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ffe4a610-aead-4300-8fef-a70aeecd3fb7.107
|
*3.10. Measurement of Eotaxin in Skin*
At 48 h after being challenged, mice were sacrificed, and the ears were removed. The ear from each mouse in each experimental group was homogenized with PBS containing 0.1% Tween 20 (100 ΐL/10 mg tissue), using a mixer mill (MM 300, Retsch, Haan, Germany) with zirconia beads (5 mm) for 2 min at 30 Hz. The homogenates were centrifuged at 12,000× *g* for 10 min. The supernatants were kept at ƺ80 °C until being assayed. The concentration of eotaxin in the supernatants was measured by a Mouse Eotaxin ELISA kit (Ray Biotech, Inc., Norcross, GA, USA). The sensitivity of the eotaxin assay was >0.01 pg/mg tissue.
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2025-04-07T04:13:04.382967
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 107
}
|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.108
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*3.11. Eosinophil Preparation*
Enriched preparations of eosinophils were obtained from the peripheral blood of IL-5-transgenic mice. Eosinophil-enriched cells were obtained by the Percoll density gradient separation method described previously [19], with modification. Briefly, isotonic Percoll was prepared using a 10× solution of Krebs Ringer PBS (KRP; 10 mM sodium phosphate buffer, pH 7.5, containing 154 mM NaCl, 6 mM KCl and 1 mM MgCl2) and diluted with KRP to achieve concentrations of 60%, 70% and 80%. In 15- mL conical polypropylene tubes (BD Falcon 352096, Becton, Dickinson and Company, Franklin Lakes, NJ, USA), 2 mL of cell suspensions of 20 to 50 × 106 cells in KRP were placed, followed by the careful layering of aliquots (2.5 mL) of each concentration of Percoll solution starting with the lowest concentration at the bottom. The tubes were spun at 1000× *g* for 20 min at room temperature. Eosinophils were extracted from the 70% to 80% Percoll fractions by removing B and T lymphocytes using anti-B220- and anti-Thy1.2-coupled Dynabeads (DYNAL A.S., Oslo, Norway). Briefly, lymphocytes bound to these beads were removed using a permanent magnet. Unbound cells were used as an eosinophil-enriched fraction. To identify eosinophils, aliquots were removed and assessed using eosinophil peroxidase staining as described previously [19]. More than 93% of the cells prepared by this method were eosinophils.
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2025-04-07T04:13:04.383026
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 108
}
|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.109
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*3.12. Chemotaxis Assay toward Eotaxin*
Eosinophils (1.3 × 106 cells/mL) were suspended in RPMI 1640 medium containing 0.1% BSA and were placed in the upper well (Transwell) of a 24-well chemotaxis chamber (KURABO Co., Osaka, Japan). Transwells with 5 ΐm pores were inserted into each well, and 4 × 105 cells in 300 ΐL of RPMI 1640 medium containing 0.1% BSA were added to the upper chamber. The lower chamber was set with 800 ΐL of RPMI 1640 medium containing 0.1% BSA and mouse eotaxin (20 ng/mL). Then, 3 or 1 ΐg of peridinin or fucoxanthin (*<sup>n</sup>* = 6) was added to each Transwell. Assay plates were incubated for 1 h at 37 °C in 5% CO2. Cells that migrated across the filter to the lower chamber were counted, and the results were expressed as the number of cells in a field of 660 ΐm × 840 ΐm. For each group, eosinophils in three fields for each well were counted, and results were reported as the mean of 18 fields (cell number ± SD).
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2025-04-07T04:13:04.383129
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 109
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|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.110
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*3.13. Statistical Analysis*
For all experiments, ANOVA was performed, and the Tukey–Kramer *post hoc* test was used to identify significantly different means. A *p*-value <0.01 was considered statistically significant.
## **4. Conclusions**
In conclusion, peridinin suppressed DTH responses in mice. Peridinin also suppressed the numbers of eosinophils in ear tissues and peripheral blood. When painted on the ears, peridinin inhibited both the migration of eosinophils toward eotaxin and the production of eotaxin in ears. However, the suppressive effect of peridinin on the production of eotaxin was not observed when administered i.p*.* A structurally related carotenoid, fucoxanthin, inhibited the migration of eosinophils toward eotaxin only *in vitro* and did not suppress the DTH response. The major structural difference between peridinin and fucoxanthin is the presence of a butenolide moiety in peridinin. The butenolide moiety of peridinin may be important for suppressing these effects on eosinophils and for the production of eotaxin. Comparison of the inhibitory effects of peridinin and other carotenoids with the butenolide moiety remains to be clarified.
As described above, peridinin may ameliorate the allergic responses in which eosinophils play a major role in inflammation responses, such as asthma or atopic dermatitis.
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2025-04-07T04:13:04.383192
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 110
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|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.111
|
**Acknowledgments**
We appreciate Shiro Ono and Hiromi Okuyama (Osaka Ohtani University) for their help in the measurement of the serum levels of cytokines. This work was supported by the Program to Disseminate the Tenure Tracking System of the Ministry of Education, Culture, Sports, Science and Technology, the Japanese Government, and by the Adaptable and Seamless Technology Transfer Program through target-driven R & D, Japan Science and Technology Agency (AS242Z03463Q).
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2025-04-07T04:13:04.383378
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1-5-2021 17:07
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 111
}
|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.112
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**Author Contributions**
Ken-ichi Onodera and Akira Tominaga planned the experiments and wrote the manuscript. Yuko Konishi, Takahiro Taguchi and Sumio Kiyoto are engaged in the experiments of this research.
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 112
}
|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.113
|
**Conflicts of Interest**
The authors declare no conflict of interest.
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{
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"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 113
}
|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.115
|
**Carotenoids in Algae: Distributions, Biosyntheses and Functions**
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2025-04-07T04:13:04.383497
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 115
}
|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.116
|
**Shinichi Takaichi**
Department of Biology, Nippon Medical School, Kosugi-cho, Nakahara, Kawasaki 211-0063, Japan; E-Mail: [email protected]; Tel.: +81-44-733-3584; Fax: +81-44-733- 3584
*Received: 2 May 2011; in revised form: 31 May 2011 / Accepted: 8 June 2011 / Published: 15 June 2011*
**Abstract:** For photosynthesis, phototrophic organisms necessarily synthesize not only chlorophylls but also carotenoids. Many kinds of carotenoids are found in algae and, recently, taxonomic studies of algae have been developed. In this review, the relationship between the distribution of carotenoids and the phylogeny of oxygenic phototrophs in sea and fresh water, including cyanobacteria, red algae, brown algae and green algae, is summarized. These phototrophs contain division- or classspecific carotenoids, such as fucoxanthin, peridinin and siphonaxanthin. The distribution of ΅-carotene and its derivatives, such as lutein, loroxanthin and siphonaxanthin, are limited to divisions of Rhodophyta (macrophytic type), Cryptophyta, Euglenophyta, Chlorarachniophyta and Chlorophyta. In addition, carotenogenesis pathways are discussed based on the chemical structures of carotenoids and known characteristics of carotenogenesis enzymes in other organisms; genes and enzymes for carotenogenesis in algae are not yet known. Most carotenoids bind to membrane-bound pigment-protein complexes, such as reaction center, light-harvesting and cytochrome *b*6*f* complexes. Watersoluble peridinin-chlorophyll *a*-protein (PCP) and orange carotenoid protein (OCP) are also established. Some functions of carotenoids in photosynthesis are also briefly summarized.
**Keywords:** algal phylogeny; biosynthesis of carotenoids; distribution of carotenoids; function of carotenoids; pigment-protein complex
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 116
}
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ffe4a610-aead-4300-8fef-a70aeecd3fb7.117
|
**1. Introduction**
Algae are classified throughout many divisions of the Kingdom Plantae. Their sizes range from single cells of picophytoplankton—the smallest of which are less than 1 μm—to seaweeds, the largest of which are more than 50 m. Attempts have been made to cultivate single-cell algae for a long time, but numbers were limited. With the recent development of culture techniques, some single-cell species can be cultured, and their characteristics, including pigments, can be studied. With the development of taxonomic technology, including DNA base sequences of 16S or 18S rRNA and some genes, algae phylogenetics has been developed.
More than 750 structurally defined carotenoids are reported from nature; land plants, algae, bacteria including cyanobacteria and photosynthetic bacteria, archaea, fungus and animals [1]. Except for animals, these organisms can synthesize many kinds of carotenoids, which are synthesized from diverse carotenogenesis pathways. These carotenoids and carotenogenesis pathways can be used as chemotaxonomic markers [2–7]. In addition, characteristics of carotenogenesis enzymes and genes are investigated. Some carotenogenesis genes have high similarity from bacteria to land plants, but some have low similarity. Some homologous genes have been proposed [8,9], but some carotenogenesis enzymes and genes, especially algae-specific ones, are not found.
In this review, the term algae refers to an oxygenic phototroph found in both seawater and fresh water, including cyanobacteria but excluding land plants. Distribution of carotenoids, carotenogenesis enzymes and pathways, and function of carotenoids in photosynthesis in algae are summarized.
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2025-04-07T04:13:04.383650
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 117
}
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ffe4a610-aead-4300-8fef-a70aeecd3fb7.118
|
**2. Distribution of Carotenoids**
Many different kinds of carotenoids were found from the algal species studied. Structures of some important carotenoids in algae are illustrated in Figure 1. Among them, approximately 30 types may have functions in photosynthesis, and others may be intermediates of carotenogenesis or accumulated carotenoids. Some carotenoids are found only in some algal divisions or classes; therefore, these carotenoids and also chlorophylls can be used as chemotaxonomic markers, and their distribution in algae is summarized in Table 1 [2–6].
Allene (C=C=C) is a unique structure in natural products, and is found mainly in carotenoids [10]; fucoxanthin in brown algae and diatoms, 19<sup>ȝ</sup>-acyloxyfucoxanthin in Haptophyta and Dinophyta, peridinin only in dinoflagellates, and 9<sup>ȝ</sup>-*cis* neoxanthin in green algae and land plants. Acetylene (C ǓC) is also a unique structure, and acetylenic carotenoids are found only in algae; alloxanthin, crocoxanthin and monadoxanthin in Cryptophyta, and diadinoxanthin and diatoxanthin in Heterokontophyta, Haptophyta, Dinophyta and Euglenophyta. Acetylated carotenoids (-O-CO-CH3), such as fucoxanthin, peridinin and dinoxanthin, are also mainly found in algae, such as Heterokontophyta, Haptophyta and Dinophyta. These carotenoids are specific to certain algal divisions and classes, and they are summarized in Table 1 based on our results [11–14] and some references [1–6].
Many cyanobacteria contain Ά-carotene, zeaxanthin, echinenone and myxol pentosides (myxoxanthophyll), while some species lack part of these and some contain additional carotenoids, such as nostoxanthin, canthaxanthin and oscillol dipentoside (Table 1, Figure 1) [13]. In addition, the carotenoid compositions of cyanobacteria are very different from those of chloroplasts in algae; consequently, during symbiosis of cyanobacteria to eukaryotic cells, carotenoids might be considerably restructured [13]. Note that since the name of myxoxanthophyll cannot specify the glycoside moieties, we have proposed the name of myxol glycosides to specify the glycosides, such as myxol 2<sup>ȝ</sup>-΅-L-fucoside, 4-ketomyxol 2ȝ-rhamnoside and oscillol dichinovoside [13,15].
Rhodophyta (red algae) can be divided into two groups based on carotenoid composition; the unicellular type contains only Ά-carotene and zeaxanthin, and the macrophytic type contains additional ΅-carotene and lutein (Table 1, Figure 1) [16]. The relationship between phylogenetics of red algae and carotenoid composition is not clear [14]. Cryptophyta also contains ΅-carotene and its acetylenic derivatives, crocoxanthin and monadoxanthin, which are only found in this division.
**Figure 1.** Structures of some carotenoids.
*Mar. Drugs* **2011**, *9*, 1101–1118
Heterokontophyta, Haptophyta and Dinophyta contain Ά-carotene and its derivatives as well as chlorophyll *c* (Table 1, Figure 1). These divisions, except for Eustigmatophyceae, which lacks chlorophylls *<sup>c</sup>*, contain unique acetylenic carotenoids of diadinoxanthin and diatoxanthin. Fucoxanthin and its derivatives are found in only four classes of Heterokontophyta (Chrysophyceae, Raphidophyceae, Bacillariophyceae and Phaeophyceae),
Haptophyta and Dinophyta. Peridinin and its derivatives are found only in Dinophyta. Fucoxanthin and peridinin have unique structures (Figure 1) and are class-specific carotenoids (Table 1).
Euglenophyta, Chlorarachniophyta and Chlorophyta contain the same carotenoids, such as Ά-carotene, violaxanthin, 9<sup>ȝ</sup>-*cis* neoxanthin [11] and lutein, as well as chlorophyll *a* and *b* with land plants (Table 1, Figure 1). Some classes contain additional carotenoids, such as loroxanthin, siphonaxanthin and prasinoxanthin, which are derivatives of lutein, and are class specific.
Note that identifications of some carotenoids were lacking because of insufficient analysis, and that some algae names were changed because of new developments in taxonomic technology and phylogenetic classification.
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{
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"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 118
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|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.119
|
**3. Carotenogenesis Pathways, Enzymes and Genes**
Carotenogenesis pathways and their enzymes are mainly investigated in cyanobacteria [13] and land plants among oxygenic phototrophs [17]. Especially in land plants, carotenogenesis pathways and characteristics of enzymes are studied in detail (Figure 2). On the other hand, algae have common pathways with land plants and also additional algae-specific pathways, which are solely proposed based on the chemical structures of carotenoids (Figure 2). Some common carotenogenesis genes in algae are suggested from homology of the known genes [8,9], but most genes and enzymes for algae-specific pathways are still unknown (Figure 2). In cyanobacteria, since carotenoid compositions are different from those in land plants and algae, the pathways and enzymes are also different from those in Figure 2, and they are shown in Figure 3. In addition, carotenogenesis enzymes and genes, whose functions are confirmed in algae, including cyanobacteria, are summarized in Table 2. Unfortunately, these enzymes are mostly from cyanobacteria and green algae (Table 2).
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2025-04-07T04:13:04.383959
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 119
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|
ffe4a610-aead-4300-8fef-a70aeecd3fb7.120
|
*3.1. Lycopene Synthesis*
## 3.1.1. Isopentenyl Pyrophosphate to Phytoene Synthesis
Isopentenyl pyrophosphate (IPP), a C5-compound, is the source of isoprenoids, terpenes, quinones, sterols, phytol of chlorophylls, and carotenoids. There are two known independent pathways of IPP synthesis: the classical mevalonate (MVA) pathway and the alternative, nonmevalonate, 1-deoxy-D-xylulose-5-phosphate (DOXP) pathway [18,19]. In the MVA pathway, acetyl-Coenzyme A is converted to IPP through mevalonate, and the enzymes and genes are well studied [20]. The pathway is found in plant cytoplasm, animals and some bacteria [18,20]. The DOXP pathway was found in the 1990s, and in this pathway, pyruvate and glycelaldehyde are converted to IPP. The DOXP pathway is found in cyanobacteria, the plastids of algae and land plants, and some bacteria [18]. Carotenoids are synthesized in plastids. Exceptionally among oxygenic phototrophs, Euglenophyceae has only the MVA pathway, and Chlorophyceae has only the DOXP pathway [18].
**Figure 2.** Carotenogenesis pathways and enzymes, whose functions are confirmed, in oxygenic phototrophs.
**Figure 3.** Carotenogenesis pathways and enzymes in cyanobacteria.
**Table 2.** Carotenogenesis genes and enzymes, whose functions are confirmed, in algae.
Red, genes and enzymes related to ΅-carotene.
Most carotenoids consist of eight IPP units. Farnesyl pyrophosphate (C15) is synthesized from three IPPs, after which one IPP is added to farnesyl pyrophosphate by geranylgeranyl pyrophosphate synthase (CrtE, GGPS) to yield geranylgeranyl pyrophosphate (C20). In a headto-head condensation of the two C20 compounds, the first carotene, phytoene (C40), is formed by phytoene synthase (CrtB, Pys, Psy) using ATP [57,58]. This pathway has been confirmed by cloning genes from two species of *Rhodobacter* (purple bacteria) and two species of *Pantoea* (previously *Erwinia*) [57–59]. Among oxygenic phototrophs, the functions of CrtE of *Thermosynechococcus elongatus* BP-1 [21], and CrtB of three species of cyanobacteria [22–24] and two species of green algae [25,26] have also been confirmed (Table 2). The *crtE* and *crtB* genes have high sequence similarity from bacteria to land plants, respectively.
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2025-04-07T04:13:04.384031
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{
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"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 120
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ffe4a610-aead-4300-8fef-a70aeecd3fb7.121
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3.1.2. Phytoene to Lycopene Synthesis
Four desaturation steps are needed in the conversion from phytoene to lycopene. Oxygenic phototrophs require three enzymes: phytoene desaturase (CrtP, Pds), Ί-carotene desaturase (CrtQ, Zds) and *cis*-carotene isomerase (CrtH, CrtISO) (Figure 2). CrtP catalyzes the first two desaturation steps, from phytoene to Ί-carotene through phytofluene, and CrtQ catalyzes two additional desaturation steps, from Ί-carotene to lycopene through neurosporene. During desaturation by CrtQ, neurosporene and lycopene are isomerized to poly-*cis* forms, and then CrtH isomerizes to all-*trans* forms. Light is also effective for their photoisomerization to all*trans* forms [34]. The functions of these enzymes have been mainly confirmed in cyanobacteria, green algae and land plants (Table 2): CrtP from *Synechocystis* sp. PCC 6803 [28], *Synechococcus elongatus* PCC 7942 [23], *Chlamydomonas reinhardtii* [29] and *Chlorella zofingiensis* [30,31], CrtQ from *Anabaena* sp. PCC 7120 (CrtQa,
*crtI*-like sequence) [32] and *Synechocystis* sp. PCC 6803 (CrtQb, plant *crtQ*-like) [33], and CrtH from *Synechocystis* sp. PCC 6803 [34,35]. The CrtP of *S. elongatus* PCC 7942 is stimulated by NAD(P) and oxygen as a possible final electron acceptor [60]. CrtQa has sequence homology with bacterial phytoene desaturase (CrtI) and CrtH, while CrtQb has sequence homology with CrtP. In addition, genes homologous to *crtQa* are not found in cyanobacteria; therefore, among oxygenic phototrophs, *Anabaena* sp. PCC 7120 is the only species to have functional CrtQa.
In contrast, the bacterial type uses only one enzyme, phytoene desaturase (CrtI), to convert from phytoene to lycopene, and the primitive cyanobacterium of *Gloeobacter violaceus* PCC 7421 uses this type of CrtI, and the homologous genes of *crtP*, *crtQ* and *crtH* are not found in the genome [22,27]; therefore, *G. violaceus* is the first oxygenic phototroph that has been shown to use this type (Table 2). These observations suggest the following evolutionary scheme for this step in the reaction: the desaturation of phytoene was initially carried out by CrtI in ancestral cyanobacteria, *crtP* and related desaturase genes were acquired, and ultimately, there was replacement of *crtI* by *crtP* [27]. Among anoxygenic phototrophs, purple bacteria, green filamentous bacteria and heliobacteria use CrtI, whereas green sulfur bacteria use CrtP, CrtQ and CrtH [61].
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2025-04-07T04:13:04.384164
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{
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 121
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ffe4a610-aead-4300-8fef-a70aeecd3fb7.122
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*3.2. Ά-Carotene and ΅-Carotene Synthesis by Lycopene Cyclases*
All carotenoids in oxygenic phototrophs are dicyclic carotenoids; Ά-carotene, ΅-carotene and their derivatives, are derived from lycopene (Figures 1 and 2). Exceptionally, myxol glycosides and oscillol diglycosides in cyanobacteria are monocyclic and acyclic carotenoids, respectively.
Lycopene is cyclized into either Ά-carotene through ·-carotene, or ΅-carotene through ·- carotene or Έ-carotene. Three distinct families of lycopene cyclases have been identified in carotenogenetic organisms [13,62,63]. One large family contains CrtY in some bacteria except cyanobacteria, and CrtL (CrtL-b, Lcy-b) in some cyanobacteria and land plants. Lycopene <sup>Ή</sup>cyclases (CrtL-e, Lcy-e) from land plants and lycopene Ά-monocyclases (CrtYm, CrtLm) from bacteria are also included. Their amino acid sequences exhibit a significant five conserved regions [39,62,64], and have an NAD(P)/
FAD-binding motif [65]. Note that Maresca *et al.* [63] divide this family into two CrtY and CrtL families. Three enzymes from Rhodophyta, *Cyanidioschyzon merolae* [38], and Chlorophyceae, *Dunaliella salina* [39] and *Haematococcus pluvialis* [40], are functionally confirmed (Table 2).
Some cyanobacteria also contain these enzymes (Table 2). *Synechococcus elongatus* PCC 7942 contains a functional CrtL [36]. *Prochlorococcus marinus* MED4 contains two lycopene cyclases (Table 2), which have sequence homology to CrtL. CrtL-b exhibits lycopene Ά-cyclase activity, while CrtL-e is a bifunctional enzyme having both lycopene Ή-cyclase and lycopene Ά-cyclase activities [37]. The combination of these two cyclases allows the production of Ά-carotene, ΅- carotene and Ή-carotene. Both enzymes might have originated from the duplication of a single gene. The characteristics of this CrtL-e are somewhat different from those in land plants [66]. In addition, the Ά-end groups of both
Ά-carotene and ΅-carotene (left half) might be hydroxylated by CrtR to zeaxanthin through Άcryptoxanthin and 3-hydroxy-΅-carotene, respectively, in *P. marinus*. *Acaryochloris marina* MBIC 11017, which produces ΅-carotene, contains only one *crtL*-like gene from genome sequence [14].
The second family of lycopene cyclases, heterodimer (*crtYc* and *crtYd*) or monomer (*crtYc-Yd*), has been found in some bacteria, archaea and fungi [62,67], but not in phototrophs.
Recently, a new family of functional lycopene cyclase, CruA, has been found in *Chlorobaculum* (previously *Chlorobium*) *tepidum* (green sulfur bacterium), and the main product is ·-carotene in *Escherichia coli*, which produces lycopene [68]. Homologous genes, *cruA* and *cruP*, have been found in the genome of *Synechococcus* sp. PCC 7002, and their main products are ·-carotene, in *E. coli*, which produces lycopene [63]. In addition, their homologous genes are widely distributed in cyanobacteria, such as *Synechocystis* sp. PCC 6803 and *Anabaena* sp. PCC 7120; however, these *cruA*- and *cruP*-like genes from both *Synechocystis* sp. PCC 6803 and *Anabaena* sp. PCC 7120 did not show the lycopene dicyclase or monocyclase activities [14]. *S. elongatus* PCC 6301 and PCC 7942, and *A. marina* MBIC 11017 contain *crtL*-, *cruA*- and *cruP*-like genes; consequently, distributions of functional lycopene cyclases (CrtL-, CruA- and CruPlike) in cyanobacteria are unknown.
Since *Synechocystis* sp. PCC 6803 and *Anabaena* sp. PCC 7120 lack *crtL*-like genes and contain non-functional *cruA*-like genes, there is a possibility to present a fourth new family of lycopene cyclases in these cyanobacteria. Further studies of distributions of functional lycopene cyclases (CrtL- and CruA-like, or others) in cyanobacteria are needed.
Distribution of ΅-carotene, that is, CrtL-e, is limited in some algae classes (Table 1). Genes and enzymes of CrtL-e are not found in algae. In some species of land plants, the characteristics of CrtL-e were investigated [66], and were shown to have sequence homology with *crtL-b*. Lycopene is first converted to Έ-carotene by CrtL-e, and then to ΅-carotene by CrtL-b. ·- Carotene produced by CrtL-b is not a suitable substrate for CrtL-e.
## *3.3. Ά-Carotene Derivatives and Their Synthesis*
## 3.3.1. Cyanobacteria
Some cyanobacteria produce zeaxanthin, and some produce both zeaxanthin and nostoxanthin (Figure 3). First, the C-3 and C-3ȝ hydroxyl groups of zeaxanthin are introduced to Ά-carotene by
Ά-carotene hydroxylase (CrtR) through Ά-cryptoxanthin. Then, the C-2 and C-2ȝ hydroxyl groups of nostoxanthin are introduced by 2,2<sup>ȝ</sup>-Ά-hydroxylase (CrtG) through caloxanthin (Table 2) [13,41–43,47]. The same enzymes, CrtR and CrtG, can also introduce hydroxyl groups to deoxymyxol and myxol to produce myxol and 2-hydroxymyxol, respectively [13,44,47]; consequently, the same enzymes are used in two pathways.
Cyanobacteria contain two ketocarotenoids, namely, canthaxanthin and 4-ketomyxol. Two distinct Ά-carotene ketolases, CrtO and CrtW, are known, and only seven enzymes are functionally confirmed in four species of cyanobacteria (Table 2) [13]. CrtO catalyzes Άcarotene to echinenone, and the
final product is canthaxanthin [22,42,45,50,51]. CrtW can introduce a keto group into Άcarotene, zeaxanthin and myxol to produce canthaxanthin, astaxanthin and 4-ketomyxol, respectively
(Figure 3) [22,27,42,50,52]; therefore, these ketolases are properly used in two pathways, Άcarotene and myxol, depending on the species [13].
The pathway and the enzymes to produce the right half of myxol 2<sup>ȝ</sup>-pentoside are still unknown (Figure 3) [13].
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3.3.2. Land Plants
In land plants, most of the carotenogenesis pathways and the functionally confirmed enzymes are known (Figure 2). Hydroxyl groups are introduced into Ά-carotene to produce zeaxanthin by
Ά-carotene hydroxylase (CrtR, CrtR-b, BCH). Epoxy groups are introduced into zeaxanthin by zeaxanthin epoxidase (Zep, NPQ) to produce violaxanthin through antheraxanthin. Under high light conditions, violaxanthin is changed into zeaxanthin by violaxanthin de-epoxidase (Vde) for dispersion of excess energy from excited chlorophylls. One end group of violaxanthin is changed to an allene group of neoxanthin by neoxanthin synthase (Nsy). Because all neoxanthin in chloroplasts has the
9<sup>ȝ</sup>-*cis* form, unknown 9ȝ-isomerase for all *trans* neoxanthin to 9<sup>ȝ</sup>-*cis* neoxanthin should be present [11].
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3.3.3. Algae
Little is known for the carotenogenesis pathways among algae, but some are proposed based on the chemical structures of carotenoids (Figure 2). Functionally confirmed enzymes are mainly reported in Chlorophyceae including *Chlorella*, *Chlamydomonas*, *Dunaliella* and *Haematococcus* for CrtB, CrtP, CrtL-b, CrtR-b [46], Zep [48], Vde [49], and CrtW (Table 2).
In the cell-free preparation of *Amphidinium carterae* (Dinophyta), 14C-labellled zeaxanthin was incorporated into allenic carotenoid of neoxanthin, and then into acetylenic diadinoxanthin and C37 peridinin (Figure 2). In addition, the three carbon atoms of C-13<sup>ȝ</sup>,14<sup>ȝ</sup>,20<sup>ȝ</sup> of peridinin were eliminated from neoxanthin (C-13,14,20) [69,70]. In organic chemistry, the C-7,8 double bond of zeaxanthin can be oxidized to the triple bond (acetylene group) of diatoxanthin [17].
Allenic carotenoids are very limited in algae. From their chemical structures, all *trans* neoxanthin might be changed to fucoxanthin, dinoxanthin, peridinin, vaucheriaxanthin and diadinoxanthin, but the pathways and enzymes are still unknown (Figures 1 and 2).
Under a stressful environment, such as high light, UV irradiation and nutrition stress, some Chlorophyceae, such as *Haematococcus*, *Chlorella* and *Scenedesmus*, accumulate ketocarotenoids, canthaxanthin and astaxanthin, which are synthesized by combining CrtR-b and Ά-carotene ketolase (CrtW, BKT) (Table 2) [53–56,71]. Note that although Ά-carotene ketolase of *Haematococcus* and *Chlorella* were named CrtO at first [53,56], they are CrtW-type not CrtOtype from amino acid sequences (Table 2).
## *3.4. ΅-Carotene Derivatives and Their Synthesis*
In *Arabidopsis thaliana*, Ά-carotene is hydroxylated mainly by the non-heme di-iron enzymes, BCH1 and BCH2 (CrtR-b), to produce zeaxanthin, while ΅-carotene is mainly hydroxylated by the cytochrome P450 enzymes, CYP97A3 for the Ά-end group and CYP97C1 for the Ά- and <sup>Ή</sup>end groups, to produce lutein [72].
Lutein and its derivatives are found only in Rhodophyta (macrophytic type), Cryptophyta, Euglenophyta, Chlorarachniophyta and Chlorophyta (Table 1), but nothing is known for hydroxylation of ΅-carotene. From the chemical structures of siphonaxanthin [12], loroxanthin, prasinoxanthin and monadoxanthin, it could be considered that they are derived from lutein, but the pathways and enzymes are still unknown (Figures 1 and 2).
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**4. Function of Carotenoids**
For photosynthesis, both carotenoids and chlorophylls are necessarily bound to peptides to form pigment-protein complexes in the thylakoid membrane. Five main kinds of the complexes described below are isolated from some algae, and the pigment compositions are investigated [73–75]. Exceptionally in cyanobacteria, myxol glycosides and some carotenoids are located in the cytoplasmic membrane for protection from high-light [76,77].
Ά-Carotene is presented in the most divisions of the reaction-center complexes (RC) and the light-harvesting complexes (LHC) of photosystem I (PSI) as well as the RC and the core LHC of photosystem II (PSII); exceptionally zeaxanthin is presented in some red algae of the LHC of PSI.
On the other hand, in the peripheral LHC of PSII, the bound carotenoids are heterogenous depending on the classes. Major carotenoids are alloxanthin (Cryptophyta); fucoxanthin (Chrysophyceae, Raphidophyceae, Bacillariophyceae, Phaeophyceae and Haptophyta); diadinoxanthin and vaucheriaxanthin (Xanthophyceae); violaxanthin and vaucheriaxanthin (Eustigmatophyceae);
peridinin (Dinophyta); diadinoxanthin (Euglenophyta); siphonaxanthin (Chlorophyceae and Ulvophyceae); and lutein, violaxanthin and 9<sup>ȝ</sup>-*cis* neoxanthin (land plants) (Figure 1) [73–75]. Ά-Carotene in both RC might have protective functions, and carotenoids in the peripheral LHC of PSII mainly might have light-harvesting functions.
The dimeric cytochrome *b*6*f* complexes of the cyanobacterium *Mastigocladus laminosus* [78] and the green alga *Chlamydomonas reinhardtii* [79] contain two Ά-carotene and two chlorophyll *a* molecules, while that of the cyanobacterium *Synechocystis* sp. PCC 6803 contains two echinenone and two chlorophyll *a* molecules [80]. These carotenoids might have protective functions.
The water-soluble peripheral LHC of peridinin-chlorophyll-protein (PCP) isolated from *Amphidinium carterae* (Dinophyta) has a trimeric structure, and the monomer contains eight peridinin and two chlorophyll *a* molecules [81]. The water-soluble orange carotenoid protein (OCP) isolated from the cyanobacterium *Arthrospira maima* forms a homodimer with two 3<sup>ȝ</sup>hydroxyechinenone molecules [82]. OCP is also found in some cyanobacteria, and its function might regulate energy dissipation from phycobilisomes to PSII [83].
The keto groups at C-8 of fucoxanthin [84], siphonaxanthin [85,86] and prasinoxanthin [87], which are found only in algae, are the single-bond *trans*-conformation for the conjugated double bond
(Figure 1). From the femtosecond time-resolved fluorescence spectroscopy of the purified carotenoids in organic solvents and the LHC in solution, these keto-carotenoids and peridinin have been found to have highly efficient energy transfer from the S1 state, not the S2 state, of carotenoids to chlorophylls. From the comparison of other structural carotenoids, these keto groups are essential for high
efficiency [88,89]. These keto-carotenoids mainly might have light-harvesting functions.
The xanthophyll cycle, also known as the violaxanthin cycle, is the cyclical interconversion of violaxanthin, antheraxanthin and zeaxanthin in green algae and land plants (Figure 2) [90]. Zep catalyzes zeaxanthin to violaxanthin through antheraxanthin during biosynthesis. Violaxanthin is found in the peripheral LHC of PSII. Under high light conditions, Vde is activated and catalyzes de-epoxidation of violaxanthin to zeaxanthin through antheraxanthin. Zeaxanthin is used for the dissipation of excess energy from excited chlorophylls. Zep from Chlorophyceae *Chlamydomonas reinhardtii* [48] and Vde from Pracinophyceae *Mantonilla squamata* [49] are functionally confirmed (Table 2). Similarly, the diadinoxanthin cycle occurs in Heterokontophyta, Haptophyta and Dinophyta, which contain diadinoxanthin and diatoxanthin (Figure 2). The enzymes of diadinoxanthin de-epoxidase and diatoxanthin epoxidase have not yet been found [9,91], but the characteristics of partially purified diadinoxanthin de-epoxidase from the diatom *Cyclotella meneghinaina* are reported [92].
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**Ana Catarina Guedes 1, Helena M. Amaro 1 and Francisco Xavier Malcata 2,3,\***
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*Received: 14 March 2011 / Accepted: 14 April 2011 / Published: 20 April 2011*
**Abstract:** Marine microalgae constitute a natural source of a variety of drugs for pharmaceutical, food and cosmetic applications—which encompass carotenoids, among others. A growing body of experimental evidence has confirmed that these compounds can play important roles in prevention (and even treatment) of human diseases and health conditions, e.g., cancer, cardiovascular problems, atherosclerosis, rheumatoid arthritis, muscular dystrophy, cataracts and some neurological disorders. The underlying features that may account for such favorable biological activities are their intrinsic antioxidant,
anti-inflammatory and antitumoral features. In this invited review, the most important issues regarding synthesis of carotenoids by microalgae are described and discussed—from both physiological and processing points of view. Current gaps of knowledge, as well as technological opportunities in the near future relating to this growing field of interest, are also put forward in a critical manner.
**Keywords:** lutein; astaxanthin; Ά-carotene; bioproduction; extraction
## **1. Introduction**
Microalgae occupy the bottom of the food chain in aquatic ecosystems; they possess the intrinsic ability to take up H2O and CO2—which, with the aid of sunlight, are converted to complex organic compounds that are subsequently kept inside or released from the cell. Those microorganisms have a worldwide distribution, and are well-adapted to survive under a large spectrum of environmental stresses— including (but not limited to) heat, cold, drought, salinity, photo-oxidation, anaerobiosis, osmotic pressure and UV exposure [1].
Microalgae combine, in a balanced way, a few properties typical of higher plants (*viz*. efficient oxygenic photosynthesis and simple nutritional requirements) with biotechnological attributes proper of microorganisms (*viz*. fast growth rates, and ability to accumulate or secrete primary and secondary metabolites). This rather useful combination has led to selection of such microorganisms for applied processes, and represents the basic rationale for the usefulness of microalgal biotechnology. Besides being currently used as feed for aquatic and terrestrial animals, the nutritional value of microalgal biomass goes well beyond—and includes use as colorant in aquaculture, and high-protein or polyunsaturated fatty acid supplement in human diets. The food, pharmaceutical and cosmetic markets have accordingly benefited from a growing array of microalgal products [2,3].
Furthermore, the large number of existing species of microalgae constitutes a unique reservoir of biodiversity, which supports potential commercial exploitation of many novel products besides vitamins, pigments and polyunsaturated fatty acids [4–6]. The key factor for their eventual economic feasibility is the possibility of operating large photobioreactors, able to handle biomass and metabolites to sufficiently high levels [7,8].
This review covers the most relevant features of a family of specialty products originated in microalgae that have already reached commercial expression—by presenting bioprocess considerations and reviewing practical applications, mainly in the food and health industries.
## **2. Cellular Location and Function**
Carotenoids constitute a class of terpenoid pigments, derived from a 40-carbon polyene
chain, which can be envisaged as their molecular backbone—indeed it provides carotenoids with distinctive molecular structures, and the associated chemical properties including light-absorption features that are essential for photosynthesis and, in general, for life in the presence of oxygen [9]. The aforementioned backbone may be complemented by cyclic groups (rings) and oxygen-containing functional groups. Hence, hydrocarbon carotenoids are denoted as carotenes as a whole, but oxygenated derivatives are known specifically as xanthophylls—with oxygen being present as –OH groups (e.g., lutein), as oxi-groups (e.g., cantaxanthin) or as a combination of both (e.g., astaxanthin) [9]. All xanthophylls synthesized by higher plants—e.g., violaxanthin, antheraxanthin, zeaxanthin, neoxanthin and lutein, can also be synthesized by green microalgae; however, these possess additional xanthophylls, e.g., loroxanthin, astaxanthin and canthaxanthin. Diatoxanthin, diadinoxanthin and fucoxanthin can also be produced by brown algae or diatoms [10].
A distinction is usually made between primary and secondary carotenoids: primary ones (*i.e*., xanthophylls) are structural and functional components of the cellular photosynthetic apparatus, so they are essential for survival [10]; whereas secondary ones encompass those produced by microalgae to large levels, but only after exposure to specific environmental stimuli (via carotenogenesis).
Xanthophylls are relatively hydrophobic molecules, so they are typically associated with membranes and/or involved in non-covalent binding to specific proteins; they are usually localized in the thylakoid membrane, whereas secondary carotenoids are found in lipid vesicles—in either the plastid stroma or the cytosol. Most xanthophylls in cyanobacteria and oxygenic photosynthetic bacteria are associated with chlorophyll-binding polypeptides of the photosynthetic apparatus [11]; however, in most green microalgae, carotenes and xanthophylls are synthesized within plastids, and accumulate therein only. Conversely, secondary xanthophylls in some green microalgae—e.g., astaxanthin in *Haematococcus* sp., accumulate in the cytoplasm; this realization raises the possibility of an extraplastidic site of carotenoid biosynthesis in that genus. Alternatively, xanthophylls synthesized in the chloroplast may be exported, and consequently accumulate in the cytoplasm [10,12,13]—so they may be found in essentially all cellular compartments.
Carotenoids perform several functions in microalgae: they are involved in light harvesting, but also contribute to stabilize the structure and aid in the function of photosynthetic complexes—besides quenching chlorophyll triplet states, scavenging reactive oxygen species and dissipating excess energy [14]. The intrinsic antioxidant activity of carotenoids constitutes the basis for their protective action against oxidative stress; however, not all biological activities claimed for carotenoids relate to their ability to inactivate free radicals and reactive oxygen species.
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**3. Practical Applications**
Several researchers have actively focused on carotenoids from microalgal sources; the major areas, in terms of actual or potential industrial applications, are food and health—and the antioxidant properties exhibited by that class of compounds constitutes at present its core interest. Pigments of microalgal origin are indeed experiencing a strong market demand: the price of microalgal Ά-carotene easily attains 700 €/kg, whereas its synthetic counterpart cannot reach more than half that figure. Natural Ά-carotene is preferred by the health market because it is a mixture of *trans* and *cis* isomers—the latter of which possess anticancer features; such a mixture can hardly be obtained via chemical synthesis [14].
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*3.1. Uses for Food and Feed Formulation*
Manufacture of carotenoids via microbiological routes has undergone a greater and greater scientific and commercial importance within the alimentary and aquaculture fields [15], especially in view of environmental and health awareness by consumers at large.
Recall that most oxidation reactions in foods are deleterious—e.g., degradation of vitamins, pigments and lipids, with consequent loss of nutritional value and development of off-flavors [16,17]. Antioxidants—which are adventitious in, or deliberately added to foods, can inhibit oxidation or slow down initiation by free alkyl radicals, as well as interrupt propagation of such free radical chains. The threshold of synthetic food additives legally permitted has been steadily decreasing, due to their suspected role as promoters of carcinogenesis, besides claims of liver and renal toxicities [18]; hence, substitution thereof by natural pigments has become common practice. One good example is the application of *Dunaliella* spp. for mass production of carotenoids aimed at a preservation role [19,20]. Another advantage of carotenoids is that they are not affected by the presence of ascorbic acid, often used as acidulant to constrain unwanted microbial growth, nor by heating/freezing cycles employed in foods with a similar goal.
On the other hand, carotenoids are particularly strong dyes, even at levels of parts per million. Specifically, canthaxanthin, astaxanthin and lutein from *Chlorella* have been in regular use as pigments, and have accordingly been included as ingredients of feed for salmonid fish and trout, as well as poultry—to enhance the reddish color of said fish or the yellowish color of egg yolk [4,21–23]. Furthermore, Ά-carotene has experienced an increasing demand as pro-vitamin A (retinol) in multivitamin preparations; it is usually included in the formulation of healthy foods, although only under antioxidant claims [24–26].
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*3.2. Uses for Health and Well-Being Promotion*
In the human being, oxidation reactions driven by reactive oxygen species can lead to protein damage and DNA decay or mutation; these may in turn lead to several syndromes, *viz*. cardiovascular diseases, some kinds of cancer and degenerative diseases, and ageing at large [17,27]. As potent biological antioxidants, carotenoids are able to absorb the excitation energy of singlet oxygen radicals into their complex ringed chain—thus promoting energy dissipation, while protecting tissues from chemical damage. They can also delay propagation of such chain reactions as those initiated by degradation of polyunsaturated fatty acids—which are known to dramatically contribute to the decay of lipid membranes, thus seriously hampering cell integrity [21].
One illustrative example is the decline of cognitive ability accompanying Alzheimer's disease, which is apparently caused by persistent oxidative stress in the brain [28]. Using transgenic mice fed with extracts from *Chlorella* sp. containing Άcarotene and lutein, Nakashima *et al.* [29] claimed significant prevention of cognitive impairment. Wu *et al*. [30] used also *Chlorella* extracts containing 2–4 mg/gDW of lutein, and reported reduction in the incidence of cancer, as well as prevention of macular degeneration [31]. Likewise, carotenoids extracted specifically from *Chlorella ellipsoidea* and *Chlorella vulgaris* inhibited colon cancer development [23]. Furthermore, astaxanthin obtained from *Haematococcus pluvialis* decreased expression of cyclin D1, but increased that of p53 and some cyclin kinase inhibitors of colon cancer cell lines [32].
Carotenoids have also the ability to stimulate the immune-system, thus being potentially involved in more than 60 life-threatening diseases—including various form of cancer, coronary heart diseases, premature ageing and arthritis [33]; this is specifically the case of canthaxanthin and astaxanthin, and other nonprovitamin A carotenoids from *Chlorella* but to a lesser degree [23]. A few epidemiological studies encompassing Ά-carotene from *Dunalliela* sp.—which contains readily bioavailable 9-*cis* and all-*trans* stereoisomers (*ca*. 40% and 50%, respectively), have indeed provided evidence of a lower incidence of several types of cancer and degenerative diseases [34]. Finally, carotenoids exhibited hyperlipidemic and hypercholesterolemic effects [19].
## **4. Industrial Production**
The worldwide demand for carotenoids was *ca*. 640 M€ in 2004, but it has been rising ever since at an average yearly rate of 2.2% [9]; Ά-carotene has specifically risen from *ca*. 175 M€ in 2004 to
*ca*. 183 M€ in 2009 [35]. A growing fraction has been accounted for by carotenoids from biotechnological sources; and Ά-carotene, as well as such xanthophylls as astaxanthin, cantaxanthin and lutein have consequently been in higher and higher demand [9]. The most famous source microalgae are *Chlorella*, *Chlamydomonas*, *Dunaliella*, *Muriellopsis* and *Haematococcus* spp.—all of which belong to the Chlorophyceae family [2]. They tend to accumulate carotenoids as an intrinsic part of their biomass, thus offering economical alternatives to chemical synthesis [36].
Among all natural sources studied to date, *Dunaliella* possesses the highest content of 9-*cis* Ά-carotene [20,34]—reaching levels up to 100 g/kgDW, [19,37,38]; Άcarotene-rich *Dunaliella* powder has been commercially exploited in many countries since the 1980s. Although many microalgae can produce xanthophylls, *H. pluvialis* is the one that accumulates them to the highest levels (e.g., asthaxanthin [10]), so it is now cultivated at large scale by several companies using distinct approaches [39]. On the other hand, *Muriellopsis* sp. holds a high lutein content (up to 35 mg Lƺ1), coupled with a high growth rate; hence, it has been exploited for commercial production of lutein [10]. Finally, *C. ellipsoidea* was reported to produce violaxanthin, together with two other minor xanthophylls, *viz*. antheraxanthin and zeaxanthin— whereas the main carotenoid in *C. vulgaris* was lutein [23]. Further pieces of related information are gathered in Table 1.
**Table 1.** Carotenoids produced by selected microalgae.
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**5. Biotechnological Processes**
Despite a few useful features already referred to above, microalgae are in general expensive to produce, so concerted efforts have been on the way toward more costefficient modes of mass cultivation.
With regard to open systems, the best choice seems to be the open shallow pond— made of leveled raceways, 2–10 m wide and 15–30 cm deep, which run as simple loops or meandering pathways; each unit may cover an area of several hundred to a few thousand m2. However, this configuration poses several problems—which restrict its use to strains that, in view of their weed-like behavior (e.g., *Chlorella*) or their ability to withstand adverse growing conditions (e.g., *Spirulina* or *Dunaliella*), can outgrow other microorganisms.
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**Table 2.** Optimal conditions of production of carotenoids by selected microalgae.
**Table 2.** *Cont.*
**AF**: air flow; **LDC**: light/dark cycle; **LI**: light irradiance; **MM**: metabolic mode; **SR**: stirring rate; **T**: temperature.
More advanced technologies have meanwhile been made available pertaining to closed systems; these provide better options for growth of most microalgal strains, by protecting the culture from contamination by unwanted (and often ill-defined) microorganisms, and allowing comprehensive and integrated control of processing conditions. Such photobioreactors are either flat or tubular, and may adopt a variety of designs and operation modes. They lead to higher volumetric productivities and an overall better quality for the biomass (or product) generated—but they are also more expensive to build and operate than their open counterparts [9].
Some microalgae exhibit unique productivity and plasticity features: when grown under distinct sets of operating conditions, they may accumulate different products to high levels; hence, careful design and control of medium composition, temperature, pH, aeration, stirring and irradiance are recommended. A few examples of optimum conditions of operation of microalgal reactors—using productivity of carotenoids as objective function, are listed in Table 2.
During microalgal cultivation, a few processing parameters can be specifically manipulated for maximum synthesis of carotenoids; the better studied cases are lutein, astaxanthin and Ά-carotene—which will be discussed below at some length**.**
## *5.1. Lutein*
The most important factors that affect lutein content in microalgae are temperature, irradiance, pH, availability and source of nitrogen, salinity (or ionic strength) and presence of oxidizing substances (or redox potential); however, specific growth rate also plays a crucial role.
High temperature favors accumulation of lutein, as happens with other carotenoids (e.g., Ά-carotene) in *Dunaliella* sp. [42], close to the limit of environmental stress; further temperature increases would thus be harmful, and eventually reduce biomass productivity.
A high irradiance level appears beneficial—but its effect depends on whether indoor or outdoor cultivation is considered; *in vitro* mimicking of all parameters that characterize outdoor operation, e.g., solar cycle and temperature fluctuation, is indeed difficult. Furthermore, the concentration of molecular oxygen outdoors cannot be manipulated, despite its interacting with illumination and temperature. Both irradiance and temperature influence the rate of lutein production, yet cultures of *Murielopsis* sp. and *Scenedesmus almeriensis* produced contradictory results; hence, these two factors should be considered in a combined, rather than independent fashion [8].
Likewise, the reported effects of pH are not consistent between batch and continuous cultivations. In the former, lutein content increased at extreme pH values, whereas the best results under continuous operation were observed at the optimum pH for growth rate. It is worth noting that pH is particularly relevant in microalgal cultures because it interferes with CO2 availability (which is essential for photosynthesis); hence, continuous supply of CO2, as a fraction of the aeration stream, and pH-controlled injection lead to different results. In general, the maximum lutein productivity is achieved at the optimum pH for biomass productivity [45].
The concentration of nitrogen in the culture medium (in the form of nitrate) does not apparently cause a significant effect upon the lutein content of biomass; however, N-limitation reduces biomass productivity, and consequently leads to poor overall lutein synthesis. Hence, nitrate should be supplied to a moderate excess—so that growth rate is not hampered, while avoiding saline stress that dramatically affects culture performance [8].
Lutein synthesis is enhanced via addition of such chemicals as H2O2 and NaClO, which behave as inducers: in the presence of Fe2+, they affect the redox state and generate stress-inducing chemical species. This induction of oxidative stress is expected because lutein holds a protection role conveyed by its antioxidant features—particularly under heterotrophic growth, where spontaneous oxidative stress is normally absent (unlike happens with phototrophic cultures)[45].
Finally, the specific growth rate affects both continuous and semicontinuous cultures: lutein tends to accumulate at low dilution rates, but not to levels sufficient to balance the decrease in biomass productivity under such circumstances. Therefore, the maximum lutein productivity is again typically attained at the optimal dilution rate for biomass production [45].
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*5.2. Astaxanthin*
Commercial production of astaxanthin by *Haematococcus* sp. has been implemented by more than one microalga company (e.g., Cyanotech and Aquasearch); they resorted to a two-stage system, consisting of a first step to produce green biomass under optimal growth conditions ("green" stage), followed by a second stage when the microalga is exposed to adverse environmental conditions to induce accumulation of astaxanthin ("red" stage) [50]. Astaxanthin productivities in large scale facilities are typically *ca*. 2.2 mg Lƺ1 [39]—even though maximum astaxanthin productivities of 11.5 mg Lƺ1 dƺ1 can be attainedat bench scale [51].
Micro Gaia, a marine biotech firm engaged in production of microalgae rich in astaxanthin, proposed a single-step, continuous manufacture process using moderate nitrogen limitation [52,53]: the biomass and astaxanthin productivities obtained were 8.0 and 0.7 mg Lƺ<sup>1</sup> dƺ1, respectively [54]. The feasibility of the latter approach for production of astaxanthin by *H. pluvialis* was tested continuous-wise in outdoor apparatuses [48]: Aquasearch Growth Modules (AGM)—*i.e*., 25,000 L enclosed, computerized photobioreactors, were combined up to three units to obtain large amounts of clean, fast growing *H. pluvialis*; they were transferred daily to a pond culture system, where carotenogenesis and astaxanthin accumulation were induced. After 5 days of synthesis, cells were harvested by gravitational settling— with a typical content of 2.5% (w/wDW) astaxanthin; a high pressure homogenizer was used to disrupt the cells, and then drying was carried out to less than 5% (w/w) moisture. The performance of AGM could be improved 2-fold within the first 9 mo of operation; and the biomass concentration increased from 50 to 90 g mƺ2, with associated productivities increasing from 9 to 13 g mƺ2 dƺ1 within the same period [39].
However, the production capacity of *H. pluvialis* was constrained by its intrinsic slow growth, low cell yield, ease of contamination by bacteria and protozoa, and susceptibility to adverse weather conditions [5]. Moreover, *H. pluvialis* cannot be efficiently cultivated in dark heterotrophic mode—so production of astaxanthin should adopt the photosynthetic mode, and thus resort to levels of irradiance (e.g., 950 μmol mƺ2 sƺ1) well beyond what would be economically reasonable [39]. Owing to its ease of culturing and high tolerance to environmental fluctuations, *C. zofingiensis* (another green microalga) has been put forward as an alternative for astaxanthin production: it grows quite fast (*ca*. three times faster than *H. pluvialis*), and accumulates significant amounts of secondary carotenoids in the dark, thus facilitating large-scale cultivation of denser biomass [47,55].
Oxidative stress induced by intense illumination has been found to play a crucial role upon astaxanthin synthesis [56]; active oxygen molecules, generated by excess photooxidation caused by high light irradiance, do apparently trigger synthesis of carotenoids as part of a cellular strategy aimed at cell protection against oxidative damage [47]. In particular, flashing light increased the rate of astaxanthin production per photon in *H. pluvialis* by at least 4-fold relative to that under continuous light sources [57]—thus proving that light quality is more important than quantity [58].
The effect of irradiance depends also on such operating variables as culture density, cell maturity (flagellates are much more sensitive than palmelloids), medium nutrient profile and light path[59]. The predominant role of light stress and nitrogen deprivation towards induction and enhancement of biosynthesis in the aplanospores of *H. pluvialis* was originally suggested in the 1950s [60]; astaxanthin accumulation comes along with growth halting, as happens in most cases of stress imposed upon microalgae [59,61]. Imamoglu *et al.* [54] compared the effect of various stress media, under high light intensities, upon astaxanthin accumulation; those authors concluded that addition of CO2 in an N-free medium, under 546 μmolphoton mƺ<sup>2</sup> sƺ1, were the best conditions for accumulation of astaxanthin—which attained *ca*. 30 mg gƺ1.
Astaxanthin may thus be efficiently produced outdoors in continuous mode, if accurate nitrate dosage is provided [48]; besides N, such oligoelements as iron play a role. This essential oligoelement takes part in assimilation of nitrate and nitrite, deoxidation of sulphate, fixation of N, and synthesis of chlorophyll [62–65]. Iron deficiency was reported to constrain microalga growth, even in rich nutrient media [64]; whereas its addition enhanced astaxanthin synthesis [66–69]. Cai *et al*. [67] further tested how iron electrovalencies and counter ions affect cell growth and accumulation of astaxanthin; 18 μmol Lƺ1 Fe2+-EDTA stimulated synthesis of astaxanthin more effectively, up to contents of 30.7 mg gƺ1; and despite the lower cell density attained (2.3 × 105 cell mLƺ1), a higher concentration (36 μmol Lƺ1) of FeC6H5O7 yielded cell density and astaxanthin production levels that were 2- and 7-fold those reached under iron-limitation.
In the "red stage" of growth, *Haematococcus* cells require only carbon as major nutrient—which this is usually supplied via directly injecting CO2 into the photobioreactor during daylight [61]. Furthermore, high irradiance provides more energy for photosynthetic fixation of C, which leads to a higher rate of astaxanthin synthesis [68];this may be further enhanced by raising the C/N ratio [69].
Finally, Chen *et al.* [70] experimented with heterotrophic conditions—using pyruvate, citrate and malate as substrates, towards synthesis of astaxanthin by *C. zofingiensis* in the absence of light. Presence of any of the aforementioned substrates above 10 mM stimulated biosynthesis of astaxanthin (and other secondary carotenoids); *ca*. 100 mM pyruvate led to yields of 8.4–10.7 mg Lƺ1 astaxanthin, which correspond to a 28%-increase.
## *5.3. Ά-Carotene*
Semicontinuous cultivation of *D. salina* at 25 °C produced 80 g mƺ3 dƺ1 biomass [42]—from which 1.25 mg Lƺ1 of Ά-carotene was recovered [71]; however, this figure could be improved up to 2.45 mg mƺ3 dƺ1 in continuous biphasic bioreactors [72]. When cultivated photoheterotrophically, a significant increase of cellular Ά-carotene content was experimentally observed: the maximum score was 70 pg cellƺ1, in a culture enriched with 67.5 mM acetate and 450 μM FeSO4 [33].
As with astaxanthin, Fe2+ plays an important role in Ά-carotene accumulation in *D. salina*; by inducing oxidative stress, those cations stimulate said synthesis, especially in the presence of a carbon source. UV-A radiation (320–400 nm) added to the photosynthetically active radiation (PAR, *i.e*., 400–700 nm) can be regarded as another stress factor during growth of, and carotenoid accumulation by *Dunalliela bardawil;* compared with cultures exposed to PAR only, addition of 8.7 W mƺ2 UV-A radiation to 250 W mƺ2 PAR stimulated long-term growth of that microalga, and led to a 2-fold enhancement in Ά-carotene accumulation by 24 d [38].
## **6. Extraction and Purification**
Although microalga-mediated synthesis of carotenoids is crucial in biotechnological production thereof, a major portion (if not most) of their cost actually lies on downstream separation—e.g., biomass drying and disruption, followed by solvent extraction and purification. Hence, these issues are addressed below, in view of their importance toward commercial scale processes.
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*6.1. Cell Disruption*
A major practical problem in using such microalgae as *Murielopsis* sp. or *S. almeriensis* is the need for cell wall disruption. This can be accomplished through a variety of ways, e.g., milling, ultrasound, microwave, freezing, thawing or chemical attack [45].
The mortar-and-pestle procedure described by Mínguez-Mosquera *et al*. [73] provides full recovery, but it cannot be scaled up to industrial practice; sonication and ball milling produce results similar to that procedure, as long as alumina is employed as disaggregating agent [45]. Ceron *et al*. [74] complemented the aluminabased cell disruption with alkaline treatment using 4% (w/v) aqueous KOH (40 °C); disaggregation and lipid expression were both facilitated.
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*6.2. Biomass Extraction*
Microalgal biomass is usually processed via solvent extraction, to render carotenoid extracts—with typical contents of 25% [45]; this can be used directly in the formulation of supplements, or undergo further multistep purification— encompassing hydrolysis to release hydroxylated carotenoids from the accompanying fatty acids, and final recrystallization to polish the product.
Obtaining a carotenoid-rich oleoresin from microalgae—dried or in wet paste form, is a more straightforward task; such extracts may then be subjected to classical processes to obtain purer lutein [45,74] that may successfully compete with that extracted from marigold.
## 6.2.1. Organic Solvent-Mediated Extraction
Solvent extraction usually resorts to hexane—and has advantages over alkaline treatment because all lutein and zeaxanthin are converted to their free forms, while carboxylic acids and chlorophylls remain in the aqueous phase [45]; this method has been optimized for *S. almeriensis* [74]. Extraction was maximized with a 1:1 (v/v) ratio of hexane to sample, and the optimal number of extraction steps was typically six— which led to 95% recovery of lutein. Less conventional solvents—e.g., ethyl lactate, have been recently proposed [76] for plant matter at large, but can in principle be applied also to microalgae.
A significant improvement would be to eliminate the drying step of microalgal biomass prior to extraction; Fernández-Sevilla *et al*. [77] have accordingly proposed a modification of a previous approach [74] that can handle wet biomass paste (*ca*. 20% DW), based on an extraction phase composed by hexane/ethanol/water and KOH—which simultaneously effects an alkaline treatment to saponify susceptible lipids and extract the intended carotenoids.
Another enhancement is the accelerated solvent extraction methodology, which uses a special type of contactor to circulate solvent at high pressure through a tightly packed bed of biomass. However, high temperatures are required (over 60 °C, and usually as high as 170 °C) to lower the viscosity of the solvent, which leads to formation of pheophorbide from the microalgal chlorophylls that are of a major toxicological concern. In any case, extraction with hexane or ethanol allows easy solvent removal afterwards, as well as high-content lutein extracts [45].
For selective extraction of free astaxanthin from red encysted *Haematococcus* sp., an alternative procedure has been designed that resorts to dodecane and methanol [75]; it consists of dodecane-mediated extraction of the crude mixture, followed by extraction with methanol. The first stage did not require previous cell harvesting, and separation of the dodecane-rich phase from the culture medium containing cell debris proceeded rapidly via plain settling. In the second stage, the free astaxanthin in the former extract was selectively solubilized in methanol along with saponification of astaxanthin esters—thus leading to a total recovery of astaxanthin above 85%.
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6.2.2. Green Solvent-Mediated Extraction
An environment-friendly downstream process using common vegetable oils was proposed by Kang *et al*. [79] for direct extraction of astaxanthin from *Haematococcus* sp. As said crude microalgal astaxanthin consists of *ca*. 70% monoesters, 25% diesters and 5% free forms, a rather lipophilic nature results, so vigorous stirring is required to gradually disrupt the cells; the oily extracts are then simply separated from the culture medium containing cell debris by gravity settling. When using olive oil, recoveries of up to 93.9% were possible [79]. Apparently, a similar method had been proposed long before by Nonomura [80], who then claimed up to 7.5% yield of lutein.
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6.2.3. Supercritical Fluid-Mediated Extraction
Classical extraction with organic solvents has attained purity degrees sufficient to meet commercial specifications for large-scale production of lutein; however, selective precipitation with supercritical CO2 constitutes a promising alternative. Note that conventional liquid extraction of carotenoids from microalgal matrices is time-consuming—as multiple extraction steps are typically required; and large relative ratios of organic solvents have to be used, which are often expensive and potentially harmful. Supercritical fluid extraction (SFE) using modified CO2 permits more straightforward purification and shorter extraction times [81].
In general, SFE is relatively rapid and efficient because of the low viscosities and high diffusivities that characterize supercritical fluids. Furthermore, extraction can be made selective by controlling solvent density; the material extracted will be recovered afterwards by simply depressurizing, thus allowing the supercritical fluid to return to its gaseous form and leaving no (or little) residual solvent in the precipitate thus originated[82]. Supercritical CO2 has so far been the most employed supercritical fluid—because it is non-flammable, non-toxic, inexpensive and relatively inert from a chemical point of view.
Previous studies demonstrated the feasibility of extracting pigments from plants using supercritical CO2—e.g., carotenoids from carrots [83] and cabbages [84]; Mendes *et al*. [85], Careri *et al*. [86] and Macías-Sánchez *et al*. [87–89] have meanwhile extended such a technique to extraction of carotenoids from *C. vulgaris*, *Spirulina platensis*, *Nannochloropsis gaditana*, *Synechococcus* sp. and *S. almeriensis*, respectively— and satisfactory results were consistently reported, as emphasized in Table 3.
**Table 3.** SFE yields of total carotenoids (including lutein), and of lutein specifically, by selected microalgae.
**Table 3.** *Cont.*
However, this mode of extraction tends to recover chlorophylls more efficiently than carotenoids, thus producing extracts with relatively poor specifications [90]. Furthermore, the cost of supercritical fluids and associated equipment make it difficult for SFE to compete with classical solvent extraction—especially because the former requires dry biomass.
The selective adsorption of lutein might constitute an alternative in terms of separation/purification, especially if specific solid phases can be used [91], coupled with contacting conveyed by expanded beds [92]; this allows raw extracts to be processed, and tolerates the presence of cell debris or other particulate matter that causes major problems in conventional preparative chromatography. Selective precipitation was also described by Miguel *et al*. [93], who proposed use of supercritical CO2 after organic solvent extraction; the first solvent (containing carotenoids) was accordingly mixed with supercritical CO2, and the conditions of pressure and temperature were duly adjusted to promote preferential precipitation of lutein. However, simple standard mixtures—rather than complex microalgal extracts have been considered, so a long way of improvement is still anticipated prior to practical use.
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6.2.4. *In Situ* Extraction
*In situ* extraction of Ά-carotene from *Dunaliella salina* was recently reported by Kleinegris *et al*. [44], using a flat-panel photobioreactor operated as a turbidostat— where the numbers of stressed cells were kept essentially constant via a continuous, well-defined level of irradiation. This two-stage system comprised an organic phase of dodecane, sparged at a rate of 286 Ldodecane Lreactorƺ1 minƺ1 that promoted formation of an emulsion in the aqueous phase; Ά-carotene was then continuously extracted from the aqueous to the organic phase, at a rate of *ca*. 2.75 mgΆ-carotene Ldodecaneƺ1 dƺ<sup>1</sup> (equivalent to 0.7 mgΆ-carotene Lreactorƺ1 dƺ1). However, this process exhibited a poor efficiency—as the yield of Ά-carotene extracted by the solvent was a mere one-tenth of that removed from the reactor via biomass overflow.
If the aforementioned carotenoid-rich biomass was extracted as well, then the overall volumetric productivity of the system would reach 8.3 mgΆ-carotene Lreactorƺ1 dƺ1; this is still below the yield attained if downstream rather than *in situ* extraction was promoted (*ca*. 13.5 mgΆ-carotene Lreactorƺ1 dƺ1) [44], so in this system simultaneous biosynthesis and extraction cannot be justified relative to the classical sequential approach.
## **7. Final Considerations**
Carotenoid production appears to be one of the most successful case studies of blue biotechnology. The rising market demand for pigments from natural sources has promoted large-scale cultivation of microalgae for synthesis of such compounds, so significant decreases in production costs are expected in coming years.
The recognized therapeutic value of some carotenoids (especially lutein) in prevention and treatment of degenerative diseases has indeed opened new avenues for development of mass production systems. Advances in knowledge of the underlying physiology, biochemistry and molecular genetics of carotenoidproducing microalgae are now urged—which would have a major impact upon development and optimization of this (and alternative) microalga-based technologies. In this regard, the genes encoding enzymes that are directly involved in specific carotenoid syntheses need in particular to be investigated—so that further development of transformation techniques will permit considerable increase of carotenoid cellular contents, and accordingly contribute to increase the volumetric productivities of the associated processes.
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**Acknowledgements**
A PhD fellowship (ref. SFRH/BD/62121/2009), supervised by author F.X.M., was granted to author H.M.A., under the auspices of ESF (III Quadro Comunitário de Apoio) and the Portuguese State. A postdoctoral fellowship (ref. SFRH/BPD/72777/2010), also supervised by author F.X.M., was granted to author A.C.G., under the auspices of ESF (III Quadro Comunitário de Apoio) and the Portuguese State. This work received partial financial support via projects OPTIC-ALGAE (PTDC/BIO/71710/2006) and MICROPHYTE (PTDC/EBB-EBI/102728/2008), both coordinated by author F.X.M., also under the auspices of ESF (III Quadro Comunitário de Apoio) and the Portuguese State.
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"author": "",
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**Synthetic Biology and Metabolic Engineering for Marine Carotenoids: New Opportunities and Future Prospects**
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**Chonglong Wang, Jung-Hun Kim and Seon-Won Kim \***
Division of Applied Life Science (BK21 Plus), PMBBRC, Gyeongsang National University, Jinju 660-701, Korea; E-Mails: [email protected] (C.W.); [email protected] (J.-H.K.)
**\*** Author to whom correspondence should be addressed; E-Mail: [email protected]; Tel.: +82-55-772-1362; Fax: +82-55-759-9363.
*Received: 14 June 2014; in revised form: 29 August 2014 / Accepted: 1 September 2014 / Published: 15 September 2014*
**Abstract:** Carotenoids are a class of diverse pigments with important biological roles such as light capture and antioxidative activities. Many novel carotenoids have been isolated
from marine organisms to date and have shown various utilizations as nutraceuticals and pharmaceuticals. In this review, we summarize the pathways and enzymes of carotenoid synthesis and discuss various modifications of marine carotenoids. The advances in metabolic engineering and synthetic biology for carotenoid production are also reviewed, in hopes that this review will promote the exploration of marine carotenoid for their utilizations.
**Keywords:** marine carotenoids; carotenoid synthesis; carotenoid modification; metabolic engineering; synthetic biology; protein engineering
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{
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"isbn": "9783039431908",
"section_idx": 144
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**1. Introduction**
Carotenoids are a class of naturally occurring pigments originated in the chloroplasts and chromoplasts of plants, algae and some photosynthetic microorganisms [1–4]. As of 2004, over 750 known carotenoids, which can be divided into xanthophylls (containing oxygen) and carotenes (pure hydrocarbons), have been isolated from natural sources [5]. These structurally diverse pigments play important biological roles in light capture, protection of cells from the damaging effects of free radicals, and synthesis of many hormones as a precursor [6–10]. Carotenoids are traditionally used as food colorants, animal feed supplements, and, very recently, as nutraceuticals and pharmaceuticals [11,12]. Over the past few decades, researches have supported that the ability of carotenoids to reduce the risk of certain cancers, cardiovascular diseases, and degenerative pathogenesis (e.g., Alzheimer and Parkinson) due to their antioxidative properties [13,14]. According to "Carotenoids: A Global Strategic Business Report" from Global Industry Analysts (GIA), the global market for carotenoids was estimated at approximately \$1.07 billion in 2010 and is projected to top \$1.2 billion by 2015 [15]. Therefore, many efforts have been made to improve the production of these natural compounds for ever-increasing demands [12,16,17].
The ocean is a complex aquatic ecosystem covering about 71% of the Earth's surface, which is around 300 times larger than the habitable volume of the terrestrial habitats on Earth. A large proportion of all life on Earth lives in the ocean. Ecologically distinct from the terrestrial ecosystem, the ocean constitutes a unique reservoir of marine biodiversity and provides a vast resource of foodstuffs, medicines, and other useful materials. As such, more than 250 novel carotenoids have originated from marine species [10], many of which show great potential in commercial applications [18]. With the advent of synthetic biology and metabolic engineering, many engineering tools including vectors, genetic controllers, and enzyme designing, have been developed for heterologous production of valuable chemicals. These tools create new opportunities for exploring marine carotenoids for food and health industries. In this review, we describe diverse and novel carotenoids from marine resources and summarize recent progresses in synthetic biology and metabolic engineering which provide great application potential for marine carotenoids.
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**2. Diversity of Marine Carotenoids**
Many carotenoids have been reported from a wide range of marine species. The advances in current technologies facilitate the elucidation of the carotenoid biosynthetic pathways and relevant enzymes from marine species, which would enable the production of important carotenoids from marine organisms.
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*2.1. Pathways and Diverse Enzymes for Biosynthesis of Carotenoids*
Biosynthetic routes to carotenoids begin with the basic building blocks isopentenyl diphosphate (IPP) and its isomer dimethylallyl diphosphate (DMAPP), although carotenoids are very diverse in chemical structure. Two distinct pathways, the 2- *C*-methyl-D-erythritol 4-phospahte (MEP) pathway and the mevalonic acid (MVA) pathway, are responsible for the synthesis of IPP and DMAPP. These two pathways have been reviewed in detail elsewhere [19–22]. IPP and DMAPP are head-to-tail condensed to generate farnesyl diphosphate (FPP) and geranylgeranyl diphosphate (GGPP) by isoprenyl diphosphate synthases (e.g., IspA of *Escherichia coli* and CrtE of *Pantoea agglomerans*) [23,24]. As shown in Figure 1, FPP and GGPP are further head-to-head condensed to produce symmetric hydrosqualene (C30) and phytoene (C40), which are dehydrogenated in a stepwise manner by desaturating enzymes representing an important branch point for pathway diversification [25,26].
**Figure 1.** Synthesis pathway of phytoene-based C40 carotenoid backbones. Most C40 marine carotenoids are modified from the backbones of ΅/Ά/·-carotenes or isorenieratene. Carotenoid structures are presented with two symmetric or asymmetric halves (Figure 2), for example, lycopene is shown as HoH'o in this review. Conjugated double bonds are shown in red.
Enzymes involved in the biosynthesis of carotenoids have been mainly investigated in carotenogenic cyanobacteria and land plants [27,28]. They are mostly associated with cytoplasmic and organelle membranes where the hydrophobic substrates of carotenogenic enzymes are located [29]. So far, very few crystal structures of carotenogenic enzymes have been elucidated because of their association with the membranes [30,31]. More than 95% of carotenoids have been characterized in nature to be phytoene-based [32], which will be extensively discussed in this review.
Phytoene synthase is positioned early in the carotenoid synthesis pathway and is responsible as a pathway gatekeeper to discriminate GGPP substrate from enormous isoprenyl diphosphates [29]. Phylogenic analysis of 20 phytoene synthases from marine organisms supports the endosymbiotic theory that plastids evolve from a cyanobacterium, which is engulfed and retained by a unicellular protist [33,34]. Cyanobacteria *Acaryochloris marina* and *Prochlorococcus marinus* are clustered with green algae and land plant tomato (Figure 2A). However, phytoene synthases still display a significant diversification by evolution. A consensus position of 24.5% (identity of 0.5%) is remained among phytoene synthases from marine algae, bacteria, Achaea and land plants. There is only a similarity of 31.9% even between the two proteobacteria phyla ΅-proteobacteria and ·-proteobacteria.
The photochemical properties of a carotenoid depend on the size of the chromophore formed by conjugated double bonds, and a C40 backbone can accumulate up to 15 conjugated double bonds [35]. Thus, six sequential desaturation steps are required to dehydrogenate colorless phytoene, which has three conjugated double bonds in the center [26]. Lycopene containing a chromophore with eleven conjugated double bonds is the direct precursor of ΅/Ά/·-carotenes or isorenieratene, the phytoene-based C40 carotenoid backbone (Figure 1). In general, oxygenic phototrophs require three enzymes, phytoene desaturase, Ί-carotene desaturase and *cis*-carotene isomerase to generate lycopene [6]. However, most bacterial phytoene desaturases are able to catalyze all three reactions [30]. There are also some organisms that disobey this general rule. Primitive cyanobacteirum *Gloeobacter violaceus* PCC 7421 uses bacterial type phytoene desaturase, and no homolog of Ίcarotene desaturase or *cis*-carotene isomerase is found in its genome [36,37]. Among anoxygenic phototrophs, green sulfur bacteria use three enzymes to catalyze desaturation, whereas purple bacteria, green filamentous bacteria, and heliobacteria use only one enzyme [38,39]. Phytoene desaturases also exhibit significant diversities among different organisms (Figure 2B). Just as phytoene synthase, green algae are clustered with tomato but they are distinguished from cyanobacteria. There is just a similarity of 23.2% among 21 proteins. It is suggested that phytoene desaturase exhibits a much faster evolution from the ancestral blueprint and higher diversities among species than phytoene synthases, which may correspond to promiscuous activities of phytoene desaturase.
**Figure 2.** Phylogenic trees of (**A**) phytoene synthases and (**B**) phytoene desaturases. Trees were built using MEGA6.0 software by Neighbor-Joining method [40]. Protein sequences were obtained from *National Center for Biotechnology Information (*NCBI).
## *2.2. Diversity of Marine Carotenoids*
Carotenogenic organisms in ocean are algae and bacteria, which possess all the genes for *de novo* synthesis of carotenoids [2–4]. Unicellular microalge *Dunaliella salina* and *Dunaliella bardawil* are rich in the orange pigment Ά-carotene (HaH'a, Figure 1) [41,42]. Two rings of Ά-carotene are often oxidized to form astaxanthin (Ha1H'a1, Figure 3) in some microalgae by Ά-carotene hydroxylase and ketolase [43], which can individually catalyze the modification of Ά-carotene to generate zeaxanthin (Ha2H'a2, Figure 3) in *Spriulina platensis* and *Spriulina maxima* [44], and canthaxanthin (Ha3H'a3, Figure 3) in *Haematococcus pluvialis*, *Clorella vulgaris* and *Colastrella striolata* [44–46]. The modifications can just occur in one ring to generate asymmetric intermediates such as Ά-cryptoxanthin (HaH'a2, Figure 3) and echinenone (HaH'a3, Figure 3). Chlorophyta *Scenedesmus almeriensis* and *Muriellopsis sp*. accumulate a large amount of lutein (Hb1H'a2, Figure 3), which is derived from ΅-carotene (HbH'a, Figure 1) [47]. Cryptophyta also synthesize ΅-carotene as well as acetylenic derivatives crocoxanthin (HbH'a4, Figure 3) and monadoxanthin (Hb1H'a4, Figure 3) [28]. Acetylenic groups are also found in Ά-carotene derivatives alloxanthin (Ha4H'a4, Figure 3) in Cryptophyta [48], and diatoxanthin (Ha2H'a4, Figure 3) and epoxy oxidized diadinoxanthin (Ha4H'a5, Figure 3) in Heterokontophyta, Haptophyta, Dinophyta, and Euglenophyta [28,49,50]. The unique acetylenic carotenoids are only found in algae. In brown algae and diatoms, acetylated and unique allenic modifications produce dinoxanthin (H'a5Ha6, Figure 3) and chain-oxidized fucoxanthin (Ha6H'a7, Figure 3) [2,51]. Some Chlorophyta species modify the methyl group of lutein to generate loroxanthin (Hb1H'a8, Figure 3) in *Scenedesmus obliquus* and *Chlorella vulgaris* [52], and siphonaxanthin (Hb1H'a9, Figure 3) in *Codium fragile* [53]. Aromatic isorenieratene (HcH'c, Figure 1) is a usual biomarker compound, which is synthesized from Ά-carotene in actinobacteria or ·-carotene (H'oHa, Figure 1) in green and purple sulfur bacteria [54,55]. ·-Carotene can also be converted to chlorobactene (HcH'o, Figure 1) and OH-chlorobactene (HcH'o1, Figure 3). Glycoside modifications generate OH-chlorobactene glucoside (HcH'o2, Figure 3) in green sulfur bacteria and myxol 2'-fucoside (Ho3H'a2, Figure 3) in Cyanophyta [54,56]. Dinophyta can synthesize C37-skeletal carotenoids such as peridinin (Hd1H'a6, Figure 3) [57]. Animals do not have pathways for *de novo* synthesis of carotenoids, but they obtain carotenoids from food and further modify carotenoids by oxidation, reduction, translocation of double bonds, cleavage of double bonds, *etc.* Peridininoriginated carotenoids such as peridininol (He1H'd1, Figure 3) and cyclopyrrhoxanthin (He2H'd2, Figure 3) have been isolated from bivalves *Crassostrea gigas*, *Paphia amabillis,* and *Corbicula japonica* [58–60]. Two unique nor-carotenoids, 2- nor-astaxanthin (Hf1H'a1, Figure 3) and actinoerythrin (Hf2H'f2, Figure 3), have been found in sea anemones *Actinia equine* and *Tealia feline* [61]. The carotenoid diversity in marine animals has been well summarized in detail elsewhere [61]. It is also worthy to note that some carotenoids are present in different stereo configurations among organisms (not covered in this review), which also greatly contributes to the diversification of carotenoids.
**Figure 3.** Diverse modifications of carotenoids. The structure shows the modification of a half carotenoid backbone. The stereo configurations are not shown in the structures. Glycosyl moieties of fucose and glucose are represented by Fuc and Glu, respectively.
*2.3. Synthesis of Some Important Marine Carotenoids and Enzymes*
Ά-Carotene as well as xanthophylls astaxanthin, zeaxanthin, lutein, and fucoxanthin are some representative marine carotenoids due to their abundance in marine organisms and their inherent antioxidant properties. Ά-Carotene is synthesized from the cyclization of lycopene, a key step in generating carotenoid diversity by lycopene cyclases, which can also lead to ΅/·-carotene formation (Figure 1). The Ά-cyclase catalyzes the symmetrical formation of two identical Άionone rings of Ά-carotene. On the other hand, ΅-carotene contains two different ring structures (Ή and Ά) formed by the action of additional Ή-cyclase with Ά-cyclase. Four distinct families of lycopene cyclases, CrtY-type Ά-cyclases in proteobacteria, CrtL Ά/Ή-cyclases in some cyanobacteria, the heterodimeric cyclases in some Grampositive bacteria and FixC dehydrogenase superfamily lycopene cyclases in *Chlorobium tepidum* and *Synechococcus sp*. PCC 7002, have been identified to date [62]. Further decorations occur via a variety of ketolation (oxidation), hydroxylation (Figure 4), which are the major causes for the diversity among carotenoids [29]. Ά-Carotene ketolase (CrtW or CrtO) adds the keto groups at the 4,4<sup>ȝ</sup>-position of the ring and Ά-carotene hydroxylase (CrtZ) adds the hydroxyl group at the 3,3<sup>ȝ</sup>-position [63]. Both enzymes are responsible for the formation of astaxanthin via zeaxanthin or canthaxanthin routes in some cyanobacteria and algae (Figure 3). Lutein formation is ascribed to the hydroxylation of ΅-carotene by cytochrome P450 enzymes in *Arabidopsis thaliana* [64], while the pathway and enzymes remain to be elucidated from marine organisms. Fucoxanthin with a unique allenic and epoxide structure is derived from zeaxanthin in brown seaweeds, diatoms and dinoflagellates. Genome analysis indicates that zeaxanthin epoxidases epoxidize zeaxanthin to form antheraxanthin (Ha2H'a5, Figure 3) and violaxanthin (Ha5H'a5, Figure 3) [65]. Two possible routes have been proposed for the synthesis of fucoxanthin from violaxanthin via neoxanthin (He1H'a5, Figure 3) or diadinoxanthin [66]. Very recently, a cytochrome P450-type carotene hydroxylase (PuCHY1) has been isolated from red alga *Porphyra umbilicalis*. The compensatory expression of PuCHY1 results in the formation of violaxanthin, neoxanthin, and lutein in *A*. *thaliana* by the Ά/Ή-hydroxylation activities [67]. Some of the carotenogenic enzymes characterized from marine organisms have been summarized in the literature [28].
**Figure 4.** Synthetic pathway of astaxanthin, lutein, and fucoxanthin from lycopene. Arrows indicate each catalysis reaction, and enzymes are shown in blue. Dashed arrows indicate hypothesized reactions. Reaction intermediates are shown in gray.
## **3. Technology Developments for Production of Carotenoids**
Over the decades, many researches have been done for the production of carotenoids. Carotenogenic pathways have been identified and manipulated in several organisms, and advances in metabolic engineering and synthetic biology have resulted in significant improved production of carotenoids including astaxanthin, zeaxanthin, and lutein.
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*3.1. Easy Colorimetric Screening of Production of Carotenoids*
Carotenoids contain chromophores absorbing visible light and appear as being yellow (e.g., Ά-carotene) to red (e.g., lycopene), which benefits carotenogenic gene mining and engineering upon carotenoid synthesis pathway. To date, many carotenoid biosynthetic genes have been cloned from plants, bacteria, and fungi based on their abilities to render different colors to the host [68–70]. This merit has been vigorously implemented for random mutagenesis, directed evolution, and proofof-principle experiments in synthetic biology. Moreover, cellular carotenoids can be easily extracted into an organic solvent and differentiated in a sensitivity of submilligrams per liter with a linear correlation between carotenoid contents and color intensity [71,72]. This provides an easy and high-throughput way to evaluate the performance of newly built synthetic circuits or methodologies for improved biosynthesis of carotenoid (Figure 5A).
**Figure 5.** Technologies and efforts to improve carotenoid production. ( **A**) Colorimetric screening of desired producer; ( **B**) Pathway engineering approaches for production improvement; ( **C**) Genetic modifications for host strain development; ( **D**) Protein engineering for enzyme and pathway improvement.
*Mar. Drugs* **2014**, *12*, 4810–4832
*3.2. Pathway Engineering for Production of Carotenoids*
Carotenoid biosynthesis emerges from the central isoprenoid pathway, either the MEP pathway or the MVA pathway, existing in all organisms [19,22]. The expression of carotenogenic genes can yield carotenoids of interest in a heterologous organism [16,73–75]. The early attempts led to the production of lycopene, Ά-carotene, and astaxanthin in *Saccharomyces cerevisiae* and *Candida utilis* by the expression of carotenogenic enzymes from *Pantoea ananatis* [74,76]. *Corynebacterium glutamicum* is a native producer of decaprenoxanthin and its glucosides, and it has been engineered to synthesize C50 carotenoids C.P.450 and sarcinaxanthin [77]. To date, there have been many exemplary illuminations to achieve high carotenoid titers from non-native producers. Carotenogenic enzymes from different sources exhibit different capacities in carotenoid biosynthesis. A two-fold higher lycopene production is obtained in *E*. *coli* by the expression of carotenogenic enzymes from *<sup>P</sup>*. *agglomerans* (27 mg/L) than from *<sup>P</sup>*. *ananatis* (12 mg/L) [78]. Metabolic engineering approaches allow the assembly of genes from different organisms for production purposes or for building new carotenoids [32,79,80]. Ά-Carotene production has been improved by hybrid expression of carotenogenic genes from *<sup>P</sup>*. *agglomeras* and *P*. *ananatis* in *E*. *coli* [81]. In another example, expression of Ά-end ketolase from *Agrobacterium aurantiacum* extends the zeaxanthin Ά-D-diglucoside pathway from *<sup>P</sup>*. *ananatis*, and synthesizes novel astaxanthin Ά-D-diglucoside and adonixanthin Ά-
D-diglucoside [29]. Generally, a sufficient precursor supply is a prerequisite for highyield production of carotenoids. Overexpression of the rate-limiting enzymes 1- deoxy-D-xylulose-5-phosphate synthase and reductoisomerase led to a 3.6-fold increase in lycopene production in *E*. *coli* when compared with the native MEP pathway for IPP and DMAPP supply [71]. Overexpression of the rate-limiting enzyme 3-hdroxy-3-methyl-glutaryl-coenzyme A (HNG-CoA) reductase of the MVA pathway from *Xanthophyllomyces dendrorhous* significantly increased Ά-carotene production in *S*. *cerevisiae* [82]. A great effort in metabolic engineering of the central carotenoid building block pathway is the introduction of a hybrid MVA pathway of *Streptococcus pneumonia* and *Enterococcus faecalis* into *E*. *coli*, which enables the recombinant host to produce 465 mg/L of Ά-carotene [83]. With more available genetic tools, microbial organisms such as *Pseudomonas putida* and *Bacillus subtilis* have also been developed as platform hosts for carotenoid production [84,85].
Carotenoids synthesis involves multiple enzymes [2]. The expression level of all the components of a multigene circuit should be orchestrated to optimize metabolic flux and to gain a high yield (Figure 5B) [86]. A random approach is screening of the best orchestra from numerous combinatorial assemblies of required genes and control elements. BioBrick™ paradigm is capable of rapidly assembling a biosynthetic pathway in a variety of gene orders from different promoters in plasmids with different copy numbers [87]. It is possible to build a hybrid carotenoid pathway wherein each enzyme possesses a right turnover number, however, BioBrick™ assembly is still not in a high throughput to create vast combinatorial expression constructs for the best combination of carotenogenic genes. Recently, several advanced assembly methods using homologous recombination, such as sequence and ligation-independent cloning (SLIC), Gibson DNA assembly and reiterative recombination, have been applied to construct multigene circuits [88–90]. These advances promise to randomize all genetic components, including genes, promoters, ribosome binding sites, and other control modules to build a large number of individual genetic circuits for screening purposes. A so-called "randomized BioBrick assembly" approach has been applied to the optimization of the lycopene synthesis pathway wherein the expression construct was designed to independently express each enzyme from its own promoter, which resulted in an increase by 30% in lycopene production [91]. A longer and more complicated pathway can be modularized into subsets, which contain pathway enzymes with similar turnover numbers. Modulating these subsets would be more convenient and efficient than regulating all components of the entire pathway for improved production [92]. By using this multivariate modular metabolic engineering (MMME) approach, recent work achieved a 15,000-fold increase in production of taxadiene, a precursor of the anti-cancer drug taxol [93]. There are also a variety of promising approaches, such as tunable promoters, tunable intergenic regions, and ribosome binding site design, which can be applied to fine tuning the expression of modules [94–96]. In the other approaches, a multi-genic operon is transcribed into a single polycistronic mRNA, and then the large transcript can be spliced to small monocistronic transcripts through post-transcriptional RNA processing such as ribozyme cleavage and *clustered regularly interspaced short palindromic repeats* (CRISPR) editing. Thus, the stability of the monocistronic transcripts can be independently modulated to differentiate the expression level of each enzyme even in a multi-gene operon. These RNA processing tools have been developed as insulating elements between operonic genes to reduce the context dependence of the genes in a polycistronic transcription unit [97]. The diffusion of pathway intermediates can decrease the effective concentrations of intermediates for following enzyme reactions and some intermediates may serve for competing pathways. By learning from Mother Nature, synthetic biologists spatially organize enzymes of the MVA pathway by protein scaffolds in *E*. *coli* to minimize diffusion limitation and achieve a 77-fold increase in mevalonic acid production [98]. The propanediol utilization machinery of *Citrobacter freundii* has been heterologously recasted in *E*. *coli* [99]. Some intermediates of carotenoid synthesis such as isoprenyl diphosphates are toxic when they accumulate over the concentration threshold [100]. To avoid the accumulation of toxic intermediates, genetic sensors can potentially be coupled with gene expression cassettes to regulate the intermediate flux in a dynamic manner. The native *E*. *coli* promoters that respond to the toxic FPP have been successfully used to dynamically regulate the amorphadiene synthesis pathway and improve the production by two-fold over common inducible promoters and constitutive promoters [101]. The Ntr regulon has been engineered to control lycopene synthesis in response to glycolytic flux dynamics, resulting in an 18-fold increase in lycopene production [102].
## *3.3. Genome Engineering for Strain Development*
For the most efficient carotenoid production, the biological system of the host organism also needs to be optimized, by, for example, redirecting cellular carbon flux to the carotenoid synthesis pathway. The *de novo* synthesis of carotenoids is initiated from acetyl-coA by the MVA pathway or glycolytic metabolites pyruvate and glycraldehyde-3-phosphate (G3P) by the MEP pathway. The direct efforts are focused on the modification of associated genes to these pathways. Deletion of pyruvate kinases PykFA can balance the availability of pyruvate and G3P for the MEP pathway, and increase lycopene production by 2.8-fold in *E*. *coli* [103]. The deletion of glucose-6-phospahte (G6P) dehydrogenase Zwf, which branches G6P to pentose phosphate pathway results in an increase by 30% in lycopene production [104]. Deletion of carbohydrate phosphotransferase system yields a seven-fold increase in lycopene production in another study [105]. Replacement of native promoters of the rate-limiting genes of the MEP pathway with the T5 promoter has been carried out for enhancement of the targeted pathway flux, which results in a 4.5-fold increase in Ά-carotene production [106].
A heterologous pathway is not just an independent entity. It communicates with the native cellular metabolism and is therefore governed by the global regulation of the host organisms. Adaptive laboratory evolution is a traditional route for strain engineering to achieve desirable industrially relevant phenotypes. Owing to the antioxidant properties of carotenoids, adaptive evolution has been successfully applied to an engineered *S*. *cerevisiae* with periodic hydrogen peroxide shocking, resulting in a three-fold increasee of Ά-carotene production. Subsequent transcriptome analysis indicates that some genes related with lipid biosynthesis and MVA pathways are up-regulated in the adopted strains [107]. It also suggests that carotenoid production can be improved by modifications (knock-out or overexpression) of distant genes, which are responsible for the overall regulation of the metabolic network or the physiological fitness of the host (Figure 5C). In a genome-wide screening of yeast deletion collection, 24 deletions exhibit significant higher carotenoid levels than the wild type. The triple deletion of *ROX1*, *YJL064W*, and *YJL062W* shows an almost four-fold increase in total carotenoid production [108]. Gene deletions of *hnr*, *yjfP*, and *yjiD* related to the improvement of lycopene production have been identified from a global transposon *E*. *coli* mutant library [109]. Other gene deletions such as *gdhA*, *cyoA*, *ppc*, *gpmA*, *gpmB*, *eno*, *glyA*, aceE, *talB*, and *fdhF* have been *in silico* identified using a stoichiometric model [110]. The triple mutation of *gdhA*, *aceE* and *fdhF* was validated to increase lycopene production by nearly 40% in *E*. *coli* over the engineered parental strain. A similar set of gene deletions *dhA*, *cyoA*, *gpmA*, *gpmB*, *icdA*, and *eno* have been also *in silico* identified using different metabolic network models [111]. Overexpression of some genes encoding global regulatory proteins AppY, Crl, RpoS, and ElbAB, oxidoreductases TorC, YdgK, and YeiA, and hypothetical proteins YedR and YhbL, result in a significant increase in lycopene production in *E*. *coli* [112]. With a profound understanding of the landscape of genome manipulation, all these knocked-out and overexpressed alleles have been combined and optimized to generate high-fitness host strains for lycopene production [113,114]. ATP and NADPH are also important cofactors for the production of carotenoids. Using engineering ATP synthesis, pentose phosphate and TCA modules, recent work has shown the highest Ά-carotene production of 2.1 g/L by a fed-batch fermentation process in *E*. *coli* [115]. The advances in synthetic biology greatly boost genome manipulation on a large scale. Multiplex automated genome engineering (MAGE) simultaneously targets many locations on the chromosome for modification in a single cell or across a population of cells by directing ssDNA to the lagging strand of the replication fork during DNA replication [116]. The modifications can cover gene inactivation, expression regulations, and so on. Aforementioned twenty genes related to lycopene production have been targeted to tune their expression using a complex pool of synthetic DNAs, and lycopene production is increased more than five-fold. A complementary method called trackable multiplex recombineering (TRMR) has been developed to simultaneously map genome modifications that affect a trait of interest, which combines parallel DNA synthesis, recombineering and molecular barcode technology to enable rapid modification of all *E*. *coli* genes in an *a priori* knowledge-independent way [117].
Metabolic engineering for the production of valuable compounds often heavily relies on plasmid-based expression of the synthesis pathway in a heterologous host. Although plasmids are easily manipulated and allow strong expression of targeted enzymes, the plasmid-based systems suffer from genetic instability such as plasmid loss, an additional antibiotic cost, and a potential risk of antibiotic marker spreading to other organisms [118]. Accordingly, chromosomal integration of the production pathway promises the host to achieve stable overproduction of the desirable chemicals including carotenoids. By Ώ-Red homologous recombination, plasmidfree engineered *E*. *coli* strain has been developed to produce lycopene and astaxanthin [119]. The expression cassettes can be integrated into different loci to increase the number of gene copies. P1 transduction usually plays a role in transfering the different alleles between host strains. Recently, an intelligent strategy called chemically inducible chromosomal evolution (ClChE) has been developed to reduce the daunting repeated one-at-a-time tasks in the chromosomal integration of target genes [88]. ClChE allows the host to acquire a high gene copy (up to 40 copies) expression of integrated pathways with increasing concentration of selective chemicals, and the increased copy number is stabilized by the removal of the *recA* gene. With this approach, lycopene production has been increased by 60% from single copy integrated strain. The ClChE strategy has been further modified to eliminate antibiotic marker for environmental safety and health issue after the evolution of the recombinant host strain [120].
## *3.4. Protein Engineering for Improvement of Carotenoid Production Enzymes*
Pathway engineering for efficient production of desired chemicals is often challenged by limitations associated with the pathway enzymes themselves, such as low turnover numbers and promiscuities generating unwanted by-products [121]. Protein engineering provides a powerful solution to improve specific activity and substrate specificity of enzymes, and even to create new activity. Methods of protein engineering include directed evolution and computer-assisted rational design (Figure 5D) [122,123]. Directed evolution is an iterative process that imitates Darwinian evolution in the laboratory to select or screen a desired phenotype from mutagenesis. Typically, error-prone polymerase chain reaction (PCR) is used to generate mutant libraries, and DNA shuffling is carried out to recombine existing mutations. It can be performed in a blind manner with limited information on target enzymes, such as structures and reaction mechanisms, but it relies on an effective screening strategy. It is practical for the evolution of carotenogenic enzymes due to the innate traits of carotenoid pigments. Six mutants ((H96L, R203W, A205V, A208V, F213L and A215T) have been isolated to improve the catalytic activity of Ά-carotene ketolase from *Sphingomonas sp*. [124]. Three mutations (L175M, M99V, and M99I) of ketolase from *Paracoccus sp*. result in the improvement if its specificity of to synthesize astaxanthin [125]. *Staphylococcus aureus* dehydrosqualene (C30) synthase has evolved to synthesize lycopene by mutation F26L or F26S [126]. DNA shuffling of phytoene desaturases from *<sup>P</sup>*. *agglomerans* and *P*. *ananatis* results in the isolation of a variant favoring the production of fully conjugated tetradehydrolycopene [127]. Rational design of proteins is based on the *in silico* simulation and the prediction using *a priori* enzyme information, which greatly liberates biologists from onerous screening task. This strategy requires adequate information to predict specific targeted amino acid mutations, which can confer desired enzyme traits [128]. Unfortunately, the limited information on carotenogenic enzymes leads to few achievements using such a method.
As aforementioned, carotenoids are derived from the central isoprenoid pathway, which is also employed to synthesize several essential and secondary metabolites in nature. The carotenoid-based colorimetric screening has been developed for evolution of other isoprenoid pathway enzymes. Mutations of GGPP synthase are hypothesized to affect the binding efficiency of the magnesium ions needed for substrate anchoring and improve its catalysis. An error-prone PCR library of *Tsuga canadensis* GPPS has been screened using the lycopene synthesis pathway as a colorimetric reporter. The GPPS variant (S239C and G295D) is created to increase levopimaradiene production with a 1.7-fold increase over the wild type in *E*. *coli* [129]. Augmentation of one pathway can tamper with other pathways, which utilize the same substrate in one organism. Based on this fact, mutagenesis libraries of terpene synthases have been screened by depigmentation of colonies due to the competition between terpene synthases and carotenoid synthases for isoprenyl diphosphates, since the weakened carotenoid color intensity indicates an improvement of terpene synthase activity [130].
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*3.5. Development of Microalgae for Carotenoid Production*
Algae are a diverse group of aquatic, photosynthetic organisms, generally categorized as macroalgae (*i*.*e*., seaweed) and unicellular microalgae. Microalgae have recently garnered interest for production of valuable chemicals including carotenoids [41,131], because they are generally regarded as safe (GRAS) for human consumption and possess the renewable-energy capturing ability of photosynthesis. Moreover, these organisms can be used for genetic manipulation and highthroughput analysis [132]. Some microalgae are also native carotenoid producers (*i*.*e*., *D. salina* for Ά-carotene and *<sup>H</sup>*. *pluvialis* for astaxanthin). The carotenoid production from microalgae is closely related to culture conditions such as illumination, pH, temperature, nitrogen availability and source, salinity, the oxidant substances, and growth rate [12,133,134]. *<sup>D</sup>*. *salina* is a model species of green microalgae which is widely cultivated outdoors for Ά-carotene production [131]. A systematic evaluation has been done to decipher the relationship between abiotic stresses (Nitrate concentration, salinity and light quality) and lutein synthesis in *D. salina* [135]. The abiotic stresses can also be applied to adaptive evolution of microalgae [136], in a similar manner to strain evolution in yeast for Ά-carotene production [107]. The freshwater microalga *Chlamydomonas reinhardtii* is the first and the best studied transformed Chlorophyte, and the nuclear genetic manipulation is easy and well established. It has been engineered with Ά-carotene ketolase from *H*. *pluvialis* to synthesize ketolutein (Hb1H'a1, Figure 3) and adonixanthin (Ha1H'a2, Figure 3) [137]. It is possible to produce diverse valuable carotenoids from marine microalgae with the development of more available genetic tools and technologies.
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**4. Opportunities and Challenges**
The vast and mysterious ocean breeds diverse marine lives and provides unexhausted foodstuffs, nutriment, and drugs for humans. Diverse carotenoids are found from marine species and show broad utilities as colorant fragrance cosmetics and pharmaceuticals. The synthetic pathway of several carotenoids has been illuminated from marine species, which could benefit engineering processes in several host organisms for the production of carotenoids such as Ά-carotene, astaxanthin, and lutein. On the other hand, carotenoids such as Ά-carotene often undergo a series of modifications in the miraculous marine ecosphere. And indeed, several novel carotenoids have been isolated during the exploration of the marine ecosphere, while their pharmaceutical potentials remain to be examined due to the limited amount of extracts. Metabolic engineering and synthetic biology allow the assembly of such a chimeric pathway in a tractable organism for the mass production of rare carotenoids and also exhibit the potential to extend the catalogs of carotenoids to non-natural carotenoids, which could accelerate the exploration of novel carotenoids. It is noted that decoded carotenoid pathways and enzymes are still limited to a few marine organisms, although the J. Craig Venter Institute with worldwide collaboration had sequenced and annotated the genomes of 177 marine microbes up until 2010. However, we believe that the developed and developing technologies will allow us to search for novel marine carotenoid pathways in the future.
## **Acknowledgments**
This work was supported by a grant (NRF-2013R1A1A2008289) from the National Research Foundation, the Intelligent Synthetic Biology Center of Global Frontier Project funded by the MSIP (2011-0031964), and a grant from the Next-Generation BioGreen 21 Program (SSAC, grant#: PJ00952003), Rural Development Administration (RDA), Korea. J.K. is supported by scholarships from the BK21 Plus Program, Ministry of Education, Science & Technology (MEST), Korea.
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**Author Contributions**
S.K. conceived the idea and held and corrected the manuscript. C.W. and J.K collected the literature, analyzed the data, and wrote the manuscript. C.W. and J.K contributed to this manuscript equally.
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{
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**Conflicts of Interest**
The authors declare no conflict of interest.
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"section_idx": 152
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**Enhanced Productivity of a Lutein-Enriched Novel Acidophile Microalga Grown on Urea**
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**Carlos Casal 1, Maria Cuaresma 2, Jose Maria Vega 3 and Carlos Vilchez 2,\***
1 CIDERTA, University of Huelva, Park Huelva Empresarial, 21007, Huelva, Spain;
E-Mail: [email protected] (C.C.)
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{
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*Received: 2 November 2010; in revised form: 18 December 2010 / Accepted: 23 December 2010 / Published: 24 December 2010*
**Abstract:** *Coccomyxa acidophila* is an extremophile eukaryotic microalga isolated from the Tinto River mining area in Huelva, Spain. *Coccomyxa acidophila* accumulates relevant amounts of β-carotene and lutein, wellknown carotenoids with many biotechnological applications, especially in food and health-related industries. The acidic culture medium (pH < 2.5) that prevents outdoor cultivation from non-desired microorganism growth is one of the main advantages of acidophile microalgae production. Conversely, acidophile microalgae growth rates are usually very low compared to common microalgae growth rates. In this work, we show that mixotrophic cultivation on urea efficiently enhances growth and productivity of an acidophile microalga up to typical values for common microalgae, therefore approaching acidophile algal production towards suitable conditions for feasible outdoor production. Algal productivity and potential for carotenoid accumulation were analyzed as a function of the nitrogen source supplied. Several nitrogen conditions were assayed: nitrogen starvation, nitrate and/or nitrite, ammonia and urea. Among them, urea clearly led to the best cell growth (∼4 × 108
cells/mL at the end of log phase). Ammonium led to the maximum chlorophyll and carotenoid content per volume unit (220 μg·m<sup>L</sup> 1 and 35 μg·mL 1, respectively). Interestingly, no significant differences in growth rates were found in cultures grown on urea as C and N source, with respect to those cultures grown on nitrate and CO2 as nitrogen and carbon sources (control cultures). Lutein accumulated up to 3.55 mg·g in the mixotrophic cultures grown on urea. In addition, algal growth in a shaded culture revealed the first evidence for an active xanthophylls cycle operative in acidophile microalgae.
**Keywords:** urea; *Coccomyxa*; extremophile microorganisms; lutein; microalgae
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**1. Introduction**
*Coccomyxa acidophila* is a novel microalgal specie isolated from Tinto River (Huelva, Spain), which is so-called the 'Red river' due to its high iron water concentration. This special feature causes the river bed to constitute an acidic environment where the pH value remains constantly between 2 and 3 along a stretch of 80 km [1]. Besides, this microalga is characterized by having important potential to accumulate high lutein concentrations, a carotenoid with powerful well-known antioxidant
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{
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properties [2,3].
Nowadays, extremophile organisms are gaining increasing interest due to their faculty to be used as renewable source of different high value compounds including carotenoids, fatty acids (PUFAs), lipids, vitamins, toxins, enzymes, *etc.* [4–6]. Furthermore, the extremophile character of these microorganisms can be a benefit for getting axenic cultures with no interference from others microalgae. In general, apart from contamination risks, one of the main problems for microalgae cultivation is the relatively high costs, which is expected to be overcome by technological advances [7]. For that reason, since some time ago, efforts are being focused on reducing the cost of elements related to microalgae cultivation. One aspect that puts up the total price of the operation of production systems is the high CO2 demand that photosynthetic microorganisms usually have. In any case, although there are currently various attempts for capturing carbon dioxide by means of algae cultures from industrial flue gases [8], one strategy aimed to reduce costs could be the replacement of the carbon source by another cheaper option.
A wide variety of nitrogen sources, such as ammonia, nitrate, nitrite and urea, can be used as nitrogen source for growing microalgae [9]. Urea (CO(NH2)2) is a smallmolecular weight polar and relatively lipid-insoluble compound which is ubiquitous in nature. This organic compound can be considered as a combined source of nitrogen and carbon and it has diverse functions. In organisms containing the enzyme urease, a nickel-dependent metalloenzyme [10] present in bacteria, fungi and plants, urea is primarily used as a source of nitrogen necessary for growth. However, since urease metabolizes urea to CO2 and ammonia, thus providing a ready source of base, metabolism of urea by urease can also enable microorganisms to respond to acid challenges [11]. On the other hand, in mammals, urea is the primary waste product of amino acid catabolism [12]. Urea is a versatile substance and its role largely depends on whether it is an end-product or can be further broken-down, and if so, the utilization of the break-down products also varies considerably, either for anabolic processes or for buffering under acidic conditions.
Previous works performed in our group with acidophile microalgae growing under mixotrophic conditions showed that urea can be a more than suitable alternative for cultivation of this microalga, showing good productivity and lutein accumulation results. Moreover, in the literature, several examples can be found where urea is shown to be an effective combined source of N and C for the production of *S. Platensis*, *Neochloris oleoabundans* and *Chlorella sp*. under different cultivation modes [7,9,13–16].
This work aimed at assessing the effect of different nitrogen sources on biomass productivity and carotenoid accumulation of *Coccomyxa acidophila*, paying special attention to the amount of accumulated lutein. In addition, the results will allow for assessing the use of nitrogen sources other than the conventional ones in growing acidophile microalgae.
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*2.1. Coccomyxa acidophila enhanced growth on urea*
It was mentioned above that acidophile microalgae have so far never been used for massive production. Massive production requires fast growing microalgal strains. Most acidophile microalgae are slow growth strains, as reported in the literature [5,17]. However, the growth of acidophile microorganisms in acidic culture media becomes advantageous for biomass production, as under such conditions the growth of other microorganisms becomes difficult. Therefore, we attempted to find culture conditions under which *Coccomyxa acidophila* cultivation is enhanced, such that growth rates and productivity values approached those of common microalgae. In such a situation, the acidophile microalga should show fastgrowth and could hopefully be grown in outdoor systems with limited risks for microbial contamination, in comparison to common microalgae.
One of the main growth conditions assayed was nitrogen source. In previous experiments, we first tested the effect of adding ammonium, nitrite or nitrate to photoautotrophically growing *Coccomyxa acidophila* cells. Growth on ammonium and nitrate resulted in the highest productivities. Unlike common microalgae, nitrite became toxic for *Coccomyxa acidophila*. In the experiments here, we also used urea as a combined source for C and N, with high CO2 concentration (5% v.vƺ1) as the main carbon source where indicated. Urea has been widely used instead of high CO2ƺ for sustaining microalgae growth and is also a cheap N source. As shown in Figure 1, urea promoted enhanced growth of *Coccomyxa acidophila*, both in terms of chlorophyll content (Figure 1A) and cell density (Figure 1B). This resulted in an increased growth rate with respect to control cultures (photoautotrophically grown on nitrate), as shown in Table 1. In addition, culture productivity was higher when the microalga was grown on urea (the highest productivity) and ammonium. Specifically, the highest productivity was reached in cultures grown on 0.67 g·L 1 urea ("control; air" in Figure 1). This urea concentration provided cultures with the same molar concentration of nitrogen than the nitrate added to control cultures. More interestingly, the best productivity values obtained from the microalgal growth on urea did not differ from those usually obtained for most of common microalgae (0.2–0.4 g·L 1·d 1). Unexpectedly, the simultaneous presence of urea and nitrate limited *Coccomyxa acidophila* growth. This will be discussed further.
**Figure 1.** Time-course of chlorophyll ( **A**) and cell density ( **B**) in *Coccomyxa acidophila* cultures grown on nitrate, urea or nitrate plus urea. Air alone or CO2 in air (5% v/v) were used as carbon source, as indicated for each culture within the Figure legend.
**Table 1.** Growth rates and productivity of *Coccomyxa acidophila* grown on different N-sources.
The previous results were obtained by means of using mixotrophic or photoautotrophic cultures, *i.e.*, either urea or nitrate were added to culture media as N sources while high CO2 concentration (5% v/v in air) was supplied as a carbon source (first set of experiments). It has also been discussed that the microalga cells could make use of urea as an additional carbon source [18,19], perhaps being one of the reasons behind the improved microalgal productivity of urea grown cultures. Low CO2 solubility at acidic pHs makes carbon uptake more difficult than at pH 7 (standard pH for most common microalgal cultures). Therefore, the supply of additional carbon in a soluble form at low pH (e.g., urea, glucose) could help to increase microalgal productivity. This raises the question of whether addition of glucose as a carbon source to cultures of an acidophila microalga should also increased microalgal productivity. Such a question was investigated by our group in previous research [1] and the answer was "no". Urea should by far allow maximum productivity in *Coccomyxa acidophila* cultures when used as a carbon source.
From the results above, *Coccomyxa acidophila* apparently prefers urea to nitrate as nitrogen source. Therefore, another question we addressed was whether such consumption preference indeed occurred. For this purpose, nitrate and urea consumption were followed in time in photoautotrophic cultures to which nitrate (control culture) or urea and nitrate (with the same molar nitrogen concentration to that used in control cultures, 22.7 mM), were added. Results are shown in Figure 2. If urea and nitrate are added simultaneously, nitrate only started to be consumed at late exponential growth phase while urea was first consumed as the only nitrogen source. A decreasing time-course trend in urea concentration is observed from the beginning of the experiment, whereas the nitrate concentration time-course trend remains stable. Inhibition of nitrate consumption by the presence of urea has been reported to occur in microalgae, though not many references dealing with the subject have been published. Cochland and Harrison [20] reported about 30% inhibition of nitrate consumption by urea in the eukaryotic picoflagellate *Microsomas pusilla*. Following consumption, assimilatory reduction of nitrate also could be inhibited. One of the first classic references was published by Smith and Thompson [21] who observed 70% nitrate reductase inhibition by urea in *Chlorella*, evidencing nitrate assimilatory reduction down regulation to be behind nitrate consumption inhibition by urea.
As already mentioned, simultaneous addition of urea and nitrate as nitrogen sources slightly limits cell growth. Merigout *et al.* [22] evidenced in *Arabidopsis* plants that urea uptake was stimulated by urea but was reduced by the presence of nitrate in the growth medium. Such conclusions from their recent study on physiological and transcriptomic aspects of urea uptake and assimilation are in good concordance with the following observations from our results: (a) urea increased *Coccomyxa acidophila* growth and (b) simultaneous presence of urea and nitrate resulted in a decreased uptake of urea and culture productivity. These observations related to nitrogen uptake regulation in *Coccomyxa acidophila* are for the first time reported in acidophile microalgae and suggest that urea uptake and assimilation patterns in extreme acidophile microalgae (living in fully urea-free environments) and plants are similar. Further experiments in nitrogen assimilatory enzymes and gene expression are currently being developed in our group.
**Figure 2.** Time course of nitrogen consumption in *Coccomyxa acidophila* cultures grown on nitrate, urea or nitrate plus urea. Air alone or CO2 in air (5% v/v) were used as carbon source, as indicated for each culture within the Figure legend. Dotted line with triangles corresponds to timecourse of nitrate consumption of cultures incubated with nitrate plus urea.
To determine whether simultaneous addition of urea and nitrate to the algal cultures has any impact on photosynthesis, relative electron transport rates were determined in each of the cultures (namely, control –nitrate ; urea; urea and nitrate, as nitrogen sources, respectively). Results are shown in Figure 3. Surprisingly, there was no nitrogen source-dependent impact on PS2 and on photosynthetic energy production chain, if the light intensity remained approximately below 150 μE·mƺ2·sƺ1. However, in urea grown cultures incubated under higher light intensities, photosynthetic energy production is shown to be clearly limited, up to the point that the electron transport chain becomes inhibited. This dramatically influences carbon assimilation and culture productivity.
According to our results, urea appears to be a suitable nitrogen source for *Coccomyxa acidophila* growth at relatively low light intensity; however, it has a dramatic impact on the photosynthetic energy production chain when exposed to high light intensity, which has never been reported for any other microalga. This is currently under study in our laboratories.
**Figure 3.** Light-dependent electron transport rates in *Coccomyxa acidophila* cultures grown on nitrate, urea or nitrate plus urea. Air alone or CO2 in air (5% v/v) were used as carbon source, as indicated for each culture within the Figure legend.
In addition, physiological responses of acidophile microalgae to urea and nitrate uptake processes anyhow differ, according to the observed pH changes in the culture media which tend to increase if nitrate (2.3 g·L 1) is the only added nitrogen source and to decrease if urea (0.67 g·L 1) is used (data not shown). So far, we have no evidence for antyport/symport mechanism details that help to elucidate the different physiological behavior.
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*2.2. Carotenoid accumulation and xanthophylls cycle activity in urea grown* Coccomyxa acidophila *cells*
*Coccomyxa acidophila* accumulates commercial value carotenoids including lutein, β-carotene and zeaxanthin (Figure 4). Besides assessment of the best nitrogen sources for biomass production, carotenoid accumulation in urea and nitrate grown cultures was also studied (Figure 5). According to the best growth conditions inferred from Figure 1, for this experiment, the carotenoid content was followed in urea grown cultures (fluidized with air) and in nitrate grown cultures (fluidized with air supplemented with 5% v/v CO2). In addition, carotenoid content was also followed in nitrogen-deprived cultures, as nitrogen depletion is a very well known carotenogenic condition for many microalgae species. In good agreement with the enhanced cell growth in urea grown cultures, total carotenoid content in the reactor also increased much more rapidly in urea grown cultures than in control cultures. Consequently, the carotenoid content of urea grown cultures ( g·mL 1) became about two-fold that of the nitrate grown cultures (control cultures), until late exponential growth phase (Figure 5). This could be due to the increased biomass production in urea grown batch cultures, therefore higher carotenoid content in the reactor is not necessarily a consequence of faster carotenoid biosynthesis. However, simple calculations of the content of specific carotenoids per cell revealed a prompt carotenoid biosynthesis enhancement (namely -carotene, lutein, zeaxanthin) in urea grown *Coccomyxa acidophila* cells, as can be inferred from the carotenoid cell content data in Figure 6; lutein by far being the most abundant carotenoid. This means that urea clearly promotes increases in lutein and β-carotene cell content, at least up to late exponential growth phase, where lack of nutrients, shading effect and stress factors change the observed trend. Besides, it can be observed that cell content of violaxanthin inversely correlates with cell content of zeaxanthin over the time course. This is the first evidence of an active xanthophylls cycle in *Coccomyxa acidophila* that converts violaxanthin into zeaxanthin by means of violaxanthin deepoxidase activity. Interestingly, nitrogen starvation did not promote carotenoid accumulation in *Coccomyxa acidophila* cultures, unlike other common microalgae including *Dunaliella*, *Haematococcus* and many others [23,24].
**Figure 4.** HPLC chromatogram showing the main carotenoids of *Coccomyxa acidophila*. AU: arbitrary units.
**Figure 5.** Total carotenoid content in *Coccomyxa acidophila* cultures grown on different nitrogen sources or under nitrogen starvation. Air alone or CO2 in air (5% v/v) were used as carbon source, as indicated for each culture within the Figure legend.
**Figure 6.** Time-course of the cell content of the indicated specific carotenoids in *Coccomyxa acidophila* cultures incubated in either nitrate or urea.
Fernández-Sevilla *et al.* [25] recently reviewed the biotechnological production of lutein. The paper includes an updated list of lutein production experiments performed on different scales using microalgae species. Table 2 shows the most relevant data of lutein productivity by microalgae and reactor type used for the production processes. Considering intracellular lutein cell content of each one of the promising species and lutein productivity in photobioreactors, *Scenedesmus almeriensis* [26], *Muriellopsis sp.* [27] and *Chlorella protothecoides* [28] emerge so far as the most efficient strains for the biotechnological production of lutein from microalgae. When incubated under standard culture conditions, *Coccomyxa acidophila* onubensis accumulates up to 6.1 mg·g 1 dry weight, which is within the upper range of lutein concentrations accumulated by the above mentioned microalgae. We are now running continuous cultures of *C. acidophila* in tubular laboratory photobioreactors in order to obtain lutein productivity data in long-term (weeks) production processes. Compared to continuous cultivation of other lutein producing species, *C. acidophila* has the practical advantage of growing well in an extremely selective culture medium at very low pH which preserves cultures from microbial contamination.
**Table 2.** Lutein productivity of lutein-enriched microalgae.
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**3. Conclusions**
The main conclusions of this manuscript are: (1) Mixotrophic cultivation on urea efficiently enhances growth and productivity of *C. acidophile*; signaling strategies towards suitable conditions definition for feasible outdoor production; (2) Urea clearly led to the fastest cell growth;
(3) Maximal lutein accumulation was found to occur in urea supplemented culture medium;
(4) In addition, algal growth in a shaded culture revealed the first evidence for an active xanthophylls cycle operative in acidophile microalgae.
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*4.1. Microorganism and cultivation conditions*
*Coccomyxa acidophila,* the algal material used in this work, was isolated from the acidic water of the Tinto River's mining area, in Huelva (Spain).
Initially, an axenic culture of the microalga was obtained by streaking it on basal agar medium at
pH 2.5. After that, isolated colonies were transferred from the solid medium to a liquid medium modified by Silverman and Lundgren [29]. *Coccomyxa acidophila* mother cultures were maintained by periodic transfers in sterile medium adjusted to pH 2.5 with concentrated H2SO4. Unless otherwise indicated, standard cultivation conditions were batch cultures grown at 25 ºC into 1 L-Roux flasks, bubbled with air containing 5% (v/v) CO2 and continuously illuminated with fluorescent lamps (Philips TLD, 30 W, 150 μE·mƺ2·sƺ1 at the surface of the flasks). In those cases where CO2 was not supplied to the cultures, it was necessary to put a carbon dioxide trap with KOH 5 M buffer for removing it from the air mix. Every day, pH was controlled and adjusted at 2.5 ± 0.1 by adding diluted HCl or NaOH.
The irradiance was measured with a quantum/photometer Licor (mod. LI 250A).
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*4.2. Dry weight measurements*
Before filtering culture samples, filters of cellulose acetate with a 0.45 μm pore size, from Sartorius (Goettingen, Germany), were washed with distilled water and dried at 80 ºC in an oven for 24 h. After that, these were weighted and used to separate cells from the medium. Five milliliter culture samples were taken, vigorously homogenated, and filtered by means of a vacuum pump. Filters containing cells were dried and kept in an oven for 24 h, after which they were weighed.
## *4.3. Measurements of fluorescence*
Optimal chlorophyll fluorescence yield measurements (Fm/Fv) were performed with a pulse amplitude modulated fluorometer (Teaching-PAM from WALZ, Effeltrich, Germany). In order to make sure that there is no reduction of the PSII primary electron acceptor QA and, therefore (consequently), all PSII reaction centers are open, cultures samples of 1 mL were previously adapted to dark conditions for 15 min [30]. After that period, a short saturating pulse of light (SP) was triggered. When necessary (e.g., low chlorophyll concentrations), the PAM modulated light (ML) had to be adjusted to higher values to obtain readings in the proper range.
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*4.4. Oxygen evolution*
In addition of fluorescence measurements, the biological activity used to check cell viability was photosynthetic activity. For these determinations, 1 mL samples of *Coccomyxa acidophila* cultures were placed into a Clark-type electrode (Hansatech, U.K.) to measure O2-evolution. Measurements were made at 25 ºC in the dark (endogenous respiration) or under saturating white light (1500 μE·mƺ2·sƺ1).
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*4.5. Analytical determinations*
Total chlorophyll and carotenoid pigments were determined spectrophotometrically after centrifuging tubes containing samples for 6 min at 13000 rpm, heating them for 1 min, and extracting cell pellets with pure methanol. Sonication by ultrasound was also applied when necessary. After that, samples were spun down again for 5 min at 5000 rpm to eliminate cellular wastes. Calculations were done using equations according to [31].
For specific carotenoid analysis and quantification, separation was performed by liquid chromatography (HPLC; Merck Hitachi) using a RP-18 column with a flow rate of 1 mL/min. The applied gradient was the following (solvent A; ethyl acetate and solvent B; acetonitrile/agua, 9:1 v/v): 0–16 min, 0–60% solvent A; 16–30 min, 60% A; 30 – 35 min, 100% A. In order to quantify, pigment standards supplied by DHI-Water and Environment (Denmark) were injected.
Nitrate was determined following the method described by Cawse *et al.* [32]. Urea was determined according to the method from Wilcox [33].
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*4.6. Statistics*
Unless otherwise indicated, all data included in figures and tables represent the average of triplicates.
## *4.7. Cell counting*
Cellular density was determined by microscopy using an Olympus CX41 in a Neubauer chamber.
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**Acknowledgements**
This work has been supported by grant AGR-4337 (Proyecto de Excelencia, Junta de Andalucía) and grant Bioándalus (Junta de Andalucía, Estrategia de Impulso a la Biotecnología de Andalucía).
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**The New Carotenoid Pigment Moraxanthin Is Associated with Toxic Microalgae**
## **Olga Mangoni 1, Concetta Imperatore 2, Carmelo R. Tomas 3, Valeria Costantino 2 ,**
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 171
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**Vincenzo Saggiomo 4 and Alfonso Mangoni 2,\***
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*Received: 22 December 2010; in revised form: 25 January 2011 / Accepted: 4 February 2011 / Published: 10 February 2011*
**Abstract:** The new pigment "moraxanthin" was found in natural samples from a fish mortality site in the Inland Bays of Delaware, USA. Pure cultures of the species, tentatively named *Chattonella cf. verruculosa*, and natural samples contained this pigment as a dominant carotenoid. The pigment, obtained from a 10 L culture of *C. cf. verruculosa*, was isolated and harvested by HPLC and its structure determined from MS and 1Dand 2D-NMR. The data identified this pigment as a new acylated form of vaucheriaxanthin called moraxanthin after the berry like algal cell. Its presence in pure cultures and in natural bloom samples indicates that moraxanthin is specific to *C. cf. verruculosa* and can be used as a marker of its presence when HPLC is used to analyze natural blooms samples.
**Keywords:** *Chattonella cf. verruculosa*; Raphidophyceae; toxic algae; carotenoids; moraxanthin
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 173
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**1. Introduction**
Phytoplankton, unicellular photosynthetic microorganisms, are ubiquitous in all aquatic environments. As primary producers, they are responsible for nearly half of the global primary production of organic carbon [1]. Photosynthesis, the process whereby energy is absorbed by pigments in algae and transformed into chemical energy, relies on the presence of energy trapping pigments. The main pigments, chlorophylls, carotenoids and phycobilins, absorb Photosynthetically Available Radiation (PAR) from 400–700 nm wavelengths [2]. However, pigments may also serve several functions including metabolic regulation, light harvesting (antenna pigments), electron donation or acceptance (in reaction centers), and photoprotection. The combination of different pigments and functions result in maximum efficiency and economy [3–6]. The kind of pigments produced and their relative proportions characterize the different phytoplankton groups.
In recent years, high performance liquid chromatography (HPLC) has been used to estimate phytoplankton composition by identifying photosynthetic pigments. Some pigments found exclusively in particular algal classes or genera may serve as useful taxonomic markers [7–13]. Such indicator pigments are termed 'finger prints'. Pigment analyses offer a valuable technique in oceanography for mapping phytoplankton populations and monitoring their abundance and composition [14– 17].
Phytoplankton blooms occur naturally in coastal waters particularly during spring and summer seasons. However, a small number of microalgae are harmful, and although each individual is small, they may occur in huge numbers known as blooms [18–21]. Among the estimated phytoplankton species, about 7% (300 species) are known to produce red tides and of those, only 2% are actually harmful or toxic [22]. In marine and brackish water environments, most toxic species belong in the Dinophyceae, but also the Diatomophyceae, Haptophyceae, Raphidophyceae, and Cyanophyceae comprise toxic species [23–28]. The algal toxins may cause damage to other flora and fauna directly or they may accumulate through the food web in e.g., shellfish or finfish, thereby causing harm to their predators including humans [29–34]. Harmful algal blooms (HABs) are an ever more frequent phenomenon expanding in coastal regions on a world scale [35–38]. These have received much attention from researchers and local regulatory authorities due to their impact on the ecosystem and human health, influencing local economic issues [39].
Monitoring of coastal waters for harmful species is costly and labor-intensive and the possibility to recognize a potentially harmful algal species by means of chemical or biochemical analyses significantly reduces the time and costs of such monitoring. The one caveat is that the analysis, pigment or biochemical, involves a species specific marker for the HAB species in question [40]. Pigment signatures in the study of HABs have been very limited, particularly in monitoring programs [33,41,42].
During the summer period of 2000, ten fish mortality events occurred from unidentified causes in the Inland Bays of Delaware, USA. During the final fishkilling event of 28 August, 2000, over two million menhaden (*Brevoortia tyrranus*) perished when a bloom of an unidentified microalgal flagellate was observed [36]. This flagellate, accompanied by the presence of a potent neurotoxin, was tentatively called *Chattonella* cf. *verruculosa* since it resembled a fish killing species found in Japan thought to be of the class Raphidophyceae. Since none of the previously described Raphidophyceae completely agreed with the molecular features (18S rDNA; 16S rDNA) [43,44], further studies are underway to define its taxonomic position.
This work describes the isolation and structural elucidation of a new pigment (**1**) found in *C. cf. verruculosa* cultures and in natural samples where this species was dominant, which has been called moraxanthin after the berry like algal cell (Figure 1). Moraxanthin, which is a new acylated form of vaucheriaxanthin (**2**), is unique to *C. cf. verruculosa*, indicating that it can be used as a marker of its presence when HPLC analyses of natural blooms are performed.
**Figure 1.** Structures of moraxanthin (**1**) and vaucheriaxanthin (**2**).
## **2. Results and Discussion**
The chromatogram of the pigments of the *C. cf. verruculosa* culture showed a major peak (Figure 2, peak 4), whose retention time (Table 1) and UV spectrum (Table 1 and Figure 3) did not fit those of any known pigments, although the UV spectrum clearly showed the pigment to be a carotenoid.
**Table 1.** Total pigments found in *C. cf. verruculosa* with relative retention times and specific absorption maxima.
*Mar. Drugs* **2011**, *9*, 242–255.
The new pigment was then isolated to determine its structure by spectroscopic (MS and NMR) means. A large-scale (10 L) culture of *C. cf. verruculosa* was grown, and harvested by continuous flow centrifugation into an algal pellet and supernatant. The algal pellet (4 g) was extracted exhaustively with MeOH, and the extract was subjected to repeated HPLC separation, yielding 1.1 mg of the pure carotenoid moraxanthin (**1**). When re-injected in the same HPLC conditions as for the chromatogram in Figure 2, the isolated compound **1** showed retention time and UV spectrum identical to that of peak 4. Compound **1** showed a [M + Na]+ pseudomolecolar ion peak in the ESI high-resolution mass spectrum at *m*/*z* 779.4879, in accordance with the formula C48 H68NaO7. Compared to the C40 carotenoid skeleton, this formula contains eight additional carbon atoms. In addition, the ESI mass spectrum also contained a peak at *m*/*z* 663 (C42 H56NaO5, [M + Na ƺ C6H12 O2]+), which can be accounted for by the in-source loss of hexanoic acid.
Most of the information used for structure elucidation came from one- and twodimensional NMR spectroscopy. The general features of the proton NMR spectrum (C6D6) resembled those of carotenoids, with several olefinic protons between Έ 6 and Έ 7, and 10 methyl singlets between Έ 1.87 and 1.08. However, one of the ten methyl signals (Έ 1.74) was part of an acetyl group, as shown by its correlation peak with the carbonyl carbon atom at Έ 169.2 in the HMBC spectrum. Other notable features of the proton NMR spectrum were (i) the AB system at Έ 5.13 and 5.07 (H2-19') of an oxymethylene group, (ii) two oxymethine protons at Έ 5.69 (H-3) and Έ 3.78 (H-3'), a methyl triplet at Έ 0.78 (H3-6'') and a methylene triplet at Έ 2.14 (H3-2''), indicative of an acyl chain and (iv) an olefinic proton singlet at Έ 6.03 (H-8), which showed correlation peaks in the HMBC spectrum with two non-protonated carbon atoms at Έ 202.1 (C-7) and 117.8 (C-6), and was therefore part of an allene system. The observed structural features were suggestive of a structure similar to vaucheriaxanthin (**2**), but a direct comparison of spectral data was hindered by the presence of the two additional acyl groups. A detailed analysis of the correlation peaks observed in the COSY, HSQC, and HMBC (Figure 4) spectra demonstrated that the planar structure of moraxanthin is indeed the same as that of vaucheriaxanthin, except for the presence of an acetyl group at position 3 and a hexanoyl group at position 19'.
In addition to all the expected geminal and vicinal couplings, the COSY spectrum revealed several proton-proton long-range couplings. Among them, the quite large W couplings of H-2 ΅ with H-4 ΅
(2.2 Hz) and of H-2' ΅ with H-4' ΅ (1.7 Hz) indicated the 1,3-diequatorial relationship of these two pairs of protons. Furthermore, the methyl protons on the polyene system showed weak correlation peaks with the olefinic protons, arising from the usual allylic 4*J*HH couplings, but also from 6*J*HH couplings (H3-19/H-12, H3-20/H-15', H3-20'/H-15) and even one remarkable 8*J*HH coupling (H3-19/H-14). To the best of our knowledge, this is the first report of a 8*J*HH coupling in a carotenoid.
The *E* configuration of double bonds at positions 11, 15, 7', and 11' was evident from the large *trans* coupling constant values of the relevant protons (see Table 2). The *E* configuration of double bonds at positions 9, 13, and 13' and the *Z* configuration of the double bond at position 9' were determined from the ROESY spectrum, displaying correlation peaks of H3-19 with H-11, H3-20 with H-15, H3-20' with H-15', and H2-9' with H-11'.
The ROESY spectrum also provided information on the relative configuration of the two terminal six-membered rings (Figure 5). The allene terminus is in the chair conformation, with the two W-coupled H-2 ΅ and H-4 ΅ protons in the equatorial orientation. The large coupling constants of H-3 with the axial H-2 Ά and H-4 Ά (Table 2) showed the former proton to be axial, and therefore on the ΅ face of the ring; as a consequence, the OAc group at C-3 must be Ά. The ROESY correlation of the methyl protons H3-19 with H-2 Ά and H-4 Ά determined the axial chirality of the allene functionality as *R*. Finally, the ROESY correlation of H3-19 with H3-18 located C-18 on the Ά face of the ring, and therefore the OH group at C-5 on the ΅ face.
As for the other terminal ring, the W coupling (1.7 Hz) of the pseudoequatorial H-2' ΅ and H-4' ΅ suggests a half-chair conformation of this ring. The *trans* relationship between the epoxide ring and the hydroxyl group was established from the ROESY correlation peaks of the two geminal methyl groups H3-16' and H3-17' with, respectively, H-3' and H-7' (Figure 5), showing that H-3' and H-7' are on opposite faces of the six-membered ring. This was confirmed by the prominent peak between the psudoaxial H-4 Ά and H3-18 in the same spectrum. The relative configuration determined for moraxanthin matches that of vaucheriaxanthin (**2**), and it may be assumed that also the absolute configuration of moraxanthin is the same as in vaucheriaxanthin.
**Figure 6.** HPLC absorbance chromatogram of natural water sample collected at the
fish-kill site of Torque Canal, Delaware on 28 August 2000 during a *C. cf. verruculosa* bloom. The arrow indicates the moraxanthin peak.
To investigate the utility of using moraxanthin as a marker for the toxic alga *C. cf. verruculosa*, natural bloom samples from the fish-kill site at Torque Canal, Delaware, collected on 28 August 2000 on Whatman GFF glass fiber filters and stored at ƺ80 °C, were extracted in methanol and subjected to HPLC analysis. The HPLC chromatogram (Figure 6) definitely showed a peak for moraxanthin in the natural sample. The moraxanthin peak clearly separated from all the other pigment peaks having no overlap with other pigments. In addition, a peak with the same retention time and absorbance characteristics was present in the HPLC chromatogram from water samples collected in 2003–2007 at various sites in Delaware's Inland Bays where *C. cf. verruculosa* blooms occurred (data not shown). This shows that the HPLC analysis may provide a simple and rapid tool for detecting harmful blooms of *C. cf. verruculosa*.
## **3. Experimental Section**
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*3.1. General experimental procedures*
ESI-MS experiments were performed on an Applied Biosystem API 2000 triplequadrupole mass spectrometer. High Resolution ESI-MS spectra were performed on a Thermo Orbitrap XL mass spectrometer. All the mass spectra were recorded by infusion into the ESI source using MeOH as the solvent. CD spectra were recorded in MeOH solution on a Jasco J-710 spectrophotometer using a 1 cm cell. 1H and 13C NMR spectra were determined in C6D6 solution on a Varian UnityInova spectrometer at 700 and 175 MHz, respectively; chemical shifts were referenced to the residual solvent signal (ΈH 7.15, ΈC = 128.0). For an accurate measurement of the coupling constants, the one-dimensional 1H NMR spectra were transformed at 64K points (digital resolution: 0.09 Hz). Homonuclear 1H connectivities were determined by COSY experiments. Through-space 1H connectivities were evidenced using a ROESY experiment with a mixing time of 500 ms. The reverse multiple-quantum heteronuclear correlation (HMQC) spectra was optimized for an average 1*J*CH of 142 Hz. The gradient-enhanced multiple-bond heteronuclear correlation (HMBC) experiment was optimized for a 3*J*CH of 8.3 Hz.
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*3.2. Plant material*
Clonal cultures of *C. cf. verruculosa* were established by single cell pipette isolation from a natural bloom sample taken at the time of a fish-kill in Torque Canal, Delaware. Individually isolated cells were grown in DYV medium [43] using sea water adjusted to a salinity of 20 to match that of the sample water. Successful isolates grown in 96 well microtiter plates were stepped up in volume eventually becoming stabilized cultures maintained in 150 mL volumes in erlenmeyer culture flasks. All cultures were maintained at 22 °C, with a fluence rate of 50 μ mol. quanta mƺ2 sƺ1 of cool white fluorescent light and a 12:12 h (LD) cycle. For this study, culture CMS TAC1050 was used and is presently deposited in the Center for Marine Science Toxic Algal Collection housed at UNCW's marine facility. This collection of harmful species is under the direction of Dr. Carmelo Tomas, Professor of Biology and Marine Biology at the CMS location who was also the isolator of the original culture. Large volume cultivation consisting of 10 L batches were grown in Bellco stirred cell system under conditions mentioned above. After a growth period of 1 month, the 10 L culture was harvested using a Sorvall RCB-2 refrigerated centrifuge equipped with a KSB (Kendro) continuous centrifuge head. A 4 g (wet weight) pellet was harvested, transferred to 15 mL cryovials and kept frozen at ƺ80 °C prior to analyses for pigments.
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*3.3. Pigment analysis*
The algal pellet (1 g) from cultures of *C. cf. verruculosa* was extracted with MeOH (3 mL), and the extract was filtered through Whatman GFF (0.45 μm). A portion of extract (500 μL) was added to 250 μL of ion-pairing solution (1M ammonium acetate), and after 5 minutes injected to the HPLC system. Assessment of the pigment composition was performed using a Hewlett-Packard HPLC 1100 Series system, equipped with a quaternary pump system and diode array detector. Pigments were separated on a temperature-controlled (20 °C) Hypersil MOS C8 reverse phase column (Sigma-Aldrich, 3 μm, 100 × 4.6 mm) according to the HPLC method of Vidussi *et al.* [45]. The mobile phases were MeOH (eluent A) and MeOH/0.5 N ammonium acetate (7:3) (eluent B). The elution gradient was kept constant at 1.0 mL/min for 20 min. The ratio of eluent B was gradually increased from 25 to 100%, and then returned to the initial proportion at the end of the elution. Chlorophylls and carotenoids were detected at 440 nm and identified by a diode array detector (Ώ = 350–750 nm, 1.2 nm spectral resolution). Standards of all the known pigments were provided by International Agency for 14C Determination (VKI Water Quality Institute) and calibration was performed according to Mantoura and Repeta [46].
## *3.4. Analysis of algal bloom*
Samples from blooms occurring in the Delaware Inland Bays were collected and returned to the laboratory or shipped by overnight courier to UNCW CMS Laboratory. Upon arrival, the sample was processed immediately. Pigment samples were taken as natural samples filtered on Whatman GFF filters and frozen immediately in liquid nitrogen and stored in a ƺ80° C freezer until extraction and pigment analyses could be performed as described above. Species contents of the sample water were determined by direct observations using a Nikon Diaphot inverted microscope. Observations of the phytoplankton included species identification, at least to genus level of live cells, cell density estimates using standard inverted microscope techniques and extraction for lipid soluble toxins in chloroform. Preserved samples (Lugol's solution) were carefully mixed, placed into a 10 or 50 mL settling chamber and allowed to settle for 24 hours prior to observation. The dominant species were identified, enumerated and used to define the phytoplankton composition.
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*3.5. Extraction and isolation of moraxanthin* (**1**)
The algal pellet (4 g) was extracted once with MeOH (40 mL). The extract was dried to give a dark green oil (104 mg) which was subjected to reversed-phase HPLC separation on a Varian Prostar 210 apparatus equipped with an Varian 325 UV detector [column: RP-18, 10 *ΐ*m, 250 × 10 mm; eluent: MeOH/H2O (9:1), flow 5 mL/min, UV detector set at 430 nm] to give partially purified moraxanthin (3.4 mg). Further reversed phase HPLC [column: RP-18, 3 *ΐ*m, 250 × 4.6 mm; eluent: MeOH/H2O (8:2), flow 1 mL/min, UV detector set at 430 nm] gave pure moraxanthin **1** (1.1 mg), whose identity was confirmed by HPLC analysis as described in Section 4.3.
## *3.6. Moraxanthin* (**1**)
Dark yellow oil; CD (MeOH; *c* 3.06ȉ10–6 M): ̇Ή471 +9.9, ̇Ή446 +11.9, ̇Ή422 +9.0; UV (EtOH): *Ώ*max nm (*Ή*): 421 (89000), 444 (129000), 472 (118000); ESI MS *m*/*z* 779 [M + Na]+; HRESIMS *m*/*z* 779.4879 [M + Na]+ (calcd. for C48 H68NaO7, 779.4857). For 1H and 13C NMR spectroscopic data, see Table 2.
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**4. Conclusions**
The newly proposed toxic species *C. cf. verruculosa* contains a new species-specific pigment, moraxanthin (**1**), whose structure was established as (3S,5 *R*,7 *R*,3'*S*,5' *R*,6'*S*)- 3-acetoxy-5',6'-epoxy-19'-(hexanoyloxy)-6,7-didehydro-5,6,5',6'-tetrahydro-Ά,Άcarotene-5,3'-diol, *i.e.*, 3- *O*-acetyl-19'- *O*-hexanoylvaucheriaxanthin. Two esterified forms of vaucheriaxanthin have been described, namely the 3- *O*-acetyl-19'- *O*octanoate and the 3- *O*-acetyl-19'- *O*-decanoate derivatives [47]. However, none of them contains the hexanoyl residue present in moraxanthin, which therefore can be easily distinguished from these known compounds on the basis of the HPLC retention time.
New harmful species have been identified and the taxonomy of other species has been revised [48]. It is usually accepted that the routine identification of phytoplankton for monitoring studies in estuaries and coastal waters requires additional methods other than traditional microscopy, which can underestimate some taxonomic groups containing fragile or poorly differentiated small cells. In conjunction with microscopy, pigment separation using HPLC has become a more widely applied method for estimating and characterizing phytoplankton biomass and community structure [6–8].
However, algal pigments usually show complex overlapping patterns with different taxa, offering only a few unambiguous markers. In our case, *C. cf. verruculosa* may be readily identified in natural samples by means of HPLC chromatograms due to the distinct peak corresponding to the species-specific pigment moraxanthin described here.
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**Acknowledgements**
The research leading to these results has received funding from the European Union's Seventh Framework Programme FP7/2007–2013 under grant agreement n° 229893 (NatPharma) and from grant No. 01504-07 awarded by U.S. Center for Disease Control and Prevention and North Carolina Department of Health and Human Services to Carmelo Tomas. Mass and NMR spectra were recorded at the "Centro Interdipartimentale di Analisi Strumentale", Università di Napoli "Federico II". The assistance of the staff is gratefully acknowledged.
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**Isolation and Analysis of the** *Cppsy* **Gene and Promoter from** *Chlorella protothecoides* **CS-41**
**Meiya Li 1,2,†, Yan Cui 1,†, Zhibing Gan 1, Chunlei Shi 1,\* and Xianming Shi 1**
Academic Editor: Graziano **Riccioni**
*Received: 25 June 2015 / Accepted: 9 September 2015 / Published:*
**Abstract:** Phytoene synthase (PSY) catalyzes the condensation of two molecules of geranylgeranyl pyrophosphate to form phytoene, the first colorless carotene in the carotenoid biosynthesis pathway. So it is regarded as the crucial enzyme for carotenoid production, and has unsurprisingly been involved in genetic engineering studies of carotenoid production. In this study, the *psy* gene from *Chlorella protothecoides* CS-41, designated *Cppsy*, was cloned using rapid amplification of cDNA ends. The full-length DNA was 2488 bp, and the corresponding cDNA was 1143 bp, which encoded 380 amino acids. Computational analysis suggested that this protein belongs to the Isoprenoid\_Biosyn\_C1 superfamily. It contained the consensus sequence, including three predicted substrate-Mg2+ binding sites. The *Cppsy* gene promoter was also cloned and characterized. Analysis revealed several candidate motifs for the promoter, which exhibited light- and methyl jasmonate (MeJA)-responsive characteristics, as well as some typical domains universally discovered in promoter sequences, such as the TATA-box and CAAT-box. Light- and MeJA treatment showed that the
*Cppsy* expression level was significantly enhanced by light and MeJA. These results provide a basis for genetically modifying the carotenoid biosynthesis pathway in *C. protothecoides*.
**Keywords:** *Chlorella protothecoides* CS-41; phytoene synthase; *Cppsy*; promoter
## **1. Introduction**
Lutein is one of more than 750 known naturally occurring carotenoids, and is synthesized by
higher plants, bacteria, fungi, and algae. Based on its molecular structure (containing oxygen), it belongs to the xanthophyll family, one of the two major carotenoid families. In the plant kingdom, lutein provides photoprotection by scavenging singlet oxygen and peroxyl radicals [1], and its bright yellow color helps plants achieve effective cross pollination. Humans cannot synthesize lutein themselves, yet it is essential for the human body. Lutein is the predominant carotenoid in the infant brain [2], and is supplemented in newborn babies in the first hours of life. Lutein can increase biological antioxidant potential and reduce the plasma concentration of total hydroperoxides. It also reduces free radical-induced damage [3]. Lutein is the main carotenoid in the human retina; hence, it has been used as a therapeutic agent for the prevention of age-related macular degeneration [4,5]. Epidemiologic data suggest that lutein plays an active role in delaying chronic diseases [6], stimulating the immune response [7], and hampering the development of cataracts and atherosclerosis [8,9]. A recent study showed that a lutein-based dye used during chromovitrectomy in humans could improve the identification and removal of the vitreous, internal limiting membrane and the epiretinal membrane [10].
As lutein has many functions, it has become increasingly important to find and create more sources of lutein production. In recent years, algae have received a great deal of attention in the production of carotenoids and proteins. Previous studies in our laboratory showed that heterotrophically cultivated *Chlorella protothecoides* CS-41 can produce considerable amounts of lutein [11]. Furthermore, optimization of the cultivation conditions, medium composition, and extraction techniques can improve lutein yields [12,13]. However, to date, there are no reports of the enhancement of lutein production by this alga using genetic modification, although genetic engineering technologies have become increasingly popular in the field of carotenoid production. The first step is to determine the genes involved in lutein biosynthesis—information that is essential for genetic modification.
It has been found that phytoene synthase (PSY) is the rate-limiting enzyme in the carotenoid biosynthesis pathway in photosynthetic organisms [14–16]. In many cases, the rate of lutein formation through the carotenoid biosynthetic pathway appears to be controlled by PSY, which catalyzes the
head-to-head condensation of two geranylgeranyl diphosphate molecules to yield phytoene—the first committed reaction in carotenogenesis. Since PSY plays a key role in the first step of carotenogenesis, it has unsurprisingly been chosen for genetic engineering studies of carotenoid production.
PSY has been extensively studied in bacteria and higher plants, but its study in algae is still in its infancy. For unicellular green algae, the *psy* gene has previously been investigated in *Chlamydomonas reinhardtii* [17], *Duanliella bardawil* [18], and *Haematococcus pluvialis* [19]. In *C. reinhardtii*, deletion of the *psy* gene resulted in a white phenotype [20]. For *Haematococcus*, *psy* was shown to be
up-regulated under stress conditions of high light and low nutrient availability [21]. Overexpression of exogenous *psy* from *D. salina* [22] or *C. zofingiensis* [23] in *C. reinhardtii* has been shown to increase the lutein content to over 1.25 and 2.2-fold, respectively.
As an efficient lutein-production alga, *C. protothecoides* CS-41 has high potential for application in the commercial production of lutein; however, its lutein biosynthesis pathway has not been well studied. Our research group previously cloned other key enzyme genes in the lutein biosynthesis pathway of this alga, such as the phytoene desaturase (*pds*) (GenBank accession No. FJ968162) [24], zetacarotene desaturase (*zds*) (GenBank accession No. GU269622) [25], and lycopene-<sup>Ή</sup>cyclase (*lyce*) (GenBank accession No. FJ752528) genes. The *psy* gene is essential for determining the complete lutein biosynthesis pathway in this alga. Therefore, in this study, the *psy* gene from the unicellular microalga *C. protothecoides* CS-41 and its promoter were isolated and analyzed. This study provides an important theoretical basis for the genetic modification of lutein biosynthesis in *C. protothecoides* CS-41, including gene sequences, expression promoter candidates, and possible regulatory environmental factors for gene expression.
## **2. Results and Discussion**
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*2.1. Cloning and Characterization of the psy Gene from C. protothecoides*
Touchdown PCR with primers YF and YR (Table 1) generated a predicted 373 bp fragment (Supplementary Figure S1, lane 1). BLAST analysis showed that the nucleotide sequence of this fragment shared about 74% and 73% identities with that of *C. reinhardtii* and *D. salina*, respectively, demonstrating that this fragment sequence is derived from a putative phytoene synthase.
**Table 1.** PCR primers and target fragments for *Cppsy.* F: forward; R: reverse; O: outer primer; I: inner primer.
With the sequence information, specific primers were designed for 5ȝ- and 3<sup>ȝ</sup>rapid amplification of cDNA ends (RACE) of the related gene. 5ȝ-RACE generated a 598 bp fragment (Supplementary Figure S1, lane 2), and 3ȝ-RACE produced an 816 bp fragment (Supplementary Figure S1, lane 3). They were displayed by sequencing as the 5ȝ and 3<sup>ȝ</sup> regions of the phytoene synthase gene of *C. protothecoides* (*Cppsy*). RT-PCR (Reverse Transcription) with a pair of primers YF1 and YR1 generated an 1143 bp fragment (Supplementary Figure S1, lane 4), which was identified as the fulllength *Cppsy* cDNA (GenBank accession No. FJ968161).
The open reading frame of *Cppsy* cDNA encoded a protein of 380 amino acid residues with a calculated molecular mass of 43.035 kDa and an isoelectric point of 6.40 (http://cn.expasy.org/
tools/protparam.html) and shared 81.7% identical sequence with *Chlorella* NC\_64A.
To characterize the corresponding gene of *Cppsy* cDNA, genomic PCR was performed. A 2488 bp fragment (Supplementary Figure S1, lane5) (GenBank accession No. GU351883) was generated and sequenced. Analysis of the obtained nucleotide sequence revealed that the product was the corresponding *Cppsy* gene.
The Southern blot analysis results indicated that there is only one *Cppsy* gene copy in *C. protothecoides* CS-41 (Supplementary Figure S2), which is different to those in higher plants. *Psy* gene replication
is common in dicot plants, such as tomato (SlPSY1 and SlPSY2), and in monocot plants, such as maize (ZmPSY1-3), rice (OsPSY1-3), and sorghum (SbPSY1-3) [16,26– 28].
Analysis of the *Cppsy* gene structure (Figure 1) revealed that it is more complicated than those of dicot and monocot plants. It consists of ten exons and nine introns. *Chlorella* has a higher intron density than other algae and higher plants; in most of the higher plants, *psy* genes always have four or five introns, but this alga has nine introns. Compared with the structure of the *psy* gene from
*C. reinhardtii* (*Crpsy*), it seems that there are two introns inserted into each of the first and second exons, and one intron inserted into the fourth exon, which makes the gene structure more complicated (Figure 1C).
**Figure 1.** Exons and introns of the *Cppsy* gene in *C. protothecoides* CS-41.The ten exons are: (1) 1 bp to 280 bp; (2) 477 bp to 576 bp; (3) 691 bp to 743 bp; (4) 913 bp to 1030 bp; (5) 1139 bp to 1180 bp; (6) 1327 bp to 1427 bp; (7) 1635 bp to 1793 bp; (8) 1957 bp to 2045 bp; (9) 2171 bp to 2233 bp; (10) 2351 bp to 2488 bp. (**A**) Intron density; (**B**) DNA structure; (**C**) The relationship between introns and exons of *Cppsy*, *Crpsy*, and *Cnpsy* genes. *Dbpsy*, *Dspsy*, *Hppsy*, *Crpsy*, *Cnpsy*, *Cppsy*, *Mzpsy*, and *Atpsy* are the *psy* genes of *Duanliella bardawil*, *Duanliella salina*, *Haematococcus pluvialis*, *Chlamydomonas reinhardtii*, *Chlorella* NC\_64A, *Chlorella protothecoides* CS-41, *Zea mays*, and *Arabidopsis thaliana*, respectively.
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*2.2. Sequence Alignment and Phylogenetic Reconstruction*
After the DNA and cDNA sequences of the *Cppsy* gene were determined, it was possible to investigate its evolutionary position among the various *psy* genes. Using MEGA 4.0 from Clustal W1.6 alignments, the phylogenetic tree of PSYs from different organisms was constructed based on their deduced amino acid sequences. It showed that *psy* was derived from an ancestor gene and later evolved into four subgroups, including higher plants, cyanobacteria, algae, and bacteria (Figure 2). According to the neighbor-joining (NJ) tree, *Cppsy* belongs to the algae group, and is more ancient than plant species (Figure 2).
The deduced amino acid sequence of *Cppsy* was submitted to NCBI for PSI-BLAST searches and the results showed that *Cppsy* has high homology with *psy* genes from other algal species, with 83% identity and 88% positives with *psy* from *Chlorella* NC\_64A. *Cppsy* was also highly similar to *psy* from *C. reinhardtii* (67% identities, 79% positives), *H. pluvialis* (63% identities, 77% positives), *D. bardawil* (68% identities, 80% positives), and *D. salina* (68% identities, 79% positives), suggesting that *Cppsy* belongs to the algae *psy* family. In the algae family, CpPSY belongs to class I of PSY according to Tran's data [29]. BlastP analysis suggested that this protein has the essential characteristics of PSY. It belongs to the Isoprenoid\_Biosyn\_C1 superfamily, and contains the consensus sequence, including three predicted substrate-Mg2+ binding sites (aspartate-rich regions) (DXXXD), 130- DELVD-134, 203-DELYD-207, and 256-DEGED-260 (Figure 3A). In other algae and higher plants, there are two (DELVD and DVGED) (Figure 3A); hence, CpPSY has one more DXXXD motif than other PSYs. The abundant 203-DELYD-207 site possibly plays an important role in the function of CpPSY, which should be studied further.
**Figure 2.** Phylogenetic tree of PSY sequences from various species. The phylogeny was derived using neighbor-joining analysis. The accession numbers of the amino acid sequences follow the taxon names. Horizontal branch lengths represent relative evolutionary distances, with the scale bar corresponding to 0.05 amino acid substitutions per site.
*Mar. Drugs* **2015,** *13*, 6620–6635
**Figure 3.** (**A**) Alignment of the selective PSY-deduced amino acid sequences from different algae produced with the GeneDoc program using Clustal W. The alignment indicates aspartate-rich regions/substrate-Mg2+ binding sites (DXXXD). The three DXXXD motifs are shown by the red boxes. Cppsy, Cvpsy, Mspsy, Olpsy, Dbpsy, Dspsy, Hppsy, Cspsy, Czpsy, Vcpsy, Ntpsy, Atpsy, and Zmpsy are the PSY of *Chlorella protothecoides* CS-41, *Chlorella variabilis*, *Micromonas* sp. RCC299, *Ostreococcus lucimarinus*, *Duanliella bardawil*, *Duanliella salina*, *Haematococcus pluvialis*, *Coccomyxa subellipsodiea* C-169, *Chromochloris zofingiensis*, *Volvox carteri* f. *nagariensis*, *Nicotiana tabacum*, *Arabidopsis thaliana*, and *Zea mays*, respectively; (**B**) Three-dimensional model structure of CpPSY. Comparative modeling was performed using homology-based
three-dimensional structural modeling. The three aspartate-rich motifs (DXXXD) are colored in orange (DELVD), yellow (DELYD), and magenta (DEGED); others are shown in green. The *N*-terminus and *C*-terminus are also shown; (**C**) High-performance liquid chromatography trace and UV spectrum of carotenoid pigments in the *E. coli* heterologous complementation system. Pigments extracted from *E. coli* cells transformed with pACCRT-E and pUC-psy together (1), pUC-psy only (2), and pACCRT-EB only (3). Absorbance was recorded at 285 nm. The peak indicated by the arrow is phytoene.
The secondary structure prediction carried out at NPS@ (https://npsaprabi.ibcp.fr/) showed that CpPSY consists of 58.68% alpha helix, 26.58% random coil, 10.79% extended strand, and 3.95% beta turn. The tertiary structure of CpPSY was constructed using homology-based modeling by Swiss-Model (Figure 3B). A total of 50 models were found. Squalene synthase (HpnC) was used as a template for molecular modeling, since the identity is the highest (30.42%). The modeled structure also showed that CpPSY consists mostly of alpha helices. The three conserved DXXXD motifs (orange DELVD, yellow DELYD, and magenta DEGED) were marked in the three-dimensional model structure (Figure 3B). It seems that the three DXXXD motifs form a circle-like structure, which could be important for enzyme activity.
All of the analysis results strongly suggest that PSY from *C. protothecoides* CS-41 is an algal phytoene synthase protein involved in the carotenoid biosynthesis pathway. Bacterial complementation assay further confirmed that this gene is functional. The expressed protein in pUC-psy could catalyze the GGPP produced by pACCRT-E (Figure 3C,1) to phytoene, similar to the function of the *crtB* gene in pACCRT-EB (Figure 3C,3).
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*2.3. Promoter Isolation and Analysis*
The promoter region of the *Cppsy* gene was cloned from *C. protothecoides* CS-41 genomic DNA. The cloned *Cppsy* promoter region was determined to be 1980 bp in length, and the sequence is shown in Figure 4. Furthermore, the cloned *Cppsy* promoter region was analyzed using the PLACE and PlantCARE databases. Several core fragments were identified, which are homologous to the *cis*-acting elements of higher plants and of great importance for the promoter functions (Figure 4). Three types of elements, which have been found to be regulated by hormones in the upstream region of some
plant genes, are present in the *Cppsy* promoter: the ABRE type (CCTGCGTGGC, CACGTG, and GCCTCGTGGC) involved in abscisic acid responsiveness; the TGACG-motif (TGACG), the *cis*-acting regulatory element involved in methyl jasmonate (MeJA) responsiveness; and the Sp1 (CCCCCGCCA and ACCCGCCATG), MNF (GTGCCCCATGCAGGTT) and Box I (TTTCAAA) types involved in light responsiveness.
The transcriptional start site (TSS) of the *Cppsy* promoter was determined by 5<sup>ȝ</sup>-RACE using total RNA extracted from *C. protothecoides*. The TSS is an adenine (A) at 34 bp upstream of the translation initiation codon. The distance between the putative TATA-box and TSS is approximately ƺ24 to ƺ28 bp, which is consistent with the distance of 32 bp ± 7 bp from previous data [30].
A previous study showed that the *psy* expression level is affected by light [31] and other biotic and abiotic stresses [16] in higher plants. Gene expression response to environmental stress is related to the regulation mechanism. To understand more about the regulation mechanism, we need to know more information about the gene, including the gene structure and regulatory domains. Here, many elements were found in the *Cppsy* promoter that belong to light-responsive elements, such as Sp1, MNF1, G-box, and chs-CMA2a. There were also some *cis*-acting elements involved in abscisic acid (ABRE) and MeJA (TGACG-motif) responsiveness. Light is one of the most important environmental factors for algae. To determine whether these motif sequences from the *Cppsy* promoter are involved in light, abscisic acid, and MeJA signaling, loss-of-function analysis needs to be carried out in future studies.
**Figure 4.** Promoter sequence of *Cppsy* from *C. protothecoides* CS-41. Numbers indicate the positions relative to transcriptional start site (TSS). The TSS is indicated as +1 and in bold; important *cis*-elements are underlined.
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*2.4. Gene Expression Response to Light and MeJA*
To investigate the effects of light and MeJA on the promoter inducibility at different time points,
we analyzed *Cppsy* mRNA expression levels after light and MeJA treatment. The results show that on treatment with light, *Cppsy* gene expression increased by up to 36 times, compared with the dark, which indicates that *Cppsy* gene expression is upregulated in response to light (Figure 5A). When treated with MeJA, *Cppsy* gene expression peaked at 10 h after treatment (Figure 5B). Therefore, the gene expression of *Cppsy* is significantly induced by treatment with light and MeJA (*p* < 0.01). These results confirm that the *Cppsy* promoter is induced by light and MeJA, and can be used as a candidate promoter element for the genetic modification of carotenoid biosynthesis in *Chlorella*, other algae, or higher plants.
**Figure 5.** The expression of *Cppsy* gene induced by light (**A**) and MeJA (**B**) at different time points in *C. protothecoides* CS-41. Data (mean ± SEM) are combined from three independent experiments. # indicates that the gene expression levels were significantly different from that at 0 h (*p* < 0.01).
## **3. Experimental Section**
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*3.1. Strains and Culture Conditions*
The microalgal strain used in this study was *C. protothecoides* CS-41 obtained from CSIRO Marine Laboratories (Hobart, Australia). They were grown in modified basal medium [32] containing 10 g·Lƺ1 glucose at 28 °C and 180 rpm, and were collected at the log phase or late log phase.
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"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 187
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*3.2. Genomic DNA and RNA Isolation*
Genomic DNA was extracted using a modified cetyltrimethylammonium bromide (CTAB) method [33]. The total RNA was isolated from *C. protothecoides* CS-41 cells at the late log phase (about 5-day incubation, with a cell density around 1 × 108 cells mLƺ1) using TRIzol® reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions.
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"isbn": "9783039431908",
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*3.3. Cloning of Full-Length Cppsy cDNA and Its Corresponding Gene*
Degenerate primers (Table 1) were designed for the amplification of a partial *Cppsy* cDNA from *C. protothecoides* CS-41. The primers were derived from the highly conserved nucleotide and amino acid sequences reported for the *psy* genes from five kinds of algae (*C. reinhardtii*, *H. pluvialis*, *D. salina*, *D. bardawil*, and *Chlorella* NC\_64A).
Sets of specific primers were synthesized based on the sequence of putative insert for 5ȝ- and 3ȝ-RACE [34]. YFO1 and YRI1 were used for 5ȝ-RACE, and YFO2 and YRI2 (Table 1) were used for 3ȝ-RACE. RACE was performed using the 5ȝ-Full RACE Kit and 3ȝ- Full RACE Core Set Version 2.0 (TaKaRa, Dalian, China) according to the manufacturer's protocol. The RACE products were gel purified and sequenced as previously described. One pair of specific primers, YF1 and YR1 (Table 1), was designed from the sequences of the 5ȝ- and 3ȝ-RACE fragments to amplify the fulllength *Cppsy* cDNA and its corresponding gene.
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*3.4. Southern Blot Analysis*
According to the *Cppsy* genomic DNA sequence, *Bam*HI, *Eco*RI, *Nco*I, *Sma*I, and *Xma*I, which showed no recognition sites in the probed region of the *Cppsy* gene, were chosen to digest the whole genomic DNA. The probe was prepared by amplifying genomic DNA with the primers psy-F and psy-R, resulting in a 552-bp fragment of the *Cppsy* gene. The digested DNA was transferred to a Hybond-N membrane (GE Healthcare, Little Chanfont, UK) by capillary transfer and hybridized with a 32P-labelled DNA probe at both low and high stringency overnight.
After hybridization, the radioactivity of the membrane was monitored using a Storm 840 Phosphor Imaging System (Molecular Dynamics, Sunnyvale, CA, USA).
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 190
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*3.5. Bioinformatics Analysis*
Comparative and bioinformatic analyses of the nucleotide sequences and deduced amino acid sequences were carried out online at http://www.ncbi.nlm.nih.gov and http://cn.expasy.org. The nucleotide sequence, deduced amino acid sequence, and open reading frame (ORF) were analyzed, and sequence comparison was conducted through database searches using BLAST programs (http://www.ncbi.nlm.nih.gov/BLAST/) and GeneDoc software. The phylogenetic analysis of *psy* from other plant species was aligned with Clustal X program version 1.83 using default parameters [35] and manual adjustments where necessary. A phylogenetic tree was constructed using MEGA (molecular evolutionary genetics analysis) program, version 4.0 [36] from Clustal W1.6 alignments. The NJ [37] method was used to construct the tree. In the NJ method, the P distance was used to analyze the amino acid sequences. A total of 1000 repetitions were performed using the bootstrap method to determine the reliability of each node of the tree. The homology-based three-dimensional structural modeling of PSY was accomplished using Swiss-Model and WebLab Viewer Lite (http://swissmodel.expasy.org).
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*3.6. Functional Complementation Experiment in E. coli*
*E. coli* JM109 (Table 2) was used as a host for complementation experiments by cotransformation of the plasmid pUC-psy with pACCRT-E (Table 2). *E. coli* JM109 harboring only plasmid pACCRT-EB (Table 2) was cultured as a positive control, and only plasmid pUC-*psy* was cultured as a negative control for PSY functional analysis. The different strains were cultivated in 100 mL LB medium containing 100 μg·mLƺ1 ampicillin and 50 μg·mLƺ1 chloramphenicol at 28 °C and 180 rpm. IPTG (1 mM) was added when the optical density at 600 nm (OD600) reached 0.5, and the culture was kept at 28 °C for 2 days. The *E. coli* cells were collected by centrifugation at 12,000 rpm and used for
high-performance liquid chromatography (HPLC) analysis.
**Table 2.** Strains and plasmids used in this study*.*
Pigments in the bacteria were extracted according to procedures described by Breitenbach *et al.* [38]. *E. coli* JM109 cells harboring different plasmids were collected by centrifugation and freeze dried. Pigment extraction was carried out in acetone (80%, v/v) using ultrasonication, and the solvent was removed by blowing with nitrogen gas. The carotenoids were then resuspended in acetone for subsequent HPLC analysis. All operations were carried out on ice under dim light to prevent photodegradation, isomerizations, and structural changes of the carotenoids. The samples were prepared for HPLC by dissolving the dried residues in 1 mL of acetone and filtered through a polycarbonate 0.22 μm filter (Millipore, Carrigtwohill, Ireland). The extracted pigments were separated on a Kromasil KR100-5C18 analytical column (250 mm × 4.6 mm, 5 μm) using an UltiMate3000 HPLC system (Thermo Fisher Scientific, Waltham, MA, USA). The procedures described by Huang [39] were used with the following modifications: the mobile phase consisted of solvent A (acetonitrile/methanol/0.1 M Tris-HCl (pH 8.0), 84:2:14, v/v/v) and solvent B (methanol/ethyl acetate, 68:32, v/v). Pigments were eluted at a flow rate of 1 mL·minƺ1 with a linear gradient from 100% solvent A to 100% solvent B over a 5 min period, followed by 25 min of solvent B. The column temperature was maintained at 30 °C and the sample volume was 20 μL. The pigments were monitored by diode array detector, and the targeted products were identified by their absorption spectra and typical retention times compared with the control.
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{
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"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
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"isbn": "9783039431908",
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*3.7. Promoter Isolation and Analysis*
The Genomic Walking Kit (TaKaRa, Dalian, China) was used to obtain promoter regions of the *Cppsy* gene from *C. protothecoides* CS-41. Based on the *Cppsy* genomic sequences, gene-specific primers were designed and are listed in Table 1. Primary and nested PCRs were performed with the *Cppsy* gene-specific primers and Genome Walking adapter primers (AP1) in the kit according to the manufacturer's instruction. The primary nested PCR products were diluted to 1:50 with distilled water for subsequent nested PCR. The nested PCR products were purified from 1.2% (w/v) agarose gel and sub-cloned into the pMD18T vector (TaKaRa, Dalian, China). The cloned vectors were then sequenced and the putative *cis*-regulatory elements were analyzed using the PLACE [40] and PlantCARE databases [41].
## *3.8. Gene Expression Response to Light and MeJA*
To analyze the light regulation pattern of the *Cppsy* gene in *C. protothecoides*, algal cells in the late log phase were cultivated in the dark for more than 2 days, then collected by centrifugation at 5000 rpm for 15 min in the darkness. The pellet was resuspended in fresh medium without glucose, and then subjected to light treatment under a light intensity of 120 μmol mƺ2·sƺ1 for different induction times
(0, 0.5, 1.0, 2.0, and 4.0 h). Each treatment was carried out with three parallel repetitions.
To analyze the MeJA regulation pattern of the *Cppsy* gene in *C. protothecoides*, algal cells in the log phase were treated with 100 μM MeJA (Sigma, St. Louis, MO, USA) diluted in dimethyl sulfoxide (DMSO) for 0, 2, 4, 6, 8 10, and 12 h. Control cultures were treated with DMSO only.
The effects of light and MeJA on the *Cppsy* gene transcripts in *C. protothecoides* were quantified
by reverse transcriptase quantitative PCR (RT-qPCR). The RT-qPCR experiment was performed in
two steps: the cDNA templates were synthesized from RNA samples using Prime Script™ Reverse Transcriptase Reagent according to the manufacturer's instructions (TaKaRa, Dalian, China) using oligo (dT) as the primer; then qPCR was conducted on an iQ Cycler (Bio-Rad, Watford, UK) using the specific primers (Table 1) and the SYBR ExScript RT-PCR kit (TaKaRa, Dalian, China). The specific primers for the corresponding genes included psyRT-F and psyRT-R for the *Cppsy* gene, and 16SRT-F and 16SRT-R for the 16S gene (Table 1). Before the qPCR expression analysis, we checked the amplification efficiency of each primer pairs, and all were well controlled between 99.83% and 101.25%.
Each qPCR measurement was carried out independently at least three times, and the mean value was used for quantification. The 2ƺ̇̇CT method was used to analyze the relative changes in gene expression, the expression of the 16S gene was used as a normalized control, and the expression of the untreated samples was used as a negative control.
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**4. Conclusions**
Carotenoid pathways in plants have been described in great detail using genetic, biochemical, and molecular data, mainly from *Arabidopsis* and other higher plants; however, this is the first study in the unicellular microalga *C. protothecoides* CS-41.
We successfully isolated and analyzed the *Cppsy* gene, which encodes the functional phytoene synthase—a vital enzyme for carotenoid biosynthesis in *C. protothecoides* CS-41—as well as its promoter. Computational analysis suggested that this protein belongs to the Isoprenoid\_Biosyn\_C1 superfamily. It contains one more substrate-Mg2+ binding site than other algae and higher plants. Analysis also demonstrated several candidate motifs for the promoter, which exhibited light- and MeJA-responsive characteristics. Light- and MeJA treatment showed that the *Cppsy* expression level was significantly enhanced by light and MeJA.
These achievements will be helpful to understand more about the regulatory mechanism of the carotenoid biosynthesis pathway in algae and the mechanisms for accumulation of lutein and other important carotenoids.
## **Acknowledgments**
This work was supported by the National 863 Program of China (2012AA101601), and the Agri-X Program of Shanghai Jiao Tong University (Agri-X2015005).
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**Author Contributions**
Chunlei Shi and Xianming Shi conceived and designed the experiments. Meiya Li, Yan Cui and Zhibing Gan performed the experiments. Chunlei Shi, Meiya Li and Yan Cui analyzed the data and prepared the manuscript.
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{
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**Conflict of Interest**
The authors declare no conflict of interest.
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{
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 196
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**Takashi Maoka**
Research Institute for Production Development, 15 Shimogamo-morimoto-cho, Sakyo-ku,
Kyoto 606-0805, Japan; E-Mail: [email protected]; Tel.: +81-75-781- 1107; Fax: +81-75-781-1118
*Received: 14 January 2011; in revised form: 16 February 2011 / Accepted: 21 February 2011 / Published: 22 February 2011*
**Abstract:** Marine animals contain various carotenoids that show structural diversity. These marine animals accumulate carotenoids from foods such as algae and other animals and modify them through metabolic reactions. Many of the carotenoids present in marine animals are metabolites of Ά-carotene, fucoxanthin, peridinin, diatoxanthin, alloxanthin, and astaxanthin, *etc.* Carotenoids found in these animals provide the food chain as well as metabolic pathways. In the present review, I will describe marine animal carotenoids from natural product chemistry, metabolism, food chain, and chemosystematic viewpoints, and also describe new structural carotenoids isolated from marine animals over the last decade.
**Keywords:** carotenoids; marine animals; metabolism; food chain; chemosystematic
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"isbn": "9783039431908",
"section_idx": 199
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**1. Introduction**
Since the first structural elucidation of Ά-carotene by Kuhn and Karrer in 1928– 1930, about 750 naturally occurring carotenoids had been reported as of 2004 [1]. Improvements of analytical instruments such as NMR, MS, HPLC, *etc.*, have made it possible to perform the structural elucidation of very minor carotenoids in nature [2–4].
Marine animals contain various carotenoids that show structural diversity [3–9]. Among the 750 reported carotenoids found in nature, more than 250 are of marine origin. In particular, allenic carotenoids, except for neoxanthin and its derivatives, and all acetylenic carotenoids originate from marine algae and animals [1].
In general, animals do not synthesize carotenoids *de novo*, and so those found in animals are either directly accumulated from food or partly modified through metabolic reactions [5–9], as shown in Figure 1. The major metabolic conversions of carotenoids found in animals are oxidation, reduction, translation of double bonds, oxidative cleavage of double bonds, and cleavage of epoxy bonds.
Up until 2001, marine animal carotenoids were reviewed by Liaaen-Jensen [5,6], Matsuno [7,8], and Matsuno and Hirao [9]. Since then, there have been no reviews of carotenoids in marine animals. The present review describes progress in the field of carotenoids in marine animals over the last decade.
## **2. Porifera (Marine Sponges)**
Characteristic carotenoids in marine sponges are shown in Figure 2. Many marine sponges are brilliantly colored due to the presence of carotenoids. Sponges are filter feeders and are frequently associated with symbionts such as microalgae or bacteria [6]. The characteristic carotenoids in sponges are aryl carotenoids such as isorenieratene (**1**), renieratene (**2**), and renierapurpurin (**3**) [6,7]. More than twenty aryl carotenoids have been reported in sponges [1]. Except for sea sponges, aryl carotenoids are found only in green sulfur bacteria [1,6]. Therefore, aryl carotenoids in sponges are assumed to originate from symbiotic bacteria [6,7]. Novel carotenoid sulfates having an acetylenic group, termed bastaxanthins (**4**), were isolated from the sea sponge *Ianthella basta* [1]. Recently, a new acetylenic carotenoid (**5**) was isolated from the marine sponge *Prianos osiros* [10]. Based on the structural similarity, bastaxanthins and compound **5** were assumed to be metabolites of fucoxanthin originating from microalgae.
**Figure 2.** Characteristic carotenoids in marine sponges.
## **3. Coelenterata (Sea Anemones)**
Astaxanthin, which originates from dietary zooplankton, was found in some jelly fish. Peridinin, pyrrhoxanthin, and diadinoxanthin were found in some corals [11]. They originate from symbiotic dinoflagellates. Unique nor carotenoids, 2-norastaxnthin (**6**) and actinioerythrin (**7**), have been reported in the sea anemones *Actinia equina* and *Tealia felina* [1] (Figure 3).
> **Figure 3.** Characteristic carotenoids in sea anemones.
## **4. Mollusca (Mollusks)**
Many chitons are herbivorous and feed on attached algae. Major carotenoids found in chitons are lutein, zeaxanthin, fucoxanthin, and their metabolites [12].
Abalone, *Haliotis discus discus*, and turban shell, *Turbo cornutus*, feed on brown and red algae. Carotenoids found in these shells are Ά-carotene, ΅-carotene, zeaxanthin, lutein, and fucoxanthin [11].
On the other hand, many sea snails are carnivores. The triton *Charonia sauliae* feeds on starfish. Therefore, astaxanthin (**8**), 7,8-didehydroastaxanthin (**9**), and 7,8,7<sup>ȝ</sup>,8<sup>ȝ</sup>-tetradehydroastaxanthin (**10**), characteristic carotenoids found in starfish, were isolated as major carotenoids in triton. Astaxanthin (**8**), originating from dietary microcrustaceans, was found to be a major carotenoid in the whelk *Buccinum bayani*. Alternatively, *Drupella fragum* preys upon corals. Thus, peridinin and diadinoxanthin are present as major carotenoids in this sea snail [11]. Carotenoids in sea snails well reflect their diet.
Canthaxanthin (**11**), (3*S*)-adonirubin (**12a**), and (3*S*,3<sup>ȝ</sup>*S*)-astaxanthin (**8a**) were found to be major carotenoids in the spindle shell *Fushinus perplexus* [13]. Furthermore, a series of carotenoids with a
4-hydroxy-5,6-dihydro-Ά-end group and/or 3,4-dihydroxy-5,6-dihydro-Ά-end (**13**– **15**) were isolated from *Fushinus perplexus* [13] (Figure 4). They were assumed to correspond to reduction metabolites of canthaxanthin (**11**), (3*S*)-adonirubin (**12a**), and (3*S*,3<sup>ȝ</sup>*S*)-astaxanthin (**8a**).
Sea slugs and sea hares also belong to Gastropoda. They are herbivorous and feed on brown and red algae. Several apocarotenoids have been reported in sea slugs and sea hares [1]. A series of
<sup>8</sup><sup>ȝ</sup>-apocarotenal and 8<sup>ȝ</sup>-apocarotenols derived from Ά-carotene, lutein, and zeaxanthin were found in the sea hare *Aplysia kurodai* [14]. They are oxidative cleavage products of the polyene chain at C-8 in C40 skeletal carotenoids [14].
Bivalves (oyster, clam, scallop, mussel, ark shell, *etc.*) contain various carotenoids that show structural diversity [3,6]. Bivalves accumulate carotenoids obtained from their dietary microalgae and modify them through metabolic reactions. Many of the carotenoids present in bivalves are metabolites of fucoxanthin, diatoxanthin, diadinoxanthin, and alloxanthin [3,6], which originate from microalgae.
> **Figure 4.** Characteristic carotenoids in sea snails.
Oxidative metabolites of diatoxanthin (**16**) and alloxanthin (**17**), such as pectenol (**18**), pectenolone (**19**), 4-hydroxyalloxanthin (**20**), and 4-ketoalloxanthin (**21**), are distributed in scallops and ark shells [3,6,7]. 8ȝ-Apoalloxanthinal (**22**), which is an oxidative cleavage product of alloxanthin, was also found in bivalves [15] (Figure 5).
A novel 3,6-epoxy derivative of diadinoxanthin (**23**), named cycloidadinoxanthin (**24**), was also isolated from the oyster [16] (Figure 5).
**Figure 5.** Metabolites of diatoxanthin, alloxanthin, and diadinoxanthin in bivalves.
Fucoxanthin (**25**) and its metabolites fucoxanthinol (**26**) and halocynthiaxanthin (**27**) were found to be widely distributed in oysters and clams [3,6,7].
Mytiloxanthin (**28**), which has a unique enol hydroxy group at C-8ȝ in the polyene chain and a
3<sup>ȝ</sup>-hydroxy-6<sup>ȝ</sup>-oxo-Ύ-end group, is a characteristic carotenoid in marine mussels and oysters [6,7]. Furthermore, three mytiloxanthin analogues containing an allenic end group (**29**), a 3,6-epoxy-end group (**30**), and a 3,4-dihydroxy-Ά-end group (**31**) were isolated from the oyster [16,17]. Compound **<sup>29</sup>**, termed allenic mytiloxanthin, was assumed to be a metabolic intermediate from fucoxanthinol to mytiloxanthin.
Some edible clams have a bright orange or red color due to the presence of carotenoids. Fucoxanthin 3-ester (**32**) and fucoxanthinol 3-ester (**33**) were found to be major carotenoids in *Mactra chinensis* [18], *Ruditapes philippinarum*, and *Meretrix petechialis* [19]*.* Amarouciaxanthin A (**34**) and its ester were also identified as major carotenoids in *Paphia amabills* and *Paphia amabillis* [20].
Other metabolites of fucoxanthin, crasssostreaxanthin A (**35**) and crassostreaxanthin B (**36**), were isolated from the Japanese oyster *Crassostrea gigas* [21]. Tode *et al.* demonstrated that crassostreaxanthin B could be converted from halocynthiaxanthin by bio-mimetic chemical reactions [22,23]. Further studies of carotenoids in marine animals revealed that crassostreaxanthin A, crassostreaxanthin B, and their 3-acetates were widely distributed in marine bivalves [16,17]. Moreover, two crassostreaxanthin A analogues, **37** and **38**, were isolated from the oyster as minor components [16,17]. Metabolic pathways of fucoxanthin in bivalves are shown in Figure 6.
**Figure 6.** Metabolic pathways of fucoxanthin in bivalves.
Bivalves also feed on dinoflagellates. Peridinin (**39**), a characteristic carotenoid in dinoflagellates with a C37-skeletal structure, and its metabolites (**<sup>40</sup>**–**43**) were also found in some bivalves. Recently, four new C37-skeletal carotenoids (**<sup>44</sup>**–**47**) were isolated from *Crassostrea gigas* [16,17], *Paphia amabillis* [20], and *Corbicula japonica* [24,25]. The metabolic pathways of peridinin in bivalves are shown in Figure 7. As well as fucoxanthin, the major metabolic conversions of peridinin in bivalves are hydrolysis of acetyl group, conversion of the allenic bond to an acetylenic bond, and hydrolysis cleavage of the epoxy ring, as shown in Figure 7.
> **Figure 7.** Metabolic pathways of peridinin in bivalves.
There are many reports on carotenoids in marine shellfish [6,7]. However, there are few
reports on the carotenoids of shellfish inhabiting brackish or fresh water [24,25]. Four new
carotenoids, corbiculaxanthin (**48**), corbiculaxanthin 3ȝ-acetate (**49**), 6- epiheteroxanthin (**50**), and
<sup>7</sup><sup>ȝ</sup>,8<sup>ȝ</sup>-didehydrodeepoxyneoxanthin (**51**), were isolated from the brackish clam *Corbicula japonica* and freshwater clam *Corbicula sandai* (Figure 8) [24,25]. 7<sup>ȝ</sup>,8<sup>ȝ</sup>-Didehydrodeepoxyneoxanthin (**51**) has an interesting structure, with both allenic and acetylenic moieties.
> **Figure 8.** New carotenoids in corbicula clams.
Carotenoids found in bivalves provide a key to the food chain as well as metabolic pathways.
Astaxanthin and its esters were found to be major carotenoids in species of octopus and cuttlefish. Their astaxanthins consisted of three optical isomers and originated from dietary zooplankton [26].
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**5. Arthropoda (Crustaceans)**
Carotenoids in the carapace of crustaceans exist as both free and esterified forms. The principal carotenoid in crustaceans is astaxanthin [6,7]. In crustaceans, astaxanthin exists as carotenoproteins such as crustacyanin, and exhibits purple, blue, and yellow colors. Many crustaceans can synthesize astaxanthin (**8**) from Άcarotene (**52**), ingested from dietary algae, via echinenone (**53**),
3-hydroxyechinenone (**54**), canthaxanthin (**11**), and adonirubin (**12**), as shown in Figure 9 [6,7].
In many crustaceans, hydroxylation at C-3 (C-3ȝ) in the 4-oxo-Ά-end group is nonestereo-selective. Therefore, astaxanthin, adonixanthin, and 3-hydroxyechinenone, having a 3-hydroxy-4-oxo-Ά-end group, present in crustaceans, are comprised of a mixture of optical isomers [6,7].
> **Figure 9.** Metabolism of Ά-carotene in crustaceans.
**(DFKFRPSRXQGFRQVLVWVZLWKRSWLFDOLVRPHUV**
Some crustaceans can convert zeaxanthin to adonixanthin (**55**) and lutein to fritschiellaxanthin (**56**) and papyrioerythrinone (**57**) [1,6,7]. Crustaceans belonging to Isopoda can introduce a hydroxy group at C-2 in the Ά-end group. This hydroxylation is also none-stereo-selective. Therefore, Ά-caroten-2-ol (**58**) in the sea louse *Ligia exotica* exists as two optical isomers [1,6,7]. Recently, two new carotenoids, 2,3<sup>ȝ</sup>dihydroxycanthaxanthin (**59**) [27] and 2,3-dihydroserythrin (**60**) [28], were isolated from the hermit crab *Paralithodes brevipes* and crawfish *Procambarus clarkii*, respectively (Figure 10).
> **Figure 10.** Characteristic carotenoids in crustaceans.
## **6. Echinodermata (Echinoderms)**
Echinenone is a well-known major carotenoid in the gonads of sea urchins and is an oxidative metabolite of Ά-carotene [6,7]. Echinenone from the gonads of sea urchins was found to have a 9ȝ*Z* configuration (**61**) [29].
Starfish are carnivorous and mainly feed on bivalves and small crustaceans. Principal carotenoids in starfish are astaxanthin (**8**), 7,8-didehydroastaxanthin (**9**), and 7,8,7<sup>ȝ</sup>,8<sup>ȝ</sup>-didehydroastaxanthin (**10**). They correspond to the oxidative metabolites of Ά-carotene, diatoxanthin, and alloxanthin, respectively. The crownof-thorns starfish *Acanthaster planci* is a large, nocturnal sea star
that preys upon coral polyps. Recently, four new carotenoids: 4- ketodeepoxyneoxanthin (**62**),
4-keto-4ȝ-hydroxydiatoxanthin (**63**), 3<sup>ȝ</sup>-epigobiusxanthin (**64**), and 7,8- dihydrodiadinoxanthin (**65**), were isolated from *A. planci* as minor components along with the major carotenoids
7,8-didehydroastaxanthin, peridininol, and astaxanthin, and several other minor carotenoids including 7,8,7<sup>ȝ</sup>,8<sup>ȝ</sup>-tetrahydroastaxanthin, diadinoxanthin, diatoxanthin, and alloxanthin [30].
3,4,3<sup>ȝ</sup>,4<sup>ȝ</sup>-Tetrahydroxypirardixanthin 4,4ȝ-disulfate, named ophioxanthin (**66**), was reported in the brittle star *Ophioderma longicaudum* [31]. Canthaxanthin and astaxanthin were found in the
gonads of sea cucumbers as major components. 5,6,5<sup>ȝ</sup>,6<sup>ȝ</sup>-Tetrahydro-Ά-carotene derivatives with 9*Z*,
9ȝ*Z* configurations, termed cucumariaxanthin (**67**), were isolated from the sea cucumber
*Cucumaria japonica* [32] (Figure 11).
Recently, zeaxanthin, astaxanthin, and lutein were identified from spiny sea-star *Marthasterias glacialis* by HPLC-PAD-atmospheric pressure chemical ionization-MS. These carotenoids showed strong cell proliferation inhibition activity against rat basophilic leukemia
RBL-2H3 cancer cell line [33].
## **7. Protochordata (Tunicates)**
As well as bivalves, tunicates are filter feeders. Carotenoids found in tunicates originate from phytoplankton such as diatoms, and are also metabolites of fucoxanthin, diatoxanthin, and alloxanthin [7,8].
Halocynthiaxanthin (**27**), an acetylenic analog of fucoxanthinol (**26**), and mytiloxanthinone (**68**), an oxidative metabolite of mytiloxanthin (**28**), were first isolated from the sea squirt *Halocynthia roretzi* [34]. They are widely distributed in various tunicates. Amarouciaxanthin A (**34**) and amarouciaxanthin B (**69**), having a unique 3-oxo-6-hydroxy-Ή-end group, were first isolated from the tunicate *Amaroucium pliciferum* [35] (Figure 12). Peridinin and its metabolites are also found in tunicates.
**Figure 11.** Characteristic carotenoids in echinoderms.
**Figure 12.** Metabolic pathways of fucoxanthin in tunicates.
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**8. Pisces (Fish)**
Many fish accumulate carotenoids in their integuments and gonads. On the other hand, Salmonidae fish peculiarly accumulate astaxanthin (**8**) in muscle. Except for catfish, carotenoids in the integuments of fish exist in an esterified form.
Astaxanthin (**8**) is widely distributed in both marine and fresh water fish. Cyprinidae fish, which inhabit fresh water, can synthesize (3S,3ȝS)-astaxanthin (**8a**) from zeaxanthin (**70**) by oxidative metabolic conversion (Figure 13). On the other hand, Perciformes and Salmonidae fish cannot synthesize astaxanthin from other carotenoids [6,7,36]. Therefore, astaxanthin present in these fish originates from dietary crustacean zooplankton. Astaxanthin in these marine fish comprises three optical isomers. Perciformes and Salmoidae fish can convert astaxanthin to zeaxanthin [36,37]. Therefore, zeaxanthin in these fish also exists as three optical isomers [38].
**Figure 13.** Metabolism of zeaxanthin in Cyprinidae and astaxanthin in Salmonidae
fish.
Tunaxanthin (**71**) is widely distributed in fish belonging to Perciformes. The bright yellow color in the fins and skin of marine fish is due to the presence of tunaxanthin. Feeding experiments involving red sea bream and yellow tail revealed that tunaxanthin (**71**) was metabolized from astaxanthin (**8**)
via zeaxanthin, as shown in Figure 14 [7,36]. Carotenoids with a 3-oxo-Ή-end group such as
<sup>Ή</sup>,<sup>Ή</sup>-carotene-3,3<sup>ȝ</sup>-dione (**72**) [37] are key intermediates in this metabolic conversion.
> **Figure 14.** Metabolism of astaxanthin in Perciformes fish.
Unique apocarotenoids, micropteroxanthins (**<sup>73</sup>**–**<sup>76</sup>**), were reported from the integuments of the black bass *Micropterus salmoides* [39]*.* They were assumed to be corresponding oxidative cleavage products of tunaxanthin, lutein, and alloxanthin.
Since 2000, there are a few reports on new structures of carotenoids from fish (Figure 15). Carotenoids with a 3,6-dihydroxy-Ή-end group, salmoxanthin (**77**), deepoxysalmoxanthin (**78**)
(from the salmon *Oncorhynchus keta*) [40], and gobiusxanthin (**79**) (from the freshwater goby
*Rhinogobius brunneus*) [41], were isolated. A series of carotenoids with a 7,8-dihydroand/or
7,8,7<sup>ȝ</sup>,8<sup>ȝ</sup>-tetrahydro polyene chain were isolated from the integuments and eggs of the Japanese common catfish *Silurus asotus* [42]. Recently, new carotenoids, 7<sup>ȝ</sup>,8<sup>ȝ</sup>,9<sup>ȝ</sup>,10<sup>ȝ</sup>tetrahydro-Ά-cryptoxanthin (**80**),
<sup>7</sup><sup>ȝ</sup>,8<sup>ȝ</sup>-dihydrodiatoxanthin (**81**), and (3*S*,6*S*,6<sup>ȝ</sup>*S*)-Ή-cryptoxanthin (**82**), were isolated from the integuments and gonads of the Japanese common catfish as minor carotenoids [43].
> **Figure 15.** New carotenoids from fish.
*Mar. Drugs* **2011**, *9*, 278–293
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**9. Mammalia (Mammals)**
There are few reports available on carotenoids in marine mammals. Only, Άcarotene and lutein were reported from the dolphin [44]. The whale is the biggest marine mammal. Whales feed on krill, which is an important dietary source of astaxanthin for marine animals. Therefore, whales might accumulate astaxanthin in the body.
Recently, absorption and metabolism of fucoxanthin (**25**) in mice was investigated. Dietary administrated fucoxanthin was converted to amarouciaxanthin A (**34**) via fucoxanthinol (**26**)
in mice [45,46] (Figure 16). This metabolic conversion was also observed in human hepatoma cell HepG2 and required NAD(P)+ as a cofactor [45].
**Figure 16.** Metabolism of fucoxanthin in mice.
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**10. Role of Carotenoids in Marine Animals and Utilization of Carotenoids for Aquaculture**
Carotenoids are not essential in the nutritional sense. However, they are beneficial for animal health. It is well-known that carotenoids have an unsubstituted Ά-end group, such as Ά-carotene, ΅-carotene, and the Ά-cryptoxanthin precursor of vitamin A in animals. Furthermore, canthaxanthin was also converted to retinol in Salmoidae fish. 3-Hydroxy carotenoids: lutein, zeaxanthin, and astaxanthin, were also reported to be precursors of 3,4-dehydroretinol (Vitamin A2) in some freshwater fish [36,47].
Many marine animals accumulate carotenoids in their integuments. Integumentary carotenoids may contribute to photoprotection, camouflage, and signaling such as breeding color.
Carotenoids have excellent antioxidative activities for quenching singlet oxygen and inhibiting lipid peroxidation. Astaxanthin supplementation in Salmonidae fish suppressed oxidative stress [48,49].
Marine animals also accumulate carotenoids in their gonads. Carotenoids are assumed to be essential for reproduction in marine animals. Astaxanthin supplementation in cultured salmon and red sea bream increased ovary development, fertilization, hatching, and larval growth [50]. In the case of the sea urchin, supplementation with Ά-carotene, which was metabolized to echinenone, also increased reproduction and the survival of larvae [51]. Carotenoids also enhance immune activity in marine animals [52,53].
Carotenoids are used for pigmentation in several aquaculture fish. Synthetic and natural astaxanthin from *Phaffia* yeast and *Haematococcus* algae is widely used for the pigmentation of salmon, trout, and red sea bream. Lutein from marigold is also used as a yellow coloration for cultured marine fish such as yellow tail and red sea bream. Zeaxanthin from spirulina is used as a red coloration for goldfish and ornamental carp.
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**11. Conclusions**
In the present review, I have described marine animal carotenoids from natural product chemistry, metabolism, food chain, and chemosystematic viewpoints and also describe new structural carotenoids isolated from marine animals during the last decade.
In plants and photosynthetic bacteria, biosynthetic roots of carotenoids were identified at the enzymatic and gene level. On the other hand, neither enzymes nor genes for the metabolism of carotenoids in animals have been clarified. Therefore, chemical, biochemical, and analytical approaches are still important to clarify carotenoids in animals.
Interesting new structural carotenoids can still be found in marine animals. The structures of these new carotenoids provide information on the function and food chain, as well as metabolic pathways in marine animals.
## **References and Notes**
Mathews-Roth, M.M., Taylor, R.F., Eds.; Plenum Press: New York, NY, USA, 1989; pp. 59–74.
compound **5** in this literature. However, this IUPAC name is un-correct. Correct IUPAC name for **5** is (3 *<sup>R</sup>*,3<sup>ȝ</sup>*R*,5*S*)-3,3<sup>ȝ</sup>,5,19<sup>ȝ</sup>-tetrahydroxy-7<sup>ȝ</sup>,8<sup>ȝ</sup>-didehydro-5,8- dihydro-Ά,Ά-caroten-8-one.
*J. Chem. Soc., Perkin Trans. 1* **2001**, 3338–3345; doi: 10.1039/B108037G.
1462–1464.
*meso*-zeaxanthin in nature. *Comp. Biochem. Physiol.* **1986**, *83B*, 121–124.
*Prod.* **2001**, *64*, 507–510. 41. Tsushima, M.; Mune, E.; Maoka, T.; Matsuno, T. Isolation of stereoisomeric epoxy carotenoids and new acetylenic carotenoid from the common freshwater
goby *Rhinogobius brunneus*.
*J. Nat. Prod.* **2000**, *63*, 960–964.
Ά,Ά-carotene, Ά-echinenone, astaxanthin, fucoxanthin, vitamin A and vitamin E on the biological defense of the sea urchin *Pseudocentrotus depressus*. *J. Exp. Mar. Biol. Ecol.* **1998**, *226*, 165–174.
© 2011 by the authors. Submitted for possible open access publication under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).
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**Carotenoids in Marine Invertebrates Living along the**
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**Kuroshio Current Coast**
**Takashi Maoka 1, Naoshige Akimoto 2, Miyuki Tsushima 3, Sadao Komemushi 4, Takuma Mezaki 5, Fumihito Iwase 5, Yoshimitsu Takahashi 6, Naomi Sameshima 6, Miho Mori 6, and Yoshikazu Sakagami 6,\***
- Kyoto 606-0805, Japan; E-Mail: [email protected]
*Sakyo-ku, Kyoto 606-8501, Japan; E-Mail: [email protected]*
3-3-138 Sugimoto, Sumiyoshi-ku, Osaka 558-8585, Japan; E-Mail: [email protected]
5 Kuroshio Biological Research Foundation, Nishidomari-560, Ootsuki-cho, Kochi 788-0333, Japan;
```
E-Mails: [email protected] (T.M.); [email protected] (F.I.)
```
*6 Faculty of Agriculture, Kinki University, Nakamachi 3327-204, Nara-shi 631-8505, Nara, Japan;*
*E-Mails: [email protected] (Y.T.); [email protected] (N.S.); [email protected] (M.M.)*
\* Author to whom correspondence should be addressed; E-Mail: [email protected]; Tel.: +81-742-43-7154; Fax: +81-742-43-1593.
*Received: 30 June 2011; in revised form: 31 July 2011 / Accepted: 8 August 2011 / Published: 22 August 2011*
**Abstract:** Carotenoids of the corals *Acropora japonica*, *A. secale*, and *A. hyacinthus*, the tridacnid clam *Tridacna squamosa*, the crown-of-thorns starfish *Acanthaster planci*, and the small sea snail *Drupella fragum* were investigated. The corals and the tridacnid clam are filter feeders and are associated with symbiotic zooxanthellae. Peridinin and pyrrhoxanthin, which originated from symbiotic zooxanthellae, were found to be major carotenoids in corals and the tridacnid clam. The crown-of-thorns starfish and the sea snail *D. fragum* are carnivorous and mainly feed on corals. Peridinin-3-acyl esters were major carotenoids in the sea snail *D. fragum*. On the other hand, ketocarotenoids such as 7,8-didehydroastaxanthin and astaxanthin were major carotenoids in the crown-of-thorns starfish. Carotenoids found in these marine animals closely reflected not only their metabolism but also their food chains.
**Keywords:** carotenoid; marine invertebrates; food chain; metabolism
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**1. Introduction**
Marine animals, especially marine invertebrates, contain various carotenoids, with structural diversity [1–4]. Interesting structural carotenoids are still being found in marine animals [4]. In general, animals do not synthesize carotenoids *de novo*, and so those found in animals are either directly accumulated from food or partly modified through metabolic reactions [2]. The major metabolic conversions of carotenoids found in marine animals are oxidation, reduction, transformation of double bonds, oxidative cleavage of double bonds, and cleavage of epoxy bonds [2,3]. Therefore, various structural varieties are found in carotenoids of marine animals [4].
We have studied carotenoids in several marine invertebrates from chemical and comparative biochemical points of view [4]. In the present study, we focused on carotenoids of the corals *Acropora japonica*, *A. secale*, and *A. hyacinthus*, the tridacnid clam (elongate giant clam) *Tridacna squamosa*, crown-of-thorns starfish *Acanthaster planci*, and small sea snail *Drupella fragum*, inhabiting the Kuroshio current coast. These animals are closely associated within the food chain. Corals and the tridacnid clam are filter feeders and are associated with symbiotic zooxanthellae (dinoflagellate algae). On the other hand, the crown-of-thorns starfish and small sea snail *D. fragum* are carnivorous and mainly prey upon corals. Therefore, carotenoids that originated from zooxanthellae are passed to starfish and small sea snails through this food chain. In the present paper, we describe the carotenoids of these marine invertebrates.
## **2. Results and Discussion**
Structural formulae of carotenoids identified from *Acropora* corals, the tridacnid clam *T. squamosa*, starfish *A. planci*, and sea snail *D. fragum* are shown in Figure 1.
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*2.1. Carotenoids of Corals and the Tridacnid Clam*
The carotenoids composition of the corals and the tridacnid clam were similar to each other (Table 1). Ά,Ά-Carotene, peridinin (including the 9<sup>ȝ</sup>*Z* isomer), pyrrhoxanthin, diatoxanthin, and diadinoxanthin were found in these animals. These carotenoid patterns resembled those of symbiotic zooxanthellae [5,6]. The results indicate that corals and the tridacnid clam directly absorb carotenoids from symbiotic zooxanthellae and accumulate them without metabolic modification. In the eggs of corals, peridinin and pyrroxanthin were present as major carotenoids. It was assumed that peridinin and pyrroxanthin play important roles in reproduction in corals, as with astaxanthin in salmonid fishes [7].
Recently, Daigo *et al.* studied carotenoids of more than 20 species of coral inhabiting reefs in Okinawa [8]. They reported that carotenoids found in these corals were not only peridinin and diadinoxanthin, that originated from symbiotic zooxanthellae, but also zeaxanthin, lutein, and, fucoxanthin, that originated from cyanobacteria, green algae, and diatoms. Cyanobacteria, green algae, and diatoms were epizoic and/or endolithic algae that grew in association with the corals. Corals accumulated carotenoids from these epizoic and/or endolithic algae [8]. However, the present study found that carotenoids in *Acropora* corals, inhabiting the Kuroshio current coast of Kochi, only consisted of those that originated from zooxanthellae. These differences might reflect the constitution of associating algae with corals.
Peridinin and pyrrhoxanthin were found to be major carotenoids in the tridacnid clam. In general, major carotenoids found in clams are fucoxanthin and its metabolites originating from diatoms [9–11]. On the other hand, neither fucoxanthin nor its metabolites were found in the tridacnid clam. This indicates that the tridacnid clam only ingested carotenoids from dinoflagellate algae. Similar results were reported in carotenoids of the bivalves, *Modiolus modiolus* and *Pecten maximus* [12].
**Figure 1.** Carotenoids identified from *Acropora* corals, the tridacnid clam *T. squamosa*, starfish *A. planci*, and sea snail *D. fragum*.
**Table 1.** Carotenoids of *Acropora* corals and the tridacnid clam *Tridacna*
*squamosa*.
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{
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 209
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*2.2. Carotenoids of the Crown-of-Thorns Starfish*
The crown-of-thorns starfish, *A. planci*, is a large, nocturnal sea star that mainly preys upon coral polyps. Like other starfish [13], 7,8-didehydroastaxanthin and astaxanthin were found to be major carotenoids along with pectenolone, 7,8,7<sup>ȝ</sup>,8<sup>ȝ</sup>tetradehydroastaxanthin, diatoxanthin, and alloxanthin (Table 2). In general, the starfish can introduce a hydroxy group at C-3 and carbonyl group at C-4 in the Άend group of carotenoids [6]. So, 7,8-didehydroastaxanthin and astaxanthin were oxidative metabolites of diatoxanthin and Ά-carotene, respectively, ingested from dietary corals. Echinenone and canthaxanthin were metabolic intermediates from Ά,Ά-carotene to astaxanthin. The acetylenic carotenoids, pectenolone, pectenol A, and pectenol B, were also metabolic intermediates from diatoxanthin to 7,8- didehydroastaxanthin. Peridinol, one of the major carotenoids in the
crown-of-thorns starfish, was converted from peridinin, which originated from corals, by hydrolysis. Furthermore, four new carotenoids; 4- ketodeepoxyneoxanthin, 4-keto-4ȝ-hydroxydiatoxanthin,
3<sup>ȝ</sup>-epigobiusxanthin, and 7,8-dihydrodiadinoxanthin, were isolated [14]. Details of the structural elucidation of those compounds were described previously [14]. In the present paper, the biosynthetic origins of these compounds are discussed (Figure 2). 4-Keto-4ȝ-hydroxydiatoxanthin was one of the metabolic intermediates from diatoxanthin to 7,8-didehydroastaxanthin. 4-Ketodeepoxyneoxanthin might be an oxidative metabolite of deepoxyneoxanthin derived from neoxanthin by deepoxydation.
3ȝ-Epigobiusxanthin might be derived from diadinoxanthin. 7,8- Dihydrodiadinoxanthin, which has a unique single bond in the 7,8-saturated polyene chain, may be a reduction metabolite of diadinoxanthin. Therefore, it was concluded that carotenoids ingested from corals were oxidatively metabolized and accumulated in the crown-of-thorns starfish.
**Table 2.** Carotenoids of the crown-of-thorns starfish *Acanthaster planci.*
**Figure 2.** Possible bioformation roots of new carotenoids in crown-ofthorns starfish.
## *2.3. Carotenoids of the Sea Snail* D. fragum
Like the crown-of-thorns starfish, the small sea snail *D. fragum* also feeds on corals. The carotenoid composition of this snail resembled that of the dietary corals (Table 3). This indicated that *D. fragum* also accumulated carotenoids from dietary corals without metabolic modification, except for the esterification of peridinin. In the present study, peridinin 3-acyl esters were fully characterized based on 1H-NMR and FAB MS spectral data. The 1H-NMR signal of H-3 (Έ 4.95), which showed 1.04 ppm downfield shift relative to the corresponding signal in peridinin [15,16], indicated that the hydroxy group at C-3 was acylated. Fatty acids esterified with peridinin were assigned as palmitic acid, palmitoleic acid, and myristic acid based on FAB-MS data. Previously, peridinol fatty acid ester was characterized by Moaka *et al*. [10] and Sugawara *et al*. [17]. However, peridinin 3-acyl esters have not yet been reported. The origin of zeaxanthin in this snail was unclear. It might have originated from associated algae such as cyanobacteria [8].
**Table 3.** Carotenoids of the sea snail *Drupella fragum*.
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*3.1. General*
The UV-Visible (UV-VIS) spectra were recorded with a Hitachi U-2001 in diethyl ether (Et2O).
The positive ion FAB-MS spectra were recorded using a JEOL JMS-700 110A mass spectrometer with
*m*-nitrobenzyl alcohol as a matrix. The 1H-NMR (500 MHz) spectra were measured with a Varian UNITY INOVA 500 spectrometer in CDCl3 with TMS as an internal standard. HPLC was performed on a Shimadzu LC-6AD with a Shimadzu SPD-6AV spectrophotometer set at 470 nm. The column used was a 250 × 10 mm i.d., 10 ΐm Cosmosil 5C18-II (Nacalai Tesque, Kyoto, Japan) with acetone:hexane (3:7, v/v) at a flow rate of 1.0 mL/min. The optical purity of astaxanthin was analyzed by chiral HPLC using a 300 × 8 mm i.d., 5 ΐm Sumichiral OA-2000 (Sumitomo Chemicals, Osaka, Japan) with *n*-hexane/CHCl3-ethanol (48:16:0.8, v/v) at a flow rate of 1.0 mL/min [18].
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 212
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*3.2. Animal Material*
The corals *A. japonica*, *A. secale*, and *A. hyacinthus*, the tridacnid clam *T. squamosa*, the crown-of-thorns starfish *A. planc*, and the sea snail *D. fragum* were collected along the Ootsuki coast, Kochi Prefecture, Japan from July to August 2009 and 2010.
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"author": "",
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"isbn": "9783039431908",
"section_idx": 213
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*3.3. Analysis of Carotenoids*
The extraction and identification of carotenoids were carried out according to our routine methods [19]. Carotenoids were extracted from living or fresh animal specimens with acetone. The acetone extract was transferred to ether-hexane (1:1) layer after the addition of water. The total carotenoid contents were calculated
employing an extinction coefficient of E1%cm = 2100 (astaxanthin) [20] for the starfish *A.*
*<sup>p</sup>lanci* and E1%cm = 1350 (peridinin) [20] for *A. japonica*, *T. squamosa*, and *D. fragum* at <sup>Ώ</sup>
max. The ether-hexane solution was evaporated. The residue was subjected to HPLC on silica gel. Carotenoid compositions were estimated by the peak area of the HPLC on silica gel with
acetone-hexane (3:7) monitored at 470 nm.
Individual carotenoids were identified by UV-VIS (ether), FAB MS, and partial 1H NMR (500 MHz, CDCl3).
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "ffe4a610-aead-4300-8fef-a70aeecd3fb7",
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"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 214
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*3.4. Identification of Carotenoids*
Identification of individual carotenoids were carried out on UV-VIS and FAB MS spectral data and compared with chromatographic property with authentic samples [19]. Optical isomer of astaxanthin in the crown-of-thorns starfish *Acanthaster planci* was analyzed by Chiral HPLC [18]. Astaxanthin fraction in *Acanthaster planci* was consisted of three optical isomers (3*<sup>R</sup>*,3<sup>ȝ</sup>*R*):(3*R*,3<sup>ȝ</sup>*S*):(3*S*,3<sup>ȝ</sup>*S*) with a ratio of 32:14:54. Furthermore, peridiniol, peridinin and 9ȝ*Z*-Peridinin were characterized by 1H NMR [15,16]. Structures of 7,8-ihydrodiadinoxanthin, 3<sup>ȝ</sup>-epigobiusxanthin, 4-keto-4<sup>ȝ</sup>hydroxydiatoxanthin, 4-ketodeepoxyneoxanthin, and deepoxyneoxanthin were fully characterized by NMR [14].
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{
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
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"author": "",
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"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 215
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*3.5. Caracterization of Peridinin-3-acyl Esters*
Peridinin-3-acyl esters. FAB-MS: *m/z* 868.5860 [M]+ (calcd. for C55H80O8, 868.5856) peridinin 3-palmitate, *m/z* 866.5698 [M]+ (calcd. for C55H79O8, 866.5703) peridinin 3- palmitolate, *m/z* 840.5550 [M]+ (calcd. for C53H76O8, 840.5547) peridinin 3-myristate; UV-VIS 455, 475 nm; 1H NMR (CDCl3), Έ 0.88 (3H, t, *J* = 7.5 Hz, CH3 in fatty acid moiety), 0.99 (3H, s, H-16), 1.07 (3H, s, H-17ȝ), 1.20 (3H, s H-17), 1.21 (3H, s H-18), 1.25 (about 24H, s, -CH2- in fatty acid moiety), 1.35 (3H, s, H-18ȝ), 1.39 (3H, s, H-16ȝ), 1.41 (1H, dd, *J* = 13, 7 Hz,H-2ȝΆ), 1.51 (1H, dd, *J* = 13, 12.5 Hz, H-4ȝΆ), 1.64 (1H, dd, *J* = 12.5, 12 Hz, H-2΅ eq), 1.79 (1H, dd, *J* = 12, 7 Hz, H-4Ά ax), 1.81 (3H, s, H-19<sup>ȝ</sup>), 2.00 (H, ddd, *J* = 13, 4, 2 Hz, H-2ȝ΅), 2.04 (3H, s, CH3CO-), 2.23 (3H, s, H-20), 2.29 (1H, overlapped, H-΅), 2.28 (2H, t, *J* = 7.5 Hz, -CH2-COOH in fatty acid moiety), 2.41 (1H, ddd, *J* = 14, 5, 1.5 Hz, H-4΅), 4.95 (1H, m, H-3), 5.37 (1H, m, H-3<sup>ȝ</sup>), 5.74 (1H, s, H-12), 6.06 (1H, s, H-8ȝ), 6.11 (1H, d, *J* = 11 Hz, H-10ȝ), 6.38 (1H, dg, *J* = 14, 11 Hz, H-11ȝ), 6.40 (1H, d, *J* = 16 Hz, H-8), 6.46 (1H, d, *J* = 11 Hz, H-14<sup>ȝ</sup>), 6.51 (1H, dd, *J* = 14, 11 Hz, H-15<sup>ȝ</sup>), 6.61 (2H, dd, *J* = 14, 11 Hz, H-11<sup>ȝ</sup>, 15<sup>ȝ</sup>).
## **4. Conclusions**
In conclusion, carotenoids found in the coral *A. japonica*, clam *T. squamosa*, starfish *A. planci*, and sea snail *D. fragum* well reflected not only their metabolism but also the food chain. The accumulation and metabolism of carotenoids that originate from zooxanthellae to the starfish through the food chain are summarized in Figure 3.
**Figure 3.** Accumulation and metabolism of carotenoids that originate from zooxanthellae to the starfish and sea snail through the food chain.
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**Acknowledgements**
We wish to thank Ryo Harada, a former student of Kinki University (now working in Torii Pharmaceutical Co., Ltd.) for his technical support.
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{
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"isbn": "9783039431908",
"section_idx": 217
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**Carotenoid Ά-Ring Hydroxylase and Ketolase from Marine Bacteria—Promiscuous Enzymes for Synthesizing**
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{
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"url": "https://mdpi.com/books/pdfview/book/3341",
"author": "",
"title": "Marine Carotenoids",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783039431908",
"section_idx": 219
}
|
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