¡Descarga tilacoide biogenesis y más Apuntes en PDF de Biología solo en Docsity!
Review
Biogenesis of thylakoid membranes☆
Anna Rast, Steffen Heinz, Jörg Nickelsen ⁎
Molekulare Pflanzenwissenschaften, Biozentrum LMU München, Großhaderner Str. 2-4, 82152 Planegg-Martinsried, Germany
a r t i c l e i n f o a b s t r a c t
Article history: Received 15 December 2014 Received in revised form 9 January 2015 Accepted 15 January 2015 Available online 20 January 2015
Keywords: Biogenesis center Thylakoid membrane Pyrenoid Lipid T-zone Assembly factor
Thylakoids mediate photosynthetic electron transfer and represent one of the most elaborate energy-transducing membrane systems. Despite our detailed knowledge of its structure and function, much remains to be learned about how the machinery is put together. The concerted synthesis and assembly of lipids, proteins and low- molecular-weight cofactors like pigments and transition metal ions require a high level of spatiotemporal coor- dination. While increasing numbers of assembly factors are being functionally characterized, the principles that govern how thylakoid membrane maturation is organized in space are just starting to emerge. In both cyanobacteria and chloroplasts, distinct production lines for the fabrication of photosynthetic complexes, in par- ticular photosystem II, have been identified. This article is part of a Special Issue entitled: Chloroplast Biogenesis. © 2015 Elsevier B.V. All rights reserved.
- Introduction
The origin of oxygenic photosynthesis dates back more than 2.4 bil- lion years. At that time, primordial cyanobacteria first succeeded in harnessing solar energy to drive the extraction of electrons from water; these are then fed into a transmembrane photosynthetic elec- tron transport chain (PET; [1]). PET takes place across specialized mem- branes named thylakoids, which represent one of the most elaborate bioenergetic machines known to date [2]. Thylakoids form an intracellu- lar membrane system which harbors most of the constituents involved in the harvesting and utilization of light energy. As a consequence of PET, the chemical energy carriers NADPH and ATP are generated, and utilized for the synthesis of carbohydrates from CO 2. Photosynthesis thus forms the basis of nature's food chains. The main protein components of thylakoids include the mobile, low- molecular-weight carriers plastoquinone and plastocyanin, which shut- tle electrons between huge multisubunit protein/pigment complexes,
i.e., photosystem II (PSII) and photosystem I (PSI), via the cytochrome b 6 f (Cytb 6 f) complex. Finally, an ATPase complex utilizes the energy stored in the proton gradient established across the thylakoid mem- brane by PET to synthesize ATP. This core molecular machinery mediat- ing PET is highly conserved from cyanobacteria to plants [3], although the light-collecting systems are more diverse, varying from soluble phycobilisomes in cyanobacteria and red algae to membrane- integrated light-harvesting (LHC) systems in plants [4]. The ultrastruc- ture of these various components has been determined at close to atom- ic resolution in recent years, providing a detailed picture of the structure and function of the PET chain [5,6]. However, despite this in-depth knowledge of their structure, the processes that guide the construction of the various photosynthetic complexes during thylakoid membrane biogenesis are only beginning to emerge. As the available structural data imply, dozens of integral and peripheral protein subunits and hundreds of organic cofactors in- cluding chlorophylls, carotenoids, cytochromes and quinones, as well as metals and other ions, have to be brought together and properly as- sembled. And of course these diverse building materials must be syn- thesized and supplied in amounts reflecting at least approximately their final stoichiometry. Then the components must be brought togeth- er in a lipid environment and assembled in a strictly determined and stepwise sequence. Thus, the entire process must be highly ordered both in space and in time. The current status of our understanding of the biogenesis of particu- lar complexes is reviewed in detail in Chapters 11–14 [7–10] of this spe- cial issue. Therefore, we focus here on how different biosynthetic pathways for thylakoid membrane constituents might be interconnect- ed to enable coordination of biogenesis in the temporal domain. In
Biochimica et Biophysica Acta 1847 (2015) 821– 830
Abbreviations: PET, photosynthetic electron transport; NADPH, nicotinamide adenine dinucleotide phosphate; ATP, adenosine triphosphate; CO 2 , carbon dioxide; Cytb 6 f, cyto- chrome b 6 f complex; PSI, photosystem I; PSII, photosystem II; LHC, light-harvesting com- plex; MGDG, monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; SQDG, sulfoquinovosyldiacylglycerol; PG, phosphatidylglycerol; PDC, pyruvate dehydrogenase complex; ACCase, acetyl-CoA carboxylase; PA, phosphatidic acid; DAG, diacylglycerol; MGS, monoglucosyldiacylglycerol synthase; CURT1, CURVATURE THYLAKOID1; THF1, thy- lakoid formation 1; POR, protochlorophyllide oxidoreductase; ChlG, chlorophyll synthase; CTM, chloroplast translation membrane ☆ This article is part of a Special Issue entitled: Chloroplast Biogenesis. ⁎ Corresponding author at: Biozentrum LMU München, Großhaderner Str. 2-4, 82152 Planegg-Martinsried, Germany. Tel.: +49 89 2180 74773; fax: +49 89 2180 9974773. E-mail address: [email protected] (J. Nickelsen).
http://dx.doi.org/10.1016/j.bbabio.2015.01. 0005-2728/© 2015 Elsevier B.V. All rights reserved.
Contents lists available at ScienceDirect
Biochimica et Biophysica Acta
j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / b b a b i o
addition, the cytological aspects of the whole process are emphasized by reviewing current findings regarding the spatial organization of thyla- koid membrane biogenesis and architecture. Despite differing signifi- cantly in structural organization, the functions of thylakoids have been highly conserved throughout evolution. Hence, the biogenesis process will be discussed in the light of recent findings in organisms ranging from cyanobacteria to vascular plants.
- Forming the matrix of thylakoid membranes — the lipids
2.1. Composition and function of the thylakoid lipidome
Thylakoids represent remarkably conserved energy-transducing membranes with a high protein content that can reach up to 80% in stacked grana regions of eukaryotic chloroplasts accumulating PSII [2]. Like the photosynthetic protein/pigment complexes that account for this high protein/lipid ratio, the lipophilic matrix in which they are embedded is unique, and has been conserved during the course of evolution. This suggests that a specific lipid environment is re- quired to ensure the stability and activity of photosynthetic com- plexes [11]. With minor deviations, thylakoids from cyanobacteria to plants con- tain four major lipid types, i.e., the galactoglycerolipids MGDG (monogalactosyldiacylglycerol) and DGDG (digalactosyldiacylglycerol), the sulfolipid SQDG (sulfoquinovosyldiacylglycerol) and the phospho- lipid PG (phosphatidylglycerol). The glycolipids comprise more than 70% of thylakoid lipids, with MGDG accounting for N50% of the total. The relative proportions of the lipid classes are stable, underlining their defined structural/functional roles which are dependent on their physicochemical properties [12,13]. These properties include the pres- ence of negatively charged headgroups in only SQDG and PG, and the small size of MGDG's headgroup, which allows it to form inverse hexag- onal structures (the so-called HII phase), unlike DGDG, SQDG and PG [14,15]. As a functional consequence, DGDG forms and stabilizes mem- brane bilayers, whereas MGDG is likely to favor curvature of thylakoids. This idea is supported by the finding that highly stacked grana regions with a high content of curved membrane regions exhibit higher MGDG/DGDG ratios than do stroma lamellae [16]. Recent analyses sup- port the critical role of the MGDG/DGDG ratio for membrane phase transitions and highlight the function of DGDG for membrane stacking via the formation of hydrogen bonds between the headgroups of adja- cent bilayers [17]. In addition to their matrix role, all glycerolipid classes have been shown to interact closely with photosynthetic protein com- plexes, in particular with PSII, as deduced from X-ray crystallographic analyses [18–20]. This suggests that individual lipids may also play crit- ical roles for the structure and function of the protein complexes. In- deed, genetic studies of mutants with defects in glycerolipid accumulation in plants and cyanobacteria have confirmed that MGDG is essential for thylakoid formation generally [21], while DGDG and PG are important for the function of PSII in particular [11,22,23].
2.2. Synthesis of lipids
The biosynthesis of thylakoid lipids follows a complex pathway which requires the production and combination of fatty acids and polar head precursors [11,24]. Fatty acid synthesis begins in the chloro- plast stroma with the conversion of pyruvate into acetyl-CoA by the py- ruvate dehydrogenase complex (PDC). Subsequently, acetyl-CoA is carboxylated to malonyl-CoA by the acetyl-CoA carboxylase (ACCase), which is considered to be the rate-limiting “committed step” in fatty acid synthesis [24]. Interestingly, recent studies of the regulation of chloroplast gene expression have revealed that the E dihydrolipoyltransferase subunit of the PDC acts as a regulator of the synthesis of the D1 protein of the PSII reaction center by localizing its mRNA to specialized biogenic membrane regions in Chlamydomonas reinhardtii as outlined below (Fig. 2B, Section 3.2 and [25]). Thus, the
moonlighting activity of a metabolic enzyme appears to provide for re- ciprocal regulation of plastid fatty acid synthesis and protein synthesis. This kind of crosstalk in the initial phase of thylakoid membrane bio- genesis would guarantee that lipid and protein syntheses are harmo- nized from the beginning to avoid unbalanced accumulation of the major classes of thylakoid constituents during chloroplast biogenesis [25]. In line with this, transcriptomic studies in Arabidopsis thaliana have recently revealed tight coordination between the expression of nuclear genes involved in later steps of both lipid and chlorophyll syntheses and genes coding for photosynthetic proteins [26]. Whether similar regulatory principles also operate in cyanobacteria is currently unknown. Fatty acid synthesis is completed within the chloroplast, and thyla- koid lipids are then synthesized via the resident prokaryotic pathway in the chloroplast or the extra-organellar eukaryotic route, which re- quires the export of fatty acids into the cytoplasm, their assembly into lipid precursors in the ER and subsequent reimport into plastids [27]. Ir- respective of their precise derivation, the lipid precursors phosphatidic acid (PA) and diacylglycerol (DAG) are conveyed to the inner envelope, and further processed by the key enzymes MGD1 and DGD1, which cat- alyze the formation of the most abundant lipids MGDG and DGDG, re- spectively. While MGD1 is localized to the inner envelope membrane, DGDG is synthesized at the outer envelope, as revealed by proteomic approaches following membrane fractionation [27–29]. Interestingly, the MGDG/DGDG ratio of membranes has a direct influence on the asso- ciation of MGD1 with the membrane and its self-organization into retic- ulated structures; this in turn suggests the existence of specialized membranous microdomains for plastid galactolipid synthesis [30]. Interestingly, recent membrane fractionation experiments have revealed that galactolipid synthesis in cyanobacteria seems to occur at both the plasma membrane and the thylakoids [31]. Forma- tion of MGDG in cyanobacteria differs from that in chloroplasts insofar as the former lack MGD1 activity. Instead, DAG is first con- verted into monoglucosyldiacylglycerol by the essential enzyme monoglucosyldiacylglycerol synthase (MGS) and then further proc- essed by an epimerase to yield MGDG [32]. In agreement with a broader distribution of lipid synthesis, MGS was found to localize to both plasma membrane and thylakoids [31].
2.3. Trafficking of lipids
One crucial but still unanswered question is how lipids are trans- ferred from the plastid envelope to the internal thylakoid membrane system during its maturation. In principle, three forms of trafficking can be envisaged, based on (i) lateral fusions between inner envelope and thylakoids, (ii) vesicular transport, or (iii) soluble lipid carriers (Fig. 1 and [33]). In algal chloroplasts and proplastids of vascular plants, thylakoids appear to develop via local invaginations of the inner envelope mem- brane [34,35]. Less frequently, connections between envelope and thy- lakoids have been observed in mature chloroplasts also, suggesting that intraplastidic lipid transfer occurs via lateral diffusion (Fig. 1A and [36]). In support of this scenario, in the A. thaliana mgd1 mutant, which does not form photosynthetically active thylakoids, invaginations of the inner envelope membrane are observed [37]. However, ultrastructural evidence for vesicle-based transfer pro- cesses has also been repeatedly obtained (Fig. 1B and [34,38–40]). Fur- thermore, inhibitors of the cytoplasmic secretory pathway similarly affect vesicle formation in isolated chloroplasts, suggesting that related systems exist in both cellular compartments [40–42]. More recently, comprehensive bioinformatic studies have provided evidence for chlo- roplast localization for several proteins that share high sequence simi- larity with known components of the vesicle-based secretory pathway in the cytoplasm [43–45]. Based on these analyses, it seems likely that a complete vesicle transport system which resembles the cytosolic COPII system is active in the chloroplast, whereas COPI and clathrin-
their mode of integration into and assembly within the membrane is a central question in chloroplast research. As the complexity of thylakoid architecture has increased during evolution, so too has the need for coordination of the assembly process (for further details see section 4 and Fig. 3). Gene expression and syn- thesis of the necessary proteins are already highly regulated (Chapters 1 – 9 [75–83]), and the same holds for the subsequent phase of complex assembly and cofactor integration (Chapters 11–14 [7–10]). Moreover, with the advent of chloroplasts after the endosymbiotic engulfment of a cyanobacterial cell, the whole process became much more complicat- ed, due to the constraints imposed by the transfer of photosynthetic genes from the chloroplast to the nucleus. As a consequence, the bio- genesis process now required the import of many protein subunits of photosynthetic complexes into the organelle, offering a new opportuni- ty for regulatory control (Chapters 15–18 [84–87]). Accumulating evi- dence indicates that most if not all of these processes are coordinated by regulatory factors which catalyze distinct assembly steps in a step- wise manner, but do not form constituents of the final photosynthetic machinery [88–91]. As such, they can serve as markers which enable one to assign specific biogenesis steps to organellar subfractions or sub- structures. At least in theory, it should therefore be possible to dissect the cytological landscape of thylakoid membrane biogenesis. Two main experimental approaches have recently been exploited to discern the design of the production line for photosynthetic membranes — the application of membrane fractionation and imaging techniques.
3.1. Lessons from membrane fractionation experiments — thylakoid heterogeneity
Cyanobacteria usually possess three different membrane systems. The outer membrane and the plasma membrane, together with the in- tervening peptidoglycan layer, form the cell envelope, which encloses the thylakoid membranes. Early cell fractionation studies using sucrose-gradient centrifugation had revealed that distinct subunits of both PSI and PSII are present not only in thylakoids but also in plasma membranes from the cyanobacterium Synechococcus elongatus [92,93]. Other subunits of PSII, including CP43 and CP47, were found exclusively in thylakoids. In line with these findings, no PSII activity (as measured by oxygen evolution) was detectable in plasma membranes, which led to the hypothesis that biogenic intermediates of photosynthetic com- plexes might accumulate in the plasma membrane [93]. This idea was
later revisited by applying a two-phase partitioning technique to sepa- rate different membrane fractions [94]. Again, in addition to being found in thylakoids, core subunits of PSII and PSI including D1, D2, Cytb559, PsaA, PsaB, PsaC and PsaD were immunologically detected – in minor amounts – in plasma membrane fractions. Moreover, substan- tial amounts of the assembly factors CtpA, Ycf3 and Ycf4 were also found in the plasma membrane, suggesting that the initial steps in the assembly of both PSII and PSI take place in plasma membranes [94]. These early assembly intermediates already contain chlorophyll and are capable of light-induced charge separation [55]. More recent membrane fractionations involving two consecutive sucrose-gradient centrifugation steps have led to the identification and characterization of a membrane subfraction that is distinct from both plasma and thylakoid membranes. Because the PSII assembly fac- tor PratA was found to accumulate in this fraction, it was named the PratA-defined membrane (PDM; [95]). A detailed characterization of the PDM revealed that it contains – besides PratA – substantial amounts of the pD1 precursor protein. Importantly, in a pratA mutant, pD1 accu- mulation in PDMs was further enhanced [95]. In addition, the D reaction-center protein and early PSII assembly factors like YCF48 and YidC, but not the thylakoid marker proteins CP47 and CP43, were de- tected, in agreement with earlier membrane fractionation studies de- scribed above. Interestingly, the protochlorophyllide oxidoreductase (POR) required for chlorophyll synthesis, and its interaction partner Pitt, as well as the chlorophyll precursor chlorophyllide a, were also found in PDMs, suggesting that the two biosynthesis pathways might intersect at PDMs [96]. A tight connection between apoprotein synthe- sis/assembly and chlorophyll synthesis was recently confirmed by the close physical linkage between the chlorophyll synthase (ChlG) and the PSII assembly factors YCF39 and YidC revealed by co-elution of these proteins together with chlorophylls during tag-based affinity pu- rification of ChlG [97,98]. However, in contrast to plasma membrane fractions described above, neither PSI subunits nor assembly factors were detected in PDMs [99]. Taken together, these data suggest that PDMs might represent a specialized membrane region where assembly of PSII is initiated, and that a tight connection between assembly and chlorophyll synthesis/insertion exists, at least in cyanobacteria (Fig. 2A). The advent of proteomic approaches then allowed the question of cyanobacterial membrane heterogeneity to be addressed more systematically [70]. The proteomes of the thylakoid and plasma
Periplasm
Cytosol
RC47 RC
Lumen RCC
TC?
Biogenesis Center
pD
PDM
OM PM
TM
PratA
Mn 2+
[Mn Mn2+ Mn2+ 4 ]
Mn 2+
Mn2+
Cytosol
Import
Stroma Pyrenoid
T-Zone
RCC1 RC47 RC
pD
? OE IE
Lumen
TM
CTM
[Mn 4 ]
A B Mn2+
Fig. 2. Organization of specific regions dedicated to PSII assembly. Comparison of a biogenesis center in Synechocystis sp. PCC 6803 (A) and a T-zone in C. reinhardtii (B). For further expla- nation see Sections 3 and 4 of the text. OM: outer membrane, PM: plasma membrane, TM: thylakoid membrane, TC: thylakoid center, PDM: PratA-defined membrane, OE: outer envelope membrane, IE: inner envelope membrane, CTM: chloroplast translation membranes. (B) is adapted from Schottkowski et al. 2012.
membranes have now been investigated in cells grown under various conditions [100–103]. These analyses revealed that cyanobacterial thy- lakoids display heterogeneity with respect to the distribution of photo- synthetic complexes [101], This also holds for plasma membranes, which have also been shown to exhibit non-uniform protein distribu- tions [100]. Recently, comparative proteomic studies have been per- formed on membrane fractions which had been purified via the two- phase partitioning technique. Many of the subunits of thylakoid mem- brane complexes, including in particular PSI and the ATP synthase were identified in the plasma membrane fraction, leading to a model which predicts transient “hemifusions” between plasma and thylakoid membranes [104]. At these sites, it was proposed, that proteins are inserted into the membrane and subsequently directed to either the plasma membrane or thylakoids [104]. Perhaps the most striking example of structural and functional membrane heterogeneity is provided by the primordial cyanobacteri- um Gloeobacter violaceus. G. violaceus does not contain an internal thy- lakoid membrane system; instead its photosynthetic complexes are inserted into the plasma membrane [105,106]. As a consequence, pho- tosynthetically active patches with a mean diameter of 140 nm can be detected by spectroscopic means within the plasma membrane of G. violaceus (section 4, Fig. 3 and [107]). When these membranes are fractionated via sucrose-gradient centrifugation, a lighter, carotenoid- rich “orange” and a heavier “green” fraction are obtained, which have related lipid compositions. However, comparison of their protein and pigment compositions revealed that the orange membranes resemble the plasma membrane, while the green moiety is more reminiscent of photosynthetically active thylakoids [107]. The accumulation of PSII assembly factors in orange membranes further suggests that spatially separated biogenic and photosynthetically active membrane microdo- mains exist even in the plasma membrane of G. violaceus. These green microdomains are likely to represent the evolutionary starting point for the development of an internal thylakoid membrane system (section 4, Fig. 3 and [107]). Chloroplast outer and inner envelopes, as well as thylakoids, can be efficiently separated by centrifugation techniques, and their respective proteomes have been extensively characterized [28,108–110]. These data revealed that lipid and carotenoid synthesis is localized to the en- velope membranes. The spatial organization of chlorophyll synthesis is, however, more complex, since it is initiated in the stroma, proceeds at the envelope and is completed at thylakoids (Chapter 24 [111,112]). In C. reinhardtii, the integration of freshly synthesized D1 protein of PSII has been shown to take place at stromal thylakoids and the protein subsequently moves towards grana regions [113]. Moreover, mem- brane fractionation experiments using sucrose gradients have revealed the existence of membranes distinct from thylakoids in which putative RNA-binding regulators of chloroplast gene expression accumulate [114]. More recently, the concept of specialized chloroplast translation membranes (CTMs) in C. reinhardtii has been further elaborated by fol- lowing the distribution of translation factors, ribosomes and assembly factors of photosynthetic complexes in sucrose gradients after floata- tion of membranous material (Fig. 2B and [115]). CTMs were found to be denser than thylakoids due to their association with ribosomes, and contained translation and assembly factors for PSII including the psbD-specific translation activator RBP40, the aforementioned psbA translation factor DLA2 and the assembly protein HCF136 [25,115, 116]. Interestingly, no PSI subunits nor PSI assembly factors were im- munologically detectable in CTMs, indicating that this fraction repre- sents a PSII-specific biogenic compartment [115]. Moreover, proteomic analysis of the eyespot has identified the PSI assembly factor YCF4, leading to speculation that this alga-specific, carotenoid-rich structure might serve as the site of PSI biogenesis in C. reinhardtii [117,118]. In vascular plants, there is currently less evidence for the existence of related biogenic membrane regions. Ribosomes are excluded from grana regions for steric reasons, but bind to non-stacked stromal
thylakoids [119]. Here, co-translational insertion of membrane proteins is assumed to take place and subsequent assembly steps are likely to occur here too. This idea is supported by the finding that stromal thyla- koid fractions are enriched for PSII assembly intermediates [120]. By employing a detergent-free, two-phase partitioning approach, the dis- tribution of PSII subunits and PSII assembly factors in spinach thylakoid subfractions, i.e., grana core, grana margin and stromal thylakoids, has recently been assessed in detail [121]. Their content of several assembly factors was clearly enhanced relative to grana regions, which is compat- ible with a biogenic function for stromal thylakoids [121].
3.2. Cytological organization of thylakoid biogenesis
With the identification of factors involved in the synthesis and as- sembly of photosynthetic complexes, their subcellular/suborganellar localization could be determined using in situ FISH/IF techniques or GFP-based live-cell imaging. Pioneering work by Zerges and coworkers showed that, upon illumination at moderate light levels, both mRNAs and ribosomes in the chloroplasts of C. reinhardtii colocalize to two or three punctate regions at the periphery of its single pyrenoid, which were named T (for translation) zones (Fig. 2B and [122]). Strikingly, as in the case of CTMs (Section 3.1), only chloroplast mRNAs coding for subunits of PSII were found to be localized to T-zones. Therefore, it was suggested that CTMs represent T-zone-related membrane material [115]. This idea is further supported by the accumulation of PSII assem- bly intermediates around the pyrenoid in a PSII assembly mutant [122]. Moreover, the DLA2 subunit of pyruvate dehydrogenase (Section 2.2) has been identified as a critical determinant for light-dependent local- ized translation of the psbA mRNA at T-zones [25]. DLA2 binds to an A-rich element in the 5′ untranslated region of the psbA mRNA, cofractionates with CTMs and colocalizes with psbA mRNA at T-zones in a light- and acetate-dependent manner [25,123]. Most importantly, localization of psbA mRNA to T-zones is lost in DLA2 RNAi lines, and D1 synthesis is affected [25]. All these findings strongly suggest that DLA2 is involved in targeted translation of the psbA mRNA at T-zones and thereby determines the starting point of the path for PSII biogenesis [25,124]. Furthermore, application of translational elongation inhibitors revealed that accumulation of mRNAs in T-zones does not require active chloroplast protein synthesis, which rules out the alternative possibility that targeting is mediated via signals in the nascent polypeptide [125]. Recently, the T-zone model for the initial steps of PSII synthesis and assembly has been expanded by incorporating the later steps of PSII maturation [115]. IF-based localization experiments have defined an extended biogenesis region which includes the region in the vicinity of T-zones forming the junctions with thylakoid lobes [115]. At these junctions TOC and TIC components of the envelope's protein import machinery were shown to accumulate upon moderate illumination. This led to the hypothesis that the integration of nucleus-encoded PSII subunits – in particular the oxygen-evolving complex (OEC) subunits and LHC proteins – into an assembling PSII core complex is spatially con- nected to the T-zone area (Fig. 2B and [115]). As depicted in Fig. 2B, this would provide a continuous assembly pathway for PSII, without release or transfer of intermediates, even when subunits originating in the cyto- plasm must be incorporated. Whether similar coordination points are uti- lized in plastids of other organisms remains to be clarified. In cyanobacteria, the situation is less complex due to the lack of organelle-based intracellular compartmentalization. Nevertheless, as already suggested by the identification of biogenic PDM fractions (Section 3.1) a distinct cellular substructure in which the initial steps of PSII biogenesis take place has recently been defined in the model or- ganism Synechocystis 6803. Transmission electron microscopy in com- bination with immunogold labeling of the PSII assembly factor PratA revealed that this PDM marker protein accumulates at so-called “thyla- koid centers”, i.e., sites where the thylakoid membranes converge to- wards the plasma membrane (Fig. 2A and [126]). Thylakoid centers
situation in C. reinhardtii, other algae might contain CTMs that either surround or traverse the pyrenoid (Fig. 3). Pyrenoids are not found in the chloroplasts of higher plants. In etioplasts, however, prothylakoids emerge from the prolamellar body, which appears as a tubular network of membranes in electron micro- graphs [65,136]. Hence, one may speculate that, during thylakoid bio- genesis in developing etioplasts, prolamellar bodies play a structural role similar to that of pyrenoids in algal chloroplasts. In red algae a wide variety of different morphotypes of plastids and thylakoids exists. For instance, some unicellular species, like the model organism Cyanidiioschyzon merolae, lack a pyrenoid [137], while others, like Erythrolobus coxiae and thallus-forming species like Porphyra sp., harbor such a structure [138,139]. Indeed, E. coxiae has a plastid con- taining a central pyrenoid surrounded by at least four plastid lobes (Fig. 3 and [139]). The pyrenoids of these red algae are penetrated by several branched thylakoids, giving rise to a network comparable to the thylakoid tubule system in C. reinhardtii. Interestingly, the vast ma- jority of secondary plastids also contain pyrenoids but, unlike those in their evolutionary ancestors, these are not penetrated by loosely orga- nized tubular networks but by paired thylakoid stacks. In the case of the diatom Phaeodactylum tricornutum, a single thylakoid stack com- posed of two thylakoid layers extends through the pyrenoid (Fig. 3 and supp. Fig. 2 of [140]). In several Euglena species the pyrenoid is, however, traversed by more than one paired thylakoid stack (Fig. 3
and [141]). It is noteworthy that, in both diatoms and euglenoids, stro- mal thylakoid stacks are normally composed of three parallel thylakoid layers (Fig. 3 and [141,142]). In addition to its putative role in thylakoid membrane biogenesis, the pyrenoid of C. reinhardtii has been shown to be involved in the chloroplast's stress response [143]. Under oxidative stress, RNA gran- ules are formed within the internal perimeter of the pyrenoid, probably to protect mRNA molecules from damage by reactive oxygen species [143]. After release of stress, mRNAs are rapidly reincorporated into polysomes in a highly dynamic manner. These findings underline the significance of the pyrenoid as an alga-specific compartment that acts as an organizer for targeted chloroplast protein synthesis [143]. In contrast, there is so far little evidence for the presence of distinct suborganellar structures dedicated to thylakoid biogenesis in higher plants. One way of addressing this question at the molecular level would be to analyze homologs of factors that are known to mark these biogenic sites in algae and cyanobacteria, in particular, the PratA protein from Synechocystis 6803. However, no proteins that show any obvious similarities with PratA have been detected by bioinformatic means. Intriguingly, two factors, namely Rep27 from C. reinhardtii and Lpa1 from A. thaliana, belong to the tetratricopeptide repeat protein (TPR) family – like PratA – and both have been shown to interact with the D1 protein based on interaction studies and phenotypic assessment of the corresponding mutants [144–146]. These studies revealed a role
Cyanobacteria
Gloeobacter violaceus Synechocystis sp. PCC 6803
Primary Plastids
Secondary Plastids
Chlamydomonas reinhardtii Arabidopsis thaliana
Phaeodactylum tricornutum Euglena sp.
Cyanidioschyzon merolae Erythrodobus coxiae
Synechoccoccus sp. PCC 7942
Fig. 3. Architectures of thylakoid membranes. Models of the diverse structures of thylakoid membranes (green) in cyanobacteria and in primary and secondary plastids. Biogenesis center and T-zones are depicted in blue, pyrenoids are shown in yellow, envelope and plasma membranes are indicated in red.
for both proteins in facilitating the integration of nascent D1 into PSII pre-complexes. Moreover, recombinant Lpa1 possesses Mn2+^ binding activity, making this factor a good candidate for a marker which might enable one to determine the localization of PSII biogenesis sites in the chloroplasts of land plants [89,126].
- The shaping of thylakoid architecture
As summarized in Fig. 3, thylakoid membrane systems can undergo substantial architectural diversification. Whereas red algae and their descendants in the “red lineage” of algae with secondary plastids have retained phycobilisome-like structures during evolution to some ex- tent, green algae and, even more prominently, vascular plants devel- oped membrane-located light-harvesting complexes [142,147]. This transition opened new opportunities for the shaping of thylakoids by removing the steric constraints associated with the assembly of the rel- atively bulky phycobilisome structure at the surface of thylakoids. This has allowed vascular plants to form an even more highly curved struc- tural element of the thylakoid membrane, i.e., the granum [4]. Although the precise ultrastructure of grana regions remains a matter of debate [65,148–150], in principle, they consist of multiple stacks of flat mem- brane discs ca. 300–600 nm in diameter, which are linked by single stro- ma lamellae. Moreover, the photosynthetic complexes are unevenly distributed between the two subsystems, with PSII and LHCII being con- centrated in grana regions whereas PSI and the ATPase are mainly found in stroma lamellae — a condition referred to as lateral heterogeneity. The physicochemical forces that drive grana formation have been at- tributed to stromal moieties of LHCII proteins, which are proposed to determine membrane stacking of adjacent thylakoid discs [151,152]. However, a novel class of thylakoid-shaping proteins, with four mem- bers named CURVATURE THYLAKOID1A-D (CURT1A-D), has recently been identified in A. thaliana [65,153]. Genetic and biochemical evi- dence strongly suggests that CURT1 proteins localize to grana margin regions. Here, they induce extreme membrane bending that results in the formation of thylakoid discs, thereby determining the architecture of a granum [153]. Their margin-specific localization makes CURT pro- teins ideal markers for this thylakoid sub-fraction in land plants, and al- lows precise tracking of these membranes during fractionation experiments [130]. Moreover, recent work has revealed that the dy- namic organization of grana margin regions is a key factor in the control of the PSII repair process after photodamage [130]. Whether CURT1 is also related to biogenic processes at stromal thylakoids is not yet known, but the phenotype of CURT1 mutants lacking grana structures revealed that these are not required for the generation of thylakoids. Surprisingly, CURT1 homologs are present not only in grana- containing land plants and green algae, but also in cyanobacteria. Future work will doubtless uncover the functional role of this class of small thylakoid membrane proteins in the shaping of thylakoids.
Competing interest statement
The authors declare no competing interests.
Acknowledgements
We apologize to any authors whose work may not have been cited owing to length restrictions. We thank P. Hardy for the critical reading of the manuscript. This work was supported by the Deutsche Forschungsgemeinschaft in the context of the Forschergruppe FOR2092, project Ni390/9-1.
References
[1] M.F. Hohmann-Marriott, R.E. Blankenship, Evolution of photosynthesis, Annu. Rev. Plant Biol. 62 (2011) 515–548.
[2] H. Kirchhoff, S. Haferkamp, J.F. Allen, D.B.A. Epstein, C.W. Mullineaux, Protein diffu- sion and macromolecular crowding in thylakoid membranes, Plant Physiol. 146 (2008) 1571–1578. [3] J.F. Allen, W.B.M. de Paula, S. Puthiyaveetil, J. Nield, A structural phylogenetic map for chloroplast photosynthesis, Trends Plant Sci. 16 (2011) 645–655. [4] C.W. Mullineaux, Function and evolution of grana, Trends Plant Sci. 10 (2005) 521 – 525. [5] N. Nelson, C.F. Yocum, Structure and function of photosystems I and II, Annu. Rev. Plant Biol. 57 (2006) 521–565. [6] S. Eberhard, G. Finazzi, F.A. Wollman, The dynamics of photosynthesis, Annu. Rev. Genet. 42 (2008) 463–515. [7] E.-M. Aro, BBA Bioenergetics Special Issue. Chloroplast Biogenesis: 11. Photosystem II assembly and repair, BBA Bioenergetics. [8] Y. Choquet, BBA Bioenergetics Special Issue. Chloroplast Biogenesis: 13. Cytb6f bio- genesis, BBA Bioenergetics. [9] D. Leister, BBA Bioenergetics Special Issue. Chloroplast Biogenesis: 14. ATPase as- sembly, BBA Bioenergetics. [10] C. Lu, BBA Bioenergetics Special Issue. Chloroplast Biogenesis: 12. PSI assembly, BBA Bioenergetics. [11] L. Boudière, M. Michaud, D. Petroutsos, F. Rébeillé, D. Falconet, O. Bastien, S. Roy, G. Finazzi, N. Rolland, J. Jouhet, M.A. Block, E. Maréchal, Glycerolipids in photosynthe- sis: composition, synthesis and trafficking, Biochim. Biophys. Acta Bioenerg. 1837 (2014) 470–480. [12] K. Gounaris, J. Barber, Monogalactosyldiacylglycerol — the most abundant polar lipid in nature, Trends Biochem. Sci. 8 (1983) 378–381. [13] P. Jarvis, P. Dormann, C.A. Peto, J. Lutes, C. Benning, J. Chory, Galactolipid deficiency and abnormal chloroplast development in the Arabidopsis MGD synthase 1 mutant, Proc. Natl. Acad. Sci. U. S. A. 97 (2000) 8175–8179. [14] C. Bottier, J. Gean, F. Artzner, B. Desbat, M. Pezolet, A. Renault, D. Marion, V. Vie, Ga- lactosyl headgroup interactions control the molecular packing of wheat lipids in Langmuir films and in hydrated liquid-crystalline mesophases, Biochim. Biophys. Acta 1768 (2007) 1526–1540. [15] A.G. Lee, Membrane lipids: it's only a phase, Curr. Biol. 10 (2000) R377–R380. [16] K. Gounaris, C. Sundby, B. Andersson, J. Barber, Lateral heterogeneity of polar lipids in the thylakoid membranes of spinach-chloroplasts, FEBS Lett. 156 (1983) 170 – 174. [17] B. Demé, C. Cataye, M.A. Block, E. Maréchal, J. Jouhet, Contribution of galactoglycerolipids to the 3-dimensional architecture of thylakoids, FASEB J. 28 (2014) 3373–3383. [18] Y. Umena, K. Kawakami, J.-R. Shen, N. Kamiya, Crystal structure of oxygen-evolving photosystem II at a resolution of 1.9 A, Nature 473 (2011) 55–60. [19] N. Mizusawa, H. Wada, The role of lipids in photosystem II, Biochim. Biophys. Acta 1 (2012) 194–208. [20] B. Loll, J. Kern, W. Saenger, A. Zouni, J. Biesiadka, Lipids in photosystem II: interac- tions with protein and cofactors, Biochim. Biophys. Acta 6 (2007) 509–519. [21] K. Kobayashi, M. Kondo, H. Fukuda, M. Nishimura, H. Ohta, Galactolipid synthesis in chloroplast inner envelope is essential for proper thylakoid biogenesis, photo- synthesis, and embryogenesis, Proc. Natl. Acad. Sci. U. S. A. 104 (2007) 17216 – 17221. [22] I. Sakurai, N. Mizusawa, H. Wada, N. Sato, Digalactosyldiacylglycerol is required for stabilization of the oxygen-evolving complex in photosystem II, Plant Physiol. 145 (2007) 1361–1370. [23] M. Hagio, I. Sakurai, S. Sato, T. Kato, S. Tabata, H. Wada, Phosphatidylglycerol is es- sential for the development of thylakoid membranes in Arabidopsis thaliana, Plant Cell Physiol. 43 (2002) 1456–1464. [24] C. Benning, Mechanisms of lipid transport involved in organelle biogenesis in plant cells, Annu. Rev. Cell Dev. Biol. 25 (2009) 71–91. [25] A.V. Bohne, C. Schwarz, M. Schottkowski, M. Lidschreiber, M. Piotrowski, W. Zerges, J. Nickelsen, Reciprocal regulation of protein synthesis and carbon metabolism for thylakoid membrane biogenesis, PLoS Biol. 11 (2013) e1001482. [26] K. Kobayashi, S. Fujii, D. Sasaki, S. Baba, H. Ohta, T. Masuda, H. Wada, Transcription- al regulation of thylakoid galactolipid biosynthesis coordinated with chlorophyll biosynthesis during the development of chloroplasts in Arabidopsis, Front. Plant Sci. 5 (2014) 272. [27] Z. Wang, C. Benning, Chloroplast lipid synthesis and lipid trafficking through ER- plastid membrane contact sites, Biochem. Soc. Trans. 40 (2012) 457–463. [28] J. Joyard, M. Ferro, C. Masselon, D. Seigneurin-Berny, D. Salvi, J. Garin, N. Rolland, Chloroplast proteomics highlights the subcellular compartmentation of lipid me- tabolism, Prog. Lipid Res. 49 (2010) 128–158. [29] N. Rolland, G. Curien, G. Finazzi, M. Kuntz, E. Marechal, M. Matringe, S. Ravanel, D. Seigneurin-Berny, The biosynthetic capacities of the plastids and integration be- tween cytoplasmic and chloroplast processes, Annu. Rev. Genet. 46 (2012) 233 – 264. [30] J. Sarkis, J. Rocha, O. Maniti, J. Jouhet, V. Vie, M.A. Block, C. Breton, E. Marechal, A. Girard-Egrot, The influence of lipids on MGD1 membrane binding highlights novel mechanisms for galactolipid biosynthesis regulation in chloroplasts, FASEB J. 28 (2014) 3114–3123. [31] T.T. Selao, L. Zhang, C. Arioz, A. Wieslander, B. Norling, Subcellular localization of monoglucosyldiacylglycerol synthase in Synechocystis sp. PCC6803 and its unique regulation by lipid environment, PLoS ONE 9 (2014) e88153. [32] K. Awai, T. Kakimoto, C. Awai, T. Kaneko, Y. Nakamura, K.-i. Takamiya, H. Wada, H. Ohta, Comparative genomic analysis revealed a gene for monoglucosyldiacylglycerol synthase, an enzyme for photosynthetic membrane lipid synthesis in cyanobacteria, Plant Physiol. 141 (2006) 1120–1127. [33] J. Jouhet, E. Marechal, M.A. Block, Glycerolipid transfer for the building of mem- branes in plant cells, Prog. Lipid Res. 46 (2007) 37–55.
[95] M. Schottkowski, S. Gkalympoudis, N. Tzekova, C. Stelljes, D. Schunemann, E. Ankele, J. Nickelsen, Interaction of the periplasmic PratA factor and the PsbA (D1) protein during biogenesis of photosystem II in Synechocystis sp. PCC 6803, J. Biol. Chem. 284 (2009) 1813–1819. [96] M. Schottkowski, J. Ratke, U. Oster, M. Nowaczyk, J. Nickelsen, Pitt, a novel tetratri- copeptide repeat protein involved in light-dependent chlorophyll biosynthesis and thylakoid membrane biogenesis in Synechocystis sp. PCC 6803, Mol. Plant 2 (2009) 1289–1297. [97] J. Knoppova, R. Sobotka, M. Tichy, J. Yu, P. Konik, P. Halada, P.J. Nixon, J. Komenda, Discovery of a chlorophyll binding protein complex involved in the early steps of photosystem II assembly in Synechocystis, Plant Cell 26 (2014) 1200–1212. [98] J.W. Chidgey, M. Linhartova, J. Komenda, P.J. Jackson, M.J. Dickman, D.P. Canniffe, P. Konik, J. Pilny, C.N. Hunter, R. Sobotka, A cyanobacterial chlorophyll synthase-HliD complex associates with the Ycf39 protein and the YidC/Alb3 insertase, Plant Cell 26 (2014) 1267–1279. [99] B. Rengstl, U. Oster, A. Stengel, J. Nickelsen, An intermediate membrane subfraction in cyanobacteria is involved in an assembly network for photosystem II biogenesis, J. Biol. Chem. 286 (2011) 21944–21951. [100] R. Srivastava, T. Pisareva, B. Norling, Proteomic studies of the thylakoid membrane of Synechocystis sp. PCC 6803, Proteomics 5 (2005) 4905–4916. [101] R. Agarwal, A. Matros, M. Melzer, H.P. Mock, J.K. Sainis, Heterogeneity in thylakoid membrane proteome of Synechocystis 6803, J. Proteome 73 (2010) 976–991. [102] F. Huang, S. Fulda, M. Hagemann, B. Norling, Proteomic screening of salt-stress- induced changes in plasma membranes of Synechocystis sp. strain PCC 6803, Prote- omics 6 (2006) 910–920. [103] L.F. Zhang, H.M. Yang, S.X. Cui, J. Hu, J. Wang, T.Y. Kuang, B. Norling, F. Huang, Pro- teomic analysis of plasma membranes of cyanobacterium Synechocystis sp. Strain PCC 6803 in response to high pH stress, J. Proteome Res. 8 (2009) 2892–2902. [104] T. Pisareva, J. Kwon, J. Oh, S. Kim, C. Ge, A. Wieslander, J.S. Choi, B. Norling, Model for membrane organization and protein sorting in the cyanobacterium Synechocystis sp. PCC 6803 inferred from proteomics and multivariate sequence analyses, J. Proteome Res. 10 (2011) 3617–3631. [105] R. Rippka, J. Waterbury, G. Cohen-Bazire, A cyanobacterium which lacks thylakoids, Arch. Microbiol. 100 (1974) 419–436. [106] M. Mimuro, T. Tomo, T. Tsuchiya, Two unique cyanobacteria lead to a traceable ap- proach of the first appearance of oxygenic photosynthesis, Photosynth. Res. 97 (2008) 167–176. [107] S. Rexroth, C.W. Mullineaux, D. Ellinger, E. Sendtko, M. Rögner, F. Koenig, The plas- ma membrane of the cyanobacterium Gloeobacter violaceus contains segregated bioenergetic domains, Plant Cell Online 23 (2011) 2379–2390. [108] N. Rolland, A. Atteia, P. Decottignies, J. Garin, M. Hippler, G. Kreimer, S.D. Lemaire, M. Mittag, V. Wagner, Chlamydomonas proteomics, Curr. Opin. Microbiol. 12 (2009) 285–291. [109] S. Simm, D.G. Papasotiriou, M. Ibrahim, M.S. Leisegang, B. Muller, T. Schorge, M. Karas, O. Mirus, M.S. Sommer, E. Schleiff, Defining the core proteome of the chlo- roplast envelope membranes, Front. Plant Sci. 4 (2013) 11. [110] M. Tomizioli, C. Lazar, S. Brugiere, T. Burger, D. Salvi, L. Gatto, L. Moyet, L.M. Breckels, A.M. Hesse, K.S. Lilley, D. Seigneurin-Berny, G. Finazzi, N. Rolland, M. Ferro, Deciphering thylakoid sub-compartments using a mass spectrometry- based approach, Mol. Cell. Proteomics 13 (2014) 2147–2167. [111] B. Grimm, BBA Bioenergetics Special Issue. Chloroplast Biogenesis: 24. Molecular and function of Tetrapyrrole biogenesis, BBA Bioenergetics. [112] U. Eckhardt, B. Grimm, S. Hortensteiner, Recent advances in chlorophyll biosynthe- sis and breakdown in higher plants, Plant Mol. Biol. 56 (2004) 1–14. [113] N. Adir, S. Shochat, I. Ohad, Light-dependent D1 protein synthesis and transloca- tion is regulated by reaction center II. Reaction center II serves as an acceptor for the D1 precursor, J. Biol. Chem. 265 (1990) 12563–12568. [114] W. Zerges, J.D. Rochaix, Low density membranes are associated with RNA-binding proteins and thylakoids in the chloroplast of Chlamydomonas reinhardtii, J. Cell Biol. 140 (1998) 101–110. [115] M. Schottkowski, M. Peters, Y. Zhan, O. Rifai, Y. Zhang, W. Zerges, Biogenic mem- branes of the chloroplast in Chlamydomonas reinhardtii, Proc. Natl. Acad. Sci. U. S. A. 109 (2012) 19286–19291. [116] C. Schwarz, A.-V. Bohne, F. Wang, F.J. Cejudo, J. Nickelsen, An intermolecular disulfide-based light switch for chloroplast psbD gene expression in Chlamydomonas reinhardtii, Plant J. 72 (2012) 378–389. [117] G. Kreimer, The green algal eyespot apparatus: a primordial visual system and more? Curr. Genet. 55 (2009) 19–43. [118] V. Wagner, K. Ullmann, A. Mollwo, M. Kaminski, M. Mittag, G. Kreimer, The phosphoproteome of a Chlamydomonas reinhardtii eyespot fraction includes key proteins of the light signaling pathway, Plant Physiol. 146 (2008) 772–788. [119] Y. Yamamoto, R.C. Ford, J. Barber, Relationship between thylakoid membrane fluid- ity and the functioning of pea chloroplasts: effect of cholesteryl hemisuccinate, Plant Physiol. 67 (1981) 1069–1072. [120] R. Danielsson, M. Suorsa, V. Paakkarinen, P.A. Albertsson, S. Styring, E.M. Aro, F. Mamedov, Dimeric and monomeric organization of photosystem II. Distribution of five distinct complexes in the different domains of the thylakoid membrane, J. Biol. Chem. 281 (2006) 14241–14249. [121] M. Suorsa, M. Rantala, R. Danielsson, S. Jarvi, V. Paakkarinen, W.P. Schroder, S. Styring, F. Mamedov, E.M. Aro, Dark-adapted spinach thylakoid protein heterogeneity offers insights into the photosystem II repair cycle, Biochim. Biophys. Acta 9 (2014) 1. [122] J. Uniacke, W. Zerges, Photosystem II assembly and repair are differentially local- ized in Chlamydomonas, Plant Cell 19 (2007) 3640–3654. [123] F. Ossenbühl, K. Hartmann, J. Nickelsen, A chloroplast RNA binding protein from stromal thylakoid membranes specifically binds to the 5′ untranslated region of the psbA mRNA, Eur. J. Biochem. 269 (2002) 3912–3919.
[124] J. Nickelsen, W. Zerges, Thylakoid biogenesis has joined the new era of bacterial cell biology, Front. Plant Sci. 4 (2013) 458. [125] J. Uniacke, W. Zerges, Chloroplast protein targeting involves localized translation in Chlamydomonas, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 1439–1444. [126] A. Stengel, I.L. Gugel, D. Hilger, B. Rengstl, H. Jung, J. Nickelsen, Initial steps of pho- tosystem II de novo assembly and preloading with manganese take place in bio- genesis centers in Synechocystis, Plant Cell 24 (2012) 660–675. [127] D. Kunkel, Thylakoid centers: structures associated with the cyanobacterial photo- synthetic membrane system, Arch. Microbiol. 133 (1982) 97–99. [128] E. Fuhrmann, J.B. Bultema, U. Kahmann, E. Rupprecht, E.J. Boekema, D. Schneider, The vesicle-inducing protein 1 from Synechocystis sp. PCC 6803 organizes into di- verse higher-ordered ring structures, Mol. Biol. Cell 20 (2009) 4620–4628. [129] M. Rütgers, M. Schroda, A role of VIPP1 as a dynamic structure within thylakoid centers as sites of photosystem biogenesis? Plant Signal. Behav. 8 (2013) e27037. [130] S. Puthiyaveetil, O. Tsabari, T. Lowry, S. Lenhert, R.R. Lewis, Z. Reich, H. Kirchhoff, Compartmentalization of the protein repair machinery in photosynthetic mem- branes, Proc. Natl. Acad. Sci. U. S. A. 111 (2014) 15839–15844. [131] M. Yoshioka-Nishimura, D. Nanba, T. Takaki, C. Ohba, N. Tsumura, N. Morita, H. Sakamoto, K. Murata, Y. Yamamoto, Quality control of photosystem II: direct imag- ing of the changes in the thylakoid structure and distribution of FtsH proteases in spinach chloroplasts under light stress, Plant Cell Physiol. 55 (2014) 1255–1265. [132] L. Minai, K. Wostrikoff, F.A. Wollman, Y. Choquet, Chloroplast biogenesis of photo- system II cores involves a series of assembly-controlled steps that regulate transla- tion, Plant Cell 18 (2006) 159–175. [133] J. Komenda, M. Barker, S. Kuvikova, R. de Vries, C.W. Mullineaux, M. Tichy, P.J. Nixon, The FtsH protease slr0228 is important for quality control of photosystem II in the thylakoid membrane of Synechocystis sp. PCC 6803, J. Biol. Chem. 281 (2006) 1145–1151. [134] D.M. Sherman, T.A. Troyan, L.A. Sherman, Localization of mmbrane proteins in the cyanobacterium Synechococcus sp. PCC7942 (radial asymmetry in the photosyn- thetic complexes), Plant Physiol. 106 (1994) 251–262. [135] I. Ohad, P. Siekevitz, G.E. Palade, Biogenesis of chloroplast membranes: I. Plastid de- differentiation in a dark-grown algal mutant (Chlamydomonas reinhardtii), J. Cell Biol. 35 (1967) 521–552. [136] J. Rosinski, W.G. Rosen, Chloroplast development: fine structure and chlorophyll synthesis, Q. Rev. Biol. 47 (1972) 160–191. [137] A. Merola, R. Castaldo, P. De Luca, R. Gambardella, A. Musacchio, R. Taddei, Revision of Cyanidium caldarium. Three species of acidophilic algae, Giorn. Bot. Ital. 115 (1981) 189–195. [138] J.C. Ackland, J.A. West, J. Scott, G.C. Zuccarello, J. Broom, Biology of Porphyra pulchella sp. nov. from Australia and New Zealand, Algae 21 (2006) 193–208. [139] J.L. Scott, B. Baca, F.D. Ott, J.A. West, Light and electron microscopic observations on Erythrolobus coxiae gen.et sp. nov. (Porphyrideophyceae, Rhodophyta) from Texas U.S.A, Algae 21 (2006) 407–416. [140] M. Peschke, D. Moog, A. Klingl, U.G. Maier, F. Hempel, Evidence for glycoprotein transport into complex plastids, Proc. Natl. Acad. Sci. U. S. A. 110 (2013) 10860 – 10865. [141] E. Kusel-Fetzmann, M. Weidinger, Ultrastructure of five Euglena species positioned in the subdivision Serpentes, Protoplasma 233 (2008) 209–222. [142] B. Lepetit, R. Goss, T. Jakob, C. Wilhelm, Molecular dynamics of the diatom thyla- koid membrane under different light conditions, Photosynth. Res. 111 (2012) 245 – 257. [143] J. Uniacke, W. Zerges, Stress induces the assembly of RNA granules in the chloro- plast of Chlamydomonas reinhardtii, J. Cell Biol. 182 (2008) 641–646. [144] L. Peng, J. Ma, W. Chi, J. Guo, S. Zhu, Q. Lu, C. Lu, L. Zhang, LOW PSII ACCUMULA- TION1 is involved in efficient assembly of photosystem II in Arabidopsis thaliana, Plant Cell 18 (2006) 955–969. [145] S. Park, P. Khamai, J.G. Garcia-Cerdan, A. Melis, REP27, a tetratricopeptide repeat nuclear-encoded and chloroplast-localized protein, functions in D1/32-kD reaction center protein turnover and photosystem II repair from photodamage, Plant Phys- iol. 143 (2007) 1547–1560. [146] D. Dewez, S. Park, J.G. Garcia-Cerdan, P. Lindberg, A. Melis, Mechanism of REP protein action in the D1 protein turnover and photosystem II repair from photodamage, Plant Physiol. 151 (2009) 88–99. [147] E. Gantt, B. Grabowski, F.X. Cunningham Jr., Antenna systems of red algae: phycobilisomes with photosystem ll and chlorophyll complexes with photosystem I, Light-Harvesting Antennas in Photosynthesis, Springer, 2003, pp. 307–322. [148] R. Nevo, D. Charuvi, O. Tsabari, Z. Reich, Composition, architecture and dynamics of the photosynthetic apparatus in higher plants, Plant J. 70 (2012) 157–176. [149] H. Kirchhoff, Architectural switches in plant thylakoid membranes, Photosynth. Res. 116 (2013) 481–487. [150] B. Daum, W. Kuhlbrandt, Electron tomography of plant thylakoid membranes, J. Exp. Bot. 62 (2011) 2393–2402. [151] R. Fristedt, A. Willig, P. Granath, M. Crevecoeur, J.D. Rochaix, A.V. Vener, Phosphor- ylation of photosystem II controls functional macroscopic folding of photosynthetic membranes in Arabidopsis, Plant Cell 21 (2009) 3950–3964. [152] B. Daum, D. Nicastro, J. Austin, J.R. McIntosh, W. Kühlbrandt, Arrangement of pho- tosystem II and ATP synthase in chloroplast membranes of spinach and pea, Plant Cell Online 22 (2010) 1299–1312. [153] U. Armbruster, M. Labs, M. Pribil, S. Viola, W. Xu, M. Scharfenberg, A.P. Hertle, U. Rojahn, P.E. Jensen, F. Rappaport, P. Joliot, P. Dormann, G. Wanner, D. Leister, Arabidopsis CURVATURE THYLAKOID1 proteins modify thylakoid architecture by inducing membrane curvature, Plant Cell 25 (2013) 2661–2678.
