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ERROR: type should be string, got "https://doi.org/10.1093/plcell/koad157\n\nTHE PLANT CELL 2023: 35: 3236–3259\n\nw\ne\ni\nv\ne\nR\n\nThe pyrenoid: the eukaryotic CO2-concentrating\norganelle\n\nShan He\n\n,1,2 Victoria L. Crans\n\n1 and Martin C. Jonikas\n\n1,2,*\n\n1 Department of Molecular Biology, Princeton University, Princeton, NJ 08540, USA\n2 Howard Hughes Medical Institute, Princeton University, Princeton, NJ 08540, USA\n\n*Author for correspondence: mjonikas@princeton.edu\n\nAbstract\nThe pyrenoid is a phase-separated organelle that enhances photosynthetic carbon assimilation in most eukaryotic algae and\nthe land plant hornwort lineage. Pyrenoids mediate approximately one-third of global CO2 fixation, and engineering a pyrenoid\ninto C3 crops is predicted to boost CO2 uptake and increase yields. Pyrenoids enhance the activity of the CO2-fixing enzyme\nRubisco by supplying it with concentrated CO2. All pyrenoids have a dense matrix of Rubisco associated with photosynthetic\nthylakoid membranes that are thought to supply concentrated CO2. Many pyrenoids are also surrounded by polysaccharide\nstructures that may slow CO2 leakage. Phylogenetic analysis and pyrenoid morphological diversity support a convergent evo-\nlutionary origin for pyrenoids. Most of the molecular understanding of pyrenoids comes from the model green alga\nChlamydomonas (Chlamydomonas reinhardtii). The Chlamydomonas pyrenoid exhibits multiple liquid-like behaviors, includ-\ning internal mixing, division by fission, and dissolution and condensation in response to environmental cues and during the cell\ncycle. Pyrenoid assembly and function are induced by CO2 availability and light, and although transcriptional regulators have\nbeen identified, posttranslational regulation remains to be characterized. Here, we summarize the current knowledge of pyr-\nenoid function, structure, components, and dynamic regulation in Chlamydomonas and extrapolate to pyrenoids in other\nspecies.\n\nIntroduction\nPhotosynthesis forms the base of the food chain in most eco-\nsystems by converting CO2 from the environment into or-\nganic carbon. At the heart of these reactions is the enzyme\nRibulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco),\nwhich assimilates CO2 into sugar precursors used to generate\nbiomass (Bracher et al. 2017). Despite its crucial role in\nphotosynthesis, Rubisco has two limitations: (1) it has a\nslow catalytic rate for an enzyme in central carbon metabol-\nism; and (2) it can also catalyze oxygenation, a wasteful reac-\ntion that uses O2 instead of CO2 (Flamholz et al. 2019). A\ntradeoff between Rubisco’s catalytic rate and specificity for\nCO2 over O2 appears to prevent the evolution or engineering\nof Rubisco to be both fast and specific (Tcherkez et al. 2006;\nSavir et al. 2010; Flamholz et al. 2019). To keep oxygenation at\n\ntolerably low levels, many plants use a specific but slow form\nof Rubisco and compensate for its slow catalytic rate by pro-\nducing a large amount of the enzyme (Raven 2013). This\nstrategy requires significant cellular resources, including up\nto 25% of total leaf nitrogen (Raven 2013).\n\nSome photosynthetic organisms overcome the limitations\nof Rubisco by using a CO2-concentrating mechanism (CCM)\nto deliver concentrated CO2 to the enzyme. This concen-\ntrated CO2 increases the turnover rate of Rubisco, and the\nhigher ratio of CO2 to O2 favors carboxylation and suppresses\noxygenation (Badger et al. 1980; Kupriyanova et al. 2023).\nThere is currently great interest in understanding how\nCCMs work, both because of their significant ecological\nrole (Ehleringer et al. 1991; Badger et al. 2006; Meyer et al.\n2017) and because engineering a CCM into crops has the\n\nReceived February 23, 2023. Accepted May 17, 2023. Advance access publication June 4, 2023\n© The Author(s) 2023. Published by Oxford University Press on behalf of American Society of Plant Biologists.\nThis is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and\nreproduction in any medium, provided the original work is properly cited.\n\nOpen Access\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3237\n\npotential to increase yields (Matsuoka et al. 2001; Kajala et al.\n2011; Hanson et al. 2016; Hennacy and Jonikas 2020; Adler\net al. 2022).\n\nCCMs are categorized into two broad classes: biochemical\nand biophysical, depending on the nature of the intermedi-\nate molecules used to concentrate CO2. Biochemical\nCCMs, which\ninclude C4, C2, and crassulacean acid\nmetabolism (CAM), transiently fix CO2 into intermediate or-\nganic molecules such as oxaloacetate and malate, from which\nconcentrated CO2\nis released in proximity to Rubisco\n(Caemmerer and Furbank 2003; Sage et al. 2012; Heyduk\net al. 2019). By contrast, in biophysical CCMs, the only inter-\n−) (Hennacy and\nmediate molecule is bicarbonate (HCO3\nJonikas 2020). Biochemical CCMs are predominantly found\nin plants and typically involve multicellular structures,\nwhereas biophysical CCMs are predominantly found in mi-\ncrobes and operate at a single-cell level (Maberly and\nGontero 2017).\n\nBiophysical CCMs differ between prokaryotes and eukar-\nyotes. Both rely on a subcellular structure whose matrix\ncontains a high concentration of Rubisco, into which concen-\n− (Wang et al. 2015; Kaplan\ntrated CO2 is released from HCO3\n2017; Hennacy and Jonikas 2020; Adler et al. 2022; Ang et al.\n2022). However, the eukaryotic compartment known as\nthe pyrenoid (1-2 µm in diameter) is much bigger than the\nbacterial Rubisco-containing compartment, the carboxysome\nin diameter). Additionally, CO2 delivery to\n(∼200 nm\nRubisco in the two structures is thought to be achieved based\non different principles: in pyrenoids, CO2 delivery is mediated\nby thylakoid membranes, as discussed below (Fei et al. 2022),\n− diffus-\nwhereas in carboxysomes, CO2 is produced from HCO3\ning directly into the carboxysome matrix (Mangan et al. 2016).\nThis review will focus on the pyrenoid.\n\nPyrenoids are found inside the chloroplasts of most eu-\nkaryotic algae (including most microalgae and many macro-\nalgae) and some species of nonvascular land plants called\nhornworts (Villarreal and Renner 2012; Meyer et al. 2017; Li\net al. 2020). The pyrenoid is typically visible under light mi-\ncroscopy as a 1–2 µm punctum within the chloroplast\n(Fig. 1A).\n\nOne of the earliest records of a pyrenoid dates from 1782\nby the Danish naturalist and scientific illustrator Otto\nFrederik Müller, who drew unnamed puncta in sketches of\nthe green alga Spirogyra (formerly Conferva jugalis) (Müller\n1782), making it one of the first scientifically documented or-\nganelles. The pyrenoid was first described in a publication in\n1803 (Vaucher 1803). The term pyrenoid was conceived in\n1882 from the Greek πυρην (pyren, kernel) (Schmitz 1882).\nFrom this point on, the pyrenoid became the focus of\nmany classic morphological studies using light and electron\nmicroscopy. Further reading on the history of pyrenoid re-\nsearch can be found in a recent review by Barrett et al. and\na book chapter by Meyer et al. (Meyer et al. 2020a; Barrett\net al. 2021).\n\nResearch on pyrenoids has recently gained momentum\nand currently has three major motivations: (1) a growing ap-\npreciation for the major role of pyrenoids in the global car-\nbon cycle; (2) prospects to engineer pyrenoids into crops\nto increase yields; and (3) the unique value of the pyrenoid\nas a model for biological phase-separated condensates, a re-\ncently discovered ubiquitous class of organelles. We discuss\neach of these motivations below.\n\nPyrenoids play a major role in the global carbon cycle, me-\ndiating approximately 30% to 40% of global CO2 assimilation\neach year (Mackinder et al. 2016). Approximately one-half of\nglobal CO2 assimilation occurs in the oceans (Field et al. 1998;\n\nFigure 1. Structure of the Chlamydomonas pyrenoid. A) The Chlamydomonas pyrenoid is visible by light microscopy. Scale bar, 1 μm. B) The\nChlamydomonas pyrenoid is composed of three major compartments: the Rubisco matrix, thylakoid tubules, and starch sheath. Scale bar,\n200 nm. C) Two-dimensional and D) Three-dimensional models of the pyrenoid showing the major compartments and protein peripheral struc-\ntures. See Table 1 for a list of the known protein components of each structure. (The circled numbers indicating the sub-pyrenoid localizations in\npanel C are coordinated with the circled numbers in Table 1).\n\nACDB\f3238\n\n|\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\nHe et al.\n\nBehrenfeld et al. 2001), and most of this assimilation is attrib-\nuted to eukaryotic algae (Flombaum et al. 2013; Rousseaux\nand Gregg 2013), nearly all of which have pyrenoids (Mann\n1996; Not et al. 2004; Thierstein and Young 2004; Meyer\nand Griffiths 2013).\n\nThere is a growing interest in enhancing yields of major\nglobal crops that do not have CCMs by engineering a CCM\ninto them (Rae et al. 2017; Hennacy and Jonikas 2020).\nAmong the various CCMs that could be engineered, the\ngreen algal pyrenoid-based CCM is a particularly promising\ncandidate for engineering into non-CCM plants as a result\nof two attractive qualities: (1) it operates at the single-cell le-\nvel, which means that leaf anatomy does not need to be en-\ngineered as would be necessary for engineering of the C4\nCCM (Kajala et al. 2011); and (2) unlike the prokaryotic\ncarboxysome-based CCM, the green algal pyrenoid-based\nCCM is natively encoded in the eukaryotic nuclear genome,\nwhich could facilitate its engineering into monocot crops\nsuch as wheat (Triticum aestivum) and rice (Oryza sativa),\nwhose prokaryotic chloroplast genomes remain challenging\nto engineer.\n\nThe pyrenoid is also of interest from a fundamental science\nperspective, as it is a phase-separated organelle (Freeman\nRosenzweig et al. 2017). Biological phase separation underlies\nthe formation of many cellular structures (Shin and\nBrangwynne 2017). The pyrenoid is one of the few phase-\nseparated organelles where the functional value of conden-\nsate formation is understood, as there is a clear functional\nand fitness cost to preventing Rubisco condensation into a\nmatrix (Meyer et al. 2012; Mackinder et al. 2016; He et al.\n2020; Fei et al. 2022). Moreover, the pyrenoid of the model\nalga Chlamydomonas (Chlamydomonas reinhardtii) is one of\nthe structurally best understood phase-separated conden-\nsates, as its phase separation was reconstituted in vitro\n(Wunder et al. 2018) and the structural basis behind this phase\nseparation was determined (He et al. 2020), making it a power-\nful system for deriving the basic fundamental principles that\nunderlie the assembly of phase-separated organelles.\n\nThe vast majority of our molecular understanding of pyr-\nenoids comes from recent studies in Chlamydomonas. As a\nwell-established model organism widely used for photosyn-\nthesis studies, Chlamydomonas benefits from a thriving com-\nmunity of researchers who have produced genome\nsequences and annotations (Merchant et al. 2007; Craig\net al. 2023), genome-wide omics data (Brueggeman et al.\n2012; Fang et al. 2012; Zones et al. 2015; Strenkert et al.\n2019), mutant libraries (Li et al. 2016; Li et al. 2019), fluores-\ncently tagged lines for gene functional analysis (Mackinder\net al. 2017; Wang et al. 2022), as well as pyrenoid proteomes\nand a pyrenoid proxiome (Mackinder et al. 2017; Zhan et al.\n2018; Lau et al. 2023). Such resources are currently lacking for\nother algal species, although recent progress has been made\ntoward developing similar tools in model diatoms such as\nhigh-efficiency transformation protocols in Phaeodactylum\ntricornutum (Miyagawa et al. 2009), stably propagated epi-\nsomes in P. tricornutum and Thalassiosira pseudonana\n\n(Karas et al. 2015), proteome analyses of mitochondria and\nplastids in T. pseudonana (Schober et al. 2019), and a fluores-\ncent protein-tagging pipeline in T. pseudonana (Nam et al.\n2022).\n\nIn this review, we discuss the basic concepts of pyrenoid\nfunction as well as the current understanding of pyrenoids\nin Chlamydomonas, other algae, and hornworts. Some as-\npects of these topics have been covered in recent reviews\n(Barrett et al. 2021; Adler et al. 2022). Our review seeks to\nprovide an update on the most recent discoveries in the field\nand discuss the evolution, biogenesis, regulation, and func-\ntion of the pyrenoid-based CCM.\n\nOperating principles and evolution\nOperating principles of the pyrenoid-based CCM\nPhotosynthetic cells can obtain their carbon from two\n−. The availability\nsources in the environment: CO2 and HCO3\nof each source can vary depending on the environment (e.g.\naquatic growth or growth on surfaces exposed to air) and\nconditions (e.g. external pH). In the aquatic environment,\n− is normally more abundant than CO2, whereas CO2\nHCO3\nmay be more available to cells growing on the surface of par-\nticles in the soil.\n\nCO2 is difficult to concentrate directly within cells because\nit is a small, uncharged molecule that rapidly leaks across\nmembranes. Current models of the pyrenoid-based CCM\nsuggest that, to overcome this issue, cells use carbonic anhy-\n−, which\ndrases to convert CO2 to the charged molecule HCO3\ncannot easily diffuse across membranes and can be directed\nto subcellular compartments via transmembrane transpor-\nters. Intercompartmental pH differences are thought to\nplay a crucial role in this process by driving the interconver-\n− in the appropriate cellular compart-\nsion of CO2 to HCO3\nments (Fig. 2) (Hennacy and Jonikas 2020; Wang and\nJonikas 2020; Fei et al. 2022).\n\nA recent study (Fei et al. 2022) provides a detailed compu-\ntational model of the Chlamydomonas CCM that is consist-\nent with all available experimental evidence. We expect that\nthe model will also be generally relevant to other algae, as\nmost algae face similar biophysical challenges and the model\nis robust over broad parameter ranges.\n\nInterestingly, this model indicates that two distinct CCM\noperating modes are feasible, which share a common core\n− is accumulated in the chloroplast\nbut differ in how HCO3\nstroma (Fei et al. 2022) (Fig. 2). At the common core of the\n− is transported into\ntwo operating modes, stromal HCO3\nthe thylakoid lumen, likely by the bestrophin-like channels\nBST1 (encoded by Cre16.g662600), BST2 (Cre16.g663400),\nand/or BST3 (Cre16.g663450), although the role of each\nBST remains unclear (Mukherjee et al. 2019). Inside specia-\nlized pyrenoid-traversing regions of the thylakoid mem-\nbranes, carbonic anhydrase 3 (CAH3, Cre09.g415700)\n− to CO2, which is driven by the low pH of\nconverts HCO3\nthe thylakoid lumen (Karlsson et al. 1998; Hanson et al.\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3239\n\nstroma, as follows. The first mode uses a passive chloroplast\nCO2 uptake strategy, where CO2 passively diffuses from the\nperiplasm across the plasma membrane via the channel\nlow CO2-inducible 1 (LCI1, Cre03.g162800) (Ohnishi et al.\n2010; Kono and Spalding 2020) and across the chloroplast\nenvelope into the chloroplast stroma (Fig. 2A). CO2 diffusing\ninto the chloroplast or leaking out of the pyrenoid is con-\n− by the low-CO2-inducible B/C (LCIB/LCIC,\nverted to HCO3\nCre10.g452800/Cre06.g307500) carbonic anhydrase complex\nin a reaction driven by the high pH in the chloroplast stroma\n(Wang and Spalding 2006; Yamano et al. 2010; Jin et al. 2016;\nKasili et al. 2023).\n\nBy contrast, the second CCM operating mode uses an ac-\n− uptake strategy. CO2 is converted to\ntive chloroplast HCO3\n− at the periplasm by the carbonic anhydrases CAH1\nHCO3\n(Cre04.g223100) and CAH2 (Cre04.g223050) (Fujiwara et al.\n− crosses the plasma\n1990; Van and Spalding 1999). HCO3\nmembrane via the transporter high light activated 3 (HLA3,\nCre02.g097800) and is then concentrated across the chloro-\nplast envelope by LCIA (Cre06.g309000), which in this model\n− pump (Fig. 2B) (Miura et al. 2004; Yamano\nis an active HCO3\net al. 2015). We note that LCIA has not been experimentally\n− across a membrane, but ac-\nshown to actively pump HCO3\ntive pumping seems likely as the model indicates that passive\n− channels across the chloroplast envelope fail to\nHCO3\nachieve an effective CCM (Fei et al. 2022). CO2 leaking out\nof the pyrenoid is recaptured by the LCIB/LCIC carbonic an-\nhydrase complex, which relocalizes to the periphery of the\npyrenoid (Wang and Spalding 2014a) to enhance the effi-\n− to\nciency of CO2 recapture and avoid conversion of HCO3\nCO2 near the chloroplast envelope, which would lead to\n− (Fig. 2B) (Fei et al.\nloss of accumulated chloroplast HCO3\n2022).\n\nThe passive CO2 uptake strategy and active HCO3\n\n− uptake\nstrategy have different performance depending on external\nCO2 concentrations and pH (Fei et al. 2022). The passive\nCO2 uptake strategy is effective and energetically efficient\nunder ambient air levels of external CO2 (0.04%, 400 ppm;\nequivalent to 10 μM cytosolic in the model) but is unable\nto deliver enough CO2 to saturate Rubisco under lower levels\nof CO2 (0.004%, also known as “very low CO2”; corresponding\nto 1 μM cytosolic CO2\nin the model). Accordingly,\nChlamydomonas appears to use the passive CO2 uptake\nstrategy under air levels of external CO2 but not under\nvery low CO2, as evidenced by the severe growth defects of\nthe lcib mutant under air levels of CO2 but not very low\nCO2 (Wang and Spalding 2006, 2014a, 2014b; Duanmu\net al. 2009; Kono and Spalding 2020). In contrast to the pas-\nsive CO2 uptake strategy, modeling suggests that the active\n− uptake strategy can be effective and energetically effi-\nHCO3\ncient under both growth conditions (Fei et al. 2022).\nIntriguingly, despite the predicted good performance of the\n− uptake strategy under air levels of CO2 in silico,\nactive HCO3\nin vivo Chlamydomonas appears to reserve this strategy only\nfor very low CO2 conditions, as evidenced by O2 evolution ex-\nperiments (Wang and Spalding 2014a; Yamano et al. 2015;\n\nFigure 2. Operating principles of the pyrenoid-based CCM. The\nChlamydomonas CO2-concentrating mechanism is shown; the basic\nprinciples are likely to apply in other species, although due to conver-\ngent evolution, the specific proteins that mediate some of the reactions\nmay be phylogenetically unrelated to those in Chlamydomonas.\nMutant phenotypes and biophysical modeling (Fei et al. 2022) support\nthe existence of two operating modes of a pyrenoid-based\n− is ac-\nCO2-concentrating mechanism, which differ based on how HCO3\ncumulated in the chloroplast stroma. A) The first mode uses a passive\nchloroplast CO2 uptake strategy, where CO2 passively diffuses across\nthe chloroplast envelope into the stroma and is converted into\n− by the LCIB/LCIC carbonic anhydrase complex. This strategy is\nHCO3\nused under low CO2 (ambient air levels of external CO2). B) The second\n− uptake strategy, which relies on\nmode uses an active chloroplast HCO3\n− into the chloroplast. This strategy is used un-\nactive pumping of HCO3\nder very low CO2.\n\n2003; Blanco-Rivero et al. 2012; Burlacot et al. 2022). This CO2\ndiffuses out of the thylakoid membranes and into the pyre-\nnoid matrix, where it is captured by Rubisco. A CO2 leakage\nbarrier is thought to slow the escape of CO2 from the pyre-\nnoid, increasing CO2 concentration and decreasing energetic\ncosts (Fei et al. 2022). In the case of Chlamydomonas, mod-\neling and experimental evidence suggest that the starch\nsheath (Toyokawa et al. 2020; Fei et al. 2022) and thylakoid\nmembrane sheets (Fridlyand 1997; Fei et al. 2022) can serve\nas CO2 leakage barriers.\n\nThe two pyrenoid-based CCM operating modes use differ-\n− in the chloroplast\n\nent strategies to accumulate HCO3\n\nAB\f3240\n\n|\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\nHe et al.\n\nTable 1. Summary of Chlamydomonas pyrenoid–specific proteins whose localization has been confirmed.\n\nProtein\nname\n\nABCF6\nCPLD2\n\nEPYC1/\nLCI5\nHDA5\nrbcL\nRBCS1\nRBCS2\nRCA1\n\nGene ID\n\nSubpyrenoid localization\n\nLocalization reference\n\nReported or predicted functions\n\nCre06.g271850\nCre03.g206550\n\nCre10.g436550\n\nCre06.g290400\nCreCp.g802313\nCre02.g120100\nCre02.g120150\nCre04.g229300\n\nMatrix\nMatrix\n\nMatrix\n\nMatrix\nMatrix\n\nLau et al. (2023)\n\n①\n① Wang et al. (2022)\n\nPredicted ATP-binding cassette family F like protein\nHomolog of the Arabidopsis XuBP phosphatase\n\nCbbY (At3g48420)\n\n①\n\nMackinder et al. (2016)\n\nRubisco linker, phase-separates with Rubisco to form\n\nthe pyrenoid matrix\n\n① Wang et al. (2022)\n①\n\nHoldsworth (1971); Mackinder\n\net al. (2017)\n\nPredicted histone deacetylase\nLarge and small subunits of the Rubisco holoenzyme,\nwhich fixes CO2 and produces 3-PG and 2-PG\n\nMatrix\n\n①\n\nVladimirova et al. (1982);\n\nRubisco activase\n\nLacoste-Royal (1987); McKay\net al. (1991); Mackinder et al.\n(2017)\n\nSTR16\n\nCre13.g573250\n\nSTR18\n\nCre16.g663150\n\n–\n–\nCAH3\nCAS1\n\nCre16.g648400\nCre02.g143635\nCre09.g415700\nCre12.g497300\n\nCYN7\nCYN20-6\nDEG8\n\nCre12.g544150\nCre12.g544114\nCre01.g028350\n\nHCF136\nPSAH\nPSBP1\n\nCre06. g273700\nCre07. g330250\nCre12.g550850\n\nRBMP1\n\nCre06. g261750\n\nTEF14\n\nCre06.g256250\n\n-\n\nCre03.g172700\n\nCGLD14\n\nCre10.g446350\n\nPNU1\n\nCre03.g183550\n\nRBMP2\n\nCre09.g416850\n\nPSBP4\n\nCre08.g362900\n\nSAGA1\n\nCre11.g467712\n\nSAGA2\n\nCre09.g394621\n\nSMC7\n\nCre17.g720450\n\nSBE3\nSTA2\n\nCre10.g444700\nCre17.g721500\n\nMatrix\n\nMatrix\n\nMatrix\nMatrix\nTubules\nTubules\n\nTubules\nTubules\nTubules\n\nTubules\nTubules\nTubules\n\nTubules\n\nTubules\n\nTubules\n\nPyrenoid center\n(reticulated region)\nPyrenoid center\n(reticulated region)\nTubules (reticulated\nregion)\nThylakoid lumen\npuncta\nPuncta surrounding the\nmatrix\nInterface between\nmatrix and starch\nsheath\nPuncta surrounding the\nmatrix\nStarch sheath\nStarch sheath\n\n⑤\n\n⑨\n\n④\n\n④\n\n④\n\n③\n③\n\n① Wang et al. (2022); Lau et al.\n\nPredicted thiosulfate sulfurtransferase containing a\n\n(2023)\n\nrhodanese domain\n\n① Wang et al. (2022); Lau et al.\n\nPredicted thiosulfate sulfurtransferase containing a\n\n(2023)\n① Wang et al. (2022)\n① Wang et al. (2022)\n②\n② Wang et al. (2016)\n\nSinetova et al. (2012)\n\n② Wang et al. (2022)\n② Wang et al. (2022)\n② Wang et al. (2022)\n\n② Wang et al. (2022)\n②\n② Wang et al. (2022)\n\nMackinder et al. (2017)\n\nrhodanese domain\n\n–\n–\nAlpha-type carbonic anhydrase\nCalcium-mediated regulator of the expression of\n\nsome CCM-related genes\n\nPredicted peptidyl-prolyl cis-trans isomerase\nPredicted peptidyl-prolyl cis-trans isomerase\nPredicted DegP-type protease, Arabidopsis homolog\nis involved in the degradation of photodamaged\nPSII reaction center protein D1\nPS II stability/assembly factor HCF136\nPSI subunit\nPredicted oxygen-evolving enhancer protein 2 of PS\n\nII\n\n②\n\nMeyer et al. (2020b)\n\nProposed to mediate pyrenoid matrix connection to\n\ntubules\n\n② Wang et al. (2022)\n\nPredicted thylakoid-luminal protein, no labeled\n\ndomains\n\n②\n\nLau et al. (2023)\n\nProtein with multiple predicted Rubisco-binding\n\nmotifs\n\n⑤ Wang et al. (2022)\n\nPSBP domain-containing protein 3, conserved in the\n\n⑤ Wang et al. (2022)\n\nPROTEIN F23H11.5, has a bifunctional nuclease\n\ngreen lineage and diatoms\n\ndomain\n\nMeyer et al. (2020b)\n\nProposed to mediate pyrenoid matrix connection to\n\ntubules\n\nMackinder et al. (2017)\n\nLuminal PsbP-like protein, Arabidopsis homolog is\n\nessential for PS I assembly and function\n\nItakura et al. (2019); Meyer et al.\n\nProposed to mediate adherence of the starch sheath\n\n(2020b)\n\nto the matrix\n\nMeyer et al. (2020b)\n\nProposed to mediate adherence of the starch sheath\n\nto the matrix\n\nLau et al. (2023)\n\nProposed to have a similar function as SAGA1\n\nMackinder et al. (2017)\nMackinder et al. (2017)\n\n–\n\n–\n\nCre09.g394547\n\nStarch sheath\n\n③ Wang et al. (2022)\n\nCre09.g415600\n\nStarch sheath\n\n③ Wang et al. (2022)\n\nConserved starch-branching enzyme\nGranule-bound starch synthase involved in amylose\nbiosynthesis and the biosynthesis of long chains in\namylopectin\n\nPredicted cyclomaltodextrin glucanotransferase/\n\ncyclodextrin glycosyltransferase\n\nPredicted glucan 1,4-alpha-glucosidase/Lysosomal\nalpha-glucosidase; has a starch-binding domain\n\n(continued)\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3241\n\nTable 1. (continued)\n\nGene ID\n\nSubpyrenoid localization\n\nLocalization reference\n\nReported or predicted functions\n\nProtein\nname\n\nLCI9\n\nLCIB\n\nCre09.g394473\n\nCre10.g452800\n\nLCIC\n\nCre06.g307500\n\n–\n\nCre09.g394510\n\nGaps between starch\nplates\nGaps between starch\nplates and tubules (very\nlow CO2)\nGaps between starch\nplates and tubules (very\nlow CO2)\nStarch-matrix interface\nand gaps between\nstarch plates\n\n⑦\n\n⑧\n\n⑧\n\nMackinder et al. (2017)\n\nContains 2 starch-binding domains; may help ensure\n\nYamano et al. (2010); Wang and\nSpalding (2014b); Mackinder\net al. (2017)\n\na close fit for adjacent starch plates\n\nBeta-type carbonic anhydrase\n\nYamano et al. (2010); Mackinder\n\nBeta-type carbonic anhydrase\n\net al. (2017)\n\n④⑦⑧ Lau et al. (2023)\n\nContains a CBM20 starch-binding domain and a\nt-SNARE domain; proposed to be involved in\nmembrane remodeling of the pyrenoid tubules;\ncould be involved in membrane remodeling of the\npyrenoid tubules\nMalate dehydrogenase\nHomolog of the Arabidopsis chloroplast division site\n\nregulator MinD1\n\nMDH1\nMIND1\n\nCre03.g194850\nCre12.g522950\n\nPyrenoid periphery\nPyrenoid periphery\n\n–\n–\n\nWang et al. (2022)\nWang et al. (2022)\n\nThe circled numbers indicating the subpyrenoid localizations are coordinated with the circled numbers in Fig. 1C.\n\nKono and Spalding 2020). A possible explanation for this ob-\nservation is that when Chlamydomonas is grown under air\n− uptake strategy may\nlevel CO2 conditions, the active HCO3\nincur additional energetic costs beyond those accounted\nfor in the model. Indeed, the model only considered energet-\nic costs in the chloroplast, and it is possible that under cer-\n− uptake strategy requires\ntain conditions the active HCO3\nenergetic input outside the chloroplast, for example to\npump HCO3\n\n− across the plasma membrane.\n\nThe operation of a pyrenoid-based CCM under either air or\nvery low CO2 is estimated to be feasible for as little as the en-\nergetic equivalent of approximately 1 ATP per CO2 fixed (Fei\net al. 2022), making it energetically inexpensive relative to the\noverall cost of CO2 fixation by the Calvin-Benson-Bassham\ncycle, which is approximately energetically equivalent to 9\nATPs per CO2 fixed (Mangan et al. 2016).\n\nThe energy for operating the CCM must ultimately come\nfrom the light reactions, which directly drive the pH differ-\nence between the thylakoid lumen and stroma and indirectly\nmaintain the pH of other compartments and drive the activ-\nities of transporters. A recent study suggested that the pro-\n− to CO2 in the\ntons needed to drive conversion of HCO3\nthylakoid lumen are produced by photosynthetic cyclic elec-\ntron flow mediated by proton gradient regulation-like 1\n(PGRL1, Cre07.g340200) and pseudocyclic electron flow re-\nsulting from O2 photoreduction mediated by flavodiiron\nand Cre16.g691800)\nproteins\n(Burlacot et al. 2022). The same study also found that\nchloroplast-to-mitochondria electron flow contributes to\nenergizing the CCM, potentially by supplying ATP to drive\ntransporters.\n\n(FLVs, Cre12.g531900\n\nRubisco fixes CO2 through carboxylation of ribulose-1,5-bi-\nsphosphate (RuBP) to produce 3-phosphoglycerate (3-PG or\n3-PGA). While some 3-PG goes on to other parts of metabol-\nism, most of it is metabolized in the Calvin-Benson-Bassham\ncycle to regenerate RuBP, allowing the cycle to continue\n\n(Calvin 1962). Interestingly, in Chlamydomonas, Rubisco is\nthe only enzyme of the Calvin-Benson-Bassham cycle found\nin the pyrenoid; all other Calvin-Benson-Bassham cycle en-\nzymes and associated regulatory proteins that have been lo-\ncalized are enriched in a region of the stroma immediately\nsurrounding the pyrenoid (Fig. 1) (Küken et al. 2018; Wang\net al. 2022). Thus, the Rubisco substrate RuBP and its product\n3-PG need to exchange efficiently between the stroma and\nthe pyrenoid. The pathway and mechanism of this exchange\nare currently unknown in any organism with a pyrenoid.\n\nPyrenoids are likely the product of convergent\nevolution\nThe predominant theory regarding the origins of pyrenoids is\nbased on historical changes in atmospheric CO2 and O2 con-\ncentrations and the evolution of polyphyletic algal lineages.\nOxygenic photosynthesis first evolved in cyanobacteria ap-\nproximately 3 billion years ago (bya) (Schirrmeister et al.\n2015) at a time when atmospheric CO2 concentrations\nwere high and O2 concentrations were low (Fig. 3). The ad-\nvent of oxygenic photosynthesis led to the Great Oxidation\nEvent approximately 2.4 bya, when atmospheric O2 concen-\ntrations first began to rise (Anbar et al. 2007). Eukaryotic al-\ngae are thought to have first evolved approximately 1.5 to 2.0\nbya (Yoon et al. 2004; Sánchez-Baracaldo et al. 2017; Strassert\net al. 2021), a time when the atmospheric CO2:O2 ratio was\nstill high and CCMs were likely not necessary for efficient\ngrowth (Raven et al. 2017). Over the course of approximately\nthe next billion years, a diverse set of algal lineages arose\nseries of endosymbiotic events\nthrough a complex\n(Falkowski et al. 2004; Reyes-Prieto et al. 2007; Keeling\n2010; Dorrell et al. 2017; Jackson et al. 2018; Strassert et al.\n2021). These different lineages can be split into two broad\ngroups, green lineage algae and red lineage algae, which are\n\n\f3242\n\n|\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\nHe et al.\n\nFigure 3. Pyrenoids appear to have convergently evolved in response to declining atmospheric CO2 levels. Approximate CO2 and O2 concentrations\nover time (Berner 2006; Whitney et al. 2011) are correlated with the phylogenetic tree of photosynthetic eukaryotes below (Strassert et al. 2021).\nBranch points correlate with the approximate timing of the divergence of different groups (see Strassert et al. 2021 and Bowles et al. 2022 for dis-\ncussions on uncertainties regarding branch points). Asterisks denote the approximate timing of the acquisition of plastids through primary (*) or\nsecondary (**) endosymbiosis (Jackson et al. 2018; Strassert et al. 2021). The blue shade highlights the proposed range for the timing of CCM evo-\nlution in different photosynthetic species (Villarreal and Renner 2012; Meyer et al. 2020a). Green and red lineages are denoted as green or purple\ntext, respectively (most dinoflagellates have red plastids with the exception of Lepidodinium sp., which have green plastids) (Kamikawa et al. 2015).\nRepresentative electron micrographs of pyrenoids are shown below cartoons of four general pyrenoid types, displaying the wide variety of morph-\nologies observed in each algal lineage and the hornworts. Roman numerals on the electron micrographs denote references to their original pub-\nlications as follows: (I) Kusel-Fetzmann and Weidinger 2008; (II) Nudelman et al. 2006; (III) Zhang et al. 2008; (IV) van Baren et al. 2016; (V)\nBorowitzka 2018; (VI) Goudet et al. 2020; (VII) Mikhailyuk et al. 2014; (VIII) Duff et al. 2007; (IX) Hall and Claus 1963; (X) Nelson and Ryan 1988;\n(XI) Ford 1984; (XII) Laza-Martínez et al. 2012; (XIII) Clay and Kugrens 1999; (XIV) Shiratori et al. 2017; (XV) Ota et al. 2007; (XVI) Schnepf and\nElbräChter 1999; (XVII) Kowallik 1969; (XVIII) Hansen and Daugbjerg 2009; (XIX) Decelle et al. 2021; (XX) Bedoshvili et al. 2009; (XXI) Bedoshvili\nand Likhoshway 2012; (XXII) Buma et al. 2000; (XXIII) Fresnel and Probert 2005. This figure was created with BioRender.\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3243\n\ndistinguished by their use of different chlorophyll accessory\npigments (Falkowski et al. 2004; Keeling 2010).\n\nDuring the time that these algal lineages were evolving,\nCO2 concentrations were trending downward (Berner and\nKothavala 2001; Berner 2006), a phenomenon accelerated\nby the evolution of land plants between 500 and 360 million\nyears ago (mya), after each algal lineage had already been es-\ntablished (Fig. 3) (Berner 1997; Morris et al. 2018). This de-\ncrease in atmospheric CO2 and the simultaneous increase\nin atmospheric O2 are thought to be the main driving forces\nfor the evolution of CCMs in aquatic microorganisms, leading\nto the theory that pyrenoids and other CCMs evolved inde-\npendently via convergent evolution (Villarreal and Renner\n2012; Rae et al. 2013; Raven et al. 2017; Meyer et al. 2020a).\nThis convergent evolution theory potentially explains why\npyrenoids first evolved, but it remains difficult to pinpoint\nthe exact timing of their origin. In the absence of concrete\nevidence from the fossil record (Knoll 1992), previous reviews\nhave estimated the most likely timeline for CCM evolution\nby considering how various factors—including fluctuating\nCO2 and O2 concentrations, temperature changes, nutrient\nlevels, and the kinetic properties of different forms of\nRubisco—would have influenced the growth advantage con-\nferred by a CCM (Griffiths et al. 2017; Raven et al. 2017; Meyer\net al. 2020a). These reviews estimate that CCMs may have\nevolved in cyanobacteria and algae approximately 300 to\n450 mya (Badger and Price 2003; Griffiths et al. 2017), likely\nwhen the atmospheric CO2 concentration was 2 to 16 times\nthe present level (Raven et al. 2017). In hornworts (discussed\nfurther below), pyrenoids are estimated to have evolved ap-\nproximately 100 mya (Villarreal and Renner 2012). These es-\ntimates all correspond\ntime when different\nphotosynthetic lineages were already established and sup-\nport the convergent evolution theory.\n\nto a\n\nThe convergent evolution theory can be tested by compar-\ning the sequences of proteins thought to perform the same\nfunctions in the pyrenoids of phylogenetically distant algal\nspecies, but this is currently difficult to do because the mo-\nlecular composition of most pyrenoids is unknown. There\nare, however, three lines of molecular evidence that support\nthe convergent evolution theory. The first\nis that\nthylakoid-luminal carbonic anhydrases of different types\nare necessary for CCM function in different lineages: the\nChlamydomonas CCM requires the alpha-type carbonic an-\nhydrase CAH3 (Karlsson et al. 1998; Hanson et al. 2003;\nSinetova et al. 2012), whereas the diatom P. tricornutum re-\nquires a theta-type carbonic anhydrase (Kikutani et al.\n2016; Matsuda et al. 2017).\n\nThe second piece of molecular evidence that supports con-\nvergent evolution is based on the different forms of Rubisco\nacross lineages. There are at least four distinct types of\nRubisco enzymes within algae and cyanobacteria, which dif-\nfer greatly in their kinetic properties and holoenzyme struc-\nture (Badger et al. 1998). The fact that pyrenoids in different\nlineages package vastly different Rubisco holoenzymes sup-\nports the theory that they evolved convergently.\n\nis\n\nto\n\nThe\n\nrelated\n\nthird piece of evidence\n\nthe\nChlamydomonas protein Essential Pyrenoid Component 1\n(EPYC1, Cre10.g436550, also known as LCI5) (Turkina et al.\n2006; Mackinder et al. 2016), which is a linker protein that\nclusters Rubisco together to form the pyrenoid matrix\n(Mackinder et al. 2016; He et al. 2020). EPYC1 is necessary\nfor the Chlamydomonas CCM, but no homologs of this pro-\ntein could be identified in algae beyond the closely related\nVolvocales (Mackinder et al. 2016), suggesting that its func-\ntion is performed by other proteins that may have conver-\ngently evolved in different algal species. Repeat proteins\nwith similar predicted properties as EPYC1 have been identi-\nfied in other algae (Mackinder et al. 2016), and work is on-\nlinker\ngoing to characterize these and other putative\nproteins.\n\nIn addition to research on the algal CCM, several studies\nhave been conducted on the evolution of CCMs in horn-\nworts, which are the only land plants known to have pyre-\nnoids. Hornworts are nonvascular plants thought to have\nbeen important in the water-to-land transition during em-\nbryophyte evolution (Qiu et al. 2006). The first hornwort pyr-\nenoids evolved approximately 100 mya (Villarreal and Renner\n2012), coinciding with a drastic decline in atmospheric CO2\nlevels (Fig. 3). The presence of a pyrenoid in hornworts is cor-\nrelated with CCM activity detected by organic isotope dis-\ncrimination and mass spectrometry analyses (Smith and\nGriffiths 1996a, 1996b, 2000; Hanson et al. 2002; Meyer\net al. 2008), suggesting that pyrenoids play a similar role in\nhornwort CCMs as they do in algal CCMs. Phylogenetic evi-\ndence and ultrastructural data suggest that hornwort pyre-\nnoids were gained and lost 5 to 6 times independently\nsince they first appeared (Villarreal and Renner 2012).\nInterestingly, the distribution of pyrenoids across the green\nlineage of algae also suggests that multiple independent\nlosses and gains have occurred (Meyer and Griffiths 2013).\nThe environmental factors favoring pyrenoid loss remain\nunclear.\n\nStructure and components\nPyrenoid morphology can vary greatly depending on the spe-\ncies (Fig. 3), but the one unifying feature of all pyrenoids is\nthe Rubisco matrix, which contains densely packed Rubisco\n(Holdsworth 1971; Borkhsenious et al. 1998). Most algae\nhave one matrix per cell, although some species have mul-\ntiple Rubisco matrices that can vary in size and shape\n(Meyer et al. 2020a). The Rubisco matrix in all observed spe-\ncies is associated with thylakoid membranes (Meyer et al.\n2017), which are thought to deliver CO2 to Rubisco (Fig. 2)\n(Hennacy and Jonikas 2020). The simplest pyrenoids consist\nof a Rubisco matrix either embedded between thylakoid\nmembranes, such as that of the coccolithophore Emiliania\nhuxleyi (Buma et al. 2000), or projecting out of the chloro-\nplast into the cytoplasm, such as that of the diatom\nAttheya ussurensis (Bedoshvili et al. 2009) (Fig. 3). In most\nspecies, the thylakoid membranes traverse the pyrenoid\n\n\f3244\n\n|\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\nHe et al.\n\nmatrix either as sheets, as in the euglenophyte Euglena carter-\nae (Kusel-Fetzmann and Weidinger 2008), or tube-like struc-\ntures, as in the chlorophyte Heveochlorella hainangensis\n(Zhang et al. 2008). In some species, Rubisco matrices lack\ntraversing thylakoids but are surrounded by polysaccharide\ndeposits that potentially act as CO2 leakage barriers, as in\nthe chlorophyte Micromonas pusilla (van Baren et al. 2016).\nThe most elaborate pyrenoid morphologies consist of all\nthree sub-structures: thylakoids traversing a Rubisco matrix\nin\nthat\nChlamydomonas (Fig. 1B). In this section, we describe what\nis known about each of the pyrenoid sub-compartments in\nChlamydomonas and give a brief introduction to what is\nknown about these structures in other species.\n\nin a starch sheath, as\n\nis encased\n\nfound\n\nThe CO2-fixing pyrenoid matrix is a phase-separated\ncondensate of Rubisco and a linker protein\nRubisco makes up approximately 90% of the protein content\nof the pyrenoid matrix (Holdsworth 1971). Based on trans-\nmission electron microscopy (TEM) images, the matrix ap-\nin several species (Holdsworth 1968;\npears crystalline\nKowallik 1969; Bertagnolli and Nadakavukaren 1970) and\namorphous in others (Griffiths 1970; Meyer et al. 2012).\n\nThe Chlamydomonas pyrenoid matrix was recently shown\nto be a liquid-like phase-separated condensate (Freeman\nRosenzweig et al. 2017). Fluorescence recovery after photo-\nbleaching experiments indicated that the matrix mixes intern-\nally on a timescale of approximately 20 seconds (Freeman\nRosenzweig et al. 2017), similar to that observed for other\nliquid-like compartments such as P granules and nucleoli\n(Bracha et al. 2019). Furthermore, the Rubisco matrix exhibits\nother liquid-like behaviors, including division by fission, and\ndissolution into the chloroplast during cell division and\nunder high CO2 conditions (>0.40% CO2). Rubisco in the\nChlamydomonas pyrenoid matrix was found by cryo-electron\ntomography (cryo-ET) to lack the long-range order character-\nistic of a crystal; instead, the distribution of Rubisco fits well\nwith a simple model for the distribution of particles in a liquid\n(Freeman Rosenzweig et al. 2017). This description of the pyr-\nenoid matrix as a phase-separated condensate likely applies to\npyrenoids in other species, as it explains observations such as\nthe spheroidal shape of most pyrenoids (Fig. 3), the rapid ap-\npearance and disappearance of pyrenoids during cell division\n(Brown et al. 1967; Retallack and Butler 1970), and their div-\nision by fission (Brown and Bold 1964; Brown et al. 1967).\n\nFor decades, only two matrix proteins were known:\nRubisco and Rubisco activase (RCA1, Cre04.g229300)\n(Holdsworth 1971; Vladimirova et al. 1982; McKay and\nGibbs 1991), and the mechanism by which Rubisco is densely\nclustered in the pyrenoid matrix was a mystery. In 2016, the\nrepeat protein EPYC1 was proposed to link individual\nRubiscos\nin Chlamydomonas\nthe matrix\n(Mackinder et al. 2016). EPYC1 localizes to the pyrenoid ma-\ntrix and is one of the most abundant proteins in the pyrenoid\n\nform\n\nto\n\nafter Rubisco (Mackinder et al. 2016; Hammel et al. 2018). In\nepyc1 mutant cells, the majority of Rubisco is dispersed in the\nchloroplast outside of the pyrenoid, indicating that EPYC1\nplays a major role in Rubisco localization to the matrix\n(Mackinder et al. 2016). Purified Rubisco and EPYC1 can\nphase-separate with each other to form liquid-like droplets\nin vitro (Wunder et al. 2018), suggesting that these two pro-\nteins are sufficient for driving the formation of the liquid-like\npyrenoid matrix.\n\nRubisco is an oligomeric holoenzyme with eight identical\nlarge subunits and eight identical small subunits. A structural\nstudy using cryo-electron microscopy (cryo-EM) found that\nEPYC1 directly binds to Rubisco on the two alpha-helices\nof each Rubisco small subunit through salt bridges and\nhydrophobic interactions (Fig. 4) (He et al. 2020). This struc-\nture is consistent with previous genetic studies showing that\nthese alpha helices are important for the formation of the\npyrenoid and for the Rubisco–EPYC1 interaction (Meyer\net al. 2012; Atkinson et al. 2019). Each Rubisco holoenzyme\nhas eight EPYC1-binding sites and each EPYC1 has five\nRubisco-binding sites (Fig. 4, C to E), allowing the two pro-\nteins to form an interdependent network that clusters\nRubisco together in the pyrenoid matrix. The low binding\naffinity (approximately 3 mM) between individual EPYC1–\nRubisco binding site is consistent with the principle that bio-\npolymer phase separation is mediated by weak multivalent\ninteractions (Li et al. 2012).\n\nin\n\nin\n\nlocalization or enrichment\n\nthe Plantae and diatoms 2\n\nBeyond Rubisco, RCA1, and EPYC1, ten additional proteins\nshow exclusive\nthe\nChlamydomonas pyrenoid matrix when examined as fluores-\ncently tagged proteins (Fig. 1; Table 1). These proteins are the\nputative S-adenosyl-L-methionine-dependent methyltrans-\nferase SMM7 (Cre03.g151650) (Mackinder et al. 2017), the\npredicted xylulose-1,5-bisphosphate (XuBP) phosphatase\n(CPLD2,\nconserved\nCre03.g206550), the putative histone deacetylase HDA5\n(Cre06.g290400), uncharacterized proteins encoded by\nCre16.g648400, Cre13.g573250, Cre16.g663150, Cre02.g143635\n(Wang et al. 2022), thiosulfate sulfurtransferase16 (STR16,\nCre13.g573250), STR18 (Cre16.g663150), and ATP-binding\ncassette F like-protein 6 (ABCF6, Cre06.g271850) (Lau et al.\n2023) (Table 1). CPLD2 is the homolog of the highly selective\nArabidopsis XuBP phosphatase AtCbbY (At3g48420), which\nconverts the Rubisco inhibitor XuBP to a non-inhibitory\ncompound that can be recycled back to the Rubisco sub-\nstrate RuBP (Bracher et al. 2015). The pyrenoid localization\nof CPLD2 suggests that it may also convert XuBP to RuBP\nin the pyrenoid. Both HDA5 and the protein encoded by\nCre16.g648400 have predicted “Rubisco-binding motifs”\n(Fig. 4) (discussed in a later section) (Meyer et al. 2020b), sug-\ngesting that they bind Rubisco in the pyrenoid matrix (Wang\net al. 2022). STR16, STR18, and the proteins encoded by\nCre13.g573250 and Cre16.g663150 are all predicted to be\nthiosulfate sulfurtransferases. In addition, STR16 and STR18\ncontain a rhodanese domain, which is predicted to function\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3245\n\nFigure 4. A common Rubisco-binding motif mediates the assembly of the major compartments of the pyrenoid. A) In Chlamydomonas, many\npyrenoid-localized proteins contain at least one Rubisco-binding motif. B) The Rubisco-binding motif mediates the assembly of the three pyrenoid\nsub-compartments. The motifs on EPYC1 link Rubisco to form the pyrenoid matrix (He et al. 2020). The motifs on the tubule-localized transmem-\nbrane proteins RBMP1 and RBMP2 are proposed to connect the Rubisco to the tubules, and the motifs on the putative starch-binding proteins\nSAGA1 and SAGA2 are proposed to mediate interactions between the matrix and the surrounding starch sheath. A Rubisco-binding motif was\nalso shown to be necessary and sufficient to target a nascent protein to the pyrenoid (Meyer et al. 2020b). C) A model illustrating how EPYC1\n(red) clusters Rubisco (blue) in the pyrenoid matrix. D) The Rubisco-binding motif of EPYC1 (red) binds to the Rubisco small subunit (dark\nblue) (He et al. 2020); other Rubisco-binding motifs in Chlamydomonas are expected to bind to the same site. E) The motif binds between two\nalpha-helices of the Rubisco small subunit.\n\nin disulfide bond formation and iron-sulfur cluster biosyn-\nthesis (Lau et al. 2023). ABCF6 is a predicted member of\nthe ATP-binding cassette F\nfamily\nthat regulates translation via binding to ribosomes (Lau\net al. 2023).\n\n(ABCF) protein\n\nPyrenoid-associated membranes likely supply\nRubisco with concentrated CO2\nThe Rubisco matrix in all known pyrenoids is in contact with a\nportion of the thylakoid membranes of the chloroplast (Meyer\net al. 2017), consistent with the idea that these membranes\n\nperform the essential function of supplying CO2 to Rubisco\n(Pronina and Semenenko 1990; Raven 2008). A broad range\nof morphologies has been observed in different species for\nthese pyrenoid-associated thylakoids (Fig. 3). Some species\nhave a single traversing membrane, some have multiple parallel\nor interconnected membranes, and some have more complex\nmorphologies, such as the undulating membranes found in\nspecies of the red algal genus Porphyridium (Nelson and\nRyan 1988) (Fig. 3). Some pyrenoids, such as in the dinoflagel-\nlate Podolampas bipes (Schnepf and ElbräChter 1999) (Fig. 3),\nhave no observed traversing membranes and instead are\n\nACDEB\f3246\n\n|\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\nHe et al.\n\nembedded between thylakoid membranes in the chloroplast.\nGiven that CO2 diffuses rapidly relative to the rate of its fixation\nby Rubisco, the exact location of CO2 release within the pyre-\nnoid likely has little effect on the distribution of CO2 within the\npyrenoid; thus, a broad range of membrane morphologies can\neffectively supply CO2 to Rubisco (Fei et al. 2022).\n\nIn Chlamydomonas, the thylakoid membranes extend into\nthe Rubisco matrix to form pyrenoid tubules whose lumina\nare continuous with the thylakoid lumen (Fig. 1, C and D)\n(Sager and Palade 1954, 1957; Ohad et al. 1967a).\nTraditional 2D TEM images have shown that thylakoid sheets\nnear the pyrenoid are directed toward the gaps of the starch\nsheath (Sager and Palade 1954, 1957; Ohad et al. 1967a). This\nobservation was corroborated by cryo-ET, which showed in\n3D that thylakoid sheets merge with each other as they\npass through gaps in the starch sheath to form cylindrical\npyrenoid tubules that traverse the Rubisco matrix (Fig. 1, C\nand D) (Engel et al. 2015). In the center of the pyrenoid,\nthe tubules converge to form a complex interconnected net-\nwork known as the reticulated region (Fig. 1, C and D) (Engel\net al. 2015; Meyer et al. 2020b).\n\nInside each Chlamydomonas tubule, there are two to eight\nsmaller tubes called minitubules (Ohad et al. 1967b; Engel et al.\n2015) (Fig. 1, C and D). Minitubules appear to provide con-\nduits from the inter-thylakoid stromal space to the pyrenoid\nmatrix (Engel et al. 2015). These minitubules have been pro-\nposed to facilitate the diffusion of small molecules such as\nRuBP and 3-PG between these two compartments (Engel\net al. 2015; Küken et al. 2018); however, their internal dia-\nmeters of approximately 3.5 ± 0.5 nm are likely too small to\nmediate a substantial flux of metabolites. Curiously, minitu-\nbules have not been observed in any species other than\nChlamydomonas (Meyer et al. 2017), although this could be\ndue to limitations in TEM imaging. The function of minitu-\nbules remains unknown, and the mechanisms by which the in-\ntricate morphology of\nthe\npyrenoid-traversing membranes is achieved is unknown in\nany organism.\n\nthe different\n\nregions of\n\nIt is notable that Chlamydomonas mutants that lack a\nRubisco matrix still have tubule networks in the canonical lo-\ncation within the chloroplast (Caspari et al. 2017; He et al.\n2020), indicating that the tubules can form in the absence\nof the matrix and suggesting that the location of the pyre-\nnoid could be determined by the tubules. However, the mo-\nlecular basis for the localization of the tubules remains\nunknown.\n\nThe delivery of concentrated CO2 to Rubisco in the matrix\nis thought to be mediated by carbonic anhydrases that con-\n− into CO2 in the lumen of the pyrenoid-traversing\nvert HCO3\nmembranes (Pronina and Semenenko 1990; Raven 2008). In\nChlamydomonas, the carbonic anhydrase that mediates\nthis key step is CAH3 (Karlsson et al. 1998; Hanson et al.\n2003). Consistent with this role, Chlamydomonas mutants\nlacking functional CAH3 have a severe growth defect when\ngrown in limiting CO2 conditions (Spalding et al. 1983;\nFunke et al. 1997; Karlsson et al. 1998) and over-accumulate\n\n− within the mutant cells (Spalding et al. 1983). CAH3\nHCO3\nlocalizes to the thylakoid lumen (Karlsson et al. 1998) and be-\ncomes enriched in the pyrenoid tubules during activation of\nthe CCM in transitions from high CO2 to limiting CO2\n(Blanco-Rivero et al. 2012; Sinetova et al. 2012) and from\ndark to light (Mitchell et al. 2014). How CAH3 relocalizes\nto the tubules remains unknown.\n\nRecent studies in Chlamydomonas have identified add-\nitional pyrenoid tubule–localized proteins that perform vari-\nous functions. Like CAH3, the Ca2+-binding protein calcium\nsensing receptor (CAS, Cre12.g497300) relocalizes to the pyr-\nenoid tubules upon CCM induction (Wang et al. 2016;\nYamano et al. 2018). CAS is a putative Rhodanese-like Ca2+-\nsensing receptor that regulates the expression of several\nCCM-related genes, including HLA3 and LCIA (Wang et al.\n2016). Upon activation of the CCM, CAS switches from being\ndispersed across the chloroplast to being associated with the\npyrenoid tubules (Wang et al. 2016). This change is accom-\npanied by an increase in Ca2+ in the pyrenoid (Wang et al.\n2016). The role of Ca2+ in the pyrenoid, the mechanism of\nCAS relocalization, and the purpose of CAS signaling in the\nCCM remain unclear.\n\nTwo other tubule-localized proteins are Rubisco-binding\nmembrane protein 1 (RBMP1, encoded by Cre06.g261750)\nand RBMP2 (Cre09.g416850) (Fig. 4). Both proteins bind to\nRubisco\nin vivo\nin vitro (Meyer et al. 2020b) and\n(Mackinder et al. 2017), suggesting that they may promote\ninteractions between the Rubisco matrix and the pyrenoid\ntubules. RBMP1 is associated with peripheral tubular regions,\nwhereas RBMP2 localizes to the central reticulated region of\nthe tubules (Meyer et al. 2020b). Intriguingly, RBMP2 con-\ntains a rhodanese domain, as do STR16, STR18, and CAS,\nbut the function of these rhodanese domains in these pro-\nteins remains to be determined. The putative roles of\nRBMP1 and RBMP2 in linking matrix to tubules also remain\nto be tested.\n\nIn addition to RBMP2, the proteins pyrenoid nuclease 1\n(PNU1, Cre03.g183550) and conserved in the green lineage\nand diatoms 14 (CGLD14, Cre10.g446350) appear to localize\nto the reticulated region of the tubules (Wang et al. 2022).\nPNU1 is a bifunctional nuclease domain-containing protein.\nAs oxidized RNA was also localized to the pyrenoid in\nChlamydomonas (Zhan et al. 2015), the pyrenoid localization\nof PNU1 suggests that the pyrenoid might be a site of oxi-\ndized RNA degradation (Wang et al. 2022). CGLD14 is con-\nserved in the green lineage and diatoms and is also named\nPSBP-domain-containing protein 3 (PPD3).\n\nMultiple components of the electron transport chain are\npresent in the Chlamydomonas pyrenoid tubules, including\nsubunits of photosystem I (PSI) (Photosystem I reaction cen-\nter subunit V [PSAG, Cre12.g560950], Photosystem I reaction\ncenter subunit H [PSAH, Cre07.g330250], Photosystem I reac-\ntion center\n[PSAK, Cre17.g724300], and\nferredoxin [FDX1, Cre14.g626700]),\nchloroplast-localized\nprotein\n(PSII)\nphotosystem\n3\n[Cre08.g362900],\nCre12.g509050],\n[PSBP3,\n\n(PsbP-like\nPSBP4\n\nsubunit K\n\nII\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3247\n\nand\n\nCre16.g651050),\n\nCre06.g261000]),\n\ncytochrome\nATP\n\nPhotosystem\nII oxygen evolution enhancer protein 3\n[PSBQ, Cre08.g372450], and Photosystem II subunit R\nb6f\n[PSBR,\n(CYC6,\nsynthase\n(ATPC, Cre06.g259900) (Mackinder et al. 2017). However,\ndespite some components of the O2-evolving PSII being pre-\nsent in Chlamydomonas pyrenoid-traversing membranes,\nother PSII components (such as subunit 1 of the\nPSII oxygen-evolving enhancer protein 1 [OEE1 or PSBO1,\nCre09.g396213] and PSII intrinsic core polypeptides D2\n[psbD, CreCp.g802329] and P5 [PSBP5, Cre09.g389578]) ap-\npear to be absent based on immunogold labeling (de Vitry\net al. 1989; McKay and Gibbs 1991). Relatedly, PSII was found\nto be inactive in the pyrenoid of the red alga Porphyridium\ncruentum based on cytochemical assays in which PSII activity\nwas detected through the production of osmiophilic difor-\nmazan upon the photoreduction of tetrazolium salts\n(McKay and Gibbs 1990, 1991). Minimizing PSII activity in\npyrenoid-traversing thylakoids may be a strategy for minim-\nizing O2 levels within the pyrenoid, which may help maintain\na high CO2 to O2 ratio around Rubisco (McKay and Gibbs\n1991). The electron transport chain components found in\nthe pyrenoid may therefore be in assembly intermediates,\nin inactive complexes undergoing repair, or may have differ-\nent functions from those found in stromal thylakoids. This\nhypothesis is supported by the pyrenoid localization of\nPSBP4, which is a homolog of the Arabidopsis PSII repair pro-\ntein PSBP-LIKE PROTEIN1 (PPL1, encoded by At3g55330)\nand\nfactors\n(Mackinder et al. 2017).\n\nfour known PSI assembly\n\ninteracts with\n\nluminal\n\nfactor 14\n\nchlorophyll\n\nfluorescence\n\nEight other proteins have recently been localized to the\npyrenoid tubules\nin Chlamydomonas: Deg protease 8\n(DEG8, Cre01.g028350), the cyclophilins CYN7 (Cre12.g544150)\nand CYN20-6 (Cre12.g544114), Photosystem II subunit P1\n(PSBP1, Cre12.g550850), the PSII stability/assembly factor\nhigh\n136\n(HCF136, Cre06.g273700), the thylakoid luminal protein\nthylakoid\n(TEF14, Cre06.g256250),\nuncharacterized proteins encoded by Cre03.g198850 (Wang\net al. 2022), and Cre03.g172700 (Lau et al. 2023) (Table 1).\nDEG8 is a predicted DegP-type protease, while CYN7 and\nCYN20-6 are two predicted peptidyl-prolyl cis-trans iso-\nmerases. The pyrenoid tubules may thus be involved in pro-\ntein folding, degradation, and/or import of new proteins into\nthe pyrenoid (Wang et al. 2022). The protein encoded\nby Cre03.g172700 is predicted to contain a long central\nalpha-helix and four “Rubisco-binding motifs” (discussed in\na later section), which might allow it to act as a potential pyr-\nenoid tether between the pyrenoid matrix and tubules along-\nside RBMP1 and RBMP2 (Lau et al. 2023).\n\nA polysaccharide sheath likely serves as a CO2\ndiffusion barrier\nPolysaccharide deposits are associated with the pyrenoids of\nsome species in every major algal lineage except the diatoms\n\nand coccolithophores (Fig. 3). In red algae and green algae,\nthe polysaccharide that makes up these deposits is starch,\nwhereas different polymers are used in other lineages, such\nas paramylon in the case of euglenoid algae (Nudelman\net al. 2006; Suzuki and Suzuki 2013; Ball et al. 2015). Green\nalgae produce starch in the chloroplast, whereas all other\nlineages produce their polysaccharide deposits in the cytosol\n(with the exception of the cryptophytes, which produce\nstarch in the periplastid) (Suzuki and Suzuki 2013; Ball\net al. 2015). Presumably because of these differences, species\nfrom all lineages except the green algae only have polysac-\ncharides associated with their pyrenoids if they have a stalked\nor bulging pyrenoid that projects into the cytoplasm (Meyer\net al. 2017) (Fig. 3). In these cases, the polysaccharide struc-\ntures are separated from the Rubisco matrix by the chloro-\nplast envelope (Meyer et al. 2017). The association of\npolysaccharide deposits with pyrenoids even when separated\nby membranes further implicates these structures in pyre-\nnoid function. In Chlamydomonas, the starch sheath is a\nshell-like structure made by curved starch granules around\nthe pyrenoid matrix (Fig. 1). Small gaps between starch plates\nallow the tubules to penetrate through into the matrix.\n\nAvailable evidence suggests that the pyrenoid polysacchar-\nide sheath, when present, serves as a barrier to slow the es-\nfrom the pyrenoid, allowing a higher\ncape of CO2\nconcentration of CO2 to be maintained in the pyrenoid\nand decreasing the energetic costs of CO2 concentration\n(Toyokawa et al. 2020; Fei et al. 2022). The most convincing\nevidence to date supporting this function comes from the\ndecreased CCM efficacy observed under very low CO2 in\nthe Chlamydomonas sta2-1 mutant (defective in starch syn-\nthase 2 [STA2, encoded by Cre17.g721500]), which has a\nthinner starch sheath but otherwise apparently normal local-\nization of key proteins (Toyokawa et al. 2020).\n\nThe pyrenoid starch sheath granules in Chlamydomonas\nare different from the stromal starch granules in their shape,\ncomposition, and the conditions under which they accumu-\nlate. The granules that make up the starch sheath are more\ncurved than stromal granules, which are globular in shape.\nThe molecular composition of starch consists of alternating\namorphous layers of amylose and crystalline layers of amylo-\npectin (Zeeman et al. 2010). Compared with stromal starch,\npyrenoidal starch has less amylose but more amylopectin\ncontent (Libessart et al. 1995; Findinier et al. 2019). Both\namylose and amylopectin have been shown to decrease O2\ngas permeability in vitro (Forssell et al. 2002), which further\nsupports the possible function of the starch sheath in slowing\ndown the escape of leaking CO2 from the matrix. Relatedly,\nthe molecular structure of starch varies depending on the al-\ngal lineage (Suzuki and Suzuki 2013; Ball et al. 2015), which\ncould have implications for the ability of starch to prevent\nCO2 diffusion in different species. In addition to differences\nin their shape and composition, pyrenoid starch sheath gran-\nules and stromal starch granules also accumulate under dif-\nferent conditions. When Chlamydomonas cells are grown in\nunfavorable conditions such as during nitrogen starvation,\n\n\f3248\n\n|\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\nHe et al.\n\nstromal starch content increases while that of pyrenoid\nstarch decreases, as starch metabolism rapidly switches\nfrom pyrenoidal to storage biosynthesis (Kuchitsu et al.\n1988; Findinier et al. 2019). However, when cells are moved\nfrom high CO2 (4%) to low CO2 (air-level), pyrenoid starch\naccumulates rapidly within hours and stromal starch is de-\ngraded (Kuchitsu et al. 1988).\n\nSeveral proteins have been implicated in the formation\nand degradation of the starch sheath in Chlamydomonas.\nThe protein StArch Granules Abnormal 1 (SAGA1, encoded\nby Cre11.g467712), which contains a putative starch-binding\ndomain, localizes to distinct puncta at the pyrenoid matrix/\nstarch interface (Figs. 1, C and D and 4B) (Itakura et al. 2019;\nMeyer et al. 2020b). Abnormally elongated and thinner\nstarch granules were observed in saga1 mutant cells, indicat-\ning that SAGA1 is required for normal starch sheath forma-\ntion (Itakura et al. 2019). A recent study suggested that\nSAGA1 is also necessary for relocalizing CAS and LCIB to\nthe pyrenoid under\nlimiting CO2 conditions and for\nCAS-dependent retrograde signaling regulation of nuclear\n− transporters (Shimamura\ngenes encoding CO2 and HCO3\net al. 2023). Another protein, bimodal starch granule 1\n(BSG1, Cre02.g091750), may be involved in the degradation\nof the starch sheath during the transition from low CO2 to\nhigh CO2 (Findinier et al. 2019).\n\nHigh-throughput studies have identified other proteins\nthat could potentially be involved in the formation of the\npyrenoid starch sheath (Table 1) (Mackinder et al. 2017;\nMeyer et al. 2020b). SAGA2 (Cre09.g394621) is a protein\nthat shares 30% sequence identity with SAGA1 and also\nhas a predicted starch-binding domain (Meyer et al.\n2020b). Like SAGA1, SAGA2 also localizes to the pyrenoid\nmatrix/starch interface (Fig. 4B), although its function is cur-\nrently unknown. Two other proteins, granule-bound STA2\n(Delrue et al. 1992; Maddelein et al. 1994) and starch-\nbranching enzyme 3 (SBE3, Cre10.g444700), localize around\nthe pyrenoid periphery,\nforming a plate-like pattern\n(Mackinder et al. 2017), which suggests that they may con-\ntribute to the biosynthesis of the starch sheath (Table 1).\nLCI9 (Cre09.g394473), which contains two starch-binding do-\nmains and\nfunction as a glucan\nto\n1,4-α-glucosidase, localizes in a mesh structure between the\ngaps of the starch sheath, suggesting that it may degrade\nstarch at the gaps between starch plates to ensure a close\nfit between adjacent plates (Mackinder et al. 2017). Further\nstudies of these starch-associated proteins are needed to\nin the biogenesis of the\nunderstand their\nChlamydomonas starch sheath.\n\nis predicted\n\nfunctions\n\nSix new pyrenoid-periphery proteins were recently identi-\nfied in Chlamydomonas: MIND1 (Cre12.g522950), malate de-\nhydrogenase 1 (MDH1, Cre03.g194850), uncharacterized\nproteins encoded by Cre09.g394547, Cre09.g415600 (Wang\net al. 2022), structural maintenance of chromosomes 7\n(SMC7, Cre17.g720450), and uncharacterized protein en-\ncoded by Cre09.g394510 (Lau et al. 2023) (Table 1). The loca-\ntion of most of these newly identified proteins relative to the\n\nstarch sheath remains unclear. Interestingly, MIND1 is a\nhomolog of the Arabidopsis chloroplast division site regula-\ntor MinD1 (At5g24020), suggesting that MIND1 could po-\ntentially play a role in coordinating pyrenoid fission or\ndissolution with chloroplast division in Chlamydomonas\n(Colletti et al. 2000; Freeman Rosenzweig et al. 2017; Wang\net al. 2022). SMC7 shows a punctate localization similar to\nthat of SAGA1 and is annotated as a member of the SMC\nfamily (Lau et al. 2023). SAGA1 and SAGA2 are also anno-\ntated as members of this family, which suggests that SMC7\nmight\nSAGA2.\nThe protein encoded by Cre09.g394510 contains a starch-\nbinding domain and localizes to the starch-matrix interface\nand the gaps between starch plates. It contains a predicted\nt-SNARE domain, which mediates vesicle fusion, suggesting\nthat it may be involved in membrane remodeling of the pyr-\nenoid tubules (Lau et al. 2023).\n\nfunction\n\nsimilarly\n\nSAGA1\n\nand\n\nto\n\nA Rubisco-binding motif mediates pyrenoid assembly\nAs previously discussed, the repeat protein EPYC1 has five\nRubisco-binding regions critical for pyrenoid matrix assembly\nin Chlamydomonas (He et al. 2020). Intriguingly, similar se-\nquences to the EPYC1 Rubisco-binding region have been iden-\ntified on many other pyrenoid-localized proteins (Fig. 4, A and\nB) (Meyer et al. 2020b). These sequences, including the\nRubisco-binding region on EPYC1, have been named\n“Rubisco-binding motifs.” The motifs on other pyrenoid pro-\nteins show similar binding affinity to Rubisco as the motifs\non EPYC1 (whose KD is approximately 3 mM) (He et al. 2020;\nMeyer et al. 2020b). Due to sequence similarity, the motifs\non other proteins are believed to bind to the same alpha-\nhelices of Rubisco small subunits as EPYC1 (Fig. 4, C–E).\n\nThe Rubisco-binding motif was shown to be necessary and\nsufficient for targeting a protein to the pyrenoid matrix\n(Meyer et al. 2020b). These observations suggest that nascent\npyrenoid proteins with copies of the motif diffuse around the\nchloroplast stroma until they encounter the matrix, where\nthey are captured by binding to Rubisco (Fig. 4B). One\nopen question is how proteins are targeted to the matrix\nwhen a full starch sheath has been assembled because stro-\nmal proteins would then not have direct access to Rubisco.\nIn addition to targeting proteins to the matrix, the\nRubisco-binding motif has been proposed to anchor the\nRubisco matrix to the pyrenoid tubules and connect\nthe starch sheath to the matrix (Meyer et al. 2020b). The\nRubisco-binding motif-containing proteins RBMP1 and\nRBMP2 localize to the tubules, suggesting that they target\na layer of Rubisco to the tubules. From there, EPYC1 may\nbe able to connect additional Rubiscos, causing the matrix\nto condense around the entire tubule network. The proteins\nSAGA1 and SAGA2 also contain Rubisco-binding motifs in\naddition to their starch-binding domains, which suggests\nthat they might link the matrix to the starch sheath\n(Fig. 4B) (Meyer et al. 2020b).\n\nThe identification of the Rubisco-binding motif and the\nhypothesis that proteins with this motif link the three\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3249\n\npyrenoid sub-compartments together in Chlamydomonas\nexplains the initially puzzling difference between the pheno-\ntypes of a mutant lacking functional EPYC1 and mutants\nwith disrupted EPYC1-binding sites on Rubisco small subu-\nnits (He et al. 2020; Meyer et al. 2020b). In a mutant lacking\nEPYC1, a minimal pyrenoid can still be observed (Mackinder\net al. 2016); however, mutants with disrupted EPYC1-binding\nsites on Rubisco small subunits lack a pyrenoid altogether\n(He et al. 2020; Meyer et al. 2012, 2020b). These findings\ncan be reconciled when considering that, in the mutant lack-\ning EPYC1, proteins other than EPYC1 that have the\nRubisco-binding motif (potentially including RBMP1 and\nRBMP2) can still bind to Rubisco and form the observed\nmuch smaller pyrenoid-like structure that still contains tu-\nbules and a starch sheath but lacks a canonical matrix.\n\nThe sequences of the Rubisco-binding motifs and their\nbinding sites on Rubisco are conserved\nin the order\nVolvocales to which Chlamydomonas belongs, but the motif\nhas not been found in any other algal lineages (Meyer et al.\n2020b). Assuming that pyrenoids convergently evolved, it is\npossible that the assembly of pyrenoids via common\nRubisco-binding motifs may broadly apply to pyrenoids\nacross the algal tree of life, although the specific sequences\nmay differ in different algal lineages (Meyer et al. 2020b).\n\nOther candidate pyrenoid components in\nChlamydomonas\nPhysical interactors of proteins that localize to the pyrenoid\nwere identified using large-scale affinity-purification mass\nspectrometry (Mackinder et al. 2017). Using this method,\n513 interactions involving 398 proteins were identified\n(Mackinder et al. 2017).\n\nIn a parallel study, Chlamydomonas pyrenoids were puri-\nfied and their proteome was analyzed, identifying 190 pro-\nteins in total (Zhan et al. 2018). Of the 190 candidate\npyrenoid proteins identified, many have confirmed or pre-\ndicted functions that are known or proposed to occur in pyr-\nenoids, such as the CCM, starch metabolism, or RNA\nmetabolism and translation. Additional proteins suggestive\nof new pyrenoid functions in tetrapyrrole and chlorophyll\nsynthesis, carotenoid metabolism, or amino acid metabolism\nwere also identified. Future work on these uncharacterized\ncandidate pyrenoid proteins will yield a better understanding\nof the biogenesis, function, and regulation of the pyrenoid.\n\nPyrenoid dynamics are likely highly regulated, but the under-\nlying regulatory mechanisms remain to be discovered.\n\nThe pyrenoid forms in response to limiting CO2 levels\nunder constant light\nWhen Chlamydomonas cells are transferred from high CO2\nto limiting CO2 under constant light conditions, the pyrenoid\nmatrix grows within one hour (Kuchitsu et al. 1991;\nRamazanov et al. 1994), presumably by relocalization of\nRubisco from the chloroplast stroma to the pyrenoid matrix.\nThe starch sheath starts to form within one hour after trans-\nfer from high to low CO2 as well and is fully formed after\nabout five hours (Kuchitsu et al. 1988; Ramazanov et al.\n1994).\n\nThe expression of many CCM-related genes, including\nthose encoding confirmed pyrenoid proteins such as\nEPYC1, STA2, and CAH3, is upregulated during the transition\nfrom high to low CO2 (Brueggeman et al. 2012; Fang et al.\n2012), which is consistent with the expansion of the pyrenoid\nmatrix and the formation of the starch sheath. The\nChlamydomonas CCM “master regulator” inorganic carbon\n(Ci) acquisition 5 (CIA5, Cre02.g096300, also known as\nCCM1) is required for the transcriptional upregulation of\nCAH3, EPYC1, STA2, LCIB, LCIC, and SMM7 in response to\nthe transition from high CO2 to low CO2 (Fang et al. 2012;\nSanthanagopalan et al. 2021), although CIA5 may also have\nother non-CCM-related functions (Moroney et al. 1989;\nMarek and Spalding 1991; Miura et al. 2004; Wang et al.\n2005; Fang et al. 2012; Redekop et al. 2022). CIA5 has zinc-\nbinding activity and was proposed to be a transcription fac-\ntor (Fukuzawa et al. 2001; Xiang et al. 2001; Kohinata et al.\n2008), but this has not been confirmed because no\nDNA-CIA5 complexes have been identified. Additionally,\nthe CIA5 regulatory mechanism is not well understood.\nCIA5 transcript and CIA5 protein levels are similar in high–\nCO2-grown and low–CO2-grown unsynchronized wild-type\ncells (Wang et al. 2005; Fang et al. 2012); thus, CIA5 activity\nregulated by posttranslational modifications\nmay be\n(Fukuzawa et al. 2001; Xiang et al. 2001; Wang et al. 2005;\nBrueggeman et al. 2012; Chen 2016). CIA5 regulates the tran-\nscription\n1\n(LCR1, Cre09.g399552), which is known to directly regulate\nthe\nLCI1\nCAH1\n(Cre03.g162800), and LCI6 (Cre12.g553350) (Yoshioka et al.\n2004).\n\n(Cre04.g223100),\n\nexpression\n\nlow-CO2\n\nresponse\n\nfactor\n\nstress\n\nof\n\nDynamics and regulation\nPyrenoids in various species are dynamic, showing noticeable\nmorphological changes under different growth conditions\nand during cell division (Brown and Bold 1964; Brown et al.\n1967; Goodenough 1970; Retallack and Butler 1970). The\nnewly reported liquid-like nature of the Chlamydomonas\npyrenoid (Freeman Rosenzweig et al. 2017) provides a new\nframework for thinking about the biophysics underlying pyr-\nenoid dissolution, condensation, and division by fission.\n\nPosttranslational modifications are thought to regulate the\nfunctions of several essential pyrenoid proteins. The Rubisco\nlinker EPYC1/LCI5 was reported to be phosphorylated under\nlow CO2 conditions but not under high CO2 (Turkina et al.\n2006), although the functional implications of this phosphor-\nylation are unknown. The relocalization of CAH3 from the\nstromal thylakoids to the pyrenoid tubules under low CO2\nconditions as well as the functions of HLA3 and LCIC under\nlow CO2 and very low CO2 have also been proposed to be\nregulated by phosphorylation (Blanco-Rivero et al. 2012;\n\n\f3250\n\n|\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\nHe et al.\n\nWang et al. 2014). LCIB is glutathionylated during acclima-\ntion to limiting CO2 (Zaffagnini et al. 2012). The effect of\nthese posttranslational modifications on the functions of\nthese CCM proteins is unclear, as is the identity of the regu-\nlatory proteins upstream of these modifications. Methylation\nmay also be involved in the regulation of pyrenoid biogenesis,\nas suggested by altered pyrenoid morphologies under low\nCO2 in mutants lacking function for the putative methyl-\ntransferase CIA6 (Cre10.g437829) (Ma et al. 2011).\n\nWhen cells are transferred from limiting CO2 to high CO2,\nthe disassembly of the pyrenoid is much slower than its assem-\nbly when cells are transferred from high CO2 to limiting CO2\n(Kuchitsu et al. 1988; Ramazanov et al. 1994). The degradation\nof the starch sheath and the dissolution of the matrix can take\ntwo to three days (Ramazanov et al. 1994). This slow degrad-\nation is consistent with the slow deactivation of the CCM,\nwhich also requires about three days when cells are moved\nfrom limiting CO2 to high CO2 (Ramazanov et al. 1994). It\nhas been suggested that CCM proteins are not rapidly de-\ngraded after the transition from low CO2 to high CO2\n(Toguri et al. 1989), whereas the synthesis of new CCM pro-\nteins stops shortly after this transition (Manuel and\nMoroney 1988). These observations have not been confirmed\nfor crucial pyrenoid proteins, and the regulatory mechanisms\nremain unclear.\n\nThe pyrenoid-based CCM is induced and\ndeactivated during the diurnal cycle\nMost studies of the Chlamydomonas pyrenoid and CCM\nhave been performed with asynchronous cultures of cells\ngrown under constant light, where the induction of the\nCCM and formation of the pyrenoid are solely determined\nby the level of CO2. However, synchronous growth under di-\nurnal cycles has revealed that the CCM is regulated during\nthe course of the day/night cycle (Mitchell et al. 2014;\nTirumani et al. 2014; Zones et al. 2015; Strenkert et al. 2019).\nChlamydomonas cells can be synchronized when grown\nunder 12-h-light/12-h-dark cycles in minimal medium, with\ncells going through one cell cycle each day (Harris 2009;\nMitchell et al. 2014; Zones et al. 2015; Strenkert et al.\n2019). The CCM is downregulated at night and fully induced\none hour before dawn (Mitchell et al. 2014). During this in-\nduction, both Rubisco and CAH3 were found to relocalize\nfrom the chloroplast to the pyrenoid based on statistical ana-\nlyses of immunogold labeling (Mitchell et al. 2014), although\nit should be noted that the original electron micrographs\nwere not provided in this study. Transcriptomics studies\nhave shown that, in cells grown under diurnal cycles, genes\nencoding the master regulator CIA5 and crucial pyrenoid\nproteins reach their highest transcript levels in the first few\nhours around the transition from dark to light (Strenkert\net al. 2019; Adler et al. 2022). However, whether CIA5 is\nalso involved in CCM activation and pyrenoid formation dur-\ning diurnal cycles is not known. The pyrenoid may grow and\nexpand during the day as cells grow (Zones et al. 2015;\n\nStrenkert et al. 2019), but this has not yet been specifically\nmeasured.\n\nPyrenoid formation can be induced by\nhyperoxia and H2O2\nHyperoxia was recently reported to induce pyrenoid forma-\n− levels (Neofotis et al. 2021).\ntion, even at high CO2 or HCO3\nThe authors reasoned that because pyrenoid formation is in-\nduced by both low CO2 and hyperoxia, a metabolite that ac-\ncumulates under both conditions may serve as a signal that\ninduces pyrenoid formation. Consistent with this idea, the\nauthors observed that hydrogen peroxide (H2O2), which is\nexpected to accumulate under both conditions, induces pyr-\nenoid formation. H2O2 is a byproduct of photorespiration,\nwhich recycles 2-phosphoglycolate, the toxic product of\nthe oxygenase activity of Rubisco (Moroney et al. 2013),\nwhich is increased under low CO2 and hyperoxia. Whether\nor\nH2O2\nindirectly (e.g. via other metabolites) has not been\ndetermined.\n\nformation\n\nregulates\n\npyrenoid\n\ndirectly\n\nstudies observed\n\nelectron microscopy\n\nThe Chlamydomonas pyrenoid matrix divides by\nfission and dissolves into the surrounding chloroplast\nduring cell division\nEarly\nthe\nChlamydomonas pyrenoid dividing by fission (Goodenough\n1970). Recent live-cell microscopy observation of pyrenoid\nmatrix dynamics using fluorescently tagged Rubisco or\nEPYC1 revealed that the matrix is inherited by fission in\nmost chloroplasts and assembled de novo in others (Fig. 5)\n(Freeman Rosenzweig et al. 2017). Approximately two-thirds\nof daughter chloroplasts inherited their matrix through\nelongation and then fission of the pyrenoid matrix from\nthe mother chloroplast (Fig. 5, A to C), whereas one of the\ndaughter chloroplasts inherited the entire matrix punctum\nin the remaining cases (Fig. 5D). When the pyrenoid divided\nby fission, matrix elongation and fission occurred toward the\nend of overall chloroplast division and took approximately\nseven minutes. A “bridge” of matrix connecting the two lobes\nwas briefly visible towards the end of fission (Fig. 5B). After\nthe bridge ruptured, the daughter pyrenoids quickly reverted\nto spherical shapes, similar to the behavior of liquid droplets\n(Fig. 5B) (Stone 1994; Yanashima et al. 2012; Freeman\nRosenzweig et al. 2017). The mechanism mediating pyrenoid\nfission during cell division is unknown.\n\nA portion of the pyrenoid matrix rapidly disperses into the\nstroma approximately 20 minutes before pyrenoid division\n(Fig. 5C, at approximately 19 minutes, the light blue through-\nout the chloroplast indicates the dispersed pyrenoid matrix).\nDuring this dispersal, small puncta of matrix often transiently\nappear throughout the stroma (Fig. 5C). The dispersal of ma-\ntrix materials may facilitate equal distribution of the pyre-\nnoid matrix to daughter chloroplasts and may also help\ndecrease the surface tension or viscosity of the matrix droplet\nto facilitate fission (Freeman Rosenzweig et al. 2017). Indeed,\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3251\n\nFigure 5. The Chlamydomonas pyrenoid exhibits liquid-like behavior during cell divisions. A) Diagram depicting the timeline and morphology of a\ntypical cell division with pyrenoid fission in Chlamydomonas (adapted from Freeman Rosenzweig et al. 2017). The time point t = 0 is the moment\nthe chloroplast division furrow passes between the daughter pyrenoids. A portion of the pyrenoid matrix disperses into the chloroplast stroma\nduring the division of the pyrenoid. The approximate timing and duration of key events are shown below the timeline. B) Diagram depicting\nthe “bridge” of matrix during pyrenoid fission. C) Diagram depicting the transient appearance of small puncta of pyrenoid matrix throughout\nthe stroma during dispersal of the matrix in some dividing cells. D) Diagram depicting the de novo formation of a daughter pyrenoid when pyrenoid\nfission fails. The lower daughter cell inherits the entire pyrenoid of the mother cell. The upper cell shows de novo pyrenoid formation with the\nappearance of one or more fluorescent puncta growing or coalescing into one pyrenoid (observed in wild-type cells expressing either\nEPYC1-Venus or Rubisco-Venus).\n\nABCD\f3252\n\n|\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\nHe et al.\n\nmany of the daughter chloroplasts that did not inherit a pyr-\nenoid through fission inherited dissolved matrix building\nblocks, from which they appeared to form a pyrenoid de\nnovo (Freeman Rosenzweig et al. 2017) (Fig. 5D). In such\ncases, multiple small Rubisco or EPYC1 fluorescent puncta\nappeared, and smaller puncta shrank whereas larger ones\ngrew until the cell contained a single pyrenoid (Freeman\nRosenzweig et al. 2017) (Fig. 5D). This behavior resembles\nOstwald ripening, a physical mechanism by which larger dro-\nplets in a phase-separated system grow by acquiring building\nblocks from smaller droplets (Hyman et al. 2014; Freeman\nRosenzweig et al. 2017; Rosowski et al. 2020). The mechan-\nisms regulating the formation of multiple puncta of matrix\nmaterial and the dispersal of the matrix remain unknown.\n\nPyrenoids in other algae are likely to leverage liquid-like\nproperties during cell division in ways similar to those ob-\nserved in Chlamydomonas (Freeman Rosenzweig et al.\n2017; Barrett et al. 2021). Indeed, both division by fission\nand the rapid disappearance and reappearance of the pyre-\nnoid matrix (which would be consistent with matrix dissol-\nution and condensation) have been observed during cell\ndivision using TEM on fixed cells in some species of the green\nalgae Tetracystis, Chlorococcum, and Bulbochaete and the dia-\ntom Donkinia (Brown et al. 1967; Retallack and Butler 1970;\nCox 1981). Additional discussions on liquid-like pyrenoid be-\nhavior during cell division can be found in the recent review\nby Barrett et al. (Barrett et al. 2021). Further studies on other\nspecies will be necessary to test the generality of this\nprinciple.\n\nThe number of pyrenoids per cell appears to be\nregulated in Chlamydomonas\nWild-type Chlamydomonas cells have only one pyrenoid.\nHowever, in mutant cells lacking SAGA1, EPYC1, or CIA6,\nmultiple pyrenoids are often observed, suggesting that wild-\ntype cells may actively work to ensure that they have a single\npyrenoid, and aspects of this regulation may be disrupted in\nthese mutants.\n\nThe most striking example of multiple pyrenoids can be\nseen in the saga1 mutant, with an average of approximately\nten matrix droplets per cell (Itakura et al. 2019). The multiple\npyrenoids in saga1 are stable without obvious changes in size\nor position in living mutant cells over the course of one\nhour. Although the molecular function of SAGA1 remains\nunclear, these observations suggest that this protein is in-\nvolved in maintaining a single pyrenoid.\n\nThe two other known cases of mutants with multiple pyr-\nenoids are epyc1 and cia6. Multiple pyrenoids were observed\nby TEM in 13% of epyc1 cells compared with 3% in wild-type\ncells (Mackinder et al. 2016). A mutant of the putative\nmethyltransferase CIA6 also exhibits multiple small pyre-\nnoids, as observed by TEM or via light microscopy (Ma\net al. 2011). TEM images showed that the morphology of\nthe pyrenoids in epyc1 and cia6 are similar, with a small ma-\ntrix and thicker starch sheath than in wild-type cells (Ma\n\net al. 2011; Mackinder et al. 2016). EPYC1 has a well-\ncharacterized function in pyrenoid matrix biogenesis; thus,\nit is possible that defects in matrix biogenesis lead to multiple\npyrenoids. The mechanism by which the number of pyre-\nnoids in a cell is regulated remains to be identified.\n\nPerspective\nThe pyrenoid provides a unique opportunity to expand our\nfundamental knowledge of liquid-liquid phase separation,\ngenetically engineer photosynthetic organisms for higher\ncrop yields, and obtain insights into critical photosynthetic\ncarbon assimilation in the oceans and fresh water, all through\nthe study of one organelle.\n\nStudies of the Chlamydomonas pyrenoid have laid the\nfoundation for exploring the molecular composition of pyr-\nenoids across the photosynthetic tree of life. Of particular\ninterest is investigating whether pyrenoids in other species\nalso exhibit\nliquid-like behavior. Additionally, exploring\nwhether Rubisco-binding motifs are a common principle\nacross all algal pyrenoids could lead to a better understand-\ning of how pyrenoids first evolved. Molecular studies of CCM\nfunction in different algae could also help reveal the minimal\ncomponents that will be necessary to create a functional pyr-\nenoid in plants. More broadly, studying the mechanisms by\nwhich Rubisco condensates associate with membrane struc-\ntures and peripheral polysaccharides could provide insights\ninto general principles by which liquid-like organelles interact\nwith other cellular structures.\n\nFuture studies in Chlamydomonas as well as in other spe-\ncies of algae will bring us closer to generating the first func-\ntional pyrenoid in vascular land plants, which will help us\nmeet increasing agricultural demands as the global popula-\ntion rises. Progress is already being made toward engineering\na Chlamydomonas-based pyrenoid into land plants with the\nsuccessful reconstitution of Rubisco-EPYC1 condensates in\nArabidopsis (Atkinson et al. 2020). The generation of thyla-\nkoid tubules that traverse these condensates, delivery of\nCO2 via these tubules, and the assembly of starch around\nthese condensates will be important next steps.\n\nIn addition, by studying the regulation of pyrenoid forma-\ntion and dynamics in Chlamydomonas as well as in dominant\nmarine species such as diatoms, we can advance our under-\nstanding of the aquatic photosynthesis that is crucial for our\necosystem and the global carbon cycle.\n\n(Princeton\n\nAcknowledgments\nWe thank Alistair McCormick (University of Edinburgh),\nMoritz Meyer\nalumni), Ned Wingreen\n(Princeton University), Debashish Bhattacharya (Rutgers\nUniversity), Ben Engel (University of Basel), Lianyong Wang\n(Princeton University), Micah\n(Princeton\nUniversity), and the two anonymous reviewers for helpful\ncomments, suggestions, and clarifications. We thank Marie\nBao, as part of Life Science Editors, for help with manuscript\n\nBurton\n\n\fThe eukaryotic CO2-concentrating organelle\n\nTHE PLANT CELL 2023: 35; 3236–3259\n\n|\n\n3253\n\nediting. This work was supported by Howard Hughes Medical\nInstitute, National Science Foundation (MCB-1935444), and\nNational Institutes of Health (R01GM140032) grants to\nM.C.J. M.C.J.\nInstitute\nInvestigator. We apologize to all colleagues whose work we\ncould not incorporate due to space constraints.\n\nis a Howard Hughes Medical\n\nAuthor contributions\nAll authors contributed to the writing and the figures of the\narticle.\n\nConflict of interest statement. None declared.\n\nBehrenfeld MJ, Randerson JT, McClain CR, Feldman GC, Los SO,\nTucker CJ, Falkowski PG, Field CB, Frouin R, Esaias WE, et al.\nBiospheric primary production during an ENSO transition. Science.\n2001:291(5513):2594–2597. https://doi.org/10.1126/science.1055071\nBerner RA. 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