The green algae CO2 concentrating mechanism and photorespiration jointly operate during acclimation to low CO2

Dao, Ousmane; Bertrand, Marie; Alseekh, Saleh; Veillet, Florian; Auroy, Pascaline; Nguyen, Phuong-Chi; Légeret, Bertrand; Epting, Virginie; Morin, Amélie; Cuiné, Stephan; Monteil, Caroline L.; Mackinder, Luke C. M.; Burlacot, Adrien; Krieger-Liszkay, Anja; Weber, Andreas P. M.; Fernie, Alisdair R.; Peltier, Gilles; Li-Beisson, Yonghua

doi:10.1038/s41467-025-60525-7

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Published: 17 June 2025

The green algae CO₂ concentrating mechanism and photorespiration jointly operate during acclimation to low CO₂

Nature Communications volume 16, Article number: 5296 (2025) Cite this article

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Abstract

Due to low availability of CO₂ in aquatic environment, microalgae have evolved a CO₂ concentrating mechanism (CCM). It has long been thought that operation of CCM would suppress photorespiration by increasing the CO₂ concentration at the Rubisco active site, but experimental evidence is scarce. To better explore the function of photorespiration in algae, we first characterized a Chlamydomonas reinhardtii mutant defected in low-CO₂ inducible 20 (LCI20) and show that LCI20 is a chloroplast-envelope glutamate/malate transporter playing a role in photorespiration. By monitoring growth and glycolate excretion in mutants deficient in either CCM or photorespiration, we conclude that: (i.) CCM induction does not depend on photorespiration, (ii.) glycolate excretion together with glycolate dehydrogenase down-regulation prevents the toxic accumulation of non-metabolized photorespiratory metabolites, and (iii.) photorespiration is active at low CO₂ when the CCM is operational. This work provides a foundation for a better understanding of the carbon cycle in the ocean where significant glycolate concentrations have been found.

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Introduction

Microalgae account for nearly half of the photosynthetic CO₂ fixation on Earth^1,2. They contribute to climate change mitigation by assimilating CO₂ and are promising candidates for the production of biobased compounds^3,4,5. Due to the low availability of CO₂ in water, most algal cells have developed biophysical CO₂ concentrating mechanisms (CCMs), thus increasing the CO₂ concentration at the catalytic site of Rubisco and favoring the carboxylase reaction at the expense of the oxygenase. Microalgal biophysical CCMs generally involve carbonic anhydrases located in different cellular compartments, bicarbonate and/or CO₂ channels or transporters, and in most cases the pyrenoid (a specific cellular compartment where Rubisco is packed in a liquid phase separated organelle and where CO₂ is concentrated)^6,7,8,9,10. As a consequence of the CCM operation, the production of 2-phosphoglycolate (2-PG) resulting from the oxygenase activity of Rubisco is diminished^11,12. Because of its inhibitory effect on the Calvin cycle, 2-PG is metabolized and recycled through photorespiration. Photorespiration consists of multiple metabolic reactions distributed across different subcellular compartments thus requiring inter-organelle communication¹³. During photorespiration, the 2-PG is converted in the chloroplast into glycolate by the 2-PG phosphatase PGP1. In vascular plants lacking a biophysical CCM, glycolate is converted into glycerate through a series of reactions in peroxisomes and mitochondria. Although photorespiration occurs in both plants and microalgae, its subcellular organization seems to differ. For example, in vascular plants lacking a biophysical CCM, photorespiration requires tight cooperation of chloroplast, peroxisome and mitochondria¹³; whereas in Chlamydomonas, photorespiration depends mainly on cooperation between chloroplast and mitochondria^14,15.

In the model green microalga Chlamydomonas reinhardtii (hereafter Chlamydomonas), part of the glycolate is transferred to mitochondria and further metabolized into glyoxylate, glycine and serine, the other part is excreted out of the cell^16,17,18. During acclimation of Chlamydomonas to low CO₂, CCM and photorespiratory genes, which share the same master regulator CIA5, are simultaneously induced^19,20,21. Such a feature is intriguing since the activity of CCM supposedly inhibits photorespiration. It was proposed that photorespiration is transiently operational during the acclimation from high to low CO₂ conditions until the CCM is fully induced^7,12, photorespiratory metabolites such as 2-PG possibly acting as signaling molecules triggering CCM induction^{12,14,22,23,24}.

Two types of CCM regimes have been described in Chlamydomonas depending on external CO₂ levels, the low-CO₂-based CCM (CO₂ air level), in which cells preferentially transport CO₂, and the very-low-CO₂-based CCM, in which specific transporters are induced, such as the ATP-binding cassette transporter High Light Activated 3 (HLA3) involved in active bicarbonate uptake^25,26. During CCM induction, mitochondria migrate from the chloroplast cup to the periphery of the cell at the vicinity of the plasma membrane²⁷. The contribution of mitochondria to the energy supply of the algal CCM has been recently evidenced, and energy trafficking between chloroplast and mitochondria was suggested to supply ATP to CCM bicarbonate transporters^28,29. Actually, energy exchange between subcellular organelles was evidenced decades ago in microalgae^30,31,32,33 and has been re-examined more recently^34,35,36,37. The exchange of reducing power between organelles may operate via “malate shuttles” composed of malate dehydrogenases (MDHs) and membrane transporters of dicarboxylic acids^38,39,40,41.

Despite the existence of several genes encoding putative chloroplast malate transporters^41,42,43, none of them has been characterized in Chlamydomonas. Based on the increased expression of Low-CO₂ Inducible 20 (LCI20) gene, which encodes a putative malate/2-oxoglutarate transporter, during acclimation to limiting CO₂^19,20,21, it was proposed that LCI20 may be involved as a malate shuttle in the energy trafficking between chloroplast and mitochondria to feed CCM bicarbonate transporters⁴⁴. By another way, photorespiration may also contribute to the energy supply to external CCM transporters^44,45. Indeed, during photorespiration NADH is produced during the mitochondrial condensation of two glycine molecules into one serine, the NADH being converted into ATP by the mitochondrial respiratory chain. Until now, the nature of metabolic pathways and the identity of proteins involved in the energy network transferring photosynthetic energy from the chloroplast towards mitochondria to feed the external CCM transporters remain largely unexplored.

In this study, we aimed at better characterizing the relationship between CCM and photorespiration in green algae. For this purpose, we a Chlamydomonas mutant deficient in LCI20 and showed that LCI20 is a malate/glutamate transporter involved in photorespiration by supplying amino groups for the mitochondrial conversion of glyoxylate into glycine. By comparing growth properties and glycolate excretion in mutants affected in photorespiration or CCM under various photorespiration regimes, we conclude that photorespiratory metabolites do not contribute to CCM induction, that glycolate excretion avoids toxicity of non-metabolized photorespiratory intermediates and that downstream glycolate metabolism, although occurring when the CCM is functioning under very low CO₂, is not required for CCM operation.

Results

LCI20 encodes a putative chloroplast malate transporter whose expression is modulated by light and CO₂ levels

Three putative chloroplast 2-oxoglutarate/malate transporters (annotated as OMT1, OMT2 and LCI20) orthologues of previously characterized Arabidopsis DiT1 and DiT2 malate transporters were identified in the Chlamydomonas genome^42,43. Previous transcriptomic analysis performed during a day-night cycle or during limiting CO₂ acclimation showed that the expression of LCI20 (but not that of OMT1 and OMT2) is strongly induced at the beginning of the light period⁴⁶ or after the transition from H-CO₂ (2%) to L-CO₂ (0.04%) or VL-CO₂ (0.01%)^19,20,21 (Supplementary Fig. 1a, b). A phylogenetic analysis revealed that LCI20, which belongs to the GreenCut2⁴⁷, is widespread in Chlorophyta, closely related to DiT2 transporters, and evolutionarily divergent from OMT family transporters (Supplementary Figs. 1c and 2). Furthermore, LCI20 has recently been localized in the chloroplast but the enrichment in the envelope was not clearly shown^48,49. Here, we reexamined the strain expressing LCI20-mVenus under the control of PSAD promoter and we can see clearly that LCI20 is localized to the chloroplast envelope (Fig. 1a and Supplementary Fig. 3).

Fig. 1: LCI20 subcellular localization, characterization and complementation of a lci20 insertional mutant. — **Fig. 1: LCI20 subcellular localization, characterization and complementation of a *lci20* insertional mutant.**

The growth of lci20 is impaired during a transition from high to very low CO₂

A mutant harboring an insertion in the sixth intron of the LCI20 locus was obtained from the Chlamydomonas library project (CLiP) collection⁵⁰ and we named the mutant here as lci20 (Fig. 1b). We confirmed the insertion of the paromomycin resistance cassette and the absence of the LCI20 transcript in lci20 (Supplementary Fig. 4a, b). We then performed genetic complementation of lci20 by expressing the full-length genomic LCI20 coding sequence driven by the PSAD promoter (Supplementary Fig. 4a, b). A LCI20 antibody was produced in this study and used to show the absence of LCI20 in the mutant and its presence in WT and the complemented lines (Fig. 1c).

The growth of lci20 was then investigated under photoautotrophic conditions in both solid and liquid cultures under various CO₂ levels (Fig. 1d, e; Supplementary Fig. 4c, d). On solid media, lci20 grew normally under H-CO₂, L-CO₂ or VL-CO₂ when cells were previously acclimated to these conditions (Fig. 1d). However, the growth of lci20 was severely impaired when transitioning from H-CO₂ to VL-CO₂ (Fig. 1e), with smaller decreases in growth observed after a transition from H-CO₂ to L-CO₂ or from L-CO₂ to VL-CO₂ (Fig. 1e). In liquid media, while lci20 grew normally in H-CO₂ or during transition from H-CO₂ to L-CO₂, its growth was reduced when expressed on the basis of cell volume during transition from H-CO₂ to VL-CO₂ (Supplementary Fig. 4c, d). Based on these results, we conclude that LCI20 is required for photoautotrophic growth when cells are subjected to a sudden and severe CO₂ limitation.

lci20 is affected in photorespiratory glycolate metabolism

To determine whether the mutant phenotype is due to the role of LCI20 in the induction of the CCM or in photorespiration, we performed growth assays on agar plates by transferring H-CO₂ grown cells into VL-CO₂ under photorespiratory (21% O₂) or non-photorespiratory (2% O₂) conditions (Fig. 2a). As a control, we used the CCM1 transcription factor knockout mutant cia5, which is defective in the transcriptional induction of CCM and of the photorespiratory pathway^11,45,51,52. cia5 grew normally under H-CO₂ but growth was completely abolished when cells were transferred to VL-CO₂, irrespective of O₂ levels (Fig. 2a), thus indicating that a defect in the CCM, but not in photorespiration, prevented growth of the cia5 mutant under VL-CO₂. Note that similar growth in both solid and liquid cultures was observed when wild-type cells were exposed to VL-CO₂ either under photorespiratory conditions (21% O₂) or non-photorespiratory conditions (2% O₂) (Supplementary Fig. 5), indicating that photorespiration is not required for CCM induction. In contrast, lci20 grew poorly after transition under photorespiratory conditions (VL-CO₂; 21% O₂), but the growth was unaffected under non-photorespiratory conditions (VL-CO₂; 2% O₂) (Fig. 2a), indicating that lci20 is defective in photorespiration rather than in CCM. To further distinguish phenotypes between photorespiration and CCM deficiency, we have analyzed the growth of two CCM mutants bsti and cah3 deficient in bestrophins (BST1-3)⁵³ and lumenal carbonic anhydrase CAH3⁵⁴, respectively (Supplementary Fig. 6). Note that BST1-3 and CAH3were reported as low CO₂ inducible genes in the same transcriptomic study that has also reported LCI20 expression^19,21. In contrast to lci20 but similar to cia5 (Fig. 2a), both bsti and cah3 were unable to grow under VLC regardless of the photorespiratory status (Supplementary Fig. 6a), indicating a defect in CCM but not in photorespiration.

Fig. 2: lci20 is affected in the photorespiratory glycolate metabolism. — **Fig. 2: *lci20* is affected in the photorespiratory glycolate metabolism.**

Glycolate is excreted into the culture medium by Chlamydomonas cells when either the CCM or the glycolate metabolism is defective^{16,18,45,55,56}. Glycolate excretion was measured after 20 h acclimation of H-CO₂ grown cells to VL-CO₂ either at 21% or 2% O₂. At 21% O₂, both cia5 and lci20 mutants showed a significantly higher excretion compared to their respective controls (Fig. 2b). Similarly, bsti and cah3 showed higher and continuous glycolate excretion during acclimation to VL-CO₂ at 21% O₂ (Supplementary Fig. 6b). Note that we did not detect glycolate excretion under H-CO₂ or VL-CO₂ at 2% O₂, conditions when photorespiration is strongly reduced (Fig. 2b). We conclude that lci20 is defective in the photorespiratory glycolate metabolism, which is triggered when Chlamydomonas cells are transferred from H-CO₂ to VL-CO₂ at 21% O₂.

Changes in abundance of photorespiration and CCM proteins during cells’ response to different CO₂ and O₂ levels

To identify possible causes for the growth defect and high glycolate excretion observed in lci20, we probed the abundance of different photorespiratory enzymes including the glycolate dehydrogenase (GYD1), hydroxypyruvate reductase (HPR1) and glycine cleavage system P (GCSP) proteins together with CCM-related proteins HLA3, low CO₂-inducible 1 (LCI1) and low CO₂-inducible C (LCIC) following a transfer from H-CO₂ to VL-CO₂ either at 21% or at 2% O₂ (Fig. 3a and Supplementary Fig. 7). We observed a lower abundance of GYD1 in lci20 compared to control strains when the transfer was performed under photorespiratory conditions (21% O₂), but no difference was observed under non-photorespiratory conditions (2% O₂) (Fig. 3a, c) indicating that the down-regulation of GYD1 in lci20 is caused by the activity of photorespiration. The abundances of other photorespiration or CCM proteins were not consistently different in all three biological replicates between lci20 and control strains (Fig. 3a and Supplementary Fig. 7).

Fig. 3: Effect of CO2 and O2 concentrations on the abundance of photorespiration and CCM proteins. — **Fig. 3: Effect of CO₂ and O₂ concentrations on the abundance of photorespiration and CCM proteins.**

In Chlamydomonas, photorespiration and CCM genes are co-regulated during acclimation to sub-optimal CO₂ and controlled by a common transcription factor CIA5. We then monitored the accumulation of photorespiratory enzymes (GYD1, HPR1 and GCSP) and of CCM-related proteins (HLA3, LCI1 and LCIC) following acclimation to different CO₂ and O₂ levels in cia5 and its wild-type control (Fig. 3b and Supplementary Fig. 8). Whereas CCM and photorespiratory proteins were induced after 20 h of acclimation to VL-CO₂ in a CIA5-dependent manner, only photorespiratory proteins were detected under H-CO₂ (Fig. 3b, Supplementary Fig. 8), indicating that photorespiratory enzymes and CCM proteins are regulated differently at the protein level. Interestingly, photorespiratory proteins GCSP and GYD1 levels were more abundant in wild-type CC125 and cia5 under VL-CO₂ at 21% O₂ compared to VL-CO₂ 2% O₂ whereas CCM related proteins were unaffected by O₂ levels (Fig. 3b, d, Supplementary Fig. 8). HPR1 abundance was only affected by O₂ levels in the cia5 (Supplementary Fig. 8b). This effect of O₂ levels on the abundance of photorespiratory proteins was not observed in the wild-type CC4533 strain (Supplementary Fig. 7b), which could indicate a feedback regulation of photorespiratory proteins.

To better understand the reasons for the growth defect observed in lci20, and further determine whether it may be caused by the observed decrease of GYD1 abundance in lci20 mutant (Fig. 3c), we have isolated and characterized two CLiP mutants of Chlamydomonas harboring insertions in introns of the GYD1 gene (Fig. 4a) and validated to be deficient in the GYD1 protein by immunoblot (Fig. 4b). Both mutants showed higher glycolate excretion than the control strain (Fig. 4c). However, the growth of both gyd1 mutants was similar to the control wild-type strain after a transition from H-CO₂ to VL-CO₂ regardless of O₂ levels (Fig. 4d, Supplementary Fig. 9). gyd1 mutants did not require H-CO₂ to grow, which is in contrast to a previous work where the H-CO₂ requiring 89 mutant (HCR89) is reported to harbor a mutation at the GYD1 locus⁴⁵. Therefore, the growth defect of lci20 observed during H-CO₂ to VL-CO₂ is unlikely due to a decrease in GYD1 abundance but rather results from the accumulation of photorespiratory metabolites downstream the GYD1 reaction step. Indeed, the lci20 mutants excreted half as much glycolate as gyd1 mutants (Figs. 2b and 4c), which likely explain why their growth is more affected during a transition from H-CO₂ to VL-CO₂ under photorespiratory conditions.

Fig. 4: Characterization of two insertional mutants of the glycolate dehydrogenase gene GYD1. — Fig. 4: Characterization of two insertional mutants of the glycolate dehydrogenase gene *GYD1.*

Photosynthesis of lci20 is affected under photorespiratory conditions

To investigate the consequence of impaired photorespiration on the photosynthetic capacity of lci20, we performed room temperature chlorophyll fluorescence measurements to assess the PSII quantum yield and PQ redox state (1-qL) in H-CO₂ grown cells and following their acclimation to VL-CO₂ (Fig. 5, Supplementary Fig. 10). Under H-CO₂, lci20 showed a slightly higher effective PSII quantum yield and lower PQ redox state compared to wild-type but was mainly similar to the complemented line (Fig. 5a, d). Following VL-CO₂ acclimation under photorespiratory conditions (21% O₂), lci20 showed a significantly reduced effective PSII quantum yield and elevated PQ redox state compared to the control strains (wild-type and complemented line) (Fig. 5b, e). These effects were suppressed under non-photorespiratory conditions (VL-CO₂, 2% O₂) (Fig. 5c, f), thus indicating that the accumulation of non-metabolized photorespiratory compounds may be responsible for the inhibition.

Fig. 5: Photosynthetic activity measured by chlorophyll fluorescence is reduced in lci20 during acclimation to VL-CO2 under photorespiratory (21% O2) but not under non-photorespiratory (2% O2) conditions. — Fig. 5: Photosynthetic activity measured by chlorophyll fluorescence is reduced in *lci20* during acclimation to VL-CO₂ under photorespiratory (21% O₂) but not under non-photorespiratory (2% O₂) conditions.

Metabolomic analyses reveal an increased accumulation of malate and glutamate in lci20

To investigate the metabolic changes occurring in lci20 and obtain clues to the metabolites transported by LCI20, we performed a metabolomic analysis in the different strains grown under photoautotrophic H-CO₂, and during transitions from H-CO₂ to L-CO₂ and from H-CO₂ to VL-CO₂ (Fig. 6; Supplementary Fig. 11; Supplementary Data 1). The metabolite profile of H-CO₂-grown lci20 cells was comparable to that of control strains (Fig. 6a Supplementary Fig. 11; Supplementary Data 1). After transition from H-CO₂ to L-CO₂ or to VL-CO₂, we observed a higher accumulation of malic acid and glutamic acid in lci20 when cells were cultivated under photorespiratory condition (21% O₂), the effect being suppressed under non-photorespiratory conditions (2% O₂) (Fig. 6b, d). Accumulation of almost all the other metabolites detected was not significantly affected in lci20 under the conditions tested (Supplementary Fig. 11; Supplementary Data 1). Based on these results, we propose that LCI20 is involved in the transport of malate and glutamate during photorespiration and that disruption of glutamate transport impairs photorespiratory glycolate metabolism at the step of glyoxylate conversion to glycine thus explaining the lci20 phenotypes observed during acclimation to VL-CO₂.

Fig. 6: Metabolomic analysis shows increased accumulation of malate and glutamate in lci20 during a transition from H-CO2 to VL-CO2 under photorespiratory but not under non-photorespiratory conditions. — Fig. 6: Metabolomic analysis shows increased accumulation of malate and glutamate in *lci20* during a transition from H-CO₂ to VL-CO₂ under photorespiratory but not under non-photorespiratory conditions.

Discussion

In this study, we demonstrate the role of LCI20 in the photorespiratory glycolate metabolism in Chlamydomonas (Fig. 7), as evidenced by a high glycolate excretion and the marked decline in growth of the lci20 mutant during a transition from H-CO₂ to VL-CO₂ at ambient O₂ levels (21%), these effects being suppressed at 2% O₂ (Figs. 1, 2). Many photorespiratory mutants have been previously reported in land plants and only very few in algae. In Arabidopsis, a defect in genes encoding photorespiratory enzymes generate a high-CO₂ requiring phenotype, which is attributed to the accumulation of non-metabolized photorespiratory intermediates¹³. In Chlamydomonas, the situation is not that clear, and this is further complicated by the occurrence of CCM in algae. A phosphoglycolate phosphatase 1 (pgp1) mutant, impaired in the conversion of 2-PG to glycolate, exhibits a growth defect under stationary conditions of limiting CO₂ levels²⁴. The high-CO₂ growth requirement of pgp1 was attributed to the inhibitory effect of non-metabolized 2-PG accumulating upon ribulose bisphosphate oxygenation²⁴. After the conversion of 2-PG into glycolate by PGP1, glycolate can follow two different fates in Chlamydomonas, it can be either excreted out of the cell or metabolized into glyoxylate by GYD1 in the mitochondria. A Chlamydomonas gyd1 mutant isolated from a screening for an H-CO₂ requirement showed a high glycolate excretion during transition from H-CO₂ to L-CO₂⁴⁵. However, the H-CO₂ requiring phenotype of this mutant was not clearly evidenced. We have shown here that impairing photorespiration at the level of GYD1 does not lead to an H-CO₂ requirement but instead to a high glycolate excretion under VL-CO₂ (Fig. 4, Supplementary Fig. 9). It was recently reported that growth of Chlamydomonas hpr1 mutant is slightly impaired during the transition from H-CO₂ to L-CO₂, accompanied by a high glycolate excretion⁵⁷. The slight growth defect of the hpr1 mutant being interpreted as a consequence of glycolate hyper-excretion⁵⁷. However, the gyd1 mutants characterized here show high glycolate excretion rates but no growth defect in any CO₂ conditions tested (Fig. 4, Supplementary Fig. 9). We therefore conclude that inhibition of photorespiration at different steps of the pathway can lead to contrasting phenotypes. While inhibition of glycolate conversion does not impair growth (i.e. in gyd1), the accumulation of intermediate metabolites such as 2-PG (i.e. in pgp1) or downstream glycolate conversion (i.e. in hpr1) is deleterious. The strong down-regulation of GYD1 observed in lci20 under photorespiratory conditions (Fig. 3a) may result from a protective mechanism aiming at limiting the toxicity of non-metabolized photorespiratory intermediates by decreasing their production while promoting glycolate excretion. Consequently, glycolate excretion together with GYD1 down-regulation could be seen as complementary protection mechanisms preventing the toxic accumulation of photorespiratory intermediates during acclimation to VL-CO₂. Such mechanisms would be particularly needed until CCM components and photorespiratory enzymes are fully induced, thus allowing photorespiratory metabolism to be tuned to the residual Rubisco oxygenase activity which remains at VL-CO₂ despite the presence of CCM.

Fig. 7: Hypothetical scheme describing the role of LCI20 during photorespiration in Chlamydomonas. — Fig. 7: Hypothetical scheme describing the role of LCI20 during photorespiration in *Chlamydomonas.*

Unlike the above-mentioned proteins (PGP1, GYD1 and HPR1), which are all involved in the core photorespiratory pathways, LCI20 turns out to be an auxiliary protein involved in the supply of amino (-NH₂) donor as glutamate for the conversion of glyoxylate into glycine (Fig. 7). In Arabidopsis chloroplasts, the export of glutamate in counter-exchange with malate is mediated by the dicarboxylate transporter DiT2.1 that is required for photorespiratory nitrogen assimilation and displays a 60% homology with the Chlamydomonas LCI20^58,59,60,61. Based on our metabolomic analysis, we suggest that LCI20 mediates the export of glutamate in counter exchange with malate across the chloroplast inner envelope (Fig. 6; Supplementary Fig. 11). Impairing glutamate export by knocking out LCI20 likely disrupts glycolate metabolism at the step of glyoxylate transamination. The higher glycolate excretion observed in lci20 (Fig. 2b) would then result from an inefficient conversion of glycolate into glyoxylate, which would in turn result in a decrease in the GYD1 abundance (Fig. 3a, c). Both glycolate and glyoxylate, which could not be determined in our metabolomic analysis, are intermediate metabolites whose accumulation has been shown to be toxic for plants^62,63. Because Chlamydomonas is unlikely to accumulate glycolate due to the existence of an excretion pathway, we conclude that the growth impairment of lci20 does not result from glycolate excretion per se, but rather from poisoning by non-metabolized glyoxylate. In line with our data, mutants deficient in the LCI20 homologues DiT2.1 and OMT1 in tobacco and Arabidopsis showed photoinhibition of PSII caused by the negative feedback of non-metabolized glyoxylate on the activation state of the Calvin cycle enzymes^60,64,65. Glyoxylate was shown to be over accumulated in omt1 mutants in both Arabidopsis and tobacco^60,64. The negative effects of glyoxylate on the activation state of Rubisco was shown in several studies^66,67. Overall, LCI20 plays a vital role during the acclimation of Chlamydomonas from H-CO₂ to VL-CO₂ by exporting glutamate from chloroplast toward mitochondria which is required for the function of photorespiratory pathway to efficiently recycle Rubisco oxygenation product 2-PG back to the Calvin cycle in the chloroplast.

The fact that lci20 growth is unaffected under steady-state L-CO₂ or VL-CO₂ could be explained in two different ways. On the one hand, we could consider that photorespiration is only transiently active before the activation of CCM during acclimation to sub-optimal CO₂ levels but is suppressed or strongly reduced upon CCM activation under steady-state conditions. A similar conclusion was drawn from the observation that the glycolate excretion is suppressed in air acclimated Chlamydomonas wild-type cells when CCM is fully induced¹⁸. However, we can also imagine that the arrest of glycolate excretion is caused by the upregulation of enzymes of the glycolate metabolic pathway both at transcript and protein levels. The later hypothesis is supported by the fact that the transaminase inhibitor aminooxyacetate (AOA), which blocks the photorespiratory pathway, induces a higher glycolate excretion in air-adapted Chlamydomonas wild-type cells with a fully activated CCM⁴⁵. Alternatively, we could consider that during photorespiration the need for LCI20-mediated -NH₂ recycling is only transient. Indeed, two -NH₂ are needed for the conversion of two molecules of glyoxylate into two glycines. One -NH₂ is likely supplied by the conversion of one serine into one hydroxypyruvate. Since the conversion of two molecules of glycine into one serine is accompanied by the release of NH₃, the later needs to be re-assimilated to balance the nitrogen budget. In Chlamydomonas, inhibition of glutamine synthetase induces excretion of NH₃ in an O₂-dependent manner therefore showing that at least part of the NH₃ produced by photorespiration is recycled within chloroplasts by the GS-GOGAT pathway⁶⁸. In Arabidopsis, the re-assimilation of photorespiratory NH₃ is mediated by the chloroplast GS/GOGAT cycle, which produces glutamate in turn re-imported into mitochondria; this process is essential for photorespiration as evidenced by the lethal phenotype of both chloroplast and mitochondrial glutamate transporters^59,69,70. In this context, the transient phenotype of lci20 may result from the induction of an alternative pathway of NH₃ reassimilation for instance through a mitochondrial isoform of the glutamate dehydrogenase⁷¹ which may compensate for the lack of LCI20 under steady state limiting CO₂ (Fig. 7).

An hypothesis often put forward regarding the physiological role of photorespiration in microalgae is that photorespiratory metabolites could be the metabolic signal responsible for the induction of CCM during acclimation to sub-optimal CO₂ levels^{11,14,22,23,24,72,73}. Our data clearly establish that both the accumulation of CCM components (Fig. 3b, Supplementary Fig. 8) and the CCM functioning (Fig. 2a) are independent of O₂ levels, thus ruling out this hypothesis. In C3 plants, it has recently been proposed that photorespiration helps to maintain cellular redox balance by generating malate in the chloroplast, with malate export to the cytosol providing NADH for nitrogen assimilation⁷⁴. Such a role in redox balance or any other physiological function of photorespiration remains to be demonstrated in microalgae.

This work paves the way for a more detailed understanding of photorespiration in microalgae and how it interacts with the CCM functioning. Significant amounts of glycolate have been found in the ocean, particularly during algal blooms^{75,76,77,78,79}, suggesting that photorespiration can be active in phytoplankton despite the occurrence of CCMs in most species^9,80,81. This apparent discrepancy could be due to the fact that CCMs from different marine species show quite different efficiencies^9,80,81. For instance, Nannochloropsis has been reported to harbour a quite leaky CCM⁸². It is also possible that algal species with an efficient CCM excrete significant glycolate amounts in particular conditions, for instance when the CO₂ concentration drops during algal blooms, or when the incident light suddenly increases. Further studies are needed to better understand how algal photorespiration contributes to the carbon footprint of oceans.

Methods

Growth conditions and strains

The lci20 (LMJ.RY0402.205944), gyd1-1 (LMJ.RY0402.206272), gyd1-2 (LMJ.RY0402.232044), cia5 (CC-2702), and wild-types CC4533 and CC125 were purchased from the Chlamydomonas resource center. Cells were cultivated in an incubation shaker (INFORS Multitron pro) maintained at 25 °C, with 120 rpm shaking and constant illumination at 80 µmol m⁻² s⁻¹ supplied by fluorescent tubes delivering white light enriched in red wavelength. Cells were grown in MOPS-buffered (20 mM MOPS, pH 7.2) minimal medium (MM) exposed to various CO₂ levels. Photorespiration was induced by transferring 2% (H) CO₂-grown cells into 0.01% (VL) CO₂ under atmospheric O₂ level (21%) and can be suppressed by 2% O₂. Growth kinetics were monitored with a Multisizer 3 Coulter counter (Beckman Coulter). For the wild-type and gyd1-1 growth assays performed in liquid culture, cells were cultivated photoautotrophically in 1-L photobioreactors operated as turbidostats. Cell density was monitored through turbidity measurement (OD 800 nm) and maintained at 0.4 O.D by addition of fresh media. The temperature was maintained at 25 °C, the pH at 7.2, at the light intensity at 125 µmol photons m⁻² s⁻¹. Cultures were bubbled with gas mixtures containing CO₂ at 4%, 0.1%, or 0.04% in air, and supplied at a constant flow rate of 0.5 L min⁻¹. The dissolved CO₂ concentration in the photobioreactor medium was measured by membrane inlet mass spectrometry⁸³ upon rapid filtration of the culture. For spot test, cells were grown in liquid culture with MM, harvested during active growth at around 10 µg mL⁻¹ of chlorophyll and resuspended in fresh MM medium to make series of dilutions to 0.5, 1, and 2 µg mL⁻¹ chlorophyll per spot. Eight-microliter drops were spotted on 1.5% of MM agar plates at pH 7.2 buffered with 20 mM MOPS and exposed to various CO₂ and light regimes. Homogeneous light was supplied by a panel of fluorescent tubes.

Protein extraction and immunoblot analysis

Total protein was extracted as previously described in ref. ²⁶. Exponentially grown cells (equivalent of 20 µg of chlorophyll) were harvested by centrifugation at 4000 g for 3 min at 4 °C. Pellets were resuspended in 200 µL of PBS with a complete protease inhibitor EDTA-free mixture tablet (Roche) by vortexing and sonicated for 15 s 30% pulses on ice using a sonicator (product number UR-21P; TOMY). 200 μL of Novex™ Nupage™ LDS buffer 2x (Invitrogen™) containing 1x reducing agent DTT was added to the solution, and the total protein was solubilized by incubation at 37 °C for 20 min. Incubated samples were subsequently centrifuged at 4000 g for 3 min. 10 μL (1X) of protein samples were loaded on Novex™ Nupage™ Bis tris 10% or tris-acetate 3-8% (Invitrogen™) gel, migrated 1 h at 190 V in Novex™ Nupage™ MOPS or tris-acetate (Invitrogen™) buffer according to protein molecular weight and transferred to nitrocellulose membrane using semidry transfer technique. Immuno-detection was performed using antibodies raised against LCI20 (1/250), GYD1 (1/500), HPR1 (1/500), GCSP (1/500), HLA3 (1/500), LCI1 (1/500) and LCIC (1/500). Antibody raised against α-Tubulin 1/2000 (Sigma-Aldrich ref. T6074) was used as control. Secondary anti-rabbit peroxidase-conjugated antibodies (Sigma-Aldrich; no. AQ132P) (1/10,000) were used for the detection with the G:BOX Chemi XRQ system (Syngene) using ECL detection reagents (GE Healthcare). Images were captured with a CCD camera equipped with a GeneSys Image Acquisition Software (Syngene).

Glycolate quantification

Excreted glycolic acid was analyzed from the supernatant after centrifugation of 1 mL culture at 4000 g for 2 min at 4 °C. The supernatant was diluted 10x in acetonitrile and analyzed using a Vanquish UHPLC/Q Exactive Plus (ThermoScientific). Glycolic acid was separated using a HILIC stationary phase (SeQuant ZIC HILIC, 100 × 2.1 mm, 3.5 µm), heated to 35 °C. A binary solvent system was used, in which mobile phase A consisted of acetonitrile: water (95:5, v/v) with 5 mM ammonium acetate and mobile phase B consisted of water:acetonitrile (95:5, v/v) with 5 mM ammonium acetate. Separations were made over a 32 min period following the gradient: 0-0.5 min: 5% B, 0.5–24.5 min: 5-95% B, 24.5–26.5 min: 95% B, 26.5–26.6 min: 95-5% B, 26.6–32 min: 5% B. The flow rate was set to 0.3 mL min^–¹ and the injection volume was 5 µL. After separation, glycolic acid was directed into the ESI source of the Q Exactive Plus (Orbitrap-mass spectrometer). The ESI source was set as following: negative mode ion spray voltage at –2.5 kV, capillary temperature at 300 °C, S-lens RF level at 50. Data was acquired using a targeted t-SIM method with glycolic acid in inclusion list (formula C₂H₄O₃, m/z 75.00877).

Measurement of chlorophyll fluorescence

Chlorophyll fluorescence measurements were performed using a Dual Pulse Amplitude Modulated Fluorometer (DUAL-PAM-100; Walz) equipped with a red LED source of actinic light. Samples (2 mL of cells at 10 µg mL^–¹ chlorophyll) were placed into a cuvette under constant stirring at room temperature (25 °C) and dark adapted for 15 min prior to measurement. Both effective PSII quantum yield and 1-qL were calculated from the light curve. The latter was obtained by increasing red actinic light stepwise every 1 min starting from 17, 110, 190, 390, 610 and 940 μmol photons m⁻² s⁻¹, each being separated by a saturating pulse 10,000 μmol photons m⁻² s⁻¹, 600 ms duration. The effective PSII quantum yield and 1-qL were calculated as ΦPSII = (F_m’-F_s)/F_m’ and qL = ((F_m’-F_s)/(F_m’-F₀))*(F₀/F_s) respectively with Fm’ the fluorescence value after saturating pulses, Fs the stationary fluorescence at each actinic light and F₀ the fluorescence value in the dark⁸⁴.

Metabolomics analysis

Cells were grown to exponential phase under photoautotrophic H-CO₂ conditions in the flask with 80 μmol photons m⁻² s⁻¹. For L-CO₂ and VL-CO₂ conditions, H-CO₂ grown cells were transferred to either L-CO₂ or VL-CO₂ levels for 20 h prior to sampling. About 60 million cell suspensions were injected into a −70 °C cold quenching solution composed of 70% methanol in water using a thermoblock above dry ice to obtain a final concentration of 35% methanol. Centrifuge tubes containing the cell suspensions were cooled in a pre-chilled cooling box to keep the sample temperature below −20 °C. Cell pellets were collected by centrifugation at 4000 g for 2 min at −10 °C. The supernatant was decanted and residual liquid together with cells carefully transferred to a 2 mL Eppendorf tube and re-centrifuged 13,000 g for 1 min at −10 °C. The pellet was flash-frozen in liquid nitrogen and lyophilized at −50 °C. The extraction and derivation of the metabolites were performed as previously⁸⁵. The analytic and quantification methods were exactly as reported in ref. ³⁷. Data is reported following recently updated metabolomics standards⁸⁶.

Statistics

All statistical tests used are noted in figure legends. One-way or two-way ANOVA using GraphPad Prism (GraphPad Software) was used to perform statistical analysis. The P-values were computed by One-way or two-way ANOVA test (uncorrected p values). Statistical significance (α = 0.05) according to p values is indicated by asterisk (*p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001 and ****p ≤ 0.0001).

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Genes studied in this article can be found on Phytozome 13 (Chlamydomonas genome v6.1 or 5.6) under the loci Cre06.g260450 [LCI20], Cre06.g295450 [HPR1], Cre06.g288700 [GYD1], Cre12.g534800 [GCSP], Cre03.g162800 [LCI1], Cre02.g097800 [HLA3], Cre06.g307500 [LCIC] and Cre02.g096300 [CIA5]. All antibodies used are available upon request to corresponding authors. Metabolites data are provided as supplementary data 1. All the other methods are reported in the Supplementary Information file . Source data are provided with this paper.

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Acknowledgements

O.D. thanks the French Atomic Energy and Alternative Energy Commission (CEA) for a PhD scholarship. We thank Shiyan Zheng for assistance in performing the genetic complementation experiment; Gaurav Kumar for technical assistance during microscopic observation; Stephanie Blangy for trials on generating antibodies against LCI20; Franziska Kuhnert for trials on the transport assays. We thank the ZoOM Microscopy facility (CEA Cadarache) and the University of York Biosciences Technology Facility for confocal microscopy access. This work is financially supported by the “L’économie Circular du Carbone” program of the CEA (CO2storage), the ANR project “AlgalCCM” (n°ANR-366 22-CE44-0023-01), the France 2030 initiative project “CO2_CMPhi” (ANR-23-PEXF-0002) and the Region Sud (“AlgalCO2” project). We acknowledge the European Union Regional Developing Fund (ERDF), the Région Provence Alpes Côte d’Azur, the French Ministry of Research and the CEA for funding the HelioBiotec platform. C.M. is grateful to the INRAE MIGALE bioinformatics facility (MIGALE, INRAE, 2020) for providing computing resources. AKL was supported by the Labex Saclay Plant Sciences-SPS (ANR-17-EUR-0007), the platform of Biophysics of the I2BC supported by the French Infrastructure for Integrated Structural Biology (FRISBI; grant number ANR-10-INSB-05). A.P.M.W. acknowledges funding by the European Union’s H2020 research and innovation and the Horizon programs (grants GAIN4CROPS, GA No. 862087 and BEST-CROP, GA No. 101082091) and the Deutsche Forschungsgemeinschaft (Cluster of Excellence for Plant Sciences (CEPLAS) under Germany’s Excellence Strategy EXC-2048/1 under project ID 390686111). S.A. and A.R.F. acknowledge the European Union’s Horizon 2020 research and innovation program, project PlantaSYST (SGA-CSA No. 739582 under FPA No. 664620) and the BG05M2OP001-1.003-001-C01 project, financed by the European Regional Development Fund through the Bulgarian ‘Science and Education for Smart Growth’ Operational Program. Both authors acknowledge the support by the Max Planck Society. A.B. acknowledges support from the Carnegie Institution for Science.

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Authors and Affiliations

Aix-Marseille Université, CEA, CNRS, BIAM, UMR7265, Institut de Biosciences et Biotechnologies Aix-Marseille, CEA Cadarache, Saint-Paul-lez-Durance, France
Ousmane Dao, Marie Bertrand, Florian Veillet, Pascaline Auroy, Phuong-Chi Nguyen, Bertrand Légeret, Virginie Epting, Amélie Morin, Stephan Cuiné, Caroline L. Monteil, Gilles Peltier & Yonghua Li-Beisson
Department of Molecular Physiology, Max Planck Institute of Molecular Plant Physiology, Potsdam-Golm, Germany
Saleh Alseekh & Alisdair R. Fernie
Center of Plant Systems Biology and Biotechnology, Plovdiv, Bulgaria
Saleh Alseekh & Alisdair R. Fernie
Centre for Novel Agricultural Products, Department of Biology, University of York, York, UK
Luke C. M. Mackinder
Department of Plant Biology, Division of Biosphere Sciences and Engineering, The Carnegie Institution for Science, Stanford, CA, USA
Adrien Burlacot
Department of Biology, Stanford University, Stanford, CA, USA
Adrien Burlacot
Université Paris-Saclay, Institute for Integrative Biology of the Cell (I2BC), CEA, CNRS, CEDEX, Gif-sur-Yvette, France
Anja Krieger-Liszkay
Institute of Plant Biochemistry, Cluster of Excellence on Plant Science (CEPLAS), Heinrich Heine University, Düsseldorf, Germany
Andreas P. M. Weber

Authors

Ousmane Dao
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Marie Bertrand
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Saleh Alseekh
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Florian Veillet
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Pascaline Auroy
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Phuong-Chi Nguyen
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Bertrand Légeret
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Virginie Epting
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Amélie Morin
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Stephan Cuiné
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Caroline L. Monteil
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Luke C. M. Mackinder
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Adrien Burlacot
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Anja Krieger-Liszkay
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Andreas P. M. Weber
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Alisdair R. Fernie
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Gilles Peltier
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Yonghua Li-Beisson
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Contributions

Y.L-B., G.P., O.D. conceived the study. Y.L-B. and G.P. supervised the work. O.D. performed most of the experiments. A.B., A.K.L. and G.P. supervised photosynthesis measurement performed by O.D. P.A., M.B., S.C. and O.D. carried out immunoblots. O.D. and V.E. isolated and characterized gyd1 mutants with contribution from A.M. C.M. performed phylogenetic analysis. M.B., F.V. and O.D. performed genetic complementation of lci20 mutant. S.A. and A.R.F. performed metabolomics. O.D., P.-C.N. and B.L. performed glycolate analysis. O.D., G.P. and Y.L-B. drafted the manuscript with contributions from L.C.M.M., A.B., A.K.L., A.R.F. and A.P.W.

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Correspondence to Gilles Peltier or Yonghua Li-Beisson.

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Dao, O., Bertrand, M., Alseekh, S. et al. The green algae CO₂ concentrating mechanism and photorespiration jointly operate during acclimation to low CO₂. Nat Commun 16, 5296 (2025). https://doi.org/10.1038/s41467-025-60525-7

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Received: 18 September 2024
Accepted: 27 May 2025
Published: 17 June 2025
Version of record: 17 June 2025
DOI: https://doi.org/10.1038/s41467-025-60525-7

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