Nitrous Oxide Emissions from Nitrite Are Highly Dependent on Nitrate Reductase in the Microalga Chlamydomonas reinhardtii

Nitrous oxide (N2O) is a powerful greenhouse gas and an ozone-depleting compound whose synthesis and release have traditionally been ascribed to bacteria and fungi. Although plants and microalgae have been proposed as N2O producers in recent decades, the proteins involved in this process have been only recently unveiled. In the green microalga Chlamydomonas reinhardtii, flavodiiron proteins (FLVs) and cytochrome P450 (CYP55) are two nitric oxide (NO) reductases responsible for N2O synthesis in the chloroplast and mitochondria, respectively. However, the molecular mechanisms feeding these NO reductases are unknown. In this work, we use cavity ring-down spectroscopy to monitor N2O and CO2 in cultures of nitrite reductase mutants, which cannot grow on nitrate or nitrite and exhibit enhanced N2O emissions. We show that these mutants constitute a very useful tool to study the rates and kinetics of N2O release under different conditions and the metabolism of this greenhouse gas. Our results indicate that N2O production, which was higher in the light than in the dark, requires nitrate reductase as the major provider of NO as substrate. Finally, we show that the presence of nitrate reductase impacts CO2 emissions in both light and dark conditions, and we discuss the role of NO in the balance between CO2 fixation and release.


Introduction
Nitrous oxide (N 2 O) is a greenhouse gas~300-fold more potent than CO 2 and considered the dominant ozone-depleting chemical emitted in the 21st century [1][2][3][4][5]. In 2020, the atmospheric N 2 O reached 333.2 ppb, which constitutes 123% of the pre-industrial (before 1750) levels, with the fastest-growing rate occurring in the past five decades [6][7][8]. N 2 O emissions are released to the atmosphere from natural (~60%) and anthropogenic sources (~40%), including oceans, soils, biomass burning, fertilizers, and several industrial activities. N 2 O emissions derived from human activities are dominated by nitrogen additions to crop plants [6,8]. In modern agriculture, the abundant supply of nitrogen fertilizers leads to excess nitrogen in the soil, and non-assimilated nitrogen can be emitted as N 2 O to the atmosphere or lost as runoffs into aquatic ecosystems, causing their eutrophication [9,10]. Nitrification and denitrification are two well-documented biochemical processes that control N 2 O emissions in terrestrial and aquatic ecosystems and are regulated by biological and environmental factors [8,11].
Bacteria and fungi are widely recognized as N 2 O producers by the scientific community [1,[11][12][13][14], but recently, plants and algae have also emerged as N 2 O emitters. In the late 1970s, Hahn and Junge already hypothesized that phytoplankton and plants could release N 2 O in the presence of nitrate (NO 3 − ) and nitrite (NO 2 − ) [15]. Several years later, this was demonstrated in microalgae [16,17] and plant leaves during photosynthesis [18][19][20][21][22]. Despite this, the intergovernmental agencies have not yet considered N 2 O emissions by plants

Nitrite Reductase Mutants (nii1) Cannot Use NO 3
− /NO 2 − for Growth but Can Reduce Them to N 2 O Chlamydomonas nii1 mutants (G1, M3, and M4) cannot reduce NO 2 − to ammonium (NH 4 + ) and, therefore, do not grow in media containing either NO 3 − or NO 2 − as the sole nitrogen (N) source (Figure 1a). The G1 strain is a deletion mutant that lacks the entire cluster of the NO 3 − assimilation genes. This cluster, located in chromosome 9, contains the genes that encode NO 3 − and NO 2 − reductases (NIA1 and NII1, respectively) and the high-affinity NO 3 − /NO 2 − transport components (NRT2.1, NRT2.2, and NAR2) [40,41]. By genetic crosses, either the NIA1 and NRT2.1-NAR2 sets of genes or only NIA1 were transferred to the G1 strain, generating the M3 and M4 mutants, respectively (see [41] for more details). As previously mentioned, Chlamydomonas cells can reduce NO 2 − to NO [37] and NO to N 2 O [26,27]; therefore, we used these mutants as model organisms to study this process in microalgae. First, we studied NO 2 − evolution in the M3 strain. NH 4 + -grown cells were washed and transferred to fresh media containing 0.1 and 1 mM NO 3 − or NO 2 − , and NO 2 − concentration in the medium was determined at different time points (Figure 1b-d). Cells exposed to 0.1 mM NO 3 − showed a stoichiometric excretion of NO 2 − after 4 h (Figure 1b), as previously reported [41]. Subsequently, extracellular NO 2 − concentration slowly decreased, being completely exhausted from the medium after 24 h. Similar depletion rates and kinetics were observed when 0.1 mM NO 2 − was added instead, but a lag of 4-6 h was observed before the concentration started to decrease.

Nitrite Reductase Mutants (nii1) Cannot Use NO3 − /NO2 − for Growth but Can Reduce Them to N2O
Chlamydomonas nii1 mutants (G1, M3, and M4) cannot reduce NO2 − to ammonium (NH4 + ) and, therefore, do not grow in media containing either NO3 − or NO2 − as the sole nitrogen (N) source (Figure 1a). The G1 strain is a deletion mutant that lacks the entire cluster of the NO3 − assimilation genes. This cluster, located in chromosome 9, contains the genes that encode NO3 − and NO2 − reductases (NIA1 and NII1, respectively) and the high-affinity NO3 − /NO2 − transport components (NRT2.1, NRT2.2, and NAR2) [40,41]. By genetic crosses, either the NIA1 and NRT2.1-NAR2 sets of genes or only NIA1 were transferred to the G1 strain, generating the M3 and M4 mutants, respectively (see [41] for more details). As previously mentioned, Chlamydomonas cells can reduce NO2 − to NO [37] and NO to N2O [26,27]; therefore, we used these mutants as model organisms to study this process in microalgae. First, we studied NO2 − evolution in the M3 strain. NH4 + -grown cells were washed and transferred to fresh media containing 0.1 and 1 mM NO3 − or NO2 − , and NO2 − concentration in the medium was determined at different time points (Figure 1b-d). Cells exposed to 0.1 mM NO3 − showed a stoichiometric excretion of NO2 − after 4 h (Figure 1b), as previously reported [41]. Subsequently, extracellular NO2 − concentration slowly decreased, being completely exhausted from the medium after 24 h. Similar depletion rates and kinetics were observed when 0.1 mM NO2 − was added instead, but a lag of 4-6 h was observed before the concentration started to decrease.  The same experiment was performed in sealed bottles, in which N 2 O emission would be retained and could be quantified. Under these conditions, similar rates of accumulation and depletion of NO 2 − were observed ( Figure 1c). However, NO 2 − depletion was induced faster than in non-sealed cultures (2 h vs. 6-8 h); therefore, NO 2 − excretion after NO 3 − reduction was not stoichiometric and reached only a concentration of 86 µM. Furthermore, as observed in non-sealed bottles, NO 2 − was exhausted before 24 h. A similar pattern was observed when cells were exposed to 1 mM NO 3 − or NO 2 − , although total depletion required longer incubations (Figure 1d).  (Figure 2). When the cells were incubated with 0.1 mM NO 2 − , N 2 O started to accumulate after 2-3 h with a rate of 3.3 ppm/h and plateaued after 21 h, reaching a final concentration of 62 ppm after 24 h (Figure 2a). In the presence of 10 mM NO 2 − , although N 2 O accumulation was also detected after 2 h of induction, the gas was released at~15-fold higher rate (51 ppm/h) than in 0.1 mM NO 2 − , and no saturation was observed after 24 h when N 2 O concentration was 864 ppm (Figure 2b). When the cells were incubated with 10 mM NO 3 − (Figure 2c), N 2 O release was delayed as expected due to the requirement to reduce NO 3 − to NO 2 − , but the production rate was boosted after 14 h (92 ppm/h), almost doubling that observed in the cells supplemented with NO 2 − . As expected, cells incubated in N-free media did not emit detectable amounts of N 2 O (Supplementary Table S1).
would be retained and could be quantified. Under these conditions, similar rates of accumulation and depletion of NO2 − were observed ( Figure 1c). However, NO2 − depletion was induced faster than in non-sealed cultures (2 h vs. 6-8 h); therefore, NO2 − excretion after NO3 − reduction was not stoichiometric and reached only a concentration of 86 µM. Furthermore, as observed in non-sealed bottles, NO2 − was exhausted before 24 h. A similar pattern was observed when cells were exposed to 1 mM NO3 − or NO2 − , although total depletion required longer incubations (Figure 1d).
To monitor N2O emissions in the headspace of the cultures, we used Cavity Ring-Down Spectroscopy (CRDS) (see Material and Methods), which allows continuous N2O measurements. The M3 cultures produced N2O in a NO2 − concentration-dependent manner and from both NO3 − and NO2 − (Figure 2). When the cells were incubated with 0.1 mM NO2 − , N2O started to accumulate after 2-3 h with a rate of 3.3 ppm/h and plateaued after 21 h, reaching a final concentration of 62 ppm after 24 h (Figure 2a). In the presence of 10 mM NO2 − , although N2O accumulation was also detected after 2 h of induction, the gas was released at ~15-fold higher rate (51 ppm/h) than in 0.1 mM NO2 − , and no saturation was observed after 24 h when N2O concentration was 864 ppm (Figure 2b). When the cells were incubated with 10 mM NO3 − (Figure 2c), N2O release was delayed as expected due to the requirement to reduce NO3 − to NO2 − , but the production rate was boosted after 14 h (92 ppm/h), almost doubling that observed in the cells supplemented with NO2 − . As expected, cells incubated in N-free media did not emit detectable amounts of N2O (Supplementary Table S1). In Chlamydomonas, N2O production may involve light-dependent and lightindependent pathways [26,27]; therefore, we additionally studied N2O production in cells incubated with NO2 − in the dark. In this condition, total N2O accumulation (270 In Chlamydomonas, N 2 O production may involve light-dependent and light-independent pathways [26,27]; therefore, we additionally studied N 2 O production in cells incubated with NO 2 − in the dark. In this condition, total N 2 O accumulation (270 ppm) and production rate (20 ppm/h) were both strongly reduced (Figure 2d), highlighting the importance of light in this process in the M3 strain.
In these experiments, the earliest N 2 O emissions were achieved during incubation with 10 mM NO 2 − , a concentration previously used by Plouviez and collaborators [26]; therefore, we set this concentration for further studies. Moreover, the kinetics and high rates of N 2 O production observed in the M3 strain led us to use this mutant as a model to study the role of other players involved in the reduction of NO 2 − to N 2 O.

Nitrate Reductase Is the Primary NO Source Involved in N 2 O Emissions from NO 2 − in the nii1 Mutants
The enzymes responsible for NO reduction to N 2 O are located in the chloroplast (FLV) [27] and mitochondria (CYP55) [42] in Chlamydomonas. However, the NO sources that feed these reactions are not well understood. In plants and algae, the cytosolic NR seems to be the main enzyme involved in NO synthesis from NO 2 − [30,31,43], although other pathways for NO synthesis have been proposed in chloroplasts [44] and mitochondria [26].
Here, we set out to elucidate the possible role of the NR-ARC complex as a NO source for the synthesis of N 2 O. First, N 2 O emissions were compared in the nii1 mutants G1 (NR − ) and M4 (NR + ) ( Figure 3a). The lack of NR led to a dramatic reduction in N 2 O accumulation after 24 h in both light (31 ppm) and dark (77 ppm) conditions, while the M4 strain behaved similarly to the M3 mutant, reaching 904 ppm after 24 h in the light and 395 ppm in the dark (Figure 3a,b).
In these experiments, the earliest N2O emissions were achieved during incubation with 10 mM NO2 − , a concentration previously used by Plouviez and collaborators [26]; therefore, we set this concentration for further studies. Moreover, the kinetics and high rates of N2O production observed in the M3 strain led us to use this mutant as a model to study the role of other players involved in the reduction of NO2 − to N2O.

Nitrate Reductase Is the Primary NO Source Involved in N2O Emissions from NO2 − in the nii1 Mutants
The enzymes responsible for NO reduction to N2O are located in the chloroplast (FLV) [27] and mitochondria (CYP55) [42] in Chlamydomonas. However, the NO sources that feed these reactions are not well understood. In plants and algae, the cytosolic NR seems to be the main enzyme involved in NO synthesis from NO2 − [30,31,43], although other pathways for NO synthesis have been proposed in chloroplasts [44] and mitochondria [26]. Here, we set out to elucidate the possible role of the NR-ARC complex as a NO source for the synthesis of N2O. First, N2O emissions were compared in the nii1 mutants G1 (NR − ) and M4 (NR + ) (Figure 3a). The lack of NR led to a dramatic reduction in N2O accumulation after 24 h in both light (31 ppm) and dark (77 ppm) conditions, while the M4 strain behaved similarly to the M3 mutant, reaching 904 ppm after 24 h in the light and 395 ppm in the dark (Figure 3a,b). Secondly, to study the potential role of ARC in N2O emission, we transferred the arc mutation to the M3 background by genetic crossing. This new strain (M3arc) showed a significant reduction in N2O accumulation after 24 h in both light (~144 ppm) and dark (~69 ppm) conditions (Figure 3a,b), suggesting that the NR-ARC complex is responsible Secondly, to study the potential role of ARC in N 2 O emission, we transferred the arc mutation to the M3 background by genetic crossing. This new strain (M3arc) showed a significant reduction in N 2 O accumulation after 24 h in both light (~144 ppm) and dark (~69 ppm) conditions (Figure 3a,b), suggesting that the NR-ARC complex is responsible for the synthesis of most of the NO that sustains N 2 O production. To confirm this idea, NO levels were measured in these four strains (M3, M3arc, M4, and G1) using the DAF-FM fluorescent probe in cells incubated for 24 h in 10 mM NO 2 − under illumination (Figure 3c). G1 and M3arc strains exhibited a pronounced reduction in fluorescence (50% and 30%, respectively) compared to their corresponding strain of reference, M4, and M3. Our results suggest that NR-ARC is the main player in NO synthesis to feed NO reductases, but also that other NR-ARC-independent pathways should be considered.
If NR is required for N 2 O production as a key NO supplier, then the exogenous addition of NO should enhance N 2 O production in the NR-lacking G1 strain. To test this hypothesis, G1 cells were incubated for 20 h with 10 mM NO 2 − in either light or dark conditions and then were exposed to NO donor (40 µM DEA-NONOate). In both conditions, an immediate burst of N 2 O emission was observed. Before NO donor addition, N 2 O was produced with a rate of 0.66 ppm/h and 4.42 ppm/h in light and dark, respectively; after NO donor supplementation, the rate increased up to 131 ppm/h in light and 150 ppm/h in the dark (Figure 3d). These results suggest that the low N 2 O emissions observed in the G1 strain are due to a limitation in NO synthesis.

Nitrite Impacts CO 2 Emissions through a NR-Dependent Process in the nii1 Mutants
NO is a signal molecule that inhibits a wide variety of processes in Chlamydomonas, including photosynthesis [45] and mitochondrial respiration [46]. Thus, taking advantage of the CRDS analyzer's functionality to quantify CO 2 , we studied CO 2 evolution to understand how NO accumulation, and indirectly N 2 O emissions, might impact central metabolism in the nii1 mutants. Under mixotrophic conditions, CO 2 emissions are mainly a result of the flux balance between CO 2 fixation (photosynthesis and Calvin-Benson-Bassham cycle) and CO 2 release by the Tricarboxylic Acid Cycle (TCA) that is fed with acetate as an exogenous carbon source, although CO 2 emissions can also be impacted by other processes such as carbon mobilization from storage compounds (i.e., starch and lipids) and, to a lesser extent, photorespiration [47][48][49] Therefore, we assayed how the different nii1 mutants were affected in CO 2 evolution.
Total CO 2 accumulation in the headspace of the cultures was monitored after 24 h of induction in the presence of 10 mM NO 2 − in light and dark conditions. In the dark, when cells cannot fix carbon, CO 2 emissions were higher in G1 (9024 ppm) than in the M4 and M3 strains (3658 ppm and 2852 ppm) (Figure 4a and Supplementary Table S1). The same experiment, carried out under illumination, showed the opposite effect: lower CO 2 emission in the G1 mutant (1153 ppm) than in the M4 and M3 strains (5188 ppm and 6151 ppm). Similar results were obtained for M3arc and M3 strains in the dark (M3arc accumulated more CO 2 , 9169 ppm, than M3, 2852 ppm) but not in the light, where they showed almost identical CO 2 accumulation (Figure 4a). We suggest that this different phenotype in the light might be a consequence of the slightly higher NO levels observed in M3arc compared to G1 (Figure 3c), as CO 2 emission patterns in light and darkness seem to be affected by NO. To test this hypothesis, the G1 cultures were supplied with a NO donor in dark and light conditions after 20 h induction in 10 mM NO 2 − . The NO addition led to a three-fold increase in the CO 2 emission rate in the light but not in the dark, where only a slight reduction was observed (Figure 4b). To further confirm whether NO reduces CO 2 emission in the dark, the M3 strain was treated with NO donor in N-free medium in the dark, and after a short incubation time (75 min) (Supplementary Figure S1). Before NO donor addition, the CO 2 emission rate was 242 ppm/h, but after NO donor addition, the CO 2 emission rate decreased to 88 ppm/h. Accordingly, the N 2 O emission rate increased from 0 to 8 ppm/h (Supplementary Figure S1).  Figure S2). In N-free medium, the atmospheric CO 2 was consumed, and almost no emission was detected after 24 h. However, CO 2 was released in the presence of NO 2 − in a concentration-dependent manner (4718 ppm and 6152 ppm in 0.1 mM and 10 mM NO 2 − , respectively). These data highlight the regulatory role of NO 2 − -derived NO in CO 2 emission levels (see Discussion Section).
ppm and 6152 ppm in 0.1 mM and 10 mM NO2 − , respectively). These data highli regulatory role of NO2 − -derived NO in CO2 emission levels (see Discussion Section

N2O and CO2 Emissions in the NO3 − /NO2 − Assimilation Wild Type Strain 6145c and nit1nit2 Mutant CMJ030
To better understand how the NO3 − /NO2 − assimilation pathway impacts N CO2 emissions, we studied the accumulation of these gases in sealed cultures of strain (6145c) and CMJ030, a mutant that cannot assimilate NO3 − and exhibits a growth on NO2 − . By genetic crossing, we demonstrated that CMJ030 is a nit1nit2 (see Supplementary Figure S3) that lacks NR activity and also NIT2, which is transcriptional factor involved in the regulation of the NO3 − /NO2 − assimilation p [36,40].
Both 6145c and CMJ030 strains accumulated much less N2O than the M3 a mutants; N2O emission reached 18 ppm in 6145c and 4 ppm in CMJ030 after 24 h light (Figure 5a,b). After normalization using chlorophyll concentration (as 614 tures double their chlorophyll content after 24 h in NO2 − ), N2O production in 614 two-fold higher than in CMJ030 (Supplementary Table S1). In the dark (where no was observed), normalized emission increased ~five-fold (Supplementary Table S1) ing characteristic kinetics with two phases of production separated by another p which N2O was not accumulated (Figure 5a,b). The lower N2O emissions observe nit1nit2 mutant further support that the NO3 − /NO2 − assimilation pathway impac synthesis in Chlamydomonas.
The low N2O production detected in these strains seems to point out that NO highly accumulated. Consequently, both strains exhibited high CO2 emissions dark and low CO2 levels in light (Figure 5c,d, and Supplementary Table S1), sug To better understand how the NO 3 − /NO 2 − assimilation pathway impacts N 2 O and CO 2 emissions, we studied the accumulation of these gases in sealed cultures of the WT strain (6145c) and CMJ030, a mutant that cannot assimilate NO 3 − and exhibits a limited growth on NO 2 − . By genetic crossing, we demonstrated that CMJ030 is a nit1nit2 mutant (see Supplementary Figure S3) that lacks NR activity and also NIT2, which is the key transcriptional factor involved in the regulation of the NO 3 − /NO 2 − assimilation pathway [36,40].
Both 6145c and CMJ030 strains accumulated much less N 2 O than the M3 and M4 mutants; N 2 O emission reached 18 ppm in 6145c and 4 ppm in CMJ030 after 24 h in the light (Figure 5a,b). After normalization using chlorophyll concentration (as 6145c cultures double their chlorophyll content after 24 h in NO 2 − ), N 2 O production in 6145c was twofold higher than in CMJ030 (Supplementary Table S1). In the dark (where no growth was observed), normalized emission increased~five-fold (Supplementary Table S1), showing characteristic kinetics with two phases of production separated by another phase in which N 2 O was not accumulated (Figure 5a,b). The lower N 2 O emissions observed in the nit1nit2 mutant further support that the NO 3 − /NO 2 − assimilation pathway impacts N 2 O synthesis in Chlamydomonas.

Discussion
Plants and algae can produce the potent greenhouse gas N2O, which can be emitted at significant amounts into the atmosphere as a result of high inputs of NO3 − /NO2 − [16,17,22]. Despite its potentially high environmental and ecological impact, the molecular mechanisms involved in N2O production by photosynthetic organisms remain largely unknown, and genetic evidence supporting N2O emissions has been only recently described in the model organism Chlamydomonas reinhardtii [26,27]. Recent works have documented the existence of two NO reductases, FLVs and CYP55, able to produce N2O when the alga is supplied with NO. Most of these experiments were performed in a Chlamydomonas nit1nit2 genetic background and demonstrated that N2O production mostly relies on FLVs in the light and on CYP55 in the dark. Another approach by Plouviez and collaborators studied the N2O production from NO2 − by Chlamydomonas strains with different genetic backgrounds for NO3 − assimilation. Their results showed that N2O production by the WT strain, able to assimilate NO3 − , occurs from NO2 − and mainly in the dark linked to CYP55. This result was supported later by Burlacot and collaborators showing that NO uptake and N2O production in the dark were much higher when WT cells were grown with NO3 − as the sole nitrogen source and reflecting the regulation of CYP55 by NO3 − metabolism. Different processes have been proposed to synthesize NO from NO2 − , the intermediary step in N2O production [17,35,44]. Plouviez et al., 2017 [26] suggest two phases in the Chlamydomonas N2O emissions by WT in the dark, an early one involving NR (3.5 h) and a late phase involving the mitochondrial COX (24 h). Here, we present and discuss new data on the NO2 − -to-N2O denitrification process in Chlamydomonas nii1 mutants and how CO2 emissions are affected in these strains.
When NO3 − /NO2 − assimilation is interrupted because of the absence of NiR activity, two main conclusions are considered: (1) NR and ARC (NOFNiR) have a vast impact on N2O emissions, and (2) this NR-dependent N2O emission is significantly higher (4.5fold) in the light than in the dark, a result in accord with Plouviez et al., 2017. Our results highlight that the NO synthesized by the cytosolic NR/ARC complex can diffuse to other organelles such as mitochondria and chloroplast, and this NO seems to regulate The low N 2 O production detected in these strains seems to point out that NO is not highly accumulated. Consequently, both strains exhibited high CO 2 emissions in the dark and low CO 2 levels in light (Figure 5c,d, and Supplementary Table S1), suggesting that 10 mM NO 2 − is not enough to alter CO 2 evolution under our experimental conditions.

Discussion
Plants and algae can produce the potent greenhouse gas N 2 O, which can be emitted at significant amounts into the atmosphere as a result of high inputs of NO 3 − /NO 2 − [16,17,22]. Despite its potentially high environmental and ecological impact, the molecular mechanisms involved in N 2 O production by photosynthetic organisms remain largely unknown, and genetic evidence supporting N 2 O emissions has been only recently described in the model organism Chlamydomonas reinhardtii [26,27]. Recent works have documented the existence of two NO reductases, FLVs and CYP55, able to produce N 2 O when the alga is supplied with NO. Most of these experiments were performed in a Chlamydomonas nit1nit2 genetic background and demonstrated that N 2 O production mostly relies on FLVs in the light and on CYP55 in the dark. Another approach by Plouviez and collaborators studied the N 2 O production from NO 2 − by Chlamydomonas strains with different genetic backgrounds for NO 3 − assimilation. Their results showed that N 2 O production by the WT strain, able to assimilate NO 3 − , occurs from NO 2 − and mainly in the dark linked to CYP55. This result was supported later by Burlacot and collaborators showing that NO uptake and N 2 O production in the dark were much higher when WT cells were grown with NO 3 − as the sole nitrogen source and reflecting the regulation of CYP55 by NO 3 − metabolism. Different processes have been proposed to synthesize NO from NO 2 − , the intermediary step in N 2 O production [17,35,44]. Plouviez et al., 2017 [26] suggest two phases in the Chlamydomonas N 2 O emissions by WT in the dark, an early one involving NR (3.5 h) and a late phase involving the mitochondrial COX (24 h). Here, we present and discuss new data on the NO 2 − -to-N 2 O denitrification process in Chlamydomonas nii1 mutants and how CO 2 emissions are affected in these strains.

When NO 3
− /NO 2 − assimilation is interrupted because of the absence of NiR activity, two main conclusions are considered: (1) NR and ARC (NOFNiR) have a vast impact on N 2 O emissions, and (2) this NR-dependent N 2 O emission is significantly higher (4.5-fold) in the light than in the dark, a result in accord with Plouviez et al., 2017. Our results highlight that the NO synthesized by the cytosolic NR/ARC complex can diffuse to other organelles such as mitochondria and chloroplast, and this NO seems to regulate processes involved in CO 2 emissions (later discussed). Despite the importance of NR as the main NO source in the nii1 mutants, the remaining NO and N 2 O levels observed in G1 cultures point out alternative NO synthesis pathways such as that involving COX, as previously reported [26].
When NO 3 − /NO 2 − assimilation is totally functional, N 2 O emissions are tremendously diminished. This result reveals that N 2 O emissions in Chlamydomonas seem to be mainly restricted to conditions in which NO 3 − /NO 2 − cannot be properly assimilated and used for growth. This might support why the WT strain emits more N 2 O in the dark, as cells need to acclimate to this condition and NO 3 − /NO 2 − assimilation is less efficient. According to this, we could expect high N 2 O emission in NO 3 − /NO 2 − -rich environments depleted of other nutrients. Therefore, growth limitation in the presence of high NO 3 − /NO 2 − concentrations should lead to high N 2 O synthesis rates. Finally, the two phases of N 2 O emission observed in dark-incubated WT cells could be attributed to NO generated by NR (first phase) and mitochondrial COX (second phase), as previously reported [17].
When NO 3 − /NO 2 − assimilation is impaired (nit1nit2 mutant), N 2 O emissions are lower than in the WT. In this genetic background, neither NR nor the regulatory NIT2 proteins are functional, and NO 2 − assimilation is slow, allowing a limited growth in this N source [50]. This residual NO 2 − assimilation is enough to avoid NO 2 − dissimilation to N 2 O. In addition, NIT2 also controls other steps in NO 3 − assimilation, including NO 3 − /NO 2 − transporters [36,40] and NO metabolism-related proteins such as AOX1 [51], THB1 and THB2 [38,39], and probably CYP55, which increases in response to NO 3 − [29]. Moreover, a putative NO 3 − -dependent regulation of the N 2 O production, mediated by NIT2, is also supported by the significant increase in the N 2 O emission rate observed in M3 cells incubated in NO 3 − compared to those incubated in NO 2 − (Figure 2c). CO 2 emissions are closely related to NO 2 − -dependent N 2 O emissions. Our results show a relationship between N 2 O and CO 2 emissions that will require further investigation to understand the metabolic adaptations of Chlamydomonas to heterotrophic and mixotrophic conditions in the presence of NO 3 − or NO 2 − . In both conditions, acetate is the main carbon source, but it is essential only in the dark to feed the TCA cycle and provide energy to the cells, releasing CO 2 [47,52,53].
This study shows that low N 2 O emissions correlate with high CO 2 release in the dark and vice versa; high N 2 O emissions correlate with less CO 2 release. The link between N 2 O and CO 2 emissions appears to be the NO signal molecule, produced mainly by the NR/ARC complex in the nii1 mutants. NO could inhibit acetate metabolism and CO 2 release, also supported by the slight inhibition of the CO 2 emission rate by NO donor.
In the light, we found the opposite correlation: low N 2 O accumulation, due to low NO synthesis, leads to a reduced CO 2 emission and vice versa. Under illumination, NO inhibits photosynthesis [45], reducing CO 2 fixation. In fact, NO supply increased by three-fold the CO 2 emissions in the light, suggesting that CO 2 fixation is very sensitive to NO. Thus, CO 2 fixation would be more active in those strains/conditions in which low NO is synthesized (low N 2 O emitted) and, therefore, lower CO 2 levels would be accumulated. The role of NO as a photosynthesis inhibitor has been described in plants and algae and has been considered a mechanism to avoid photo-damage in algae under nutrient deprivation [45,54,55]. Nitrogen- [56] or sulfur-starved [57] Chlamydomonas cells accumulate NO, which causes the degradation of the cytochrome b 6 f complex and Rubisco by the FtsH and Clp proteases. More recently, transcriptomic analyses reported the molecular mechanisms underlying the acclimation of Chlamydomonas after NO supply [45]. Among the regulated process, NO decreases photosynthesis, respiration, N availability, and induces NO scavenging (THB1, THB2, FLVB, and CYP55).
The contributions of plants and algae to the N 2 O atmospheric budget are not being considered by the expert panels, even when increasing reports support their participation in this process, and the high input of nitrogen fertilizers is the primary cause [8,17,22]. Our data shed light on the mechanisms involved in the N 2 O synthesis and highlight the nii1 mutants as good models to study the molecular bases of the N 2 O emission in photosynthetic organisms. Moreover, the NR role on N 2 O emission raises an important link between NO 3 − assimilation and dissimilation, making of this enzyme a good candidate for future studies in order to acquire a better understanding on those environmental conditions that promote NO 3 − dissimilation over assimilation.

Strains and Growth Conditions
The strains used in this study are listed in Supplementary  [58] (obtained from the Chlamydomonas Library Project (CLiP), https://www.chlamylibrary.org).
All the cell cultures were performed in TAP medium (Tris, Acetate, Phosphate) [59] in a chamber (AlgaeTron AG 230, Photon System Instruments, Drásov, Czech Republic) at 25 • C, with continuous agitation (120 rpm) and illumination (light intensity 130 µmol photons·s −1 ·m −2 ). When indicated, cell cultures were transferred to dark in the same chamber.
Cells were grown in TAP medium with NH 4 + as a nitrogen source (8 mM NH 4 Cl) (pre-cultures). At the exponential phase of the culture, cells were harvested by centrifugation (2 min at 3000× g), washed twice with nitrogen-free TAP and transferred to new media containing the indicated nitrogen sources. The initial chlorophyll concentration was adjusted to 9-10 µg mL −1 .
For unsealed flask, Erlenmeyer flasks covered with foil paper were used. The same flasks were hermetically sealed with screw caps (sealed flasks), and a syringe was used to collect samples from the culture.

Chlorophyll, NO 2 − , and Cell Counting Measurements
Samples of 1 mL were centrifuged at 15,000× g for 5 min, and the supernatant (cell-free medium) and the pellet were separately frozen at −20 • C. NO 2 − was quantified in the cell-free medium using the Griess reagents according to Snell and Snell (1949) [60]. For chlorophyll concentration, the pellet was resuspended in 1 mL ethanol and incubated for 3 min, at room temperature. Afterwards, the samples were centrifuged and the chlorophyll concentration in the supernatant was quantified as previously described [61]. Cell quantification of liquid cultures was determined using the Sysmex Microcellcounter F-500 cell counter.

NO Measurements
Cells cultures (25 mL) were induced in media with 10 mM NO 2 − during 24 h. Then, 2 µM of DAF-FM (4,5-Diaminofluorescein) was added and incubated for 1h. An amount of 200 µL of the culture was used for NO quantification in a fluorescence spectrophotometer (Varioskan Lux, Thermo scientific, Waltham, MA USA) using OptiPlate Black Opaque 96-well Microplate (PerkinElmer, Waltham, MA USA). The excitation and emission wavelengths for the NO indicator were 485 and 515 nm, respectively. Data are represented as arbitrary fluorescence units.

Determination of N 2 O and CO 2 Emissions
N 2 O and CO 2 were simultaneously quantified by using a Cavity Ring-Down Spectroscopy (CRDS) analyzer (PICARRO G2508). For this purpose, we used 1 L bottles (DURAN TM ) that were hermetically sealed with screw caps (GL 45 with 2 or 3 connectors) both from DWK Life Sciences (Mainz, Germany). The bottles were set with 250 mL liquid culture medium and 750 mL headspace (gas phase). The CRDS analyzer was connected to the bottle through a combined inlet and outlet Teflon tubes (2.5 m in length). The outlet tube extracted the sample from the headspace of the bottle (0.3 L/min), and the inlet tube returned the sample into the gas phase of the cultures, passing the air through a 0.22 µm PVDF filter (Dualex TM -Plus; Merck, Darmstadt, Germany) to avoid culture contamination.

Genetic Crosses
Genetic crosses were performed according to [62] and the random spore plating method. Then, 100 segregants were analyzed, and several of them were chosen for further experiments.