., Burd H., Kaznadzey D., Haselkorn R., Galperin M.Y. The cyanobacterial genome core and the origin of photosynthesis. Proc. Natl. Acad. Sci. U. S. A. 2006;103:13126–13131. doi: 10.1073/pnas.0605709103. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 108.Petrov A.S., Bernier C.R., Hsiao C.L., Norris A.M., Kovacs N.A., Waterbury C.C., Stepanov V.G., Harvey S.C., Fox G.E., Wartell R.M., Hud N.V., Williams L.D. Evolution of the ribosome at atomic resolution. Proc. Natl. Acad. Sci. U. S. A. 2014;111:10251–10256. doi: 10.1073/pnas.1407205111. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 109.Granick S. Speculations on the origins and evolution of photosynthesis. Ann. N. Y. Acad. Sci. 1957;69:292–308. doi: 10.1111/j.1749-6632.1957.tb49665.x. [DOI] [PubMed] [Google Scholar]
• 110.Mauzerall D. Light, iron, Sam Granick and the origin of life. Photosynth. Res. 1992;33:163–170. doi: 10.1007/BF00039178. [DOI] [PubMed] [Google Scholar]
• 111.Mulkidjanian A.Y., Bychkov A.Y., Dibrova D.V., Galperin M.Y., Koonin E.V. Origin of first cells at terrestrial, anoxic geothermal fields. Proc. Natl. Acad. Sci. U. S. A. 2012;109:E821–E830. doi: 10.1073/pnas.1117774109. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 112.Franz, H.B., P.R. Mahaffy, C.R. Webster, G.J. Flesch, E. Raaen, C. Freissinet, S.K. Atreya, C.H. House, A.C. McAdam, C.A. Knudson, P.D. Archer, J.C. Stern, A. Steele, B. Sutter, J.L. Eigenbrode, D.P. Glavin, J.M.T. Lewis, C.A. Malespin, M. Millan, D.W. Ming, R. Navarro-Gonzalez, and R.E. Summons, Indigenous and exogenous organics and surface-atmosphere cycling inferred from carbon and oxygen isotopes at Gale crater. Nat Astron, 2020. DOI: 10.1038/s41550-019-0990-x. [DOI]
• 113.Lu A.H., Li Y., Ding H.R., Xu X.M., Li Y.Z., Ren G.P., Liang J., Liu Y.W., Hong H., Chen N., Chu S.Q., Liu F.F., Wang H.R., Ding C., Wang C.Q., Lai Y., Liu J., Dick J., Liu K.H., Hochella M.F. Photoelectric conversion on Earth’s surface via widespread Fe- and Mn-mineral coatings. Proc. Natl. Acad. Sci. U. S. A. 2019;116:9741–9746. doi: 10.1073/pnas.1902473116. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 114.Mendler K., Chen H., Parks D.H., Lobb B., Hug L.A., Doxey A.C. AnnoTree: visualization and exploration of a functionally annotated microbial tree of life. Nucleic Acids Res. 2019;47:4442–4448. doi: 10.1093/nar/gkz246. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 115.Ward L.M., Cardona T., Holland-Moritz H. Evolutionary implications of anoxygenic phototrophy in the bacterial phylum Candidatus Eremiobacterota (WPS-2) Front. Microbiol. 2019;10:1658. doi: 10.3389/fmicb.2019.01658. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 116.Sievers F., Wilm A., Dineen D., Gibson T.J., Karplus K., Li W.Z., Lopez R., McWilliam H., Remmert M., Soding J., Thompson J.D., Higgins D.G. Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Mol. Syst. Biol. 2011;7:539. doi: 10.1038/msb.2011.75. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 117.Castresana J. Selection of conserved blocks from multiple alignments for their use in phylogenetic analysis. Mol. Biol. Evol. 2000;17:540–552. doi: 10.1093/oxfordjournals.molbev.a026334. [DOI] [PubMed] [Google Scholar]
• 118.Guindon, S., J.F. Dufayard, V. Lefort, M. Anisimova, W. Hordijk, and O. Gascuel, New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Syst. Biol., 2010. 59: 307–21. DOI: 10.1093/sysbio/syq010. [DOI] [PubMed]
• 119.Lefort V., Longueville J.E., Gascuel O. SMS: smart model selection in PhyML. Mol. Biol. Evol. 2017;34:2422–2424. doi: 10.1093/molbev/msx149. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 120.Anisimova M., Gascuel O. Approximate likelihood-ratio test for branches: a fast, accurate, and powerful alternative. Syst. Biol. 2006;55:539–552. doi: 10.1080/10635150600755453. [DOI] [PubMed] [Google Scholar]
• 121.Huson D.H., Scornavacca C. Dendroscope 3: an interactive tool for rooted phylogenetic trees and networks. Syst. Biol. 2012;61:1061–1067. doi: 10.1093/sysbio/sys062. [DOI] [PubMed] [Google Scholar]
• 122.Gascuel O. BIONJ: an improved version of the NJ algorithm based on a simple model of sequence data. Mol. Biol. Evol. 1997;14:685–695. doi: 10.1093/oxfordjournals.molbev.a025808. [DOI] [PubMed] [Google Scholar]
• 123.Gouy M., Guindon S., Gascuel O. SeaView version 4: a multiplatform graphical user interface for sequence alignment and phylogenetic tree building. Mol. Biol. Evol. 2010;27:221–224. doi: 10.1093/molbev/msp259. [DOI] [PubMed] [Google Scholar]
• 124.Kumar S., Stecher G., Li M., Knyaz C., Tamura K. MEGA X: molecular evolutionary genetics analysis across computing platforms. Mol. Biol. Evol. 2018;35:1547–1549. doi: 10.1093/molbev/msy096. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 125.Stothard P. The sequence manipulation suite: JavaScript programs for analyzing and formatting protein and DNA sequences. BioTechniques. 2000;28:1102–1104. doi: 10.2144/00286ir01. [DOI] [PubMed] [Google Scholar]
• 126.Lartillot N., Lepage T., Blanquart S. PhyloBayes 3: a Bayesian software package for phylogenetic reconstruction and molecular dating. Bioinformatics. 2009;25:2286–2288. doi: 10.1093/bioinformatics/btp368. [DOI] [PubMed] [Google Scholar]
• 127.Lepage T., Bryant D., Philippe H., Lartillot N. A general comparison of relaxed molecular clock models. Mol. Biol. Evol. 2007;24:2669–2680. doi: 10.1093/molbev/msm193. [DOI] [PubMed] [Google Scholar]
• 128.Kishino H., Thorne J.L., Bruno W.J. Performance of a divergence time estimation method under a probabilistic model of rate evolution. Mol. Biol. Evol. 2001;18:352–361. doi: 10.1093/oxfordjournals.molbev.a003811. [DOI] [PubMed] [Google Scholar]
• 129.Li W., Godzik A. Cd-hit: a fast program for clustering and comparing large sets of protein or nucleotide sequences. Bioinformatics. 2006;22:1658–1659. doi: 10.1093/bioinformatics/btl158. [DOI] [PubMed] [Google Scholar]
• 130.Ashkenazy H., Penn O., Doron-Faigenboim A., Cohen O., Cannarozzi G., Zomer O., Pupko T. FastML: a web server for probabilistic reconstruction of ancestral sequences. Nucleic Acids Res. 2012;40:W580–W584. doi: 10.1093/nar/gks498. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 131.Hanson-Smith V., Kolaczkowski B., Thornton J.W. Robustness of ancestral sequence reconstruction to phylogenetic uncertainty. Mol. Biol. Evol. 2010;27:1988–1999. doi: 10.1093/molbev/msq081. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 132.Yang Z.H. PAML 4: phylogenetic analysis by maximum likelihood. Mol. Biol. Evol. 2007;24:1586–1591. doi: 10.1093/molbev/msm088. [DOI] [PubMed] [Google Scholar]
• 133.Kumar S., Stecher G., Tamura K. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol. Biol. Evol. 2016;33:1870–1874. doi: 10.1093/molbev/msw054. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 134.Ago H., Adachi H., Umena Y., Tashiro T., Kawakami K., Kamiya N., Tian L.R., Han G.Y., Kuang T.Y., Liu Z.Y., Wang F.J., Zou H.F., Enami I., Miyano M., Shen J.R. Novel features of eukaryotic Photosystem II revealed by its crystal structure analysis from a red alga. J. Biol. Chem. 2016;291:5676–5687. doi: 10.1074/jbc.M115.711689. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 135.Yu L.J., Suga M., Wang-Otomo Z.Y., Shen J.R. Structure of photosynthetic LH1-RC supercomplex at 1.9 Å resolution. Nature. 2018;556:209–213. doi: 10.1038/s41586-018-0002-9. [DOI] [PubMed] [Google Scholar]
• 136.Gisriel C., Sarrou I., Ferlez B., Golbeck J.H., Redding K.E., Fromme R. Structure of a symmetric photosynthetic reaction center-photosystem. Science. 2017;357:1021–1025. doi: 10.1126/science.aan5611. [DOI] [PubMed] [Google Scholar]
• 137.Jordan, P., P. Fromme, H.T. Witt, O. Klukas, W. Saenger, and N. Krauss, Three-dimensional structure of cyanobacterial Photosystem I at 2.5 Å resolution. Nature, 2001. 411: 909–17. DOI: 10.1038/35082000. [DOI] [PubMed]
• 138.Toporik H., Li J., Williams D., Chiu P.L., Mazor Y. The structure of the stress-induced photosystem I-IsiA antenna supercomplex. Nat. Struct. Mol. Biol. 2019;26:443–449. doi: 10.1038/s41594-019-0228-8. [DOI] [PubMed] [Google Scholar]
• 139.Jia Y.T., Dewey G., Shindyalov I.N., Bourne P.E. A new scoring function and associated statistical significance for structure alignment by CE. J. Comput. Biol. 2004;11:787–799. doi: 10.1089/1066527042432260. [DOI] [PubMed] [Google Scholar]
• 140.Kumar S., Stecher G., Suleski M., Hedges S.B. TimeTree: a resource for timelines, timetrees, and divergence times. Mol. Biol. Evol. 2017;34:1812–1819. doi: 10.1093/molbev/msx116. [DOI] [PubMed] [Google Scholar]
• 141.Shih P.M., Ward L.M., Fischer W.W. Evolution of the 3-hydroxypropionate bicycle and recent transfer of anoxygenic photosynthesis into the Chloroflexi. Proc. Natl. Acad. Sci. U. S. A. 2017;114:10749–10754. doi: 10.1073/pnas.1710798114. [DOI] [PMC free article] [PubMed] [Google Scholar]

Supplementary Materials