Next Article in Journal
Reorganization of Protein Tyrosine Nitration Pattern Indicates the Relative Tolerance of Brassica napus (L.) over Helianthus annuus (L.) to Combined Heavy Metal Treatment
Next Article in Special Issue
Corn Responsiveness to Azospirillum: Accessing the Effect of Root Exudates on the Bacterial Growth and Its Ability to Fix Nitrogen
Previous Article in Journal
Influence of Citrus Scion/Rootstock Genotypes on Arbuscular Mycorrhizal Community Composition under Controlled Environment Condition
Previous Article in Special Issue
Exogenous Carbon Compounds Modulate Tomato Root Development
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Plants
Volume 9
Issue 7
10.3390/plants9070903
plants-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Chlamydomonas reinhardtii, an Algal Model in the Nitrogen Cycle

by
Carmen M. Bellido-Pedraza
1,
Victoria Calatrava
1,†,
Emanuel Sanz-Luque
1,
Manuel Tejada-Jiménez
1,
Ángel Llamas
1,
Maxence Plouviez
2,
Benoit Guieysse
2,
Emilio Fernández
1,* and
Aurora Galván
1,*
1
Departamento de Bioquímica y Biología Molecular, Campus de Rabanales y Campus Internacional de Excelencia Agroalimentario (CeiA3), Edif. Severo Ochoa, Universidad de Córdoba, 14071 Córdoba, Spain
2
School of Food and Advanced Technology, Massey University, Private Bag, 11222 Palmerston North, New Zealand
*
Authors to whom correspondence should be addressed.
†
Current address: Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA.
Plants 2020, 9(7), 903; https://doi.org/10.3390/plants9070903
Submission received: 22 June 2020 / Revised: 12 July 2020 / Accepted: 13 July 2020 / Published: 16 July 2020
(This article belongs to the Special Issue Plant Nitrogen Assimilation and Metabolism)
Browse Figures
Review Reports Versions Notes

Abstract

:
Nitrogen (N) is an essential constituent of all living organisms and the main limiting macronutrient. Even when dinitrogen gas is the most abundant form of N, it can only be used by fixing bacteria but is inaccessible to most organisms, algae among them. Algae preferentially use ammonium (NH4+) and nitrate (NO3−) for growth, and the reactions for their conversion into amino acids (N assimilation) constitute an important part of the nitrogen cycle by primary producers. Recently, it was claimed that algae are also involved in denitrification, because of the production of nitric oxide (NO), a signal molecule, which is also a substrate of NO reductases to produce nitrous oxide (N2O), a potent greenhouse gas. This review is focused on the microalga Chlamydomonas reinhardtii as an algal model and its participation in different reactions of the N cycle. Emphasis will be paid to new actors, such as putative genes involved in NO and N2O production and their occurrence in other algae genomes. Furthermore, algae/bacteria mutualism will be considered in terms of expanding the N cycle to ammonification and N fixation, which are based on the exchange of carbon and nitrogen between the two organisms.

Graphical Abstract

1. Introduction

Nitrogen (N) is an essential macronutrient that supports life in all living beings. This nutrient is an elemental constituent of biomolecules, such as nucleic acids, proteins, chlorophylls, cofactors, and signal molecules, among others.
In nature, living organisms can use different N reservoirs, and the biotic and abiotic conversions among different N forms constitute the biogeochemical N cycling. These N reservoirs and N flux in terrestrial and marine ecosystems have been recently reviewed [1]. Ammonium (NH4+) bound into rocks and sediments is the largest N reservoir (1.8 × 1010 Tg nitrogen). However, this NH4+ has a minimal impact on the annual N cycling because it is accessible only upon erosion [1]. The largest available N reservoir for organisms is dinitrogen gas (N2) with 3.9 × 109 Tg, which represents 79% of the atmospheric air. However, due to the reduced reactivity of the triple bonded N2, only a small group of organisms having the dinitrogenase enzyme complex can use it as N source [1,2,3]. The global and usable N-reservoirs are organic nitrogen (9 × 105 Tg), nitrate (NO3−) (6 × 105 Tg), and nitrous oxide (N2O) (2000 Tg) [1,2,3,4]. The ammonia (NH3) reservoir estimated in the marine ecosystem is 340–3600 Tg, but it is unknown in the terrestrial ecosystem [1]. Other global inorganic N sources such as nitrite and nitric oxide are considered minor reservoirs.
N compounds exhibit different oxidation states ranging from −3 (organic N and NH3/NH4+) to +5 (NO3−), with intermediate values in compounds such as hydrazine (N2H4) (−2), hydroxylamine (NH2OH) (−1), N2 (0), nitrous oxide (N2O) (+1), NO (+2), nitrite, NO2− (+3), and nitrogen dioxide (NO2) (+4). Bacteria can catalyze many redox reactions involving the above compounds, which can be used to provide nitrogen for growth and redox energy to fuel cells. Bacterial species have been classically classified according to the reactions that they mediate within the N cycle (nitrification, denitrification, fixation, ammonification, anammox, assimilatory, and dissimilatory processes). Notwithstanding, DNA genome sequencing has revealed the set of genes present in each microorganism and highlighted the enormous metabolic flexibility of some of the organisms studied, making it almost impossible to classify them using only the classical divisions [1,2]. Moreover, it is expected that further research will unravel new N-transforming reactions and enzymes that will emphasize the metabolic flexibility of many of these organisms [1].
Prokaryotic organisms exhibit an enormous versatility to metabolize most of the N compounds, and they are the main players for most of the transformations in the N cycle. In addition, enzymes required for N2 fixation, such as nitrogenase, are exclusively found in prokaryotes or bacteria involved in symbiosis/mutualism with other eukaryotic organisms. For instance, N2-fixing bacteria from the order Rhizobiales establish symbiotic interactions with legume plants, whose crops provide almost 20% of food protein in the world [5]. Other examples are found in marine ecosystems, where a symbiotic interaction between the haptophyte algae and the most widespread N2-fixing cyanobacterium ‘Candidatus Atelocyanobacterium thalassa’ (UCYN-A) has been described [6,7]. In the lab, the N2-fixing bacterium Azotobacter chroococcum provides N to the alga Chlamydomonas and promotes its growth, whereas Chlamydomonas fixes CO2 and provides organic carbon to the bacterium. This symbiotic interaction has reported benefits for the industry by increasing lipid production in the microalga [8].
Anthropomorphic activities have affected the natural equilibrium of the N cycle for more than a century. The industrial N2-fixation into ammonia (Haber-Bosch process) and the use of this ammonia as fertilizers has dramatically improved crop productivity but, at the same time, it has increased the reactive N load by more than 120%, which includes all biologically active, photochemically reactive, and radiatively active N compounds in the atmosphere and biosphere [1,2,9,10]. This increase in the N load comprises a higher global amount of reduced inorganic forms of N (NH3 and NH4+), oxidized inorganic forms (as NOx, HNO3, N2O, and NO3−), and organic compounds (such as urea, amines, and proteins), in detriment to the unreactive N2 gas. One of the main unwanted effects of this imbalance in the N cycle is the transfer of reactive N from terrestrial to coastal systems and ground waters, causing algal blooms and the eutrophication of these ecological niches [9,10]. In addition, it is estimated that 3–5% of the N used in fertilizers is converted into N2O, a greenhouse gas that is 310 times more potent than CO2 and considered the dominant ozone-depleting pollutant emitted in the 21st century [1,11,12,13]. The principal substrate for N2O synthesis is NO, which is mainly reduced by bacterial NO reductases but also can be reduced by some eukaryotic enzymes that carry out nitrate dissimilatory reactions.
Nitrogen is a macronutrient for plants, algae, and bacterial growth. Nitrate is the most oxidized form of nitrogen and highly abundant in aquatic habitats. The nitrate assimilation pathway (NO3− → NO2− → NH4+ → amino acids) is quantitatively the most important redox process, supporting almost 20% of marine algal growth [1,4]. Dissimilatory nitrate reduction pathways such as dissimilatory nitrate reduction to ammonium (DNRA), denitrification to N2O, and total denitrification to N2 are also processes mediated by eukaryotes in aquatic systems. DNRA is described in diatoms as an adaptation and survival process to darkness and anoxia, where the intracellular NO3− is used as an electron acceptor instead of O2 [14,15]. On the other hand, a complete denitrification process (NO3− → NO2− → NO → N2O → N2) has been described in some species belonging to the foraminifera group, which represent the only eukaryotic organisms able to perform complete denitrification to N2 [16,17,18]. Other foraminifer species perform only partial nitrate denitrification to N2O [17,18,19]. The genes involved in the foraminifer denitrification pathway have been recently identified [20]; however, the genes encoding the enzymes responsible for the first and last steps of this pathway, the dissimilatory nitrate reductases and nitrous oxide reductase, are missing in most of the foraminifer genomes but found in the associated bacteria. Partial nitrate denitrification to N2O has also been well documented in fungi [21,22,23,24,25] and recently described in the green microalga Chlamydomonas reinhardtii, where a NO reductase (CYP55) [26] and flavodiiron proteins (FLVs) [27] catalyze the reduction of NO to N2O in dark and light conditions, respectively. These two proteins constitute the molecular evidence that supports that microalgae contribute to N2O emissions, something known for decades [28]. In this review, we analyze the metabolic flexibility of microalgae to carry out different steps of the N cycle with special attention to the denitrification pathway.

2. The Role of Chlamydomonas reinhardtii in the N Cycle

Traditionally the genus Chlamydomonas corresponds to unicellular, biflagellate, green microalgae and encompasses more than 500 species. However, current phylogenetic studies have reevaluated the taxonomy of Chlamydomonas into three species: C. reinhardtii, C. incerta, and C. schloesseri [29,30]. Today, all the unequivocally identified C. reinhardtii cells have been isolated from soil habitats suggesting a preference for rich nutrient environments. Chlamydomonas spp., other than C. reinhardtii, have been isolated from very different environments such as acid mine drainage, temperate marine, Antarctic ice, and even air [29]. C. reinhardtii, named as Chlamydomonas throughout, has been developed as a powerful model organism useful to study essential biological processes but also biotechnological applications [31].
Nitrogen assimilation has been widely studied in Chlamydomonas and the pathways for the assimilation of different forms of nitrogen, inorganic (nitrate, nitrite, and ammonium) as well as organic (urea, amino acids, and purines), have been identified [32,33,34]. However, besides the classical N assimilation pathways that have been described in axenic Chlamydomonas cultures, recent advances on nitrogen-transforming reactions have revealed that this microalga can also carry out both an ammonification process by mutualistic interactions with bacteria and a denitrification process that releases N2O to the environment (Figure 1, Table 1).

2.1. Ammonification and Mutualism with Bacteria for N Scavenging from Amino Acids and Peptides

Chlamydomonas cells scavenge nitrogen from amino acids in three ways: transport, deamination/ammonification, and mutualistic interaction with bacteria [41]. Arginine is the only amino acid that is efficiently transported into the cell from the surrounding media and then used as carbon and nitrogen sources [42]. L-leucine and L-cysteine is also transported, but by passive diffusion [43]. However, most of the proteinogenic amino acids, including arginine, but also di- and tri-peptides, can be extracellularly deaminated, producing NH4+ and the corresponding keto acids. This ammonification process is mediated by the periplasmic L-amino acid oxidase (LAO1) (step 7, Figure 1, and Table 1), which is induced upon inorganic N deprivation [41,44,45,46,47]. Interestingly, only the NH4+ generated by the LAO1 enzyme is transported inside the cell by high-affinity ammonium transporters (AMT) and used by the alga, but the carbon skeletons remain in the medium [47]. It has been hypothesized that such carbon skeletons could be used for establishing mutualistic interactions with specific bacteria in the surrounding medium. These interactions depend on whether the released keto acid can meet the particular carbon requirements of a specific bacterium [41]. Besides ammonium and the keto acid, the ammonification process mediated by LAO1 also generates H2O2, which has bactericidal properties [48]. The eco-physiological role of H2O2, if any, is not clear but its production could repel other microorganisms to avoid competition for the organic N and, simultaneously, promote bacterial cell death making more N available. Alternatively, H2O2 synthesis could also serve as a mechanism to select specific H2O2-tolerant bacterial species in order to restrict the mutualistic interactions. The role of H2O2 in algal-bacterial mutualistic interaction has been previously reported [49].
Chlamydomonas LAO1 homologs have been found in other algal species, including Rhodophyta, Alveolata, Heterokonta, Haptophyta, and Dinophyta species, but not in other chlorophytes [41].
Despite the promiscuous LAO1 activity to deaminate amino acids and oligopeptides, Calatrava and collaborators [50] have reported that this enzyme cannot use proline and some oligopeptides as substrate. Interestingly, when proline is the only N source in the media, Chlamydomonas can establish a strict and specific mutualism with Methylobacterium species [50]. Under this situation, the bacteria mineralize the amino acid (or peptide) releasing ammonium that can be incorporated by the alga. In return, Chlamydomonas, which fixes CO2 to satisfy its carbon demand, produces C-compounds as glycerol that can be secreted and used as a C source for the bacterium.
Another step in which Chlamydomonas could participate in the N cycle is the use of N2, as described from co-cultures with the nitrogen-fixing anaerobic bacteria Azotobacter chroococcum. In this situation, the bacteria supply the nitrogen source and CO2 to the alga by nitrogen fixation, meanwhile, the alga supplies carbohydrates and O2 via photosynthesis to the bacteria [8].

2.2. Nitrate Denitrification

The nitrate flux toward its assimilation is the primary nitrogen pathway that occurs in Chlamydomonas as well as in eukaryotic organisms containing an assimilatory nitrate reductase (NR). The nitrate assimilation appears to occur via a relatively simple linear pathway involving (1) nitrate uptake by specific nitrate transporters (i.e., NRT2.1/NAR2), (2) cytosolic nitrate reduction by a typical eukaryotic nitrate reductase (NR), (3) nitrite transport into the chloroplast by specific transporters (i.e., NAR1.1), (4) nitrite reduction to ammonium by a typical plant-like ferredoxin-dependent nitrite reductase (NIR), and finally (5) the ammonium incorporation into carbon skeletons by the GS/GOGAT cycle that synthesizes glutamate [33,34].
Besides its assimilatory role, the NR is a moonlighting protein that interacts with different partners to carry out alternative reactions. In photosynthetic organisms, NR is considered one of the principal nitric oxide (NO) sources [35] and also participates, as shown in Chlamydomonas, in the NO oxygenation to convert this signaling molecule back to nitrate. These reactions, in which NR is central and acts with different partners, can be called the NO3-/NO3- cycle (steps 1, 4, and 5 in Figure 1). Moreover, the intracellular NO can also be the substrate for NO reductases producing N2O in mitochondria and chloroplasts [1]. Thus, NR seems to be a cross-point among three different pathways: nitrate assimilation, a partial nitrate-denitrification to NO and the contaminant gas N2O, and the NO oxygenation to nitrate.

2.2.1. The NO3−/NO3− Cycle

The eukaryotic NR is a homo-dimeric protein that contains three prosthetic groups in each subunit (FAD, Heme b557, and molybdenum cofactor –Moco-). Nitrate reduction to nitrite is a redox reaction that occurs across a mini electron transport chain (mini-ETC) from NAD(P)H to nitrate through three prosthetic groups (NAD(P)H → FAD → heme → Moco → NO3−). However, as mentioned above, NR can bind to other protein partners and detour electrons at different levels of this mini-ETC. When the partner protein is the molybdoprotein NOFNIR (NO forming nitrite reductase), electrons flow until the heme cofactor, where they reduce the NOFNIR that converts nitrite to NO [35,51]. When the partner is the truncated hemoglobin 1 (THB1), electrons go out of the mini-ETC at the FAD cofactor level and reduce the THB1 protein, which transforms NO into nitrate [33]. These three reactions constitute the named NO3−/NO3− cycle.
Nitric oxide is a signal molecule that regulates a high number of physiological processes in mammals, plants, and algae [52]. In Chlamydomonas, NO regulates N assimilation at the transcriptional and posttranslational level. NO is a negative regulator for the expression and activities of the high-affinity nitrate/nitrite transporters, the NR, and the high-affinity ammonium transporters [33,53]. As a signal molecule, NO is expected to have a temporal effect, and therefore, its synthesis and degradation have to be tightly controlled.
This NO3−/NO3− cycle could work for regulating nitrate assimilation without N loss. When the conditions are optimal, nitrate goes toward its assimilation up to glutamate. If the conditions are not optimal (i.e., α-ketoglutarate is deficient) and nitrite accumulates, NO is synthesized and stops the nitrate uptake and reduction. However, although NO is a gas and can diffuse out of the cells, the NR-THB complex could convert the NO into nitrate, avoiding the N loss.

2.2.2. NO Synthesis and Reduction: A Prokaryote Pathway

In plants and plant-like organisms, NO is produced by different mechanisms that use different substrates (i.e., NO2− or arginine) in different intracellular compartments such as cytosol [35,54,55], mitochondria [26,56], chloroplast [57,58], and peroxisomes [59]. The oxidation of arginine to NO and citrulline by the NO synthase (NOS) is a well-known reaction in mammals, bacteria, and some algae [60,61]. NOS-like reactions have been described in land plants (peroxisomes) [62] and Chlamydomonas [63], but extensive genome analyses have ruled out the existence of NOS in photosynthetic organisms except for some microalgae [60,61]. However, NO production from NO2− is well documented to occur in the cytosol, by NR [54,55] or NR/NOFNIR [35], and also in mitochondria [26,56] and chloroplast by the electron transport chains [57].
Besides NO conversion into nitrate, NO can also be reduced by NO reductases to N2O. This denitrification process implies the loss of nitrogen to the atmosphere as a potent greenhouse gas. This pathway (NO2− → NO → N2O) has been well defined in both prokaryotes [1] and eukaryotes (fungi). In fungi, denitrification involves NIRK (copper-containing nitrite reductase) (NO2− → NO) and the cytochrome P450, CYP55 (NO → N2O), both acquired from bacteria [22]. For fungal CYP55, a horizontal gene transfer from bacteria is suggested, but for NIRK, a proto-mitochondrial origin has been proposed.
The Chlamydomonas genome contains two genes encoding for both proteins, NIRK and CYP55 (Table 1). CYP55 appears to be responsible for N2O production in the dark [26,27]. Accordingly, CYP55 is located in mitochondria [38] and its transcript is accumulated in the dark [64] and in the presence of nitrate [38]. The dark NO reduction attributed to CYP55 activity has also been shown to be stronger when cells are grown in the presence of nitrate [27]. Although found in the mitochondrial proteome, CYP55 is also predicted to be targeted to the chloroplast [27]. Thus, it is possible that the protein is targeted to both organelles. In this case, CYP55 could be able to use electron donors from photosynthetic origin. Like CYP55, NIRK is also up-regulated in the dark [64] and in nitrate-containing media (Calatrava, unpublished data). In silico analyses strongly suggest that NIRK could also be localized in the mitochondria (Table 1), pointing out that Chlamydomonas seems to have a complete and operative NO2− → NO → N2O pathway in the mitochondria.
The second type of protein shown to reduce NO to N2O, in Chlamydomonas, corresponds to FLVs (flavodiiron proteins) [27]. Flavodiiron proteins belong to a singular family of oxygen and/or nitric oxide reductases widespread in prokaryote and eukaryote organisms. This protein family has been classified into eight classes (A to H) according to the presence of modular domains [65,66]. Class A FDPs are the most dominant class in the family and only contain the two core domains (Fe-Fe and FMN) [65,66]. Some examples for this class A show a preference for O2 as revealed for the protozoal and archaeal enzyme [67,68,69,70], or they may have a dual activity toward NO and O2 as in M. thermoacetica or D. gigas [71,72]. Class B FDPs contain an additional C-terminal rubredoxin domain and, because of that, were named flavorubredoxins (FIRD) or NORV (from the encoding gene in E. coli) [65]. In E. coli, FLVB/FIRD/NORV was shown to have significant preference for NO over O2 and confers resistance to NO under anaerobic conditions [73,74]. Class C FDPs have a C-terminal flavin-binding domain, predicted to have a flavin reductases activity accepting electrons from NAD(P)H [65,66]. This class C is present in cyanobacteria, the protozoan Paulinella chromatophora, green algae, mosses, lycophytes, and gymnosperms, but absent in angiosperms [65]. Chlamydomonas FDPs correspond to class C (also called Flv), which forms a distinct cluster that falls into two groups. One of these groups (FlvA) has the ligand binding diiron site strictly conserved, while in the other (FlvB) the ligand binding sites are not conserved. The Chlamydomonas genome contains two genes, FLVA and FLVB, encoding both groups of flavodiiron proteins. Moreover, the Chlamydomonas FLVs could work as a heterodimer as shown by different insertional mutants of the FLVB gene that result in the loss of expression of both FLVB and FLVA proteins [71]. The study of these mutants revealed that FLVs participate in a Mehler-like reduction of O2, providing an alternative valve for electrons during the dark to light transition in the chloroplast. Thus, FLVs are critical for the algal growth under fluctuating light regimes but not in continuous light [75]. These mutants were also useful to prove the FLVs role as a NO reductase, being the major enzyme responsible for NO reduction to N2O under conditions of light in the presence of O2 in Chlamydomonas. In addition, in vivo experiments suggest that Chlamydomonas FLVs shows a higher affinity for NO than for O2 [27]. Therefore, Chlamydomonas has inherited two types of N2O-producing proteins, FLVs in the chloroplast and CYP55 in the mitochondria.
Besides CYP55 and FLVs, the Chlamydomonas genome also contains four genes encoding for hybrid cluster proteins (HCPs) [39], which are metalloproteins characterized by the presence of an iron-sulfur-oxygen cluster considered to be unique in biology [25]. The analysis of these protein sequences and their phylogenetic analyses support that the HCP family has emerged from a single gene of an alpha-proteobacterium with subsequent duplications in Chlamydomonas [39]. Like for CYP55 and NIRK, both nitrate and darkness contribute to the strong up-regulation of the HCPs in Chlamydomonas growing under oxic conditions [39]. In dark anoxia, the HCP4 transcript shows the stronger induction [76], and the studies performed in mutants obtained by gene silencing suggest a role in regulating fermentative and nitrogen metabolism pathways [77]. Concerning their cellular localization, HCP1, 3, and 4 are predicted to be in the chloroplast [39], but HCP2 is located in the mitochondria, as confirmed in mito-proteome analysis [38]. In bacteria, the function(s) of HCP is debated. Initially, they were considered hydroxylamine reductases, as they can use this substrate to produce ammonium in vitro, but the low affinity for hydroxylamine has questioned this physiological role [78]. Later, Wang and collaborators demonstrated that E. Coli HCP is a high-affinity nitric oxide (NO) reductase and proposed it as the major enzyme for reducing NO to N2O under physiologically relevant conditions [25]. The in vivo high affinity NOR activity is also demonstrated in Methylobacter tundripaludum, where NO mediates a response to hypoxia, quorum sensing, the secondary metabolite tundrenone, and methanol dehydrogenase [79]. What makes the history of HCP attractive is that it can function as a protein S-nitrosylase with a role in the NO sensing pathway. Thus, HCP is auto-S-nitrosylated and afterward results in the assembly of an HCP interactome, including enzymes that generate NO, synthesize SNO-proteins, and propagate SNO-based signaling mediating the NO-mediated stress responses [80]. The function(s) of HCP in Chlamydomonas as NOR and or protein-S-nitrosylase has to be solved as well as its relationship with nitrate metabolism. Seth and collaborators propose that HCPs oxidize NO to NO+, which is the form required for S-nitrosylation [80]. However, the redox potentials of both HCP and HCR (HCP reductase, also required for HCP function) do not fit with an oxidase activity and Lis et al. have proposed that HCPs might exhibit reductase activity and perform the following two-electron reduction reaction, NO3− + 4H+ + 2e− → NO+ + 2H2O [39].

3. N2O Production and Enzymes in Other Microalgae

For decades, N2O emission has been detected from lakes, coasts, and oceans, ecosystems where microalgae are mainly present [81], although the contribution of microalgae to these N2O emissions has been mostly neglected and attributed to the prokaryotic organisms present in these environments [28,82,83,84]. However, the correlation between algal bloom events, triggered by the excess of N from anthropogenic origins, and N2O emissions suggests that these microorganisms may be partly responsible for this production [1,28,83]. Importantly, many microalgae are currently cultivated at a large scale for commercial and biotechnological purposes. Therefore, the potential contribution to N2O emissions of these microalgae should be considered [26,28]. Microalgae cultures, supplied with NO3− as the sole N source, have been estimated to emit N2O in amounts 2.5 to 14 times more than from natural vegetation [85,86]. Although the emission of N2O by microalgae depends on many environmental factors, the major ones seem to be the species of microalga and the N source [26,27,87]. While N2O is produced on nitrate or nitrite, its production is prevented on ammonium [28].
The existence of a full NO2− → N2O dissimilation pathway in Chlamydomonas made interesting the analysis of other microalgae genomes. Public algal databases were searched for homologous proteins to the Chlamydomonas NIRK, possibly involved in NO synthesis and CYP55, FLVs, HCP as potential NO reductases. Those microalgae containing at least one of these proteins were selected. A total of 33 microalgae were considered, 23 Chlorophyta and 10 non-Chlorophyta species. Considering there are around one hundred microalgae genomes available, it can be assumed that almost 33% of microalgae contain at least one of these proteins. Phylogenetic trees of these proteins were constructed to provide a broader picture of the distribution of these proteins in the tree of life consistent with previous reports for FLV [65] (Figure S1), HCP [39] (Figure S2), CYP55 [88] (Figure S3), and NIRK [21] (Figure S4). These data were correlated to the production of N2O by the microalgae (Table 2) and discussed.
From the 33 selected microalgae, only three Chlorophyta species (Chlamydomonas reinhardii, two subspecies of Scenedesmus obliquus, and Chlorella sorokiniana) contained a full set NIRK, CYP55, FLVA, FLVB, and HCP. In other words, they contain a prokaryote NO synthetase and redundant potential NO reductases. This represents almost 9% of the microalgae in Table 2. In addition, these three microalgae have been reported to produce N2O [26,27,89].
For the rest of microalgae in Table 2, an incomplete NO2− → N2O pathway or the potential to produce NO (NIRK) or reduce NO to N2O through FLVs, CYP55, or HCP reductases was observed. The molecular evidence for these NO reductases is restricted to a few microalgae and comes from a recent study by Burlacot and collaborators [27]. Both CYP55 and FLVs were responsible for producing N2O under two different conditions, CYP55 in the dark and FLVs in the light. Therefore, microalga strains lacking CYP55 and/or FLV also are unable to produce N2O under its particular condition. However, whether HCP is or is not a NO reductase is unsolved in microalgae.
As previously reported, FLVs are present only in Chlorophyta and in approximately 74% of the selected chlorophytes in Table 2. These species contain both FLVA and FLVB, suggesting that these proteins may function as a heterodimer [75]. The phylogenetic tree suggests that FLVs from chlorophytes and early-diverged land plants are related to cyanobacteria (Figure S1), most likely derived from the endosymbiotic gene-transfer (EGT) of both genes from the chloroplast to the nuclear genome in the green lineage, after the divergence of the red algae and glaucophytes. Otherwise, both genes may have been transferred to the nucleus of the ancestral archaeplastidan and subsequently lost in red algae and glaucophytes.
As previously reported, HCP genes, although not present land plants, are highly represented in Chlorophyta; most of the chlorophytes harbor more than one copy of this gene [39]. In Chlamydomonas reinhardtii, the four HCP gene copies are phylogenetically related and located in the same chromosome, which suggests that these genes were recently duplicated [39]. In Table 2 it is shown that 69% of the chlorophytes harbor HCP. Moreover, putative HCP orthologs are also found in nonchlorophytes, including three Nannochloropsis spp. (Eustigmatales), the Guillardia thetha and the Stramenopiles Phaeodactylum tricornutum and Thalassiosira pseudonana (Table 2, Figure S2). Supported by further phylogenetic analyses, an endosymbiotic origin of the microalgal HCP genes from mitochondria has been proposed [39].
Although less abundant, CYP55 is in almost 39% of the Chlorophyta in Table 2 and Figure S3. It was also found only in one of other non-chlorophytes (Thalassiosira pseudonana), but it corresponds to a truncated sequence and it was not included in the phylogenetic tree. The most closely related to microalgal CYP55 are fungal members, which are hypothesized to have been acquired from Actinobacteria [22]. In fungi, CYP55, known as p450nor (cytochrome p450 NO reductase), is considered as a biomarker for N2O production, since it is detected in all fungal isolates producing N2O from NO2− [24]. Because Actinobacterial are not considered canonical denitrifying bacteria, the prevalent hypothesis for the origin and evolution of CYP55/p450nor is its acquisition from Actinobacteria and next evolution to a new role in denitrification [22]. Recently, a new hypothesis suggests that fungal p450nor has a role in secondary metabolism rather than denitrification and that N2O is a minor bioproduct of the secondary metabolism [88].
The NO-forming NIRK, which shares a common bacterial origin in both fungi and microalgae [21], is poorly represented in chlorophytes (3 out of 33) (Table 2, Figure S4), but it is present in the genome of the red alga Galdieria sulpharia, the haptophyte Emiliana huxleyi, and in Excavata. However, the NO reductase activity by this protein in these species is still unknown.

4. Concluding Remarks and Future Perspectives

N homeostasis in ecosystems relies on the balance between N fixation and N release to the atmosphere in gaseous forms (mainly N2 and, to a lower extent, N2O and NO). In-depth studies of the reactions mediated by isolated organisms, as well as those carried out by consortiums, will help us to understand how organisms keep the equilibrium of the biogeochemical N cycle and will allow us to forecast putative imbalances.
Chlamydomonas reinhardtii performs a high number of N-transforming reactions. Quantitatively and preferentially are those devoted to inorganic nitrogen (NH4+, NO3−, NO3−) assimilation for biomass production. The ammonification/deamination of amino acids and peptides constitutes a poor N supply but leads to bacterial mutualistic interactions for C/N exchange. NO3− offers additional N-transforming reactions, such as the NO3−/NO3− cycle, and dissimilation through a typical prokaryotic pathway NO2− → NO → N2O by means of apparently redundant NO reductases.
Algae, playing essential roles in aquatic ecosystems in carbon an N cycles, were claimed for a long time to produce the potent greenhouse gas N2O. This point was recently further confirmed by molecular evidence with CYP55 and FLVs in Chlamydomonas reinhardtii. This review opens new perspectives for future investigations of the full or partial dissimilation pathway (NO2−→ NO → N2O) by algae, where NIRK, CYP55, FLVs, and HCPs may be involved. In fact, some green microalgae seem to share the dissimilating full pathway NO2− → NO → N2O with Chlamydomonas. However, partial pathways seem to be prevalent in microalgae.
Understanding this NO2− → NO → N2O pathway operating in microalgae as complete or partial routes will be important to understand the production and regulation of either a signal molecule as is NO or the loss of nitrogen as the contaminant gas N2O.

Supplementary Materials

The following are available online at https://www.mdpi.com/2223-7747/9/7/903/s1, Figure S1. Evolutionary relationships of putative FLV proteins, Figure S2. Evolutionary relationships of putative HCP proteins, Figure S3. Evolutionary relationships of putative CYP55 proteins, Figure S4. Evolutionary relationships of putative NIRK proteins.

Author Contributions

Conceptualization, A.G., E.F. and C.M.B.-P.; writing—Original draft preparation, A.G.; writing—Review and editing C.M.B.-P., V.C., E.S.-L., M.T.-J., Á.L., M.P., B.G., A.G. and E.F.; funding acquisition, A.G. and E.F. All authors have read and agreed to the published version of the manuscript.

Funding

This work was funded by MINECO (Grant BFU2015-70649-P), the European FEDER program, the Plan Propio de la Universidad de Córdoba, and the U.E.INTERREG POCTEP-055_ALGARED_PLUS5_E.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Kuypers, M.M.M.; Marchant, H.K.; Kartal, B. The microbial nitrogen-cycling network. Nat. Rev. Microbiol. 2018, 16, 263–276. [Google Scholar] [CrossRef] [PubMed]
  2. Stein, L.Y.; Klotz, M.G. The nitrogen cycle. Curr. Biol. 2016, 26, R94–R98. [Google Scholar] [CrossRef] [Green Version]
  3. Zerkle, A.L.; Mikhail, S. The geobiological nitrogen cycle: From microbes to the mantle. Geobiology 2017, 15, 343–352. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  4. Gruber, N.; Galloway, J.N. An Earth-system perspective of the global nitrogen cycle. Nature 2008, 451, 293–296. [Google Scholar] [CrossRef] [PubMed]
  5. Burris, R.H.; Roberts, G.P. Biological nitrogen fixation. Annu. Rev. Nutr. 1993, 13, 317–335. [Google Scholar] [CrossRef] [PubMed]
  6. Harding, K.; Turk-Kubo, K.A.; Sipler, R.E.; Mills, M.M.; Bronk, D.A.; Zehr, J.P. Symbiotic unicellular cyanobacteria fix nitrogen in the Arctic Ocean. Proc. Natl. Acad. Sci. USA 2018, 115, 13371–13375. [Google Scholar] [CrossRef] [Green Version]
  7. Muñoz-Marin, M.D.C.; Shilova, I.N.; Shi, T.; Farnelid, H.; Cabello, A.M.; Zehr, J.P. The transcriptional cycle is suited to daytime N2 fixation in the unicellular cyanobacterium Candidatus Atelocyanobacterium thalassa (UCYN-A). mBio 2019, 10. [Google Scholar] [CrossRef] [Green Version]
  8. Xu, L.; Cheng, X.; Wang, Q. Enhanced lipid production in Chlamydomonas reinhardtii by co-culturing with Azotobacter chroococcum. Front. Plant Sci. 2018, 9, 741. [Google Scholar] [CrossRef]
  9. Erisman, J.Z.; Sutton, M.A.; Galloway, J.; Klimont, Z.; Winiwarter, W. How a century of ammonia synthesis changed the world. Nat. Geosci. 2008, 1, 636–639. [Google Scholar] [CrossRef]
  10. Galloway, J.N.; Townsend, A.R.; Erisman, J.W.; Bekunda, M.; Cai, Z.; Freney, J.R.; Martinelli, L.A.; Seitzinger, S.P.; Sutton, M.A. Transformation of the nitrogen cycle: Recent trends, questions, and potential solutions. Science 2008, 320, 889–892. [Google Scholar] [CrossRef] [Green Version]
  11. Ravishankara, A.R.; Daniel, J.S.; Portmann, R.W. Nitrous oxide (N2O): The dominant ozone-depleting substance emitted in the 21st century. Science 2009, 326, 123–125. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  12. Crutzen, P.J.; Moiser, A.R.; Smith, K.A.; Winiwarter, W. N2O release from agro-biofuel production negates global warming reduction by replacing fossil fuels. Atmos. Chem. Phys. 2008, 8, 389–395. [Google Scholar] [CrossRef] [Green Version]
  13. Davidson, A. The contribution of manure and fertilizer nitrogen to atmospheric nitrous oxide since 1860. Nat. Geosci. 2009, 2, 659–662. [Google Scholar] [CrossRef]
  14. Kamp, A.; Stief, P.; Knappe, J.; de Beer, D. Response of the ubiquitous pelagic diatom Thalassiosira weissflogii to darkness and anoxia. PLoS ONE 2013, 8, e82605. [Google Scholar] [CrossRef] [PubMed]
  15. Kamp, A.; de Beer, D.; Nitsch, J.L.; Lavik, G.; Stief, P. Diatoms respire nitrate to survive dark and anoxic conditions. Proc. Natl. Acad. Sci. USA 2011, 108, 5649–5654. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  16. Risgaard-Petersen, N.; Langezaal, A.M.; Ingvardsen, S.; Schmid, M.C.; Jetten, M.S.; Op den Camp, H.J.; Derksen, J.W.; Pina-Ochoa, E.; Eriksson, S.P.; Nielsen, L.P.; et al. Evidence for complete denitrification in a benthic foraminifer. Nature 2006, 443, 93–96. [Google Scholar] [CrossRef]
  17. Piña-Ochoa, E.; Hogslund, S.; Geslin, E.; Cedhagen, T.; Revsbech, N.P.; Nielsen, L.P.; Schweizer, M.; Jorissen, F.; Rysgaard, S.; Risgaard-Petersen, N. Widespread occurrence of nitrate storage and denitrification among Foraminifera and Gromiida. Proc. Natl. Acad. Sci. USA 2010, 107, 1148–1153. [Google Scholar] [CrossRef] [Green Version]
  18. Bernhard, J.M.; Casciotti, K.L.; McIlvin, M.R.; Beaudoin, D.J.; Visscher, P.T.; Edgcomb, V.P. Potential importance of physiologically diverse benthic foraminifera in sedimentary nitrate storage and respiration. J. Geophys. Res. 2012, 117, 1–14. [Google Scholar] [CrossRef] [Green Version]
  19. Kamp, A.; Hogslund, S.; Risgaard-Petersen, N.; Stief, P. Nitrate storage and dissimilatory nitrate reduction by eukaryotic microbes. Front. Microbiol. 2015, 6, 1492. [Google Scholar] [CrossRef] [Green Version]
  20. Woehle, C.; Roy, A.S.; Glock, N.; Wein, T.; Weissenbach, J.; Rosenstiel, P.; Hiebenthal, C.; Michels, J.; Schonfeld, J.; Dagan, T. A Novel eukaryotic denitrification pathway in foraminifera. Curr. Biol. 2018, 28, 2536–2543. [Google Scholar] [CrossRef] [Green Version]
  21. Kim, S.W.; Fushinobu, S.; Zhou, S.; Wakagi, T.; Shoun, H. Eukaryotic nirK genes encoding copper-containing nitrite reductase: Originating from the protomitochondrion? Appl. Environ. Microbiol. 2009, 75, 2652–2658. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  22. Shoun, H.; Fushinobu, S.; Jiang, L.; Kim, S.W.; Wakagi, T. Fungal denitrification and nitric oxide reductase cytochrome P450nor. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2012, 367, 1186–1194. [Google Scholar] [CrossRef] [Green Version]
  23. Maeda, K.; Spor, A.; Edel-Hermann, V.; Heraud, C.; Breuil, M.C.; Bizouard, F.; Toyoda, S.; Yoshida, N.; Steinberg, C.; Philippot, L. N2O production, a widespread trait in fungi. Sci. Rep. 2015, 5, 9697. [Google Scholar] [CrossRef] [Green Version]
  24. Higgins, S.A.; Welsh, A.; Orellana, L.H.; Konstantinidis, K.T.; Chee-Sanford, J.C.; Sanford, R.A.; Schadt, C.W.; Loffler, F.E. Detection and diversity of fungal nitric oxide reductase genes (p450nor) in agricultural soils. Appl. Environ. Microbiol. 2016, 82, 2919–2928. [Google Scholar] [CrossRef] [Green Version]
  25. Wang, J.; Vine, C.E.; Balasiny, B.K.; Rizk, J.; Bradley, C.L.; Tinajero-Trejo, M.; Poole, R.K.; Bergaust, L.L.; Bakken, L.R.; Cole, J.A. The roles of the hybrid cluster protein, Hcp and its reductase, Hcr, in high affinity nitric oxide reduction that protects anaerobic cultures of Escherichia coli against nitrosative stress. Mol. Microbiol. 2016, 100, 877–892. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  26. Plouviez, M.; Wheeler, D.; Shilton, A.; Packer, M.A.; McLenachan, P.A.; Sanz-Luque, E.; Ocana-Calahorro, F.; Fernandez, E.; Guieysse, B. The biosynthesis of nitrous oxide in the green alga Chlamydomonas reinhardtii. Plant J. 2017, 91, 45–56. [Google Scholar] [CrossRef] [Green Version]
  27. Burlacot, A.; Richaud, P.; Gosset, A.; Li-Beisson, Y.; Peltier, G. Algal photosynthesis converts nitric oxide into nitrous oxide. Proc. Natl. Acad. Sci. USA 2020, 117, 2704–2709. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  28. Plouviez, M.; Shilton, A.; Packer, M.A.; Guieysse, B. Nitrous oxide emissions from microalgae: Potential pathways and significance. J. Appl. Phycol. 2019, 31, 1–8. [Google Scholar] [CrossRef]
  29. Sasso, S.; Stibor, H.; Mittag, M.; Grossman, A.R. From molecular manipulation of domesticated Chlamydomonas reinhardtii to survival in nature. eLife 2018, 7. [Google Scholar] [CrossRef]
  30. Proschold, T.; Marin, B.; Schlosser, U.G.; Melkonian, M. Molecular phylogeny and taxonomic revision of Chlamydomonas (Chlorophyta). I. Emendation of Chlamydomonas Ehrenberg and Chlor´omonas Gobi, and description of Oogamochlamys gen. nov. and Lobochlamys gen. nov. Protist 2001, 152, 265–300. [Google Scholar] [CrossRef]
  31. Salome, P.A.; Merchant, S.S. A series of fortunate events: Introducing Chlamydomonas as a reference organism. Plant Cell 2019, 31, 1682–1707. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  32. Fernandez, E.; Galvan, A. Nitrate assimilation in Chlamydomonas. Eukaryot. Cell 2008, 7, 555–559. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Sanz-Luque, E.; Chamizo-Ampudia, A.; Llamas, A.; Galvan, A.; Fernandez, E. Understanding nitrate assimilation and its regulation in microalgae. Front. Plant. Sci. 2015, 6, 899. [Google Scholar] [CrossRef] [Green Version]
  34. Fernandez, E.; Galvan, A. Inorganic nitrogen assimilation in Chlamydomonas. J. Exp. Bot. 2007, 58, 2279–2287. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Chamizo-Ampudia, A.; Sanz-Luque, E.; Llamas, A.; Ocana-Calahorro, F.; Mariscal, V.; Carreras, A.; Barroso, J.B.; Galvan, A.; Fernandez, E. A dual system formed by the ARC and NR molybdoenzymes mediates nitrite-dependent NO production in Chlamydomonas. Plant Cell. Environ. 2016, 39, 2097–2107. [Google Scholar] [CrossRef]
  36. Düner, M.; Lambertz, J.; Mugge, C.; Hemschemeier, A. The soluble guanylate cyclase CYG12 is required for the acclimation to hypoxia and trophic regimes in Chlamydomonas reinhardtii. Plant J. 2018, 93, 311–337. [Google Scholar] [CrossRef] [Green Version]
  37. Sanz-Luque, E.; Ocaña-Calahorro, F.; de Montaigu, A.; Chamizo-Ampudia, A.; Llamas, A.; Galvan, A.; Fernandez, E. THB1, a truncated hemoglobin, modulates nitric oxide levels and nitrate reductase activity. Plant J. 2015, 81, 467–479. [Google Scholar] [CrossRef]
  38. Gerin, S.; Mathy, G.; Blomme, A.; Franck, F.; Sluse, F.E. Plasticity of the mitoproteome to nitrogen sources (nitrate and ammonium) in Chlamydomonas reinhardtii: The logic of Aox1 gene localization. Biochim. Biophys. Acta 2010, 1797, 994–1003. [Google Scholar] [CrossRef] [Green Version]
  39. van Lis, R.; Brugiere, S.; Baffert, C.; Coute, Y.; Nitschke, W.; Atteia, A. Hybrid cluster proteins in a photosynthetic microalga. FEBS J. 2020, 287, 721–735. [Google Scholar]
  40. Atteia, A.; Adrait, A.; Brugiere, S.; Tardif, M.; van Lis, R.; Deusch, O.; Dagan, T.; Kuhn, L.; Gontero, B.; Martin, W.; et al. A proteomic survey of Chlamydomonas reinhardtii mitochondria sheds new light on the metabolic plasticity of the organelle and on the nature of the alpha-proteobacterial mitochondrial ancestor. Mol. Biol. Evol. 2009, 26, 1533–1548. [Google Scholar] [CrossRef] [Green Version]
  41. Calatrava, V.; Hom, E.F.Y.; Llamas, A.; Fernandez, E.; Galvan, A. Nitrogen scavenging from amino acids and peptides in the model alga Chlamydomonas reinhardtii. The role of extracellular L-amino oxidase. Algal Res. 2019, 28, 101395. [Google Scholar] [CrossRef]
  42. Kirk, D.L.; Kirk, M.M. Carrier-mediated uptake of arginine and urea by Chlamydomonas reinhardtii. Plant Physiol. 1978, 61, 556–560. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  43. Zuo, Z.; Qingqing, R.; Chen, K.; Yang, L.; Cheng, Z.; Peng, K.; Zhu, Y.; Bai, Y.; Wang, Y. Study of amino acids as nitrogen source in Chlamydomonas reinhardtii. Phycol. Res. 2012, 60, 161–168. [Google Scholar] [CrossRef]
  44. Vallon, O.; Bulte, L.; Kuras, R.; Olive, J.; Wollman, F.A. Extensive accumulation of an extracellular L-amino-acid oxidase during gametogenesis of Chlamydomonas reinhardtii. Eur. J. Biochem. 1993, 215, 351–360. [Google Scholar] [CrossRef] [PubMed]
  45. Schroda, M.; Vallon, O. Chaperones and Proteases. In The Chlamydomonas Sourcebook, 2nd ed.; Harris, E.H., Stern, D.B., Witman, G.B., Eds.; Academic Press: Oxford, UK, 2009; pp. 671–729. [Google Scholar]
  46. Schmollinger, S.; Muhlhaus, T.; Boyle, N.R.; Blaby, I.K.; Casero, D.; Mettler, T.; Moseley, J.L.; Kropat, J.; Sommer, F.; Strenkert, D.; et al. Nitrogen-sparing mechanisms in Chlamydomonas affect the transcriptome, the proteome, and photosynthetic metabolism. Plant Cell 2014, 26, 1410–1435. [Google Scholar] [CrossRef] [Green Version]
  47. Muñoz-Blanco, J.; Hidalgo-Martinez, J.; Cardenas, J. Extracellular deamination of L-amino acids by Chlamydomonas reinhardtii cells. Planta 1990, 182, 194–198. [Google Scholar] [CrossRef] [PubMed]
  48. Kasai, K.; Ishikawa, T.; Nakamura, T.; Miura, T. Antibacterial properties of L-amino acid oxidase: Mechanisms of action and perspectives for therapeutic applications. Appl. Microbiol. Biotechnol. 2015, 99, 7847–7857. [Google Scholar] [CrossRef] [PubMed]
  49. Hunken, M.; Harder, J.; Kirst, G.O. Epiphytic bacteria on the Antarctic ice diatom Amphiprora kufferathii Manguin cleave hydrogen peroxide produced during algal photosynthesis. Plant Biol. 2008, 10, 519–526. [Google Scholar] [CrossRef]
  50. Calatrava, V.; Hom, E.F.Y.; Llamas, A.; Fernandez, E.; Galvan, A. OK, thanks! A new mutualism between Chlamydomonas and methylobacteria facilitates growth on amino acids and peptides. FEMS Microbiol. Lett. 2018, 365. [Google Scholar] [CrossRef]
  51. Chamizo-Ampudia, A.; Sanz-Luque, E.; Llamas, A.; Galvan, A.; Fernandez, E. Nitrate reductase regulates plant nitric oxide homeostasis. Trends Plant. Sci. 2017, 22, 163–174. [Google Scholar] [CrossRef]
  52. Leon, J.; Costa-Broseta, A. Present knowledge and controversies, deficiencies, and misconceptions on nitric oxide synthesis, sensing, and signaling in plants. Plant Cell Environ. 2020, 43, 1–15. [Google Scholar] [CrossRef] [PubMed]
  53. de Montaigu, A.; Sanz-Luque, E.; Macias, M.I.; Galvan, A.; Fernandez, E. Transcriptional regulation of CDP1 and CYG56 is required for proper NH4+ sensing in Chlamydomonas. J. Exp. Bot. 2011, 62, 1425–1437. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Rockel, P.; Strube, F.; Rockel, A.; Wildt, J.; Kaiser, W.M. Regulation of nitric oxide (NO) production by plant nitrate reductase in vivo and in vitro. J. Exp. Bot. 2002, 53, 103–110. [Google Scholar] [CrossRef] [PubMed]
  55. Yamasaki, H.; Sakihama, Y. Simultaneous production of nitric oxide and peroxynitrite by plant nitrate reductase: In vitro evidence for the NR-dependent formation of active nitrogen species. FEBS Lett. 2000, 468, 89–92. [Google Scholar] [CrossRef] [Green Version]
  56. Gupta, K.J.; Stoimenova, M.; Kaiser, W.M. In higher plants, only root mitochondria, but not leaf mitochondria reduce nitrite to NO, in vitro and in situ. J. Exp. Bot. 2005, 56, 2601–2609. [Google Scholar] [CrossRef]
  57. Jasid, S.; Simontacchi, M.; Bartoli, C.G.; Puntarulo, S. Chloroplasts as a nitric oxide cellular source. Effect of reactive nitrogen species on chloroplastic lipids and proteins. Plant Physiol. 2006, 142, 1246–1255. [Google Scholar] [CrossRef] [Green Version]
  58. Tewari, R.K.; Prommer, J.; Watanabe, M. Endogenous nitric oxide generation in protoplast chloroplasts. Plant Cell Rep. 2013, 32, 31–44. [Google Scholar] [CrossRef]
  59. Corpas, F.J.; Del Rio, L.A.; Palma, J.M. Plant peroxisomes at the crossroad of NO and H2O2 metabolism. J. Integr. Plant Biol. 2019, 61, 803–816. [Google Scholar] [PubMed] [Green Version]
  60. Foresi, N.; Mayta, M.L.; Lodeyro, A.F.; Scuffi, D.; Correa-Aragunde, N.; Garcia-Mata, C.; Casalongue, C.; Carrillo, N.; Lamattina, L. Expression of the tetrahydrofolate-dependent nitric oxide synthase from the green alga Ostreococcus tauri increases tolerance to abiotic stresses and influences stomatal development in Arabidopsis. Plant J. 2015, 82, 806–821. [Google Scholar] [CrossRef]
  61. Jeandroz, S.; Wipf, D.; Stuehr, D.J.; Lamattina, L.; Melkonian, M.; Tian, Z.; Zhu, Y.; Carpenter, E.J.; Wong, G.K.; Wendehenne, D. Occurrence, structure, and evolution of nitric oxide synthase-like proteins in the plant kingdom. Sci. Signal. 2016, 9, re2. [Google Scholar] [CrossRef] [Green Version]
  62. del Rio, L.A.; Corpas, F.J.; Barroso, J.B. Nitric oxide and nitric oxide synthase activity in plants. Phytochemistry 2004, 65, 783–792. [Google Scholar] [CrossRef] [PubMed]
  63. Gonzalez-Ballester, D.; Sanz-Luque, E.; Galvan, A.; Fernandez, E.; de Montaigu, A. Arginine is a component of the ammonium-CYG56 signalling cascade that represses genes of the nitrogen assimilation pathway in Chlamydomonas reinhardtii. PLoS ONE 2018, 13, e0196167. [Google Scholar] [CrossRef] [PubMed]
  64. Hemschemeier, A.; Duner, M.; Casero, D.; Merchant, S.S.; Winkler, M.; Happe, T. Hypoxic survival requires a 2-on-2 hemoglobin in a process involving nitric oxide. Proc. Natl. Acad. Sci. USA 2013, 110, 10854–10859. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. Folgosa, F.; Martins, M.C.; Teixeira, M. The multidomain flavodiiron protein from Clostridium difficile 630 is an NADH:oxygen oxidoreductase. Sci. Rep. 2018, 8, 10164. [Google Scholar] [CrossRef]
  66. Romao, C.V.; Vicente, J.B.; Borges, P.T.; Frazao, C.; Teixeira, M. The dual function of flavodiiron proteins: Oxygen and/or nitric oxide reductases. J. Biol. Inorg. Chem. 2016, 21, 39–52. [Google Scholar] [CrossRef]
  67. Seedorf, H.; Dreisbach, A.; Hedderich, R.; Shima, S.; Thauer, R.K. F420H2 oxidase (FprA) from Methanobrevibacter arboriphilus, a coenzyme F420-dependent enzyme involved in O2 detoxification. Arch. Microbiol. 2004, 182, 126–137. [Google Scholar] [CrossRef]
  68. Smutna, T.; Goncalves, V.L.; Saraiva, L.M.; Tachezy, J.; Teixeira, M.; Hrdy, I. Flavodiiron protein from Trichomonas vaginalis hydrogenosomes: The terminal oxygen reductase. Eukaryot. Cell 2009, 8, 47–55. [Google Scholar] [CrossRef] [Green Version]
  69. Di Matteo, A.; Scandurra, F.M.; Testa, F.; Forte, E.; Sarti, P.; Brunori, M.; Giuffre, A. The O2-scavenging flavodiiron protein in the human parasite Giardia intestinalis. J. Biol. Chem. 2008, 283, 4061–4068. [Google Scholar] [CrossRef] [Green Version]
  70. Vicente, J.B.; Gomes, C.M.; Wasserfallen, A.; Teixeira, M. Module fusion in an A-type flavoprotein from the cyanobacterium Synechocystis condenses a multiple-component pathway in a single polypeptide chain. Biochem. Biophys. Res. Commun. 2002, 294, 82–87. [Google Scholar] [CrossRef]
  71. Silaghi-Dumitrescu, R.; Coulter, E.D.; Das, A.; Ljungdahl, L.G.; Jameson, G.N.; Huynh, B.H.; Kurtz, D.M., Jr. A flavodiiron protein and high molecular weight rubredoxin from Moorella thermoacetica with nitric oxide reductase activity. Biochemistry 2003, 42, 2806–2815. [Google Scholar] [CrossRef]
  72. Rodrigues, R.; Vicente, J.B.; Felix, R.; Oliveira, S.; Teixeira, M.; Rodrigues-Pousada, C. Desulfovibrio gigas flavodiiron protein affords protection against nitrosative stress in vivo. J. Bacteriol. 2006, 188, 2745–2751. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  73. Justino, M.C.; Vicente, J.B.; Teixeira, M.; Saraiva, L.M. New genes implicated in the protection of anaerobically grown Escherichia coli against nitric oxide. J. Biol. Chem. 2005, 280, 2636–2643. [Google Scholar] [CrossRef] [Green Version]
  74. Gardner, A.M.; Helmick, R.A.; Gardner, P.R. Flavorubredoxin, an inducible catalyst for nitric oxide reduction and detoxification in Escherichia coli. J. Biol. Chem. 2002, 277, 8172–8177. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  75. Chaux, F.; Burlacot, A.; Mekhalfi, M.; Auroy, P.; Blangy, S.; Richaud, P.; Peltier, G. Flavodiiron proteins promote fast and transient O2 photoreduction in Chlamydomonas. Plant Physiol. 2017, 174, 1825–1836. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  76. Mus, F.; Dubini, A.; Seibert, M.; Posewitz, M.C.; Grossman, A.R. Anaerobic acclimation in Chlamydomonas reinhardtii: Anoxic gene expression, hydrogenase induction, and metabolic pathways. J. Biol. Chem. 2007, 282, 25475–25486. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  77. Olson, A.C.; Carter, C.J. The involvement of hybrid cluster protein 4, HCP4, in anaerobic metabolism in Chlamydomonas reinhardtii. PLoS ONE 2016, 11, e0149816. [Google Scholar] [CrossRef] [Green Version]
  78. Overeijnder, M.L.; Hagen, W.R.; Hagedoorn, P.L. A thermostable hybrid cluster protein from Pyrococcus furiosus: Effects of the loss of a three helix bundle subdomain. J. Biol. Inorg. Chem. 2009, 14, 703–710. [Google Scholar] [CrossRef] [Green Version]
  79. Yu, Z.; Pesesky, M.; Zhang, L.; Huang, J.; Winkler, M.; Chistoserdova, L. A Complex interplay between nitric oxide, quorum sensing, and the unique secondary metabolite tundrenone constitutes the hypoxia response in Methylobacter. mSystems 2020, 5. [Google Scholar] [CrossRef] [Green Version]
  80. Seth, D.; Hess, D.T.; Hausladen, A.; Wang, L.; Wang, Y.J.; Stamler, J.S. A Multiplex enzymatic machinery for cellular protein s-nitrosylation. Mol. Cell 2018, 69, 451–464LO. [Google Scholar] [CrossRef] [Green Version]
  81. Bange, H.B.; Arevalo-Martiniez, D.L.; Paz, M.; Farias, L.; Kaiser, J.; Kock, A.; Law, C.S.; Rees, A.P.; Rehder, G.; Tortell, P.D.; et al. A harmonized nitrous oxide (N2O) ocean observation network for the 21st century. Front. Mar. Sci. 2019, 6. [Google Scholar] [CrossRef]
  82. Cohen, Y.; Gordon, L.G. Nitrous oxide in the oxygen minimum of the eastern tropical North Pacific: Evidence for its consumption during denitrification and possible mechanisms for its production. Deep Sea Res. 1978, 25, 509–524. [Google Scholar] [CrossRef]
  83. Smith, C.J.; DeLaune, R.D.; Patrick, W.H. Nitrous oxide emission from Gulf Coast wetlands. Geochim. Cosmochin. 1983, 47, 1805–1814. [Google Scholar] [CrossRef]
  84. Mengis, M.; Gächter, R.; Wehrli, B. Sources and sinks of nitrous oxide (N2O) in deep lakes. Biogeochemistry 1997, 38, 281–301. [Google Scholar] [CrossRef]
  85. Guieysse, B.; Plouviez, M.; Coilhac, M.; Cazali, L. Nitrous Oxide (N2O) production in axenic Chlorella vulgaris microalgae cultures: Evidence, putative pathways, and potential environmental impacts. Biogeosciences 2013, 10, 6737–6746. [Google Scholar] [CrossRef] [Green Version]
  86. Smeets, E.M.; Bouwman, L.F.; Stehfest, E.; Vuuren, D.P.; Posthuma, A. Contribution of N2O to the greenhouse gas balance of first-generation biofuels. Glob. Chang. Biol. 2009, 15, 1–23. [Google Scholar] [CrossRef]
  87. Bauer, S.K.; Grotz, L.S.; Connelly, E.B.; Colosi, L.M. Reevaluation of the global warming impacts of algae-derived biofuels to account for possible contributions of nitrous oxide. Bioresour. Technol. 2016, 218, 196–201. [Google Scholar] [CrossRef]
  88. Higgins, S.A.; Schadt, C.W.; Matheny, P.B.; Loffler, F.E. Phylogenomics reveal the dynamic evolution of fungal nitric oxide reductases and their relationship to secondary metabolism. Genome Biol. Evol. 2018, 10, 2474–2489. [Google Scholar] [CrossRef] [Green Version]
  89. Weather, P.J. N2O Evolution by green algae. Appl. Environ. Microbiol. 1984, 48, 1251–1253. [Google Scholar] [CrossRef] [Green Version]
  90. Florez-Leiva, L.; Tarifeño, E.; Cornejo, M.; Kiene, R.; Farias, L. High production of nitrous oxide (N2O), methane (CH4) and dimethylsulphoniopropionate (DMSP) in a massive marine phytoplankton culture. Biogeosci. Discuss. 2010, 7, 6705–6723. [Google Scholar] [CrossRef]
  91. Fagerstone, K.D.; Quinn, J.C.; Bradley, T.H.; De Long, S.K.; Marchese, A.J. Quantitative measurement of direct nitrous oxide emissions from microalgae cultivation. Environ. Sci. Technol. 2011, 45, 9449–9456. [Google Scholar] [CrossRef] [PubMed]
Plants 09 00903 g001 550
Figure 1. Nitrogen-transforming reactions by Chlamydomonas reinhardtii. NO3− assimilation (steps 1,2,3); NO3−/NO3− cycle, (steps 1,4,5); NO3− dissimilation (steps 4,6) and ammonification from amino acids and peptides (step 7). Transporters for ammonium (AMT), nitrate (NRT), and arginine are indicated. Mutualism with bacteria for N/C exchange is also shown.
Figure 1. Nitrogen-transforming reactions by Chlamydomonas reinhardtii. NO3− assimilation (steps 1,2,3); NO3−/NO3− cycle, (steps 1,4,5); NO3− dissimilation (steps 4,6) and ammonification from amino acids and peptides (step 7). Transporters for ammonium (AMT), nitrate (NRT), and arginine are indicated. Mutualism with bacteria for N/C exchange is also shown.
Plants 09 00903 g001
Table
Table 1. Enzymes for nitrogen-transformation reactions in Chlamydomonas.
Table 1. Enzymes for nitrogen-transformation reactions in Chlamydomonas.
Step Enzyme Gene IDRegulationLocalizationRef.
1NO3– + 2e– + 2H+→ NO2– + H2O NRCre09.g410950NO3–Cytosol[33]
2NO2– + 6e– + 8H+ → NH4+ + 2H2O NIRCre09.g410950NO3– [33]
3NH4+ + 2e– + α-ketoglutarate + ATP→
Glutamate + ADP + Pi		GS/GOGAT [33]
4NO2– + e– + 2H+ → NO + H2O
NR/NOFNIR
NirK		Cre09.g395950/Cre09.g389089
Cre08.g360550		NO3–
Dark		Cytosol
Mitochondria		[35]
[36]
5NO + 2H2O → NO3– + 3e– + 4H+NR/THB1Cre09.g410950/Cre14.g615400NO3–Cytosol[37]
62NO + 2e– + 2H+ → N2O + H2O CYP55
FLVA
FLVB
HCP1
HCP2
HCP3
HCP4		Cre01.g007950
Cre12.g531900
Cre16.g691800
Cre09.g391650
Cre09.g393543
Cre09.g393506
Cre09.g391450		NO3–/Dark
-
-
NO3–/Dark
NO3–
NO3–/Dark
NO3–/Dark/anaerobiosis		Mitochondria*
Chloroplast
Chloroplast
Chloroplast
Mitochondria*
Chloroplast
Chloroplast		[38]
[27]
[27]
[39]
[38]
[39]
[40]
7L-Amino acid +H2O+ O2 → NH4+ + α-keto acid + H2O2LAO1Cre12.g551353Nitrogen starvationPeriplasm[35]
Steps 1 to 7 correspond to those in Figure 1. Gene identification (Phytozome). * Protein indentified in the mitoproteome. NR (Nitrate Reductase), NIR (Nitrite Reductase), GS (Glutamine Synthetase), GOGAT (Glutamine Oxoglutarate Aminotransferase), NOFNIR (NO Forming Nitrite Reductase), NirK (Copper-Containing Nitrite Reductase), THB1 (Truncated Hemoglobin 1), CYP55 (Cytochrome P450), FLV (Flavodiiron Proteins), HCP (Hybrid Cluster Protein) and LAO1 (L-Amino Acid Oxidase).
Table
Table 2. Microalgal production of N2O and putative NIRK proteins, involved in NO formation, and putative HCP, CYP55, and FLV, involved in NO reduction to N2O.
Table 2. Microalgal production of N2O and putative NIRK proteins, involved in NO formation, and putative HCP, CYP55, and FLV, involved in NO reduction to N2O.
MicroalgaeNIRKHCPCYP55FLVAFLVBN2O/Ref.
Chlamydomonas reinhardtiiPNW79625.1 XP_001694756.1
XP_001694571.1
XP_001694671.1
XP_001694454.1		XP_001700272.1 XP_001692916.1 PNW71243.1 + [26,27]
Chlamydomonas eustigma GAX74118.1 GAX73888.1 GAX73438.1
Chlorella variabilis XP_005851548.1XP_005848845.1XP 005849742.1 XP 005849474.1+ [27]
Chlorella sorokinianaPRW57688.1PRW58172.1PRW58571.1 (A)PRW59370.1PRW58882.1
PRW60486.1PRW58572.1 (B)
PRW58278.1
Scenedesmus obliquus UTEX 393SZX67391.1ID:15469ID:435SZX62872.1SZX63966.1+ [89]
ID:3042 SZX76545.1SZX639665.1
Scenedesmus obliquus EN0004ID:579459ID:571596ID:324854ID:321778/332768XP002948938.1+ [89]
ID:618262 ID:342393
Nannochloropsis salina NSK 004142 + [90,91]
Nannochloropsis oceanica CCMP1779 599317
Nannochloropsis gaditana XP 005853562.1 − [27]
Gonium pectorale KXZ41847.1KXZ42279.1KXZ51271.1KXZ42718.1
KXZ44628.1
Coccomyxa subelipsoidea XP 005643451.1 XP 005643449.1 + [27]
Tetrabaena socialis PNH09786.1
Raphidocelis subcapitata GBF99305.1GBF95475.1GBF88523.1GBF98453.1
GBF96263.1
Volvox carteri XP_002952683.1 XP 002947563.1XP 002948938.1
Yamagishiella unicocca ARO50069.1
Micractinium conductrix PSC73563.1 PSC76351.1 PSC76929.1
PSC72373.1
PSC68490.1
Micromonas pusilla XP 003057013.1XP 003057495.1
Micromonas commoda XP 002502419.1 XP 002502038.1
Monoraphidium neglectum SAG 48.87 XP013900867.1ID:2893
ID:2894
Bathycoccus prasinos XP007514084.1XP 007514099.1
Bathycoccus lucimarinus XP001416100.1XP001416098.1
Ostreococcus tauri XP003075211.2XP003075215.2
Klebsormidium nitens GAQ79906.1 GAQ89021.1
Galdieria sulphurariaXP_005707361.1 − [27]
Porphyridium purpureum − [27]
Phaeodactylum tricornutum ID:12416 − [27]
Thalassiosira pseudonana ID:32716ID:263399a − [27]
Guillardia theta CCMP2712 ID:152637
Andalucia godoyiKAF0852161.1
Phaeocystis antarctica CCMP1374ID:684
Emiliana huxleyiXP 005783766.1
Reciprocal BLAST was performed using the Chlamydomonas homolog of each protein as a query. Green background highlights microalgal species that belong to the Chlorophyta phylum. For those algal species that did not show in a positive hit using the current available genome sequences, the gene coding for the corresponding queried protein was considered to be absent. Protein accession numbers correspond to those indicated in Figures S1–S4, from NCBI or Phycocosm (protein references obtained from Phycocosm are indicated as follows “ID: (ref number)”. “a” indicates partial sequence. N2O/Ref., positive or negative (+/−) N2O production by the microalgae and empty when it is unknown.

Share and Cite

MDPI and ACS Style

Bellido-Pedraza, C.M.; Calatrava, V.; Sanz-Luque, E.; Tejada-Jiménez, M.; Llamas, Á.; Plouviez, M.; Guieysse, B.; Fernández, E.; Galván, A. Chlamydomonas reinhardtii, an Algal Model in the Nitrogen Cycle. Plants 2020, 9, 903. https://doi.org/10.3390/plants9070903

AMA Style

Bellido-Pedraza CM, Calatrava V, Sanz-Luque E, Tejada-Jiménez M, Llamas Á, Plouviez M, Guieysse B, Fernández E, Galván A. Chlamydomonas reinhardtii, an Algal Model in the Nitrogen Cycle. Plants. 2020; 9(7):903. https://doi.org/10.3390/plants9070903

Chicago/Turabian Style

Bellido-Pedraza, Carmen M., Victoria Calatrava, Emanuel Sanz-Luque, Manuel Tejada-Jiménez, Ángel Llamas, Maxence Plouviez, Benoit Guieysse, Emilio Fernández, and Aurora Galván. 2020. "Chlamydomonas reinhardtii, an Algal Model in the Nitrogen Cycle" Plants 9, no. 7: 903. https://doi.org/10.3390/plants9070903

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop