The production of zeaxanthin from microalgae is attractive and varies depending on microalgae species and cultivation factors. The favorable light condition for zeaxanthin varies in microalgae species. The zeaxanthin content of *C. zofingiensis bkt1* (5.69 ± 0.45 mg/g DW) was strongly enhanced by high light as compared with that of low light control (0.34 ± 0.04 mg/g DW) [33]. Under high light, combined with N-starvation, C. *zofingiensis bkt1* can accumulate 7.00 ± 0.82 mg/g DW. However, high light is not always preferred for microalgae to accumulate zeaxanthin. A mutant *D. salinazea1* can accumulate 5.9 mg/g DW of zeaxanthin under continuous low light (100 μmol photons m−2 s−1), higher than that transferred to high light (4.18 mg/g DW) [43]. As *D. salina* grows well in hypersaline medium, cultivation of this strain with seawater is potentially profitable. Results of satisfactory zeaxanthin production from microalgae are summarized in Table 2 with details of contents and productivities.

Additionally, downstream processing of zeaxanthin needs more attention, such as extraction and separation. Solvent systems have different efficiencies in zeaxanthin extraction from microalgae, and the maximum extraction of zeaxanthin (30.2 mg/g DW) was in chloroform: methanol (1:2) [41]. Pressurized liquid extraction was also applied to extract zeaxanthin from *C. ellipsoidea* [41]. More importantly, zeaxanthin shares similar chemical structures to lutein. Huang et al. developed an efficient method for the separation of zeaxanthin and lutein by ultra-high performance liquid chromatography (UHPLC) equipped with a Waters YMC Carotenoid C30 column [33].

#### 2.2.5. Fucoxanthin

Fucoxanthin is a natural carotenoid that occupies nearly 10% of total natural carotenoid production [62]. The global production of fucoxanthin was about 500 tons in 2016 and was estimated to increase further with an annual rate of 5.3% [63]. As shown in Figure 2, it has a unique molecular structure with an allenic bond, a conjugated carbonyl, a 5,6-monoepoxide and acetyl groups. Fucoxanthin and its derivatives have shown potential anti-cancer [64], anti-inflammatory [65] and anti-obesity effects [46].

**Figure 2.** The chemical structure of (*all-E*)-fucoxanthin.

Currently, the main natural sources of fucoxanthin rely on some macroalgae species, such as *Laminaria japonica* and *Undaria pinnatifid*, which are common in Asian diets for iodine supplement [66]. However, the fucoxanthin contents in these macroalgae are relatively low (0.02–0.58 mg/g fresh weight) and not feasible for commercialization. Microalgae grow faster than macroalgae with higher productivity of carotenoids. Fucoxanthin is a primary light-harvesting carotenoid that transfers energy to LHC with high efficiency (>80%) [67] and may also protect microalgae from high light with its internal antioxidant property [47]. Up until now, research about producing fucoxanthin from microalgae is still relatively lacking.

Microalgae are potential candidates for fucoxanthin production. Fucoxanthin mainly exists in heterokont and haptophyte groups of algae (>20,000 species, especially in Synurophyceae (up to 26.6 mg/g DW), diatoms (up to 21.67 mg/g DW) and Prymnesiophyceae (up to 18.23 mg/g DW) [45]. Under low light intensity, carbon skeletons are diverted into carotenoids, especially fucoxanthin in *Isochrysis zhanjiangensis*, which can produce 23.29 mg/g DW of fucoxanthin [46]. The marine diatom *Odontella aurita* can accumulate 18.47 mg/g DW of fucoxanthin in a nitrogen-replete medium under low light [47]. Moreover, the purified fucoxanthin from *O. aurita* is (*all-E*)-fucoxanthin, which owns strong antioxidant ability and bioavailability. Maria et al. isolated a novel strain *Mallomonas sp.* (Synurophyceae) with the highest known content of fucoxanthin at 26.6 mg/g DW [45]. The highest ever reported fucoxanthin productivity was found in *Tisochrysis lutea* with 9.81 mg/L/day under batch culture with continuous chemostat dilution [49].

Diatom *Phaeodactylum tricornutum* also has a high fucoxanthin content at 16.33 mg/g DW [50]. As *P. tricornutum* is a model system for investigations and its genome has been sequenced, further exploitations of key genes related to fucoxanthin, fucoxanthin hyperaccumulating mutants and genetic modifications of diatoms for fucoxanthin production are expected in the near future. Results of satisfactory fucoxanthin production from microalgae are summarized in Table 2 with details of contents and productivities.

#### **3. Salt-Stress Treatment for Carotenoid Production from Microalgae**

Microalgae have flexible metabolic systems, which means that external environmental changes will affect biochemical components of microalgal cells. Stress factors, such as high light intensity, salinity, nitrogen starvation and high/low temperatures, will provoke metabolic changes in microalgae and lead to different cell compositions.

Though carotenoid production could be boosted by several environmental abiotic stress factors, salt-stress treatment is much more attractive because it is cheap, easy to operate and has great commercialization value for seawater culture of microalgae. Here, we elucidate the contributions of salinity in microalgal culture and production of carotenoids and summarize the current strategies of salt-stress treatment in carotenoid production.

#### *3.1. Microalgal Responses to Salt Stress*

Salt stress is one of the major abiotic stresses decreasing productivity of agriculture all over the earth [68,69]. Salinity inhibits plant growth, development, seed germination and yields. In contrast, plants have developed strategies to adapt to high salt concentrations, such as regulating stress hormones and growth hormones to balance growth and stress responses [69]. Currently, research on salt stress is more in-depth and comprehensive in higher plants, but there are a few studies on microalgae.

Due to the fast growth rate of microalgae and their richness of high-value products, large-scale microalgae cultivation is imperative at present [6]. Microalgae are widely distributed in the ocean and freshwater, while salt stress is still a threat to the large-scale production of freshwater microalgae. As freshwater resources are becoming scarce today, the cultivation of freshwater microalgae with seawater is of great significance. On the other hand, appropriate salt stress shows promoting effects on secondary metabolite accumulation (such as secondary carotenoids and lipids) [70]. Therefore, it is of great significance to study the response of microalgae to salt stress. The possible responses in microalgae cells caused by salt stress are summarized in Figure 3.

**Figure 3.** Possible mechanisms of responses of microalgae induced by salt stress.

Salt stress can affect microalgae on multiple levels, as physiological characteristics, gene expression and metabolic pathways. Salt stress can reduce cell growth, decrease chlorophyll content, inhibit photosynthesis and cause morphological changes in microalgae [9]. Under salt stress, the cell wall of *H. pluvialis* thickens, the cell volume becomes larger and gradually becomes immobile cysts [10]. These morphological changes are related to a series of in-depth signaling and downstream changes in both genetic and metabolic aspects.

#### 3.1.1. Early Signaling

Ca2+ is considered as a universal second messenger for the primary stress signals [71], acting in real-time in response to the imbalance of cell ion homeostasis under NaCl stress in plants [72]. Similar to plants, salt stress can elicit a temporary increase in cytosolic Ca2+ concentration ([Ca2+]cyt), which can regulate the activity of downstream effector proteins, such as calmodulin (CaM), Ca2+-dependent protein kinases and CaM-dependent protein kinases [73]. Moreover, the decarboxylation of glutamate to γ-aminobutyric acid (GABA) was regulated by the Ca2+/CaM protein.

GABA is a recently identified endogenous signaling molecule in plants, participating in cell growth and enhancing abiotic stress tolerance [74]. It is a non-protein amino acid derived from the decarboxylation of glutamate, and exogenous GABA can enhance plant resistance to abiotic stress by activating the GABA bypass pathways and TCA cycle. Under salt stress, GABA helps to maintain C/N balance and even acts as a scavenger of toxic ROS [75]. It is also related to nitric oxide (NO) accumulation under stress conditions as it amplifies NO stress signaling [74].

NO is an important molecule involved in plant growth, development and tolerance to abiotic stress. It plays important roles in resistance to drought, temperature (high or low), UV-B and heavy metal stress [76]. NO can also act as a signal in activating antioxidant enzyme defense against oxidative stress induced by salt stress [76]. By applying NO donors, such as sodium nitroprusside, plants under stress conditions showed restoration of chlorophyll to recover their damaged photosynthetic system [77].

Reactive oxygen species (ROS) are also second messengers induced by salinity stress, which is also associated with Ca2+ signaling [73]. ROS is a stress indicator in response to abiotic stress of microalgae, which can regulate cell growth and metabolites synthesis. Toxic ROS will lead to lipid peroxidation, membrane deterioration, DNA and protein damage [68]. To eliminate excessive ROS, antioxidant enzymes (such as superoxide dismutase and catalase) and antioxidants (such as carotenoids) in microalgae are two essential mechanisms evolved by organisms [78].

#### 3.1.2. Downstream Signaling

Salinity-induced signals may then have effects on the gene expression of microalgae cells, which is related to salt concentrations [79]. Low salt concentration can promote the growth of some microalgae, such as cultures of *Scenedesmus sp.* [80], *Botryococcus braunii* [81] and *H. pluvialis* [79]. Treatment of *H. pluvialis* with low-dose NaCl (12.5 mg/L) showed a promoting effect on biomass concentration (28% higher) and productivity (from 0.15 d−1–0.22 d−1). These effects were related to upregulation of growth-related genes, such as *rbcL*, *rbcS* and nitrate reductase gene (*NR*).

Under high-dose salt stress, both ionic and osmotic homeostasis need to be maintained. As for the halotolerant microalga *D. salina*, osmoregulation under osmotic stresses can be divided into two mechanisms. One is to maintain intracellular Na+ and K+ concentrations by the plasma membrane electron transport (redox) system, as Na+-ATPase and K+ carriers, and the other is to regulate glycerol concentrations inside to maintain the water potential inside and outside of cells [82]. Accumulation of glycerol as a soluble substance is a strategy for microalgae to keep osmotic homeostasis under salt stress, which has been demonstrated in *C. reinhardtii* [83] and *C. zofingiensis* [15]. Salt stress can also upregulate the genes related to starch catabolism (like pyruvate kinase, PK) and downregulate the genes for gluconeogenesis (like phosphoenolpyruvate carboxykinase, PEPCK), providing more building blocks for storage lipids (fatty acids and triacylglycerol, TAGs) and carotenoids in *C. zofingiensis* [15]. Salt-induced acetyl-coenzyme A carboxylase (ACCase) expression for fatty acid synthesis was also demonstrated in *Chlamydomonas* sp. [84], *Chlorella* sp. [85] and *Nitzschia* sp. [86].

Omics approaches like genomic, transcriptomic and metabolomic are also applied to reveal the changes occurring under salt stress. By analyzing differential expressed genes (DEGs) under stress, transcription factors (TFs), such as myeloblastosis (MYB), WRKY and basic helix–loop–helix (bHLH), are demonstrated to play important roles in regulating gene expressions under salinity [68]. Responses of microalgae to both low-dose and high-dose salinity are shown in Table 3.

**Table 3.** Overview of the effects of salinity on microalgae at different doses.

#### *3.2. Salt Stress Strategies for Carotenoid Accumulation*

Carotenoid production could be affected by environmental abiotic stress. Under salt stress, carotenoids will accumulate and protect cells as antioxidants to increase the surviving possibility of microalgae. In addition, optimal salt condition varies among different microalgal species. After NaCl treatment (1%, 0.17 M) for 10 days, the astaxanthin content of *H. pluvialis* climbed from 3.53 mg/g to 17.7 mg/g [87]. *C. zofingiensis* CCAP 211/14 can tolerate moderate NaCl concentration of 100 mM, and a significant enhancement of astaxanthin content was observed under 200 mM NaCl treatment [88]. The optimal condition to obtain the highest amount of fucoxanthin (79.40 ± 0.95 mg/g DW) of *Tisochrysis lutea* was determined as 36.27 g/L salt addition [64].

As the optimal conditions for cell growth are commonly different from those for secondary metabolite accumulation, two-stage cultivation has been widely applied to microalgae cultivation, a potent strategy to balance cell growth and metabolite accumulation. Acidophilic eukaryotic microalga *Coccomyxa onubensis* can endure moderate salt stress and serves as a potential resource for lutein production. Its growth rate and biomass productivity were significantly boosted under salt treatment (100 mM NaCl). By adding 500 mM NaCl, the lutein content was significantly enhanced by 47% to 7.80 mg/g DW, though the cell growth was inhibited [89]. Thus, cultures containing 100 mM salt can be applied at the first stage for higher biomass, then extra salt was added to induce lutein accumulation at the second stage.

Moreover, salt stress can also be coupled with other treatments to increase the carotenoid accumulation in microalgae. Light induction and nutrient starvation are widely used methods to boost carotenoid production. Combined with high light, salinity treatment increased the astaxanthin yield of *C. zofingiensis* 7.53-fold compared with the control [70]. *D. salina* was able to adapt to NaCl ranging from 0.05 to 5.5 M and the β-carotene content of *D. salina* achieved 13% of DW under salt stress, combined with high light at high temperature under nutrient deficiency [40].

The addition of chemicals can also boost carotenoid accumulation under salt stress. Photocatalyst TiO2 can enhance zeaxanthin accumulation under salt stress in *Coelastrella sp.* by increasing oxidative stress [42]. The maximum zeaxanthin (13.2 ± 4.4 mg/g DW) was achieved under high salinity (3% NaCl) and N-starvation at 40 ◦C after TiO2 treatment. An 0.25 mM amount of γ-aminobutyric acid (GABA) could facilitate astaxanthin productivity by 3.24-fold in *H. pluvialis* under high light with salinity treatment (2 g/L) [90]. Melatonin (MT) addition enhanced the expression of carotenogenic genes of *H. pluvialis* and induced astaxanthin accumulation by 1.20-fold under N-starvation and salt stress (1 g/L) [91]. NaCl treatment could also amplify the effect of linoleic acid (LA) on boosting astaxanthin accumulation in *Chlorella sorokiniana*, and LA could increase astaxanthin content by 1.25-fold in

the presence of 20% NaCl (*w*/*v*) [92]. Strategies with salinity treatment to boost carotenoid accumulation in microalgae are summarized in Table 4.

**Table 4.** Salt treatment and salinity-combined conditions for carotenoids production.

In summary, moderate salt stress can increase biomass productivity and induce accumulation of carotenoids, depending on microalgae species, while strict stress will be toxic and even fatal to microalgae. As both high cell density and carotenoid productivity are desirable for biotechnological goals, an optimal salt concentration should be chosen cautiously according to different cultivation goals and microalgae species. Two-stage or multi-stage cultivation and strategies combining salinity with other stress factors also have potential in microalgae cultivation for carotenoids.

#### **4. Potential Applications of Internet of Things (IoT) in Carotenoids Production**

The Internet of Things (IoT) is a huge ecosystem that connects things, machines and humans, anytime and anywhere. As the third wave of the world information industry, the IoT has attracted thriving research interest from various fields, such as the food supply chain [93], healthcare applications [94] and precision agriculture [95]. The number of IoT devices worldwide is predicted to reach 75 billion by 2025 [93]. Considering its successful experiences in agriculture and its advantages as a real-time, efficient and intelligent comprehensive data system of works, the IoT also shows competitiveness in microalgae biorefinery [96].

Microalgae biorefinery can be divided into two consecutive categories, the upstream processing and downstream processing [97]. The upstream processing mainly focuses on the high-density cultivation of selected microalgae and the regulation and optimization of the cultivation process, such as choosing the suitable trophic mode and light conditions. The downstream process mainly includes the harvest of microalgae, the separation and extraction of key products and subsequent processing to make microalgae-derived products. The IoT, as an ubiquitous, huge network, can be perfectly combined with traditional biotechnologies in microalgae biorefinery [96]. As shown in Figure 4, both upstream and downstream events are connected and collected, and all information will then be processed by the IoT platform. Then, how can the IoT be applied in the production of microalgae-derived carotenoids?

**Figure 4.** Characteristics of IoT and their potential utilizations in microalgae biorefinery. UP, upstream processing; DOWN, downstream processing.

For the upstream processing of carotenoid production, sensors are indispensable for data monitoring. Traditional detection techniques (such as measurement of biomass, cell size, etc.) are time-consuming, prone to causing culture contamination and cannot get data in time. In comparison, online sensors can monitor microalgae growth to gain real-time data, calculate unmeasured variables and even predict growth models.

For example, *H. pluvialis* is relatively sensitive to culture environment; its growth and cell composition can go far away from expectations without careful tuning. Possible contamination during sampling can disturb its growth. In addition, nutrient deficiency could drive it into a cyst structure and cause it to begin accumulating astaxanthin and other secondary metabolites [10]. In this case, innovative online sensors can monitor its growth in several aspects. Optical density (OD) sensors [98,99] can determine real-time biomass and dissolved oxygen (DO), CO2 sensors [100] can screen metabolic activity, pH sensors and nitrate and phosphorous sensors [101] can guarantee a suitable environment and nutrient sufficiency; these can all be applied to ensure high-density cultivation of *H. pluvialis*. Moreover, the IoT can also act as the base of a decision support system. By combining the data from sensors with simulation models, the IoT platform can predict growth trends and provide cultivation suggestions to maximize biomass production [102]. Afterwards, automation units of the IoT are essential to achieve condition debugging at the right time after data analysis, such as a suitable feeding strategy [103]. As for cultivation with salt-stress treatments, the IoT will facilitate picking the right timepoint to conduct salinity pressure to adequate microalgal cell density.

For the downstream processing of carotenoid production, it is particularly critical to distinguish microalgal cells from other contaminants in the culture medium. Since traditional turbidimeters detect both microalgal cells and pollutants, the measurement results are often not accurate enough [104]. As fluorescence only comes from chlorophyll pigment of alive cells, convenient chlorophyll fluorescence sensors have been designed and widely applied in in situ detection of microalgal cells with high efficiency and accuracy [105,106].

Take the astaxanthin production from *C. zofingiensis* as an example. *C. zofingiensis* can synthesize astaxanthin in adversity and store it in lipid droplets [55]. As it can take up glucose for rapid growth, it is more likely to be contaminated by bacteria during the actual cultivation process. Chlorophyll fluorescence sensors can visibly separate microalgae from bacteria and cell debris, which can be applied in quantification of microalgae. Additionally, bioproducts content can be non-invasively detected *in situ*, such as with a triacylglycerides (TAG) observer [107]. As for cultivation with salt-stress treatments, the IoT will facilitate in distinguishing cell debris under salt stress to accurately harvest and quantify microalgal

cells, and a TAG observer can also display astaxanthin content to achieve preliminary evaluation of productivities. The IoT, as an intelligent system, can also choose the best extraction method after the target product has been separated, which saves time while ensuring purity and extraction efficiency.

#### **5. Future Prospects**

#### *5.1. Genetic Modifications of Microalgae for Salt Tolerance and Carotenoid Accmulation*

Microalgae are widespread and large in number, and only a small part of them have been studied. Therefore, there are still numerous novel microalgae species that may be able to tolerate salinity and accumulate carotenoids. Here, we mainly focus on three methods to get expected microalgae strains: (1) adaptive evolution, (2) random mutagenesis and (3) targeted genetic engineering.

Different to strain isolation, adaptive evolution can obtain an expected strain with a target phenotype. Adaptive evolution has been widely utilized for strain improvement to stress tolerance in various microalgae species, such as *C. reinhardtii* for 200 mM salt tolerance [108] and *Chlorella sp.* for high phenol concentration [109]. It was performed for improving the tolerance of a freshwater strain *Chlorella sp.* AE10 to 30 g/L salt for 138 days (46 cycles). The genes of the resulting strain *Chlorella sp.* S30 related to Calvin–Benson cycle, C4-dicarboxylic acid cycle and crassulacean acid metabolism (CAM) pathways were upregulated, which was beneficial for CO2 fixation under salt stress [85]. However, salinity acclimation of microalgae involves different functional genes and numerous pathways, which makes the exploitation of specific genes in salt tolerance more difficult.

Chemical mutagens, such as N-methyl-N'-nitro-N-nitrosoguanidine (MNNG) and ethyl methane sulfonate (EMS), are effective in mutant generation and may lead to mutants with enhanced carotenoid accumulation [110]. When genes in the carotenoid synthesis pathway (shown in Figure 1) are mutated, carotenoid profiles in microalgae may change. By chemical mutagenesis with EMS, carotenoid content in *Coelastrum sp.* C1-G1 was increased about 2-fold over its mother strain [39]. The Astaxanthin-overproduction strain *H. pluvialis* MT 2877 is a mutant by MNNG, which produced 4-fold astaxanthin over the WT strain [111]. *C. zofingiensis bkt1*, a chemical mutagen by MNNG, can accumulate high amounts of three essential carotenoids, e.g., zeaxanthin (7.00 ± 0.82 mg/g), lutein (13.81 ± 1.23 mg/g) and β-carotene (7.18 ± 0.72 mg/g) under different cultivation conditions [33], which can serve as a competent option for large-scale cultivation. *Chlorella sp.* was irradiated by 137Se- γ ray and domesticated with a seawater culture medium (salinity 3% wt.) under 15% CO2 stress. The biomass yield of the mutant was increased by 25% with 54.9% DW of lipids [112]. Although random mutagenesis could lead to novel traits, unpredictable traits may also show up, and the mutated genes still need further verification, such as gene sequencing, knockout and retro-complementation.

Targeted genetic engineering could generate specific insertions, deletions or substitutions in the host genome while avoiding random results [113]. Up until now, a lot of genes related to salt responses have been recognized, such as *nhaP* encoding a Na+/H+ antiporter, *codA* encoding choline oxidase to synthesize compatible solutes and *Dps* encoding DNA-binding proteins (Dps). Their introduction or overexpression strains have shown enhanced salt-tolerant abilities and potentials for seawater cultivation, which have been comprehensively reviewed [114].

Genetic engineering of microalgae for carotenoid hyperaccumulation is now a mature technology [115]. Genetic engineering of the green algae *C. zofingiensis* with a modified norflurazon-resistant endogenous *pds* gene resulted in up to 54.3% higher astaxanthin [116]. Transformation and expression of *dxs* and *psy* genes in *P. tricornutum* can increase fucoxanthin content by 2.4-fold and 1.8-fold, respectively [117]. *H. pluvialis* can also act as a gene donor. Transgenic *D. salina* with *bkt* and *chyb* genes from *H. pluvialis* was capable of astaxanthin biosynthesis with a better tolerance to high light [118].

At present, several microalgal genomes sequences are available in the public databases, such as *C. reinhardtii*, *Chlorella variabilis*, *Nanochloropsis graditana*, *C. zofingiensis*, etc. [119]. This offers worthy opportunities for precise engineering technology. Novel molecular tools have been applied to modify microalgal traits, including advanced genome editing systems, such as clustered regularly interspaced palindromic repeats and associated proteins (CRISPR-Cas), transcription activator-like effector nuclease (TALEN), zinc-finger nuclease (ZFN) and RNA interference (RNAi) [10,116]. For example, knocking down zeaxanthin epoxidase gene (*zep*) in C. *reinhardtii* by using DNA-free CRISPR-Cas9 RNP-mediated mutagenesis can increase zeaxanthin content by 56-fold [120].

#### *5.2. Co-Production of Carotenoids with Value-Added Products from Microalgae*

Lipids and several carotenoids (such as astaxanthin) are secondary metabolites of microalgae under adversity, which means certain stimulating conditions may promote the accumulation of two products simultaneously. In *H. pluvialis*, astaxanthin esterified by FAs is stored in lipid bodies, where TAGs are also positioned. Additionally, the biosynthesis of FAs and TAG has been demonstrated to be correlated with astaxanthin biosynthesis in *H. pluvialis* [121]. By adding 6 μM Cu2+, the astaxanthin and lipid content was increased by 66.87 and 34.99%, respectively, with carotenogenic and lipogenic genes being upregulated at transcriptional level [122].

Microalgae also contain biofunctional unsaturated fatty acids. Omega-3 polyunsaturated fatty acids (Omega-3 PUFA) from microalgae, such as eicosapentaenoic acid (EPA. C20:5) and docosahexaenoic acid (DHA, C22:6), have been proved to have various health effects with dietary intake, lowering the risks of neurodegenerative diseases and cardiovascular disorders [123]. *Nannochloropsis oceanica* oil is rich in EPA, while its zeaxanthin content (30.2 mg/g DW) is relatively high among microalgae species [41], making it a potential species for co-production of zeaxanthin and EPA. *O. aurita* also contains high concentrations of EPA (28% of total fatty acids), which has been confirmed for large-scale culture in open ponds [47], making it competent for producing both fucoxanthin (18.47 mg/g DW) and EPA.

In addition, co-production of fucoxanthin and DHA is feasible in microalgae. The marine algae genus *Isochrysis* is rich in DHA, and it allows easy extraction of DHA and fucoxanthin as it has no cell wall. Sun et al. screened 16 strains of *Isochrysis* for DHA and fucoxanthin production, and *Isochrysis* CCMP1324 in semi-continuous cultivation excels with productivities as 9.05 and 7.96 mg/L/day of DHA and fucoxanthin, respectively [124]. *I. zhanjiangensis* accumulated both high content of fucoxanthin (23.29 mg/g DW) and stearidonic acid (SDA, 17.16% of total fatty acid), which is the common precursor of EPA and DHA [46].

Additionally, both cultivation strategies and efficient extraction methods are worth noting. As zeaxanthin and lipid are soluble in organic solvents, choosing an efficient extraction system, such as hexane: methanol (3:2), can realize the best extraction efficacy for both EPA (36.4 mg/g DW) and zeaxanthin (28.2 mg/g DW), which also avoid the usage of toxic chloroform [41]. As most fucoxanthin was mostly in the hydroalcoholic phase (over 99%), lipid and fucoxanthin can be separated by this biphasic system after the same extraction process with pure ethanol solvent [50].

The residual biomass after carotenoid extraction of microalgae can be further processed to harvest other by-products. Under nutrient starvation, *D. salina* accumulated β-carotene and triglycerides up to 30–60% DW [125]. After carotenoid extraction, the residuals can be employed for co-production of biodiesel after transesterification. In addition, the carbohydrate in *D. salina* can also be employed for bioethanol production [126].

#### **6. Conclusions**

In this review, we comprehensively introduced health-promoting carotenoids in microalgae, summarized the carotenoid biosynthetic pathways and discussed microalgal responses to salt stress. Advances in applying the Internet of Things (IoT) in modern biotechnology and co-production of carotenoids with other high-value products were also considered in detail. Through in-depth research on the effects of salt stress on microalgae, it is believed that the cultivation of freshwater microalgae in seawater will no longer be an obstacle in the future. Moreover, it is also important to exploit more potential salt-tolerant microalgae species for high-yield production of biofunctional carotenoids.

**Author Contributions:** Conceptualization, resources and supervision, F.C. and J.H.; writing original draft preparation, Y.R. and J.D.; writing—review and editing, H.S., J.H. and F.C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the National Natural Science Foundation of China (Project 31471717), Key Realm R&D Program of Guangdong Province (No. 2018B020206001), Guangdong Province Zhujiang Talent Program (No. 2019ZT08H476) and the Science and Technology Innovation Commission of Shenzhen (No. KQTD20180412181334790).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

## *Review* **Microalgae as Sustainable Biofactories to Produce High-Value Lipids: Biodiversity, Exploitation, and Biotechnological Applications**

**Tomásia Fernandes 1,2 and Nereida Cordeiro 1,2,\***

**Abstract:** Microalgae are often called "sustainable biofactories" due to their dual potential to mitigate atmospheric carbon dioxide and produce a great diversity of high-value compounds. Nevertheless, the successful exploitation of microalgae as biofactories for industrial scale is dependent on choosing the right microalga and optimum growth conditions. Due to the rich biodiversity of microalgae, a screening pipeline should be developed to perform microalgal strain selection exploring their growth, robustness, and metabolite production. Current prospects in microalgal biotechnology are turning their focus to high-value lipids for pharmaceutic, nutraceutic, and cosmetic products. Within microalgal lipid fraction, polyunsaturated fatty acids and carotenoids are broadly recognized for their vital functions in human organisms. Microalgal-derived phytosterols are still an underexploited lipid resource despite presenting promising biological activities, including neuroprotective, antiinflammatory, anti-cancer, neuromodulatory, immunomodulatory, and apoptosis inductive effects. To modulate microalgal biochemical composition, according to the intended field of application, it is important to know the contribution of each cultivation factor, or their combined effects, for the wanted product accumulation. Microalgae have a vital role to play in future low-carbon economy. Since microalgal biodiesel is still costly, it is desirable to explore the potential of oleaginous species for its high-value lipids which present great global market prospects.

**Keywords:** sustainability; industrial valorization; carbon dioxide fixation; biological activities; polyunsaturated fatty acids; phytosterol; carotenoids

#### **1. Introduction**

Microalgae use light energy and inorganic nutrients to produce oxygen and biomass rich in a diversity of value-added compounds (Figure 1) [1]. They can thrive in almost all environments and can be found in oceans, brackish water, freshwater, rocks, and soils [2]. Microalgal biotechnological historical data go back to the Aztec population, who harvested *Arthrospira* from lake Texcoco for food purposes [2,3]. *Arthrospira* has also been collected by local people surrounding Lake Chad and consumed as a nutritional supplement called "dihe" [2]. *Nostoc* species have been used by the Chinese as a food delicacy and for their properties for hundreds of years [2].

*Arthrospira platensis*, *Aphanizomenon flosaquae* var. flosaquae, *Chlorella luteoviridis, Chlorella pyrenoidosa*, *Chlorella vulgaris*, and *Auxenochlorella protothecoides* have been on the market as a food or food ingredient and consumed to a significant degree before 15 May 1997 in the European Union market; thus, its access to the market is not subject to the Novel Food Regulation (EU) 2015/2283 [4]. Dried *Tetraselmis chuii*, *Odontella aurita*, and astaxanthin-rich oleoresin from *Haematococcus pluvialis* are microalgal products approved

**Citation:** Fernandes, T.; Cordeiro, N. Microalgae as Sustainable Biofactories to Produce High-Value Lipids: Biodiversity, Exploitation, and Biotechnological Applications. *Mar. Drugs* **2021**, *19*, 573. https://doi.org/ 10.3390/md19100573

Academic Editor: Carlos Almeida

Received: 23 September 2021 Accepted: 11 October 2021 Published: 14 October 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

as a novel food and, as the microalgae listed previously, are within the list of microalgae that can be commercialized in the EU [5]. Through Figure 2a, it is possible to visualize that in most European countries, algae farmers mainly produce microalgal species belonging to Cyanobacteria and Chlorophyta phyla, except for Belgium (BE), Norway (NO), and Sweden (SE). This is consistent with the microalgae approved for human consumption which belong to Chlorophyta and Cyanophyta phyla, except for *Odontella aurita*, which belongs to Bacillariophyta phylum.

**Figure 1.** Uncovering the potential of a microalga as a biofactory. C—carbohydrates; P/E—protein/enzymes; L—lipids; V—vitamins; Pig.—pigments.

**Figure 2.** Microalgae production in Europe: (**a**) relative abundance of microalgae at phylum level produced by algae farmers (AT—Austria; BE—Belgium; BG—Bulgaria; CH—Switzerland; CZ—Czech Republic; DE—Germany; DK—Denmark; EE— Estonia; EL—Greece; ES—Spain; FO—Faroe Islands; FR—France; GR—Greenland; HU—Hungary; IC—Iceland; IE—Ireland; IT—Italy; LV—Latvia; NL—the Netherlands; NO—Norway; PT—Portugal; SE—Sweden; UK—the United Kingdom). (**b**) Number of algae farmers against main microalgae phyla and diversity of species exploited. (**c**) Diversity of Chlorophyta species produced by algae farmers. Based on EMODnet database [6].

Health, energy, and human nutrition are the three main applications for microalgal products [7]. However, the energy production from microalgae is experiencing a slow growth compared to other segments [7]. Through Figure 3a, it is possible to observe that both patents and research activities presented an exponential increase between 2004 and 2014. This trend continued for research activities, in contrast to patent publications. These waves of microalgae-related research and development activities are mainly associated with energy-driven trends, and having as the driving force the high crude oil price [8]. After 2015, this driving force was lost due to the development and popularization of electric cars [8]. Figure 3b shows that research activities were mainly focused on the biological sciences before 1990s. From this decade forward, biotechnology-applied microbiology appeared in the main research categories meeting the slight increase in patent and research publications.

**Figure 3.** Patenting and research activity in the microalgal field: (**a**) numbers of microalgae-related patents and research publications against publication years; (**b**) timeline with the main research activities categories according to Web of Science. The information used to construct these plots using "microalgae" as topic can be found in Espacenet [9] and Web of Science databases [10].

Through Figure 2b, it is possible to visualize that the divisions comprising the greatest diversity of microalgal species industrially exploited are Chlorophyta (18) and Bacillariophyta (10) phyla. According to Griffiths et al. [11] most microalgal species considered for biofuel production are either Chlorophyta or Bacillariophyta, which may explain the previous observation. In the north-west European algae strategic initiatives, the bioenergy (e.g., biodiesel) market is the most mentioned. The energy demand is growing worldwide, especially in the rapidly developing countries such as China and India [12]. Biofuels currently account only 1.9% of global transport fuel consumption and are projected to achieve a threefold increase over the next 20 years [13]. Brazil's ethanol, derived from agricultural crops, is the most price-competitive biofuel in the world, which reflects the large investment of governmental agencies in research and technology that allowed the improvement of production processes, which in turn lowered biofuel manufacturing costs [14]. However, the food or fuel controversy derived from using agricultural crops for biofuel prompt the search for alternative sources as microalgae. Although these microorganisms have several advantages such as that they have a rapid growth rate, an ability to readily adapt to a wide range of climatic conditions, and that they do not compete for arable lands, the high initial capital investment and high biofuel production costs makes its exploitation for bioenergy purposes not feasible [15]. According to Moshood et al. [13], an economically efficient policy assistance may be required to overcome the main challenges found for microalgal biofuel production.

Ongoing efforts have been performed to reduce microalgal production costs; these include the evaluation of reactor design approaches, the development of biorefinery ap-

proaches, and the use of low-cost inputs for microalgal production. The use of wastewater for microalgal cultivation can offset the cost of freshwater and nutrients delivering environmental benefits such as the recycling of resources and reducing the nutrient discharges responsible for eutrophication in water bodies [16,17]. Numerous studies have evaluated the feasibility of microalgae cultivation for lipid production using wastewater from different sources: municipal [18–20], industrial [21], mining activity [22], landfill leachate [23], agricultural [24], and fish farm effluents [25]. Nevertheless, further research is needed to solve the drawbacks of the simultaneous microalgal biomass/product production and wastewater treatment, namely the everchanging chemical content of wastewater; the fact that not all metal ions and contaminants can be reduced using microalgae; and the need of wastewater pre-treatment (e.g., anaerobic digestate) to reduce organic compounds concentration, water turbidity, and low transparency, which affect microalgal growth [21,26].

According to the International Energy Agency [27], global energy-related carbon dioxide (CO2) emissions remained at 31.5 Gt, despite the decline in 2020. CO2 contributes up to 68% of the total greenhouse gases, the accumulation of which in the atmosphere has been considered as the main driver of climate changes [28]. Presenting higher carbon dioxide fixation efficiencies than terrestrial plants, microalgae have a pivotal role to play in future low-carbon economy [28,29]. In the literature, it is estimated that producing 280 t of microalgal dry biomass per ha per year using 9% of the incoming solar energy fixes roughly 513 t of CO2 [30]. In this sense, microalgae are often called "sustainable biofactories" due to their dual potential to mitigate/bioremediate atmospheric CO2 and produce a wide array of high-value compounds, which can be further enhanced through induced changes in its growth conditions.

Current prospects of algal biotechnology are turning their focus to high-value lipids production, namely polyunsaturated fatty acids (PUFA), which can be used in dietary supplements, functional food, pharmaceutical, and infant formula segments [31]. Some companies already produce ω3-PUFA with microalgal origin as dietary supplements or food ingredients (Oceans Alive (USA), Blue Biotech (Germany), Flora Health (USA)— *Nannochloropsis*; InnovalG (France)—*Odontella*) [32]. The global market size for ω3-PUFA in 2020 was estimated at US \$16.2 billion, and its 2027 value projections are expected to reach US \$36.9 billion at a compound annual growth rate (CAGR) of 12.5% over the forecast period of 2020–2027 [33].

Within Chlorophyta phylum, it is possible to observe that most algae farmers have been focusing on the production of oleaginous microalgae species, namely from *Chlorella*, *Tetraselmis*, *Botryococcus*, *Scenedesmus* genera, which are more specialized for biofuel production [7]. *Chlorella* sp., *Haematococcus pluvialis*, *Tetraselmis* sp., and *Chlorella vulgaris* are on the first line of microalgal species produced by algae farmers (Figure 2c). These microalgae have in common its versatility which allows to apply them in food, feed, energy production, and as a source of high-value molecules, which may reflect the efforts of algal farmers to target more than one market [34–36]. Furthermore, species belonging to *Haematococcus* and *Dunaliella* genera are mainly used for pigments exploitation, namely for astaxanthin and *β*-carotene production, respectively [7]. Protein is on the first line of development in human nutrition and health sectors, followed by pigments and lipids [7]. Accounting for the large number of oleaginous species already produced by algae farmers and the potentialities of microalgal lipids for high-value market, this review outlines the potential of microalgal high-value lipids for dietary supplements, cosmetics, and pharmaceutics, along with its health-promoting activities and optimization strategies.

#### **2. Species Selection and Exploitation**

Microalgae present a rich biodiversity comprising 40,000–50,000 described species with an estimation of nearly 800,000 existing species [37]. Chlorophyta comprises 6952 algal species [38], and Bacillariophyta is the most diversified group within microalgae, with more than 10,000 diatom species being described [39]. Despite this great diversity, only a few microalgal species have been exploited for biotechnological applications, with

only 18 species of Chlorophyta and 10 species of Baccilariophyta phyla being produced by European algae farmers (Figure 2b). A smaller number of species is recorded for the other phyla: Cyanobacteria, Ocrophyta, Haptophyta, Rhodophyta, Euglenozoa, Charophyta, Miozoa, and Prasinodermatophyta.

The industrial production of microalgae heavily depends on biomass productivity, which is the most significant factor that can reduce the production cost levels [40]. To increase microalgae productivity, several strategies have been developed, namely the exploitation of the cultivation conditions to direct the metabolism towards desired product accumulation, selection and breeding of strains with increased biomass productivity, and genetic modification [40].

The selection of the right microalga considering specific culture conditions and desired product content could be performed through an exhaustive screen of scientific data through databases and literature, followed by physiological data collection (Figure 1). With respect to the product synthesized by microalgae, they can be applied in several fields from bioenergy (low-value market) to cosmetics and pharmaceutics (high-value market).

When exploiting the great biodiversity of microalgae as new natural sources of highvalue phytochemicals, a screening pipeline should be developed including the following key aspects: *(i) growth*—for the successful improvement and progression of microalgaebased industries, the discovery and improvement of new fast-growing strains is essential [41]. Thus, the study of the maximum growth rate, maximum cell density, and dry cell biomass at different growth conditions, along with their amenability for heterotrophic/mixotrophic growth, are critical features to predict the feasibility of microalgae for large-scale production [42]; *(ii) robustness*—the selected microalgae should withstand variable local climatic conditions and be resistant to possible infections (e.g., other algae strains, grazers, bacteria, or viruses) in order to prevent large-scale crashes [43]. Nevertheless, strategies have been developed to combat and prevent contamination in microalgae cultivation such as the use of extreme conditions to create an unfavorable environment for the competitive organisms of the microalgae [44,45], and the use of allelopathic approaches to control microalgae cultivation [46]; *(iii) metabolite production*—in the production of microalgae for food and health purposes, the potential toxicity (phycotoxins) of some species as well as desired product (high-value products) accumulation should be considered.

#### *2.1. Phycotoxins*

Phycotoxins are causative agents of seafood-borne poisoning syndromes (e.g., ciguatera fish poisoning) in humans [47]. From the wide microalgal diversity, only around 200 species are health-threatening, with the main toxic microalgae belonging to dinoflagellates and diatoms groups [48]. Moreover, contamination of microalgae-based products resultant from unsuitable location of cultivation ponds (e.g., inflows of effluents containing pollutants) is a concern for human health [49]. Therefore, standard guidelines provided by international regulatory organizations, such as the Food and Drug Administration (United States of America) and European Food Safety Authority (Europe), guarantee that microalgae-based industries operate in conformity with safety requirements [48]. Through Table S1, is possible to have a brief insight on the regulations and directives applied in the European Union.

#### *2.2. High-Value Products*

Microalgae can be produced targeting different fields of application such as food, feed, pharmaceutical, personal care, and biomedicine (Figure 1). This versatility is derived from its ability to synthesize a multiplicity of metabolites distributed among pigments, vitamins, carbohydrates, lipids, and proteins/enzymes. Moreover, several biological properties attributed to microalgae, namely anti-inflammatory [50,51], anti-pyretic [51], and anticancer [52] activities, have shown its potential to high-value compound production.

Using the Cosmetic Ingredient Database (CosIng) [53], it is possible to see that several substances with microalgal origin have already been authorized, namely the oils from *Odontella aurita*, *Nannochloropsis oceanica*, *Chlorella minutisssima*, *Chlorella protothecoides*, and *Haematococcus pluvialis*. These can be incorporated into cosmeceuticals functioning as skin-conditioning emollient, skin protection, and anti-oxidants in cosmetic products [53]. Moreover, when analyzing research activities performed on microalgal production for food and health purposes, two main groups of metabolites stand out: fatty acids and carotenoids (Figure 4). These are mainly related to the lipid-soluble fraction of microalgae.

**Figure 4.** Concept's network obtained with VOSviewer software [54] for the research on "microalgae AND food AND health" in Web of Science database [10].

#### **3. Polyunsaturated Fatty Acids (PUFA) Exploitation from Autotrophic Microalgae**

PUFA are broadly known for their vital functions in human organisms [55]. For instance, docosahexaenoic acid (DHA, C22ω3) is enriched in human milk, plasma, and sperm, and along with arachidonic acid (AA, C20:4ω6) is concentrated in the membrane lipids of gray matter and in the visual elements of retina [56]. There is a lot of evidence that an adequate supply of these fatty acids improves visual acuity and infant cognitive development [56]. Moreover, several therapeutic properties have been attributed to the consumption of these fatty acids such as reduced risk of arthritis and cardiovascular diseases [57]. In contrast to microalgae, mammals do not have the ability to convert oleic acid (C18:1ω9) to the precursors of long-chain polyunsaturated fatty acids (LC-PUFA) biosynthesis pathway, and they poorly synthesize C20-C22 PUFA from dietary linoleic acid (LA, C18:2ω6) and *α*-linolenic acid (ALA, C18:3ω3).

#### *3.1. PUFA—Synthesis by Microalgae*

LC-PUFA biosynthesis pathways by microalgae are initiated by Δ12 desaturation of C18:1ω9, producing LA, which might be further desaturated by a ω3-desaturase generating ALA (see Figure 5) [58]. The ω3-pathway is initiated with the Δ6 desaturation of LA and leads to the synthesis of ω3-LC-PUFA eicosapentaenoic acid (EPA, C20:5ω3) and DHA, whereas the ω6-pathway initiates with the Δ6 desaturation of ALA and produces the ω6-LC-PUFA, AA [58]. However, some EPA-producing eustigmatophytes, such as *Nannochloropsis* sp. and *Monodus subterraneus*, are thought to preferentially synthesize EPA via the ω6-pathway by the action of a ω3-desaturase, which catalyzes the conversion of AA to EPA [58,59].

**Figure 5.** Biosynthesis of long-chain polyunsaturated fatty acids in microalgae [55,58,60]. ΔxD— "front-end" desaturase, adds a double bond at position x from carboxyl end; ωyD—"methyl-end" desaturase, adds a double bond at position y from methyl end; E—elongase, which catalyzes carbon chain extension; PKS—polyketide synthase.

As with other organisms, the microalgal fatty acid composition is known to vary among the different phylogenetic groups [58]. In Figure 5, is possible to visualize an alternative route for LC-PUFA biosynthesis: Δ8 desaturase pathway. According to Khozin-Goldberg [55], this pathway is known to exist in some microalgae, namely in haptophytes *Isochrysis galbana, Pavlova salina*, and *Emiliana huxleyi*, and the Euglenophyte *Euglena gracilis*. For the DHA-producing haptophyta *Isochrysis galbana*, a gene encoding a C22-Δ4 polyunsaturated fatty acid specific desaturase has been isolated and characterized [61]. The Trebouxiophyceae *Lobosphaera incisa* is a rare case in which AA is the major product of LC-PUFA biosynthesis in microalgae [55].

#### *3.2. PUFA Role in Human Health*

From microalgal lipids, PUFA are the most studied for their pharmacological potential. In human health, C20-C22 PUFA play important roles in many physiological and pathological processes [57]. Moreover, most of PUFA health-benefits are due to their key roles as lipid mediators in inflammatory processes and as important compounds for growth and development. EPA and DHA are parent compounds of specialized pro-resolving lipid mediators (protectins, resolvins, and maresins) that act as inflammatory brakes and promote the return of the affected site to homeostasis (Figure 6) [62,63]. As with ω3-PUFA, AA-derived lipoxins and their carbon-15 position epimers have beneficial effects on inflammation and resolution [62].

**Figure 6.** Lipid mediators of inflammatory process derived from arachidonic acid (AA), eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA) [63,64]. EETs—epoxyeicosatrienoic acids; HETEs—hydroxyeicosatetraenoic acids; PG prostaglandin.

> EPA competitively inhibits the utilization of AA by cyclooxygenasse/lipoxygenases to less pro-inflammatory mediators [65]. In contrast with eicosanoid products from AA (prostaglandin E2, thromboxane A2, and leukotriene B4), EPA-derived eicosanoids (thromboxane A3, and leukotriene B5) are weak inducers of inflammation and have attenuated platelet-aggregating and vasoconstriction abilities [65]. In mammalian cells, ω6- and ω3-fatty acids are not interconvertible because they lack ω3-desaturase; therefore, their balance in the diet is important [65]. In the Western diet, the greatest amounts of ω6-fatty acids (ω6:ω3 of 20:1) drive higher levels of AA eicosanoids products (e.g., prostaglandins, thromboxanes, leukotrienes, and hydroxy fatty acids), causing an imbalance between pro- and anti-inflammatory molecules and shifting the physiological state to one that is proinflammatory, prothrombotic, and proaggregatory [65]. Thus, a balanced intake of ω6 and ω3-fatty acids is crucial.

#### *3.3. Microalgae—PUFA Enhancement Strategies*

Although the largest share of the EPA/DHA oil market comes from wild fish, the declining fish stocks and susceptibility to contamination by pollutants (such as mercury) have turned the attention of PUFA exploitation to microalgae [66]. In this sense, the oxidative stability, sustainability, suitability for vegetarians, and the absence of fishy taste/smell are some of the advantages that make microalgae a feasible source for PUFA commercialization [66]. However, the immature production process is one of the weaknesses that must be surpassed for making PUFA exploitation from microalgae a feasible process [66].

New competitors of PUFA exploitation from microalgae are DHA producers belonging to thraustochytrids (*Traustochytrium* sp. and *Aurantiochytrium*) [66]. These unicellular heterothrophic organisms have already been authorized for food/feed/nutraceuticals and are marketed by DSM/Evonik and Source-Omega companies [66]. However, some disadvantages from the exploitation of these organisms for ω3-production (mostly produce DHA; the production chain increases pressure on arable land—since heterotrophic organisms need a sugar input to grow, and produces CO2) are opportunities for the production of ω3-fatty acids from microalgae [66].

Since the microalgae composition is known to vary with growth conditions, the study of strategies for microalgae PUFA enhancement is crucial to overcome the challenge of the undeveloped production process of phototrophic PUFA exploitation from microalgae. Although several strategies have been proposed to enhance microalgae lipids production, most of these have been projected for biofuel production. However, nowadays, more attention has been given towards PUFA production. Conventional approaches for enhancing microalgae lipid accumulation include nutrient stress (e.g., alterations in nitrogen, phosphorus, and carbon supply) or changes in cultivation conditions (e.g., light, and temperature) [67]. The main advantage of nutrient stress is its easy applicability at both lab and large-scale cultivation, while cultivation conditions such as light have high operational costs and are not easy to control within open cultivation systems [67]. Within nutrient regime alterations, nitrate limitation is a commonly employed strategy to enhance microalgal lipids quantity [67].

As essential constituent of proteins, nucleotides, vitamins, and coenzymes, any changes in the nitrogen source and concentrations can trigger growth changes and biochemical remodeling in microalgae species [68–70]. Although the study of Huang et al. [71] had the purpose of improving lipids properties of the microalgal strains *Tetraselmis subcordiformis* SHOU-S05, *Nannochloropsis oculata* SHOU-S14, and *Pavlova viridis* SHOU-S16, the insights on the fatty acid composition at different nitrogen concentrations enables to see some trends with respect to PUFA accumulation. Therefore, towards high nitrogen supplementations, the lipid content decreased, whereas the PUFA proportion increased for *N. oculata* and *P. viridis*. For *T. subcordiformis*, the highest PUFA percentage was registered at lower nitrogen supplementations as with the highest lipid content. As with Sukenik [72], this seems to suggest that the growth conditions for the maximization of PUFA production are similar to the conditions required to maximize biomass production for *N. oculata* and *P. viridis*.

Other cultivation conditions affecting microalgal growth and chemical diversity are the salinity, light intensity, and photoperiod. Mitra et al. [73] studied the effects of these factors in *Nannochloropsis gaditana* CCNM1032 strain, and concluded that the most positive factor for fatty acid enhancement was the photoperiod. In this study, maximal EPA productivities were achieved at 60 μmol photons m−2 s−1 and at a photoperiod regime of 18 h:6 h (light:dark); this observation was made for 1L cultures.

Two-stage cultivation and combined nutrient and abiotic stresses are novel approaches used to enhance the microalgal biochemical composition. In the two-stage cultivation, microalgae are first grown to gain higher biomass and are then exposed to different cultivation conditions to trigger the accumulation of desired product content. For *Nannochloropsis gaditana* IMTE1, a two-stage cultivation was studied by Xiao et al. [74]. This microalgal strain was firstly grown in batch culture for 6 days, washed, and then transferred for a chemostat culture with a fixed dilution rate and adjusted nitrate concentrations. In these cultivation conditions, the highest biomass (897.10 mg dw L−1; dw—dry weight) and EPA content (2.62% dw) were obtained at high nitrogen supplementations.

When combining nutrient and abiotic stresses, it is important to know the importance of each factor for the desired product accumulation, as well as its synergistic effects [67]. With the purpose of optimizing ω3-fatty acid production by *Pavlova lutheri*, Carvalho and Malcata [75] studied the combined effects of the dilution rate, light intensity, and CO2 concentration under continuous mode. The optimum conditions for EPA and DHA

production were found in cultures supplied with 0.5% CO2, at a dilution rate of 0.297 d−1 and a light intensity of 120 μE m−2 s−1 [75]. Other combined nutrient and abiotic stresses are summarized in Table 1 for the following strains: *Chlamydomonas reinharditii* CC124, *Nannochloropsis gaditana* CCNM1032, *Phaeodactylum tricornutum* CS-29C, and *Chaetoceros muelleri* CS-176. The major disadvantage of the enounced novel approaches is that largescale trials are required.

**Table 1.** Some strategies used for microalgae lipid enhancement and its impact on polyunsaturated fatty acids accumulation.

PUFA—polyunsaturated fatty acids; DHA—docosahexaenoic acid; EPA—eicosapentaenoic acid; TFA—total fatty acids; dw—dry weight.

#### **4. Sterols as an Underexploited Lipid Resource from Microalgae**

When linking microalgae with food and human health, the predominant terms are fatty acids and carotenoids (Figure 4). Nevertheless, some microalgae are high-level producers of phytosterols, which have been playing a key role in the functional food market [79,80]. In addition, mixtures of phytosterols can function as skin conditioning in cosmetic products (creams and lipstick), and in pharmaceutics, they are gaining interest for the production of therapeutic steroids [53,81].

Plant-derived phytosterols have been added to food products for their ability to reduce serum cholesterol levels and prevent coronary heart diseases [79,82]. Cyanophya *Nostoc commune* var. *sphaeroides* lipid extracts have been found to inhibit the expression of key regulatory genes involved in cholesterol and fatty acid biosynthetic pathways; this property could contribute to lower serum cholesterol as well as triglyceride concentrations [83]. Phytosterols occur in four common forms: as free sterols (FS), as fatty acid esters (sterol is esterified to fatty acid; SE), as steryl glycosides (bound to sugar with a glycosidic bond; SG), and as acylated steryl glycosides (sugar moiety is acylated with a fatty acid; ASG) [84]. Due to the poor solubility of free phytosterols, major phytosterol/phytostanol products being marketed are in their conjugated forms (SE, SG, and ASG) as it is easier to add them into food products [84]. Table 2 summarizes information with respect to some brands that commercialize phytosterol fortified food products (yoghurts, spreads, soft cheese, and drinks), and other phytosterol-based products (supplements and paste). Although the raw materials used for phytosterols isolation for the food industry are tall oil (fatsoluble by-product obtained from trees) and vegetable oil, the scarcity of land resources has pulled the attention of scientific and industrial communities towards the search for new sustainable natural sources of phytosterols [85,86]. The concentration of phytosterols in different vegetable oils was estimated by Yang et al. [87] and ranged between 142.64 (camellia oil) and 1891.82 (rice bran oil) mg 100 g−1.

\* Foods with plant sterols and commercialized include emulsified sterols (ES200); fine particle sterols (FP100); granular phytosterols (FP300); steryl esters (SE-C100); water dispersible steryl esters (WDSE-33).

The potential of Chlorophytes *Dunaliella tertiolecta* and *Dunaliella salina* as sources of phytosterols was studied by Francavilla et al. [94]; in its study, the highest yields of total sterols were 1.3% and 0.89% dw for *D. tertiolecta* and *D. salina*, respectively. Another promising microalgal strain for phytosterol production, investigated by Ahmed et al. [79], was *Pavlova lutheri*, with the phytosterol content reaching up to 5.1% dw. Comparing these values with the ones previously mentioned for vegetable oils, it is possible to recognize that microalgae have the potential to become a useful alternative source of phytosterols to use as functional ingredients. Microalgal-derived phytosterols are mainly distributed among four groups: 4-desmethyl-Δ5-sterols; 4-desmethyl-Δ7-sterols; 4-methylsterols; and dihydroxylated sterols [80], with 4-desmethyl-Δ5-sterols being the predominant phytosterol in microalgae [80,95]. According to Moreau et al. [84], 4-desmethyl sterols and stanols have been shown to inhibit the uptake of cholesterol from the intestine, resulting in a decrease of serum cholesterol levels. This once again strengthens the potential of using microalgae-phytosterols as a novel industrial application.

#### *4.1. Sterol Synthesis by Microalgae*

For microalgae, there are two distinct and compartmentalized pathways for isoprenoid synthesis; these are (i) the mevalonic acid (MVA) pathway, in the cytosol; and (ii) the 1-deoxy-D-xylulose-5-phosphate/2-C-methyl-D-erythritol-4-phosphate (DOXP/MEP) pathway, in the plastid [80,96]. In general, microalgae possess both DOXP/MEP and MVA pathways [96]. Exceptions include Cyanophytes, Chlorophytes, as well as the Bacillariophyta *Haslea ostrearia*, and the Rhodophyta *Cyanidioschyzon merolae*, which produce sterols only from the MEP/DOXP pathway [97,98]. Figure 7 shows a generalized overview of sterol biosynthesis pathway for microalgae. Therefore, sterol biosynthetic pathway can be split into three main stages: (i) biosynthesis of isoprenoid precursors—isopentenyl pyrophosphate (IPP) and its isomer dimethylallyl pyrophosphate (DMAPP); (ii) biosynthesis of polyprenyl pyrophosphates such as farnesyl pyrophosphate (FPP); and (iii) squalene (precursor of all phytosterols) formation—dimerization of FPP [97]. According to Sasso et al. [97], there is evidence for the transport of IPP and polyprenyl pyrophosphates, such as FPP and geranyl pyrophosphate (GPP), across plastid membranes. Therefore, in algae such as Chlorophytes, in which sterols are only produced through MEP/DOXP pathway, IPP synthesized in the plastids could be exported to cytosol for the formation of sterols [99].

**Figure 7.** Generalized sterol biosynthesis pathway for microalgae [96,97].

#### *4.2. Microalgae-Derived Phytosterols Biological Activities*

Phytosterols synthesized by microalgae have shown interesting biological activities including neuroprotective, anti-inflammatory, anti-cancer, neuromodulatory, immunomodulatory, and apoptosis inductive effects. From Table S2, it is possible to visualize that the predominant biological activity studied for microalgal-derived phytosterols was antiinflammatory. The anti-inflammatory potential of phytosterols can be assessed in vitro through determination of nitric oxide (NO), prostaglandins (PG) and cytokines production, and/or expression of nitric oxide synthase (iNOS), and cyclooxygenase (COX-2), after cell treatment with inflammation stimulation agents such as concanavalin A (Con A) and lipopolysaccharide (LPS) [100–104]. Promising results were obtained for a sterol rich fraction of *N. oculata* which was found to inhibit NO production, and down-regulate LPSstimulated protein levels of inducible iNOS and COX-2 in a dose-dependent manner [102]. Moreover, a study performed for *D. tertiolecta*, testing several mixtures of phytosterols, showed that ergosterol and a mix of ergosterol and 7-dehydroporiferasterol suppressed a highly pro-inflammatory cytokine (tumor necrosis factor alpha; TNF-α), a pleiotropic cytokine (interleukin (IL)-6), and increased the levels of an anti-inflammatory cytokine (IL-10), showing the anti-inflammatory potential of both sterols and suggesting that phytosterol anti-inflammatory properties might depend on the existence of a synergistic effect of these molecules [101].

Besides anti-inflammatory activity, the ability of phytosterols to cross the blood– brain barrier and act as acetylcholinesterase enzymes inhibitors has sparked the attention of neurodegenerative diseases research [105]. Fagundes et al. [105] studied the neuroprotective potential of *Phormidium autumnale* phytosterol-rich extracts and determined the anti-cholinergic, antioxidant, and anti-inflammatory capacities. In this study, the phytosterol-rich extract demonstrated higher in vitro neuroprotective activity than nonenriched extract, exhibiting a moderate–high anticholinergic potential, and showing to be an effective lipoxygenase inhibitor. This was further supported through molecular docking simulation, which showed the specificity of stigmasterol interaction with acetylcholinesterase active sites. Moreover, a previous study has found in vivo neuromodulatory activity of *D. tertiolecta*—derived phytosterols in selective brain areas of rats [85].

Through Table S2, it is possible to visualize that the biological studies with respect to phytosterols were mainly performed in vitro presenting some limitations, namely, these types of studies only give partial information on bio-functionality, and a lack of systemic factors [106]. However, they provide fast and inexpensive screening of bioactivities, have high sensitivity, and are easy to perform, manage, and interpret [106]. Moreover, it is crucial to highlight that most of the studies summarized in Table S2 are mainly performed in sterol-rich fractions and that future research targeting the potential of the different types of microalgae-derived phytosterols, including their functional activity and synergistic effects, is crucial for gaining in-depth knowledge of microalgae sterols potential.

#### *4.3. Strategies for Sterol Enhancement*

To boost phytosterol accumulation in microalgae, it is crucial to understand its trigger mechanisms. Table 3 summarizes some strategies already employed for inducing microalgae phytosterol accumulation. For Haptophyta *P. lutheri*, the effects of nutrient-induced changes, salinity, ultraviolet-C (UV-C) radiation, and sampling days have been assessed aiming phytosterol production [79,82]. From these variables, the most effective were UV-C radiation and sampling days [79,82]. Although the UV-C radiation equipment is simple and easy for operation and maintenance, this physical stressor has a great disadvantage for large-scale production, which is the significant cell damage connected with UV-C radiation mutagenic factor (as it attacks an organism's deoxyribonucleic acid (DNA)) [107–109]. The mutagenic factor also poses a concern since little is known about the stability of modified algal strains or whether they can potentially take any environmental risks [110].

**Table 3.** Some strategies evaluated for microalgae sterol enhancement.

**Table 3.** *Cont.*

According to Ahmed et al. [79] and Ahmed and Schenck [82], *P. lutheri* did not increased its sterol content when subjected to peroxide hydrogen input, variations in nitrogen and phosphorus concentrations, and changes in salinity. Still, previous studies have described salinity-induced changes as an effective tool to induce phytosterol accumulation in microalgae. Francavilla et al. [94] studied the effect of different salinity concentrations for two Chlorophytas (*D. tertiolecta* and *D. salina*) using a different approach from the study performed to Haptophyta *P. lutheri*, and good yields of total sterols were observed at a lower salt concentration [94]. Thus, the different observations made in both studies might be derived from (i) species-specific differences and/or (ii) the different approaches used to impose salinity-induced changes.

Besides the growth conditions, the specific growth phase at which microalgal biomass is harvested can influence lipid yields and composition for specific purposes [70,114]. The effect of growth phase on the sterol content of dinoflagellate species (*Prorocentrum donghaiense*, *Prorocentrum minimum*, *Karenia mikimotoi*) in batch cultures was assessed by Chen et al. [112]. In this study, the sterol content of two microalgae was susceptible to changes in the growth phase (from exponential to stationary phase), with the greatest increase being verified for dinosterol (168%) in *P. minimum*, and for brassicasterol (423%) in *K. mikimotoi*. When assessing product accumulation over the growth phase, it is also important to take into consideration that there are some growth phases in which the production of stress-associated molecules (e.g., carotenoids, lipids) is still balanced with good amounts of growth-associated ones (e.g., proteins) [115]. This point is especially important for co-exploitation of high-value microalgae products.

Interactive effects of nutrient and abiotic factors have been reported to enhance the production of phytosterol in some freshwater microalgal species [111,116]. For instance, varying light intensities in both high- and low-phosphorus environments was shown to affect sterol accumulation for Chlorophyta *Scenedesmus quadricauda* and *Chlamydomonas globosa*, and Bacillariophyta *Cyclotella meneghiniana*, with sterol contents increasing with light intensity under high phosphorus [111]. Another strategy applied by Piepho et al. [116] was varying the temperature with phosphorus, as well as the silicate supply. For this study, *C. meneghiniana* increased its sterol content from low to high temperature, and this was even higher in the high-phosphorus treatment. Simultaneous effects of nutrient supply and abiotic factors in microalgae highlight the importance of investigating more than just one environmental factor when inducing the accumulation of a desired product for a field of application.

#### **5. Carotenoids**

Some studies have explored the co-production of carotenoids with lipids [117] and fatty acids [118,119] in microalgae. Carotenoids are lipid-soluble compounds synthesized by microalgae and can be divided into carotenes (hydrocarbons) and xanthophylls (oxygenated hydrocarbons) [34,120]. As structural components of light-harvesting complexes, these compounds play key roles within microalgal cells, namely in the protection against excess irradiance, chlorophyll triplets, and reactive oxygen species [121]. Through Figure 8, it is possible to visualize that the global market for carotenoids was USD 1.7 billion in 2020. In microalgae, astaxanthin, *β*-carotene, and lutein are among the key carotenoids with high market potential [34,120,122]. Astaxanthin presents the highest value for the global market size, USD 663.89 million in 2020, and value projections for 2027, USD 977.74 million (Figure 7). *Haematococcus pluvialis* and *Dunaliella salina* are the most popular microalgal species exploited for the commercial production of astaxanthin and *β*-carotene, respectively [34]. In *Haematococcus*, astaxanthin is majorly esterified to palmitic acid (C16:0) and unsaturated fatty acids of the C18 family (C18:1, C18:2, and C18:3) [120,123].

**Figure 8.** Carotenoids global market prospects—market size in 2020, 2027 value projection, and compound annual growth rate (CAGR). The values presented for carotenoids and lutein CAGR correspond to the forecast period of 2020–2027, while for astaxanthin and *β*-carotene, CAGR values correspond to the forecast period of 2021–2027. Data for carotenoids and lutein were collected from StrategyR [33], whereas data for astaxanthin and *β*-carotene were collected from Global Market Insights [124].

Carotenoids are extensively used in food, feed, nutraceuticals, and cosmetics [125]. The consumption of a diet rich in carotenoids is often associated with positive effects on skin health, cancer, cardiovascular, neuronal, and gastrointestinal protection, and vision and immune system enhancement [125]. As with essential fatty acids and phytosterols, carotenoids cannot be synthesized by humans, which, in turn, must obtain these through their diet [125]. The beneficial effects of carotenoids to human health are thought to

be derived from its potent anti-oxidant activity and the provitamin A activity of some carotenoids, which can be converted to retinal (e.g., *α*- and *β*- carotenes) [125].

Currently, astaxanthin is the best biological antioxidant, presenting a free-radical scavenging capacity 65 times more powerful than ascorbic acid (vitamin C) and 54 times stronger than *β*-carotene [126]. This carotenoid is best known for its use in aquaculture, namely for giving the pinkish-red color of salmonids, shrimps, lobsters, and crayfishes. In aquaculture this carotenoid is also used for its positive impacts on organisms' immunesystem and fertility [127]. Astaxanthin has a wide range of applications beyond aquaculture, namely in the food, cosmetic, and pharmaceutical industries [126]. Astaxanthin has several health-promoting properties, and it is used for anti-tumor therapies and prevention, treatment of neural damage interrelated with age-related macular degeneration, Alzheimer and Parkinson diseases [127].

Although natural astaxanthin has a notably higher antioxidant capacity and safety for human consumption, synthetically derived astaxanthin has a low production cost [128]. This makes astaxanthin from natural sources only account for less than 5% of the commercialized astaxanthin [128]. Given the metabolic plasticity of microalgae and its high growth rates, efforts have been displaced for reducing the costs of naturally derived astaxanthin. From astaxanthin producers, *H. pluvialis* is one of the richest sources of natural astaxanthin, accumulating up to 4% of astaxanthin on a dry weight basis. Table 4 summarizes different strategies already applied to *H. pluvialis* strains for prompting astaxanthin accumulation [128].

**Table 4.** Strategies applied for enhancing astaxanthin production for *Haematococcus pluvialis* strains.

Car—carotenoids; N—nitrogen; SA—succinic acid.

As can be seen in Table 4, most studies concerning astaxanthin accumulation by *H. pluvialis* have an induction stage from green- to red-phase. This distinction is associated with *H. pluvialius* cellular morphologies. Therefore, macrozooids (zoospores), microzooids, and palmella cellular morphologies are usually called "green-phase cells", while hematocysts

(aplanospores) are referred as "red-phase cells" [126]. The first predominate in favorable growth conditions, whereas the second occur under unfavorable environmental or culture conditions [126]. Since *H. pluvialis* accumulate large amounts of carotenoids, especially astaxanthin, under the red-phase, stress inputs such as nitrogen depletion and high light intensity are often applied to *H. pluvialis* to prompt astaxanthin production [126].

Several factors have already being studied for *H. pluvialis* in both green- and red-phases with the aim to prompt astaxanthin production, namely the effect of high light intensity exposure time, growth phase, cultivation mode, and the use of alternative substrates such as succinic acid, ethanol, and walnut shell extracts, as detailed in Table 4. Nonetheless, commercial production of astaxanthin from *H. pluvialis* still faces challenges such as the high operating costs of the mass cultivation of microalgae [128].

Besides *H. pluvialis* (5.8–22.7 mg g−1 dw), there are other astaxanthin-producing microalgae, such as *Scenedesmus vacuolatus* (1.5–2.7 mg g−1 dw), *Scotiellopsis oocystiformis* (6.4–10.9 mg g−1 dw), *Chlorella zofingiensis* (3.5–6.8 mg g−1 dw), *Neochloris wimmeri* (5.1–19.3 mg g−1 dw), and *Protosiphon botryoides* (12.9–14.3 mg g−1 dw).

The carotenoids database [134] at the access date of 8 October 2021 presented information on 1204 natural compounds distributed among 722 organisms from all domains of life. Xantophylls can be classified based on their chemical modifications as hydroxyl (lutein, zeaxanthin), epoxide (violaxanthin, neoxanthin), keto (astaxanthin), or carbonyl groups (canthaxanthin, capsanthin) [135]. Besides their health benefits and industrial applications, carotenoids can be useful biomarkers for microalgal distinction. Table 5 summarizes the main carotenoids and xantophylls that can be found for each algal phylum.

**Table 5.** Main carotenoids and xanthophylls of the algal phyla [136].

As can be seen in Figure 7, in microalgae, carotenoids are biosynthesized using the isoprenoid precursor's IPP and DMAPP, which are further condensed to yield geranyl pyrophosphate. This molecule is elongated yielding geranylgeranyl pyrophosphate, which, in turn, can be dimerized to generate phytoene, the precursor of carotenoids [97]. Carotenoid accumulation often occurs when microalgal is exposed to some stress factors (e.g., nitrogen deficiency [137], ultraviolet-A radiation [118], salinity [138,139]) and growth is arrested [34]. Thus, for prompting carotenoids overproduction, for industrial scale, a two-stage cultivation strategy is often applied to first obtain higher biomass and then to trigger carotenoids accumulation [34].

#### **6. Lipid Characterization**

Carbon fixed by microalgae can be allocated into diverse metabolites. This distribution is dependent on microalgae metabolic and cellular organization, which in turn varies across distinct phylogenetic classes [140]. Thus, to improve microalgal biomass/product accumulation, several questions must be solved, namely how flux to desired product accumulation is controlled, and which interactions occur between and within metabolic pathways [141]. Metabolic profiling is a promising approach for identifying and quantifying the intracellular metabolic fluxes of microalgae, under different cultivation conditions [129].

Studies monitoring chemical diversity often screen a specific biological activity, a targeted compound class or individual molecule [142]. This constrains the development of microalgae-based industries once it overshadows the diversity of compounds produced by microalgae [143]. Several methods have been developed to allow a quantitative and simultaneous analysis of many groups of metabolites on complex mixtures by capillary electrophoresis mass spectroscopy (CE-MS), gas chromatography–mass spectroscopy (GC-MS), liquid chromatography–mass spectroscopy (LC-MS), nuclear magnetic resonance spectroscopy (NMR), and Fourier transform ion cyclotron resonance–mass spectroscopy (FTICR-MS) [144,145]. These strategies allow to save time, with respect to laborious isolation and quantification procedures, and give prompt information for complex mixtures, constituting a major advantage for microalgal-based industries [145].

Lipidomic comprises the detailed identification and quantification of lipid classes [146]. GC-MS has been widely used for the determination of the fatty acid compositions of microalgal lipids, which, in turn, are the basis of microalgae lipidomic studies [146]. However, the challenge with microalgal lipidomic studies lies in addressing its vast complexity and chemical heterogeneity [147]. GC-MS methods are recognized by their high detection sensitivity, accuracy, and excellent reproducibility; nevertheless, sample pretreatment (e.g., hydrolysis, derivatization) needs to be performed for samples [148]. A trimethylsilyl 2,2,2 trifluoro-N-trimethylsilylethanimidate (BSTFA) derivatization with GC-MS has been used as a simple, fast, low-cost, and powerful tool to gain in-depth knowledge on unknown but relevant lipids [149]. Through this non-target approach, it is possible to simultaneously analyze fatty acids, sterols, monoglycerides, aliphatic alcohols, glycosyl sterols, and other lipid-soluble molecules, such as *α*-tocopherols, without prior knowledge of the sample composition [150].

#### **7. Conclusions**

Until this stage of algal development, the production of bioenergy from microalgae is still not feasible. Thus, to take advantage of already exploited oleaginous microalgal species, algae farmers are turning their focus towards high-value lipids production for health and food sectors. Nevertheless, the great biodiversity of microalgae remains to be explored, holding back the opportunities that come from the wide diversity of compounds amongst microalgae taxa. Within lipids, the already recognized ω3-PUFA health-promoting properties are turning the focus of microalgal-based industries towards autotrophic ω3- PUFA production. However, more knowledge on the production strategies to prompt ω3-PUFA accumulation is lacking. Phytosterols are an unexplored lipid resource, which could pose an opportunity to explore them as food additives, as functional food, or dietary supplements. Lipid-soluble compounds carotenoids are the most extensively used in food, feed, nutraceuticals, and cosmetics. For high-value lipid exploitation, increasing the knowledge on new, simple, and cost-effective strategies to increase the production of these molecules is necessary. Additionally, the use of high-throughput methods that allow the identification and quantification of a wide array of lipid components are needed. The promising biological activities of microalgal-derived phytosterols show that the future of microalgal high-value lipids should not be restricted to fatty acids.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/md19100573/s1: Table S1. Insight on the legislation available in the European Union to regulate the food market, Table S2. Biological activities studied for microalgae-derived phytosterols in cell culture experiments and animal models.

**Author Contributions:** Conceptualization, T.F. and N.C.; writing—original draft preparation, T.F.; writing—review and editing, T.F. and N.C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was partially supported by the Foundation for Science and Technology-FCT, through UIDB/04423/2020 and UIDP/04423/2020, and by the European Territorial Cooperation Programme PCT-MAC 2014-2020, through the REBECA-CCT (MAC/1.1.B/269) project. Tomásia Fernandes was financially supported by a doctoral grant from the Regional Agency for the Development of Research, Technology, and Innovation of Madeira-ARDITI, Project M1420-09-5369-FSE-000002.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

### *Review* **Antioxidant Compounds from Microalgae: A Review**

**Noémie Coulombier 1,\*, Thierry Jauffrais 2 and Nicolas Lebouvier 3**

**\*** Correspondence<EMAIL_ADDRESS>
**Abstract:** The demand for natural products isolated from microalgae has increased over the last decade and has drawn the attention from the food, cosmetic and nutraceutical industries. Among these natural products, the demand for natural antioxidants as an alternative to synthetic antioxidants has increased. In addition, microalgae combine several advantages for the development of biotechnological applications: high biodiversity, photosynthetic yield, growth, productivity and a metabolic plasticity that can be orientated using culture conditions. Regarding the wide diversity of antioxidant compounds and mode of action combined with the diversity of reactive oxygen species (ROS), this review covers a brief presentation of antioxidant molecules with their role and mode of action, to summarize and evaluate common and recent assays used to assess antioxidant activity of microalgae. The aim is to improve our ability to choose the right assay to assess microalgae antioxidant activity regarding the antioxidant molecules studied.

**Keywords:** reactive oxygen species; ascorbic acid; glutathione; tocopherols; phenolic compounds; carotenoids

T.; Lebouvier, N. Antioxidant Compounds from Microalgae: A Review. *Mar. Drugs* **2021**, *19*, 549. https://doi.org/10.3390/md19100549

Academic Editor: Carlos Almeida

**Citation:** Coulombier, N.; Jauffrais,

Received: 14 September 2021 Accepted: 24 September 2021 Published: 28 September 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

#### **1. Introduction**

The demand for natural products isolated from microalgae has increased over the last decade and has drawn attention from the food, cosmetic and nutraceutical industries. Microalgae are eukaryotic unicellular cells that combine several advantages for the development of biotechnological applications: high biodiversity, photosynthetic yield, growth, productivity and a metabolic plasticity that can be orientated using culture conditions [1,2]. Some of these metabolites are molecules of interest such as pigments (e.g., carotenoids), polyunsaturated fatty acids (PUFAs, e.g., the omega-3 or -6 fatty acids), polysaccharides, vitamins and sterols which can be introduced as dietary supplements in human nutrition and animal feed e.g., [3,4]. In addition, most of them are bioactive molecules with antiinflammatory, antibacterial, anti-UV, antifungal, anticancer, and/or antioxidant activities which may bring added value to cosmetics, nutraceuticals or food products e.g., [5–9].

The demand for natural antioxidants as an alternative to synthetic antioxidants has increased [6,10]. Indeed, many synthetic antioxidants (e.g., butylated hydroxyanisole (BHA), butylated hydroxytoluene (BHT)) are considered to have a carcinogenic and/or toxic effect on animal models [11–14]. Although, most natural antioxidants currently available on the market are derived from terrestrial plants, microalgae are being more and more considered as a potential source of natural antioxidant compounds by the food industry [15–17] and by the cosmetic and nutraceutical industries [4,18].

Regarding the wide diversity of antioxidant compounds and mode of action combined with the diversity of ROS, this review first covers a global presentation of antioxidant molecules with their role and mode of action, to finally summarize and evaluate common and recent assays used to assess antioxidant activity of microalgae. The aim of this review is to improve our ability to choose the right assay to assess microalgae antioxidant activity regarding the antioxidant molecules studied. It also emphasizes and discusses the potential use of microalgae by the food industry for their antioxidant activity.

#### **2. Antioxidant and Reactive Oxygen Species (ROS)**

An antioxidant is defined as "a substance that, when present at low concentrations compared with those of an oxidizable substrate, significantly delays or prevents oxidation of that substrate" [19]. Antioxidant molecules produced by microalgae are used to protect the cell against reactive oxygen species (ROS) produced in response to biotic or abiotic stressors. Indeed, irradiance, UV, temperature, pH, metals, and nutrient can directly influence the production of antioxidant molecules in response to their availability, either through an excess or a limitation [7,20–26].

Antioxidants used for ROS detoxification have enzymatic and nonenzymatic origins with intracellular or extracellular mode of action (e.g., singlet O2 quencher, radical scavenger, electron donor, hydrogen donor, peroxide decomposer, enzyme inhibitor, gene expression regulation, synergist, and metal-chelating agents) [27].

In microalgae, ROS are produced by electron transport chains in chloroplasts and mitochondria, by the activity of some enzymes such as peroxidases and oxidases and also by the activity of some photosensitizers such as the chlorophyll [28]. The reactive oxygen species are therefore essentially generated in the chloroplasts and mitochondria but also in the peroxisomes [29]. More generally, ROS refer to O2 derivatives that are more reactive than O2 itself. This includes free radicals that contain at least one unpaired electron, as well as nonradical molecules [30]. Briefly, the activation of O2, in its stable state triplet oxygen (3O2), takes place (i) either by a transfer of energy large enough to reverse the spin of one of the electrons, which leads to the formation of singlet oxygen (1O2), or (ii) by an electron transfer that leads to the sequential reduction of 3O2 to superoxide radical (O2 −•), hydrogen peroxide (H2O2) and hydroxyl radical (OH•).

In plants and algae, singlet oxygen 1O2 is produced under high light by chloroplasts in the reaction center of the photosystem II (PSII) and to a lower extent in the antenna complex [31]. In the antenna complex, triplet-excited chlorophyll (3Chl \*) is formed from singlet-excited chlorophyll (1Chl \*) by intersystem conversion [32]. The chlorophyll in the triplet state has a longer lifespan than in the singlet state and can react with 3O2 to form the highly reactive 1O2 [33]. The singlet oxygen is responsible for extensive cell damage (e.g., protein, lipid and nucleic acid oxidation, chloroplasts and thylakoids membranes disruption and photoinhibition) around the production area [34,35]. The reaction center of PSII is thus particularly threatened. The superoxide radical (O2 −•) generation takes place in the chloroplast during photosynthesis, in the mitochondria during oxidative phosphorylation and in cell membranes through the activity of the NADPH oxidase [30]. The superoxide radical is poorly reactive because it lacks the ability to modify macromolecules and is quickly transformed into hydrogen peroxide (H2O2) [34]. However, its protonated form is the precursor of much more reactive radicals [30]. The hydrogen peroxide is formed by disproportionation of the O2 −• a redox reaction that can be spontaneous or catalyzed by the superoxide dismutase (SOD). The hydrogen peroxide is also poorly reactive; however, it remains particularly toxic, as it can cross membranes, diffuse throughout the cell and oxidize sulfhydryl groups, causing the deactivation of essential enzymes [36]. It can also react with DNA and more specifically with some transition metals (e.g., iron and copper) inducing the formation of highly reactive hydroxyl radicals by the Haber–Weiss reaction [36,37]. The hydroxyl radical (OH•) is formed in the same cell compartments as the H2O2, i.e., in the stroma of the chloroplasts using the H2O2 generated by the photosystems, but needs the presence of reduced metal of transition [30]. The hydroxyl radicals can induce lipid peroxidation, protein and nucleic acid denaturation. In addition, there are no enzymes that can detoxify these radicals; in excess, it might lead to cell death [38], and lipid peroxidation may also generate other very reactive free radicals (e.g., the perhydroxyl HO2 •, alkyl radical, reactive aldehydes malondialdehyde

(MDA) and 4-hydroxy-2-nonenal (HNE)) [33,35]. Thus, the lipid-rich membranes and their functions are particularly affected by lipid peroxidation mainly through a decrease in membrane fluidity, an increase in their permeability and by enzyme, protein, ion channel and membrane receptor inactivation, which could lead to cell damage [33].

#### **3. The Antioxidants Molecules of Microalgae**

#### *3.1. Ascorbic Acid*

Ascorbic acid or vitamin C (**1**) is one of the most abundant water-soluble antioxidants synthesized by plants (Figure 1). It is mainly present in the cytosol and chloroplasts where it can directly neutralize superoxide and hydroxyl radicals as well as singlet oxygen by electron transfer, in addition to its role in the detoxification of hydrogen peroxide during the ascorbate-glutathione cycle [39]. Ascorbic acid is also involved in the protection of the photosynthetic apparatus through its participation in the regeneration of carotenoids of the xanthophyll cycle (cofactor of violaxanthin de-epoxidase) and α-tocopherol linked to membranes [39]. It has been shown that ascorbate can also have a pro-oxidant action by the reduction of transition metals (Fe3+ to Fe2+ and Cu2+ to Cu+) which can reduce hydrogen peroxide to hydroxyl radical by the Fenton reaction [40].

**Figure 1.** Molecular structure of ascorbic acid, glutathione, tocopherols and phenolic compounds.

#### *3.2. Glutathione*

Glutathione (**2**) is a water-soluble tripeptide (L-γ-glutamyl-L-cysteinylglycine) present in all cellular compartments that play a crucial role in the antioxidant response (Figure 1). In addition to its role as a cofactor in the neutralization of hydrogen peroxide by glutathione peroxidase and in the regeneration of ascorbate in reduced form via the ascorbateglutathione cycle, glutathione can directly deactivate superoxide and hydroxyl radicals

as well as singlet oxygen. In addition, like ascorbate, glutathione participates in the regeneration of α-tocopherol in its reduced form [37].

#### *3.3. Tocopherols*

Tocopherols or vitamin E are fat-soluble molecules only synthesized by photosynthetic organisms and located in the lipid bilayers of membranes, mainly in those of chloroplasts [41]. The name "vitamin E" groups together four natural forms of tocopherols (α-, β-, γ- and δ-) (**3a**–**d**) to which are added the four forms of tocotrienols (α-, β-, γ- and δ-) (**4a**–**d**) (Figure 1). Tocopherols and tocotrienols consist of a chromanol ring and a hydrophobic phytyl side chain, tocotrienols differing from tocopherols by the presence of three double bonds on the side chain [41].

Tocopherols and tocotrienols have the capacity to neutralize lipid peroxyl radicals by giving a hydrogen atom from the hydroxyle group of the chromanol ring, thus making it possible to stop the chain reaction of lipid peroxidation [41]. The reaction results in the formation of a hydroperoxide, which can be neutralized by the action of glutation peroxidase, and of a tocopheroxyl radical (for tocopherols) or tocotrienoxyl (for tocotrienols), which are less reactive. Tocopherols and tocotrienols can then be regenerated by the action of ascorbate and glutathione at the interface of the membrane and cytosol or by coenzyme Q (UQH2) in the membrane [41]. Tocopherols can also deactivate singlet oxygen by two mechanisms: a physical quenching by charge transfer and a chemical reaction resulting in the formation of tocopherol quinone by irreversible opening of the chromanol ring [42].

#### *3.4. Phenolic Compounds*

Phenolic compounds are a large family of molecules: more than 8000 phenolic structures have been described to date in the plant kingdom [43]. These molecules contain at least one aromatic ring carrying one or more hydroxyl groups (Figure 1). The main families of compounds are phenolic acids, tocopherols described above, flavonoids and tannins as well as stilbenes and lignans [43]. Phenolics are an important class of antioxidants in higher plants and macroalgae but have only recently been studied in microalgae. However, the total content of phenolic compounds has been shown to contribute to the antioxidant activity of microalgae extracts [10,44–47]. The main molecules identified to date in microalgae are phloroglucinol (**5**) and phenolic acids derived from hydroxybenzoic acid (**6**) and hydroxycinnamic acid (**7**). Several studies have also shown the presence of weak concentrations of flavonoids e.g., [8,47–54]. All of these molecules are found in higher plants where their concentration is generally higher than in microalgae [55].

Phenolic acids can neutralize ROS primarily by hydrogen atom transfer. The antioxidant activity of the different molecules is directly linked to their chemical structure such as the number of hydroxyl groups or their position on the aromatic cycle [55]. The reaction results in the formation of a phenoxyl radical which is stabilized by the delocalization of the single electron around the aromatic ring (resonance stabilization). Phenolic acids also have the ability to inactivate radicals by monoelectronic transfer, and some can chelate the transition metals involved in the Fenton reaction thus preventing the formation of the highly reactive hydroxyl radical [55,56].

Among the pigments, we can also mention marennine, a blue-green pigment produced by *Haslea ostrearia*, which shows particularly interesting anti-free radical and antioxidant properties [57].

#### *3.5. Carotenoids*

Carotenoids are the most common pigments in nature, and more than 750 molecules have been described in algae, higher plants, bacteria and fungi [58] (Figure 2). They are fat-soluble molecules belonging to the terpenoids family containing a central chain with a system of conjugated double bonds, which can carry cyclic end groups. Carotenoids are separated into two groups: carotenes which contain only carbon and hydrogen atoms, and xanthophylls which contain at least one oxygen atom (hydroxyl, epoxy, ketone functions, for example) [59].

**Figure 2.** Molecular structure of carotenoids.

Carotenoids are mainly present in the pigment-protein complexes of the thylakoid membrane, but certain species of microalgae can also accumulate carotenoids (β-carotene (**8**) and astaxanthin (**9**)) in lipid globules located in the stroma of the chloroplast or in the cytoplasm [60]. Some carotenoids are only found in specific classes of algae and so be used as chemotaxonomic markers [58].

The role of carotenoids is on the one hand to transfer light energy to chlorophylls and on the other hand to protect the photosynthetic system by deactivating ROS and preventing their formation [61]. The first photoprotection mechanism involves xanthophylls associated with the antennal complexes of the PSII which allows the dissipation of an excess of light energy without damage, according to a series of reactions called the "xanthophyll cycle" [62]. In excess light, violaxanthin (**10**) is converted to antheraxanthin (**11**) and then to zeaxanthin (**12**) by de-epoxidation provided by violaxanthin de-epoxidase, which uses ascorbate as cofactor. This enzyme, bound to thylakoids in the lumen, is activated by an acidic pH, an excess of proton in the lumen signaling that the light energy absorbed exceeds the capacity of the electron transport chain. The de-epoxidation of violaxanthin to zeaxanthin is a very rapid phenomenon on the order of a few minutes and reversible at low light intensity or in darkness by the action of zeaxanthin epoxidase. Zeaxanthin, unlike violaxanthin, can deactivate 1Chl\* by dissipating its energy by heat [63]. This nonphotochemical quenching (NPQ) mechanism decreases the lifespan of 1Chl\* and therefore prevents the formation of 3Chl\* and then singlet oxygen in the PSII. In addition, by dissipating the excess energy, the possibilities of reducing O2 to the superoxide radical O2 −• in the PSI are minimized (less electron leakage in the transport chain) [32]. The violaxanthin cycle takes place primarily in chlorophytes. There is an alternative xanthophyll cycle, with similar photoprotective functions, in certain classes of microalgae (heterokonts, haptophytes, euglenophytes and dinophyceae) for which diadinoxanthin (**13**) is converted to diatoxanthin (**14**) [62].

At high light intensity, the probability of 3Chl\* formation is high despite the action of the xanthophyll cycle [32]. In antenna complexes, carotenoids are located near chlorophylls and can thus quickly neutralize 3Chl\* by triplet–triplet transfers before they react with 3O2 to form 1O2 [32]. Carotenoids can also directly deactivate singlet oxygen if it is formed [64]. This ability to deactivate 1O2 is particularly important in the reaction center of PSII where there are no carotenoids in close proximity to the special pair of chlorophylls which can change to the triplet state and then react with the 3O2 without that the reaction is not neutralized beforehand by the carotenoids [32]. Carotenoids therefore deactivate the 1O2 that is formed in the reaction center, thus protecting the photosynthetic system from oxidative damage. The deactivation of 3Chl\* and of 1O2 results in the formation of triplet carotenoids (3CAR\*) which de-excite without damage by dissipating the excess energy absorbed in the form of heat and can again intervene in a deactivation cycle [32].

Carotenoids are considered to be the most efficient molecules in deactivating 1O2 owing to their system of conjugated double bonds. Thus, the greater the number of conjugated double bonds is, the more effective the carotenoid will be [64]. Carotenoids also have the ability to react with free radicals through three mechanisms: hydrogen atom transfer, monoelectronic transfer and adduct formation [65].

The interactions between carotenoids and free radicals are complex. Indeed, many parameters are involved, such as the nature of the radical, the polarity of the reaction medium, the partial pressure of oxygen, the interactions with other antioxidants, such as ascorbate or tocopherols, and the concentration and structure of the carotenoid (number of conjugated double bonds, presence and types of oxygen functions, presence of end groups, cis- or trans-configuration, etc.) [65]. Carotenoids can, for example, react with a peroxyl radical (ROO•), which is added to the polyene chain of the carotenoid forming an adduct ROO-CAR• which can react with another peroxyl radical forming a nonradical product ROO-CAR-OOR, thus allowing one to break the reaction chain of lipid peroxidation. This phenomenon takes place at low partial pressure of oxygen; however, at higher partial pressure, the ROO-CAR• radical can react with 3O2 to form a ROO-CAR-OO• radical which acts as a pro-oxidant and could in this case contribute to the spread of lipid peroxidation [65,66].

#### *3.6. Miscellaneous Antioxidants*

There are other more specific antioxidant molecules produced by certain microalgae: Mycosporins-like amino acids (MAA) form a family of thirty-five molecules. They are colorless, water-soluble molecules found in a wide variety of marine organisms [67]. In microalgae, the most abundant MAAs are mycosporin-glycine (**15**), porphyra 334 (**16**), shinorin (**17**), asterina-330 (**18**), palythene (**19**) and palythine (**20**) [68,69] (Figure 3). The

main function of these molecules is UV protection, but some of them have also been shown to have antioxidant properties. In particular, they can inhibit lipid peroxidation and neutralize singlet oxygen and certain free radicals [67].

**Figure 3.** Molecular structure of other miscellaneous molecules with antioxidant activity.

Polysaccharides are polymers composed of osidic units linked to glycosidic bonds attached to the cell wall or released into the medium (exopolysaccharides) [70]. Several polysaccharides derived from microalgae have shown antioxidant activity against free radicals; however, this in vitro activity remains quite low [71–75].

Phycobiliproteins are water-soluble pigments participating in the photosynthesis of certain groups of microalgae. They are composed of a protein and a chromophore called phycobilin particularly effective at absorbing red, orange, yellow and green light, which is not optimally absorbed by chlorophyll a [76]. There are four different structures: phycoerythrobilin (**21**), phycourobilin (**22**), phycocyanobilin (**23**) and phycoviolobilin (**24**) (Figure 3). They can neutralize ROS and chelate or reduce ferrous ions [77].

#### **4. Common and Recent Assays Used to Evaluate Antioxidant Activity of Microalgae**

Many antioxidant assays have been developed with different types of reactions to highlight the wide variety of antioxidant molecules and ROS, which act with different mechanisms. It is important to note that there is no single ideal test, and it is necessary to use several tests with different mechanisms of action to evaluate the whole antioxidant capacity of an extract or molecule [7,78–80].

The majority of the assays are based on the two main mechanisms of action of antioxidants (AH) to deactivate radicals (X•):

Hydrogen atom transfer (or HAT):

$$\text{AH} + \text{X}^{\bullet} \rightarrow \text{A}^{\bullet} + \text{XH}$$

These reactions are generally fast; they are completed in seconds to minutes. The effectiveness of the antioxidant is determined by its ability to give a hydrogen atom (homolytic dissociation energy); therefore, the weaker the A-H bond, the more effective the antioxidant is [80].

Single electron transfer (SET):

$$\begin{array}{rcl} \text{AH} + \text{X}^{\bullet} & \rightarrow \text{AH}^{\bullet+} + \text{X}^{-} \\\\ \text{AH}^{\bullet+} & \stackrel{\text{H}\sharp\text{O}}{\longleftrightarrow} \text{A}^{\bullet} + \text{H}\_{3}\text{O}^{+} \\\\ \text{X}^{-} + \text{H}\_{3}\text{O}^{+} & \rightarrow \text{XH} + \text{H}\_{2}\text{O} \end{array}$$

These reactions are slower than hydrogen transfer reactions. The reaction is pH dependent, and the effectiveness of the antioxidant is mainly determined by its ionization potential. In general, the ionization potential decreases with increasing pH leading to an increase in the ability to donate an electron by deprotonation [80].

Other methods can also be used to evaluate the capacity of antioxidants to chelate transition metals or to inhibit the lipid peroxidation chain reaction. The most commonly used methods to evaluate the antioxidant activity of microalgae are presented in Table 1, and the most relevant results to assess antioxidant activity of microalgae extracts by in vitro chemical methods are presented in Table 2. Some cell-based antioxidant activity assays are presented, although few results using microalgae are found in the literature (Table 3). In addition, there does not seem to be any specific assays to evaluate antioxidant activity of microalgae on an animal model. Indeed, in most cases, microalgae are administrated to animals by food with a defined period and dosage; the testing animals are then sacrificed, and common in vitro chemical antioxidant activity assays (TBARS mostly) are used on animal tissues or blood by comparing with animals that did not consume microalgae (Table 4).

color is measured at 632 nm.

In

**Table**

**1.**

*Cont.*

43 and 56% inhib.

> (vi) nitric oxide

scavenging activity

 μg

#### *Mar. Drugs* **2021**, *19*, 549

**Table 2.** *Cont.*

**Table 2.** *Cont.*

**Table 2.** *Cont.*

 activity, CHCl3:

**Table 3.**

Antioxidant

 activity evaluation

 of microalgae

 extracts by cellular assays (AA: ascorbic acid, Ac: acetone, CAA: cellular antioxidant

**Table 4.** *Cont.*

#### **5. Antioxidant Activity of Microalgae**

The studies included in this part have been selected using Scopus and Google Scholar databases, using the terms "microalgae" in combination with "antioxidant", "antioxidant activity", "antioxidant capacity" or "antioxidant properties" as keywords. The research was limited to publication with an impact factor higher than 0.5, published until 2020. The studies have been selected based on these criteria: studies using in vitro (Table 2) or in cellular assays (Table 3) reporting the antioxidant activity of crude extract of eukaryotic microalgae. Studies focusing in the antioxidant activity of specific purified metabolite(s), or antioxidant enzyme activity have not been considered.

In addition, we have included a nonexhaustive selection of studies evaluating the antioxidant activity of microalgae in different animal models (Table 4).

The main publications evaluating the antioxidant activity of crude microalgal extracts by in vitro chemical tests are presented in Table 2. In these studies, more than two hundred *strains* of microalgae were evaluated. The most studied genera are *Chlorella* (29 *strains*), *Scenedesmus* and *Tetraselmis* (14 *strains*) (Supplementary Materials Figure S1).

The most commonly used assays to evaluate the antioxidant activity of microalgae are the DPPH (36 studies out of 52 referenced), ABTS (20 studies) and FCA assays (13 studies) (Supplementary Materials Figure S2). Overall, the results are very heterogeneous depending on the species of microalgae studied and the tests used to measure antioxidant activity. The protocols of the assays vary from one study to another, notably in terms of the extraction method, the solvents used, the reaction time and the concentrations tested. In addition, the results are expressed in different ways, making it difficult to compare the results. For example, for the DPPH assay, the results are expressed in percentage of inhibition for a given concentration (at different concentrations according to the studies), in IC50, in equivalent trolox (per unit of weight of extract or per unit of dry weight) or in equivalent ascorbic acid. Finally, the use of a reference product as a point of comparison is not systematic, and the choice of the reference product is not always relevant according to the assay used.

Nevertheless, several studies highlight the potential of microalgae as a source of antioxidants:

*Chloromonas* sp. and *Botryidiopsidaceae* sp. (ethanolic extracts) show a strong ability to neutralize DPPH radicals (IC50 of 0.97 and 1.53 μg mL−1, respectively) and ABTS (IC50 of 0.95 μg mL−1 and 1.79 μg mL−1) similar to vitamin C [118,119]. The ABTS assay also revealed interesting activities of *Scenedesmus obliquus* (IC50 of 41 μg mL−1, [96]), *Haematococcus pluvialis* (activity up to 1974 μmol TE g−1 extract for supercritical H2O extraction, [114]) and *Dunaliella salina* (activity up to 1118 μmol TE g−1 extract with hexane extraction, [83]). Interesting results are also obtained with the DPPH assay for *Galdieria sulphuraria*, *Ettlia carotinosa*, *Neochloris texensis*, *Chlorella minutissima*, *Chlorella vulgaris*, *Schizochytrium limacinum*, *Stichococcus bacillaris* and *Crypthecodinium cohnii* with inhibition percentages between 89% and 95% with aqueous or methanolic extracts at concentrations of 250 μg mL−1 [106].

Natrah et al. [84] showed that *Chaetoceros calcitrans*, *Scenedesmus quadricauta*, *Isochrysis galbana*, *Chlorella vulgaris*, *Nannochloropsis oculata*, and *Tetraselmis tetrahele* had a strong ability to inhibit lipid peroxidation with inhibition percentages ranging from 88% to 98% for methanolic extracts at 80 μg mL−1 with the TBARS assay and between 88.4 and 97% for extracts at 200 μg mL−1 with the FTC assay (Ferric ThioCyanate assay, indirect measurement of the quantity of hydroperoxides formed during the first stages of lipid oxidation). The ability of the genera *Tetraselmis* to inhibit lipid peroxidation is confirmed by Coulombier et al. [21] who have obtained an IC50 up to 3,4 μg mL−1 with a methanoldichloromethane extract. *Euglena tuba* also seems to be an interesting species for its ability to inhibit lipid peroxidation (IC50 with TBARS assay = 42 μg mL−1) and to neutralize the superoxide radical (IC50 = 5.2 μg mL−1, [49]). Some species show good ability to neutralize superoxide radical such as *Chaetoceros* sp. (1029 μmol TE g−1 dichloromethane extract), *Nannochloropsis* sp. (3224 μmol TE g−1 methanol extract), *Chlorella stigmatophora*

and *Phaeodactylum tricornutum* (IC50 of 48.37 and 68.61 μg mL−1 with aqueous extracts, [87]). Chloroform and methanol extracts of *Chaetoceros* sp. also show interesting results with the FRAP assay (610 and 492.50 μmol TE g−1, [86]). Good results are also obtained with the TAC assay with IC50 below 100 μg mL−1 for methanolic extracts of *Chlorella vulgaris* and *Chlamydomonas reinhardtii* [47].

The antioxidant activity of the genus *Chlorella* has been demonstrated by several authors with different antioxidant assays. In addition to the results obtained with the DPPH, TBARS, FTC, TAC, and superoxide radical neutralization assays presented above, Aremu et al. [44,48] obtained IC50 up to 25 μg mL−1 with the β-carotene bleaching assay for *Chlorella minutissima* and *Chlorella* sp. and Plaza et al. [81] showed activities up to 1008 μmol TE g−1 of *Chlorella vulgaris* extract with the ORAC assay.

Overall, few links are made between these antioxidant activities and the metabolites involved. Still, correlations have been shown with carotenoid content [44,83], phenolic compound content [44,106] including flavonoids [47] and gallic acid and vitamin E content [114].

Despite cellular assays potentially giving more biological relevant information, as they take into account the bioavailability and metabolism of the tested compounds, we found only four studies using cellular assays to determine antioxidant activity of microalgae extract (Table 3). Those studies use different antioxidant cellular assays and different cell models (mouse fibroblast, macrophage or lymphoma cells and human liver cancer cell line).

Chloroform, methanol, acetone and 70% ethanol extracts of *Chaetoceros calcitrans* showed high nitric oxide scavenging activity in mouse macrophage with IC50 values of 3.46, 3.83, 15.35 and 17.94 μg mL−1, respectively, that is closed to reference compounds (IC50 of 4.7 and 6.1 μg mL−1 for quercetin and curcumin, [88]). This strong inhibitory activity of nitric oxide was attributed to the carotenoid content of *Chaetoceros calcitrans* (fucoxanthin, astaxanthin, violaxanthin, zeaxanthin, canthaxanthin and lutein). Karawita et al. [92] showed that *Pediastrum duplex* extract has a good protective effect against DNA damage induced by hydrogen peroxyde exposure (Comet assay). Indeed, a decrease of 80% of DNA damage on mouse lymphoma cells was measured with *Pediastrum* extract at 100 μg mL−1 compared to control with no microalgae extract. Good antioxidant activity was also measured with CAA (cellular antioxidant activity) and CLPAA (cellular lipid peroxidation antioxidant activity) assays on human liver cancer cell line with *Ostreopsis ovata* and *Alexandrium minutum*; however, both species extracts showed toxicity in cytotoxicity assay [91].

Similarly to cellular assays, the evaluation of the antioxidant activity of microalgae extracts by in vivo experimentations are limited compared to in vitro assays (Table 4). Those studies used different antioxidant in vitro assays couple with other physiological measurement, such as antioxidant enzyme activity, on various animal models (e.g., shrimps, chicken, catfish, rats, turbots or trouts, Table 4) to assess the effect of microalgae. The microalgae (*Schizochytrium* sp., *Chlorella vulgaris*, *Amphora coffeaformis*, *Schizochytrium limacinum*, *Acutodesmus obliquus*, *Nannochloropsis* spp., *Tetraselmis chuii* and *Botryococcus braunii*) were mostly included in the animal feed as dry microalgae with a percentage of inclusion mainly going from 1–10% or as a molecule equivalent of given antioxidant compounds. The results are variable depending on species from no effect of the microalgae tested [120,123] to a decrease in oxidative stress measurements such as the malondialdehyde or hydrogen peroxide content [124–129] or a decrease in DNA damage [123]. In most cases, it seems that the inclusion of microalgae directly in the fed has a positive effect on the animal physiology, which is promising regarding further used of microalgae in the food industry either in human or animal nutrition as functional ingredients. It also raises the question of the bioavailability of an antioxidant compound in the algal matrices and thus of the digestibility of the microalgae tested.

#### **6. Applications in the Food Industry**

In the food industry, antioxidants are used for human and animal nutrition as functional ingredients to provide nutritional benefits to a product (e.g., orange juice enriched with vitamin C), and as preservatives to extend the shelf life of foods and beverages to prevent their degradation by oxidation [130,131]. The use of antioxidant ingredients in food products intended for humans is highly regulated by country-specific laws owing to their potential toxicity. In the European Union, there is a list of authorized antioxidant additives, some of which may be of natural origin such as vitamin C (E300-E304), vitamin E (E306-E309), guaiac resin (E314) and rosemary extract (E392). Certain carotenoids are also authorized as dyes but can have an antioxidant role such as β-carotene (E160a), lycopene (E160d), lutein (E161b), violaxanthin (E161e), zeaxanthin (E161h), canthaxanthin (E161g) or astaxanthin (E161j) [130]. For foods and ingredients that were not significantly consumed before 1997, such as most microalgae, the "Novel Food" regulation framework was to be applied in Europe [132]. New microalgae on the market must obtain this authorization; however, to receive it, it has to be demonstrated that the product does not present any risk in terms of safety for human health [133] as some microalgae are known to produce phytotoxins [134–136]. In addition, and beyond the regulatory framework, to be of interest to the food industry, an antioxidant should not affect the color, smell and taste of the food and should be effective at low concentrations (0.001–0.01%), be easily usable, stable during processing and storage and be inexpensive [130,131]. The use of microalgae may thus be regarded as promising additive for human food, livestock feed and shelf life; however, it greatly depends on the microalgae productivity and nutrient compositions in protein, carbohydrates, lipids, vitamins, and antioxidants, which also strongly depend on species, mode of cultivation and culture medium composition e.g., [7,21,22]. Currently, around 10 species of microalgae or microalgae extract are authorized for human consumption in Europe as a food or food ingredient [137].

For livestock feed, antioxidant additives are subject to authorization before going on the market, an authorization that remains only valid ten years. On the other hand, raw materials are not subject to authorization, but a contribution of microalgae as an antioxidant in animal feed could only be considered as a raw material if it also provides proteins, minerals, fats, fibers, energy or carbohydrates [138]. Microalgae presents growth rate and dietary value of interest (e.g., polyunsaturated fatty acids, vitamins, pigments, polysaccharides) for livestock feed or aquaculture feed either fish, live feed and shellfish applications (e.g., in Table 4). Indeed, in aquaculture, the polyunsaturated fatty acids (PUFAs) eicosapentaenoic acid (DHA) and docosahexanoic acid (DHA) are of nutritional importance in aquafeeds and are hitherto ensured by inclusion of fish oil in aquafeeds. However, this resource is limited, and microalgae offer an alternative to fish oil. In addition, microalgae are not only seen as a source of PUFAs but also as source of other metabolites of interest such as pigments, polysaccharides, vitamins (e.g., vitamin E and C) and sterols which are introduced as dietary supplements for dietetic and therapeutic purposes [3,129]. In terms of applications, antioxidant molecules (asthaxanthin, lutein, β-carotene) carotenoids are produced by a wide variety of microalgae (see Table 2).

#### **7. Conclusion**

Antioxidant molecules from microalgae are more and more considered as a potential source of natural antioxidant compounds by the food, the cosmetic and nutraceutical industries as they may bring benefits to their products.

However, it is very crucial to assess properly the antioxidant activity of an algal extract owing to the wide diversity of antioxidant compounds and the mode of action combined with the diversity of ROS involved. This review highlights the lack of standardization between extractions procedures used to assess antioxidant activity from microalgae matrices, and more disturbingly, it highlights the inappropriateness between the assay used and the molecules studied. These often hamper the comparison between studies and bring the authors to false or incorrect interpretation of their results.

Therefore, although all the assays have their merits and demerits, the appropriate selection of a given assay was to be made based on the mode of action of a studied molecule in front of the principle and mechanism of an assay, especially in vitro assays. In addition to the need of normalization of the extraction procedures and to the appropriate use of an assay, we conclude that it is crucial to combine many assays to assess microalgae full antioxidant activity.

This review also highlights that microalgae are rich in antioxidant molecules with more or less potent activities, which can be used as an ingredient in food, cosmetic and nutraceutical industries. In addition, research publications are available on modern in vitro chemical methods, but application on cellular assays and in vivo experimentations are still lacking. There is a need to develop models to improve our ability to assess the activity of antioxidant molecule on these kinds of models to further improve industrial adaption and application.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/md19100549/s1, Figure S1. Top 15 microalgae genus studied for their antioxidant activity, Figure S2. In vitro chemical assays used to evaluate the antioxidant activity of microalgae crude extracts.

**Author Contributions:** N.C. built the bibliographic database. N.C., N.L. and T.J. wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was funded by the Province Nord, the Province Sud, the Government of New Caledonia and the Comité Interministériel de l'Outre-Mer (CIOM) through the AMICAL (Aquaculture of Microalgae in NewCALedonia) 1 and 2 research programs.

**Institutional Review Board Statement:** Not applicable.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

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