Next Article in Journal
Diversity of Bacterial Secondary Metabolite Biosynthetic Gene Clusters in Three Vietnamese Sponges
Next Article in Special Issue
Characterization and Cytotoxic Activity of Microwave-Assisted Extracted Crude Fucoidans from Different Brown Seaweeds
Previous Article in Journal
Two New Alkaloids and a New Butenolide Derivative from the Beibu Gulf Sponge-Derived Fungus Penicillium sp. SCSIO 41413
Previous Article in Special Issue
Gold Nanoparticles Synthesized by an Aqueous Extract of Codium tomentosum as Potential Antitumoral Enhancers of Gemcitabine
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Marine Drugs
Volume 21
Issue 1
10.3390/md21010028
marinedrugs-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

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

A Review on a Hidden Gem: Phycoerythrin from Blue-Green Algae

by
Hui Teng Tan
1,
Fatimah Md. Yusoff
2,3,*,
Yam Sim Khaw
1,
Nur Amirah Izyan Noor Mazli
1,
Muhammad Farhan Nazarudin
1,
Noor Azmi Shaharuddin
4,
Tomoyo Katayama
5 and
Siti Aqlima Ahmad
4,*
1
Laboratory of Aquatic Animal Health and Therapeutics, Institute of Bioscience, Universiti Putra Malaysia, Serdang 43400, Selangor, Malaysia
2
Department of Aquaculture, Faculty of Agriculture, Universiti Putra Malaysia, Serdang 43400, Selangor, Malaysia
3
International Institute of Aquaculture and Aquatic Sciences, Universiti Putra Malaysia, Port Dickson 71050, Negeri Sembilan, Malaysia
4
Department of Biochemistry, Faculty of Biotechnology and Biomolecular Sciences, Universiti Putra Malaysia, Serdang 43400, Selangor, Malaysia
5
Graduate School of Agricultural and Life Sciences, The University of Tokyo, 1-1-1, Yayoi, Bunkyo, Tokyo 113-8657, Japan
*
Authors to whom correspondence should be addressed.
Mar. Drugs 2023, 21(1), 28; https://doi.org/10.3390/md21010028
Submission received: 10 November 2022 / Revised: 21 December 2022 / Accepted: 24 December 2022 / Published: 29 December 2022
(This article belongs to the Special Issue Researches in High-Value Algal Products: From Extraction to Application)
Browse Figures
Versions Notes

Abstract

:
Phycoerythrin (PE) is a pink/red-colored pigment found in rhodophytes, cryptophytes, and blue-green algae (cyanobacteria). The interest in PE is emerging from its role in delivering health benefits. Unfortunately, the current cyanobacterial-PE (C-PE) knowledge is still in the infant stage. It is essential to acquire a more comprehensive understanding of C-PE. This study aimed to review the C-PE structure, up and downstream processes of C-PE, application of C-PE, and strategies to enhance its stability and market value. In addition, this study also presented a strengths, weaknesses, opportunities, and threats (SWOT) analysis on C-PE. Cyanobacteria appeared to be the more promising PE producers compared to rhodophytes, cryptophytes, and macroalgae. Green/blue light is preferred to accumulate higher PE content in cyanobacteria. Currently, the prominent C-PE extraction method is repeated freezing–thawing. A combination of precipitation and chromatography approaches is proposed to obtain greater purity of C-PE. C-PE has been widely exploited in various fields, such as nutraceuticals, pharmaceuticals, therapeutics, cosmetics, biotechnology, food, and feed, owing to its bioactivities and fluorescent properties. This review provides insight into the state-of-art nature of C-PE and advances a step further in commercializing this prospective pigment.

1. Introduction

Phycoerythrin (PE) is a colored, water-soluble pigment–protein complex from the phycobiliprotein (PBP) family and is predominantly found in rhodophytes, cryptophytes, and cyanobacteria [1,2]. It is a 240 kDa oligomeric chromoprotein with an intense pink/red color [3]. This PBP is a light-harvesting accessory pigment that absorbs wavelengths that chlorophyll cannot [4]. In terms of energy transfer, PE is the first PBP to absorb light and emits energy to phycocyanin (PC) and allophycocyanin (APC), consecutively [5]. PE is characterized by its original spectrum properties, such as fluorescence emission [6]. On the other hand, previous PBP research was focused on PC, and it was only recently that the scientific community realized that PE is endowed with antioxidant potential due to its ability to scavenge free radicals [7,8,9]. This allows them to prevent or delay the onset of potentially fatal diseases, such as cancer, heart disease, and hepatotoxicity. The potent antioxidant activity promotes human health and boosts the immune response against infections [10]. These bioactive compounds also resist photooxidative damage, which would otherwise have an adverse impact on crucial cellular machineries, such as proteins, lipids, and DNA [11,12]. In comparison to PC and APC, the unique fluorescent properties of PE have made it a useful tool in the diagnosis and treatment of tumors [13]. PE has gained prominence not only as pharmaceuticals, nutraceuticals, cosmetics, and colorants but also as biologically active compounds, which will be described in this review.
PE is generally classified into three types based on different spectral properties and the photosynthetic organisms that produce them—namely, B-phycoerythrin (B-PE), C-phycoerythrin (C-PE), and R-phycoerythrin (R-PE) from Bangiophyceae (primitive filamentous Rhodophyta), cyanobacteria, and complex Rhodophyta, respectively [14,15]. B-PE and R-PE from red algae have been well-studied and widespread in their use across various industries [13,16]. B-PE became a valuable candidate as an analytical reagent and in fluorescent technologies, owing to its large absorption coefficient and high fluorescence efficiency (high quantum yield and high stokes shift) [14]. In contrast, C-PE derived from cyanobacteria received relatively little attention due to public concern about cyanotoxins.
In recent years, previous reviews have reported on the scientific progress and challenges of microalgal pigments [17,18,19]. Instead of a detailed report on C-PE, the existing publications focused on the general characteristics and features of cyanobacterial PBP, particularly C-PC, while overlooking the specific strains, characteristics, and upstream and downstream processes specifically optimized for C-PE as well as the specific applications of C-PE [20,21]. Spirulina platensis is the most commonly used strain since it has been recognized as an effective multiple-product-producing strain for decades [20]. Nevertheless, there is a growing need to explore the cyanobacterial world in order to discover new potential strains for C-PE production. Furthermore, there is a dearth of exclusive reports on C-PE, maximum C-PE valorization, and the possibility of bolstering novel strains for C-PE production. Hence, this review discusses C-PE and its various benefits and applications. It also presents an abridged overview of a decade-long journey by highlighting research areas from scientific literature published on C-PE production. This review also featured a SWOT (strengths, weaknesses, opportunities, and threats) analysis to determine the internal and external factors and obstacles associated with C-PE commercialization.

The Odyssey of Phycoerythrin Research

The necessity of developing naturally derived bioactive compounds capable of preventing diseases motivated researchers to explore various algae or plant sources in pursuit of novel and sustainable pigment supplies. This sector experienced massive development in the 1970s when public awareness shifted toward health safety and a predilection for naturally derived pigments over chemically synthesized pigments [22]. This has resulted in a growing interest in studying cyanobacteria as a potential source due to the possibilities of commercial applications in several fields. More specifically, PBP extracted from cyanobacteria has been studied and reported to have anticancer, antioxidant, anti-inflammatory, and antimicrobial properties, making it a priority for industrial production in various lucrative sectors, such as pharmaceuticals, foods and feeds, cosmetics, and nutraceuticals [16]. Aside from the well-known PC, PE, with its excellent fluorescent features, has progressively become a research focus. The advent of PE as a fluorescent marker for macromolecules and cells was followed by its widespread use in high-sensitivity fluorescence technologies [14]. Most PE sources are limited to red algae, especially Porphyridium sp., which have long been studied and commercially exploited for R-PE [23,24]. However, cyanobacteria can become an alternative source. C-PE may enter the mainstream due to their fast and easy cultivation, complementary chromatic adaptation (CCA) ability, and greater resistance to bacterium contamination [3,25].

2. Structure of C-PE

Each PE comprises two different polypeptides (denoted as α and β), with each subunit containing a protein backbone and a phycobilin bound by a covalent bond (Figure 1) [16]. The cpeA gene encodes the α subunit of PE, while the cpeB gene encodes the β subunit [26]. The three types of PE (C-PE, R-PE, and B-PE) differ in the number and type of chromophores attached, resulting in different absorption maxima. Unlike R-PE and B-PE, C-PE has only one absorption maximum, at about 564 nm (Figure 2) [27]. C-PE is produced mainly by cyanobacteria and is further subdivided into C-PE-I and C-PE-II [28]. There are four forms of phycobilins in cyanobacteria, each with a different color: phycocyanobilin (PCB) in blue, phycourobilin (PUB) in yellow, phycoerythroblin (PEB) in red, and phycoviolobilin (PVB) in violet [29]. These phycobilins have been reported to be one of the major components responsible for the bioactivity of PE [16]. Each form of phycobilin has a system of conjugated double bonds that give it a unique spectral characteristic that can be further modified by interactions between the subunits and the chromophore [30]. Marine and freshwater or soil cyanobacteria have different phycobilin contents. PE from freshwater and soil cyanobacteria carry PEB chromophores solely, with absorption and fluorescence emission spectra resembling the spectral properties of C-PE-I [31]. C-PE-I carried five PEBs (αβ) that were covalently bonded to the cysteine (Cys) residue (α-84, α-140 or β-143, β-84 or β-82, β-50 or β-61, and β-155 or β-159) [21]. C-PE-I is a disk-shaped hexamer formed by two trimeric (αβ)3 through face-to-face aggregation. It is typically found in the form of (αβ)6 LR, in which a rod-linker polypeptide (LR) is bound to the central hole of the two stacking trimers. Unlike the B-PE and R-PE subunits, the LR of C-PE-I does not carry phycobilins [31]. C-PE-II, found in marine cyanobacteria, such as Synechocystis sp. and Synechococcus sp., has five PEB chromophores in the same position as C-PE-I and a PUB molecule attached to α-75 cysteine [21]. Phycoerythrocyanin, an alternate form of PE, may be found in some cyanobacteria [16]. It exists in trimeric (αβ)3 or hexameric (αβ)6 forms, each containing a PVB chromophore at the α-84 residue and two PCB linked to the β-84 and β-153 residues [16,21]. When some cyanobacteria acclimate to light intensity and quality variations, the chromophore composition of C-PE can be modified through the CCA process [32]. PE has a quantum yield of up to 0.98 and a molar coefficient of up to 2.4 × 106. Since it has superior spectral characteristics, PE outperforms the commonly used fluorescent dyes (fluorescein and rhodamine) and is used as a fluorescent probe [13].

3. C-PE Producer Species

The current commercial PE is derived from red algae, either microalgae or macroalgae. Such production is governed by their species and metabolism. In different species, the R-PE content might range from 0.14 mg g−1 to 54.3 mg g−1 [34,35]. Palmaria palmata has the highest content among the studies [34]. R-PE production is, therefore, considered insufficient to satisfy global market demand. Aside from that, red algae such as Porphyridium, Ceramium, and Porphyra, contained a large amount of polysaccharide, which formed a gel (carbohydrate), making the purification of PE procedures tedious and laborious [36]. The high cost of cultivation and production spurs researchers to improve the production process or look for a new source of PE. Cyanobacteria are a great source of PBP because they are fast-growing microorganisms with high PBP contents, particularly in PC content. Unlike most algae, cyanobacteria could thrive in open ponds under adverse conditions [13]. The unique features of CCA and nitrogen fixation ability have considerably reduced cultivation costs and made cyanobacterial growth easier [3,16]. To date, numerous cyanobacteria species, including Tolypothrix sp. (660 mg g−1), Nostoc sp. (125.11 mg g−1), Anabaena sp. (102 mg g−1), and Pseudanabaena sp. (92.57 mg g−1) have been reported to have higher C-PE content than red algae (Figure 3) [37,38,39,40]. These species have been extensively studied for C-PE synthesis and optimized for improved productivity in the literature.
On the other hand, the biosynthesis of PE in a heterologous host is an efficient method for producing recombinant PE. The co-expression of multiple genes for chromophore and apoprotein synthesis, as well as chromophore attachment, is essential for the production of recombinant PBP [41]. Recombinant PBP has been explored since 1980 and is mostly expressed in Escherichia coli. However, there are still some limitations (i.e., the inefficiency of chromophorylation) in recombinant PBP that need to be resolved before it can be utilized commercially. Chen et al. (2022) have summarized the characteristics and research literature on recombinant PBP. The vast advancements in genetic manipulation technologies, such as the CRISPR (clustered regularly interspaced short palindromic repeats), make the development of a stable PBP pathway in the host more feasible and would enhance PBP production [13].

4. Cultivation Methods

Cyanobacteria can be cultivated in open systems (tanks/ponds or raceways) or closed systems (photobioreactors). The high risk of contamination is a critical issue that impedes the mass production of C-PE in open systems [3]. Enclosed photobioreactors are recommended as a preferable choice for pure, large-scale C-PE production since they allow for consistent light provision, temperature control, and carbon dioxide distribution as well as reduced contamination risk [41].

4.1. Cultivation Mode

Cyanobacteria cultivation can be carried out in a variety of modes, including heterotrophic, photoautotrophic, and mixotrophic settings. Heterotrophic mode is usually performed in dark or harsh environments where cyanobacteria need to acclimate to the environment by consuming available organic compounds (i.e., acetate, glucose) as a carbon source [42]. Although the organic compound promoted the growth of cyanobacteria, it also increased the potential for contamination, resulting in health risks within food industries [43]. Photoautotrophic cyanobacteria use natural sunlight as an energy source and atmospheric air as a carbon dioxide supply. As a result, when the light intensity is too high (photo-inhibition) or too low (photo-limitation), the biomass is affected and reduced [3]. Mixotrophic cultivation combines heterotrophic and photoautotrophic modes; both are performed in the presence of sunlight and carbon sources. Specifically, it is a dual-limiting system in which low light intensities or low organic carbon substrate concentrations can limit cell growth. At the same time, high light intensities or high carbon substrate concentrations can also suppress cell growth [43]. Previous research has shown that cyanobacteria cultivated in a mixotrophic environment are more resistant to light stress than photoautotrophs [44]. However, not all cyanobacteria can grow in mixotrophic or heterotrophic conditions [3].

4.2. Light

Light condition is the major factor that affects C-PE accumulation. This condition involves not just light intensity but also light color and photoperiod. It has generally been documented that there is a preference for low to medium light intensity for C-PE production [16]. This is related to the role of PE in increasing the range of light absorption in the photosynthetic system. Thus, in low light-intensity conditions, the cyanobacteria will stimulate PE production to capture more light and energy for growth.
In terms of light color, previous research has found that cyanobacteria cultivated under green light accumulate more C-PE compared to other light spectra (Table 1) [38,45,46]. Under green light, the PSII overexcites and induces a transition to State II, resulting in more excitation energy transfer from the PSII to the PSI. This transfer enables PE to get the green light and use it to activate the photochemistry of both photosystems (PSI and PSII) [47]. Hence, the light color changed the composition of phycobilisomes via CCA. Hsieh Lo et al. (2019) have discussed the mechanism of CCA, but further study is needed to explore it in detail [3]. Further, Mishra et al. (2012) postulated that C-PE production could be stimulated under blue light since the light absorption PE ranges from 450 to 570 nm, allowing it to capture more blue light (460–475 mm) [46,48]. On the other hand, photoperiod affects the balance of heterotrophic and photoautotrophic metabolisms. Although scientific literature has shown that a dark period is always required for phycobiliprotein production, the optimal photoperiod for C-PE accumulation has yet to be determined [49,50]. Literature has shown that a photoperiod of 16 hours of light and 8 hours of darkness was the ideal condition for PBP synthesis [16,50]. Nevertheless, most research on C-PE yield used a photoperiod of 12 hours of light and 12 hours of darkness in their study, despite the fact that this parameter has not been optimized for C-PE yet [4,38,40,46,51].

4.3. Temperature, pH, and Salinity

Temperature is another major factor influencing cyanobacterial growth and C-PE productivity since it directly impacts cell membrane fluidity, nutrient availability, and absorption [3,49]. The ideal temperature for target pigment synthesis varies with different cyanobacterial species or strains, depending on their adaptability and tolerance [3]. Extremely high temperatures denature enzymes or proteins, inhibiting growth and decreasing C-PE production, whereas extremely low temperatures will cause certain cyanobacterial metabolism to malfunction [54]. Previous studies reported that temperature in the range of 30–36 °C was ideal for the growth of most cyanobacteria [49]. For example, the optimal temperature of Anabaena sp. was reported at 30 °C (Table 1) [38]. Studies to optimize temperature, focusing on C-PE production, still received little attention.
pH fluctuations affect nutrient solubility and bioavailability, the transportation of molecules across cytoplasmic membranes, enzyme activity, photosynthetic electron transport, and cell physiology [39,53]. Most cyanobacteria have been found to prefer an alkaline environment for growth and phycobiliprotein production. Extreme pH levels will change the charge on protein, eventually leading to protein denaturation [39]. The ideal pH for cyanobacteria growth is strain-dependent, ranging from 6.0 to 10.0 and never below a pH of 3.0 or exceeding a pH of 12.0 [16]. However, the study of the optimal pH for high C-PE production is still limited to a few strains (Table 1). The neutral pH of 7.0 is reported to be ideal for high C-PE accumulation in Nostoc sp. and Spirulina platensis, while a pH of 8 is suitable for Anabaena sp. and Nodularia sphaerocarpa [39,48,52,53].
Salt stress decreases plant growth and metabolite production, which are usually related to lower photosynthesis [55]. The optimal salinity depends on the strain. Halophilic cyanobacteria can thrive in hypersaline environments, but high salinity can inhibit the photosystem and electron transport chain in non-tolerant strains [53,56]. Furthermore, excessive sodium ions can cause phycobilisome detachment from the thylakoid membrane, reducing photosynthetic activity and nutrient absorption [49,57]. Pagels et al. (2019) compiled a few pieces of research that looked into the effect of salinity on the growth and phycocyanin production of marine cyanobacteria [16]. Yet, there is a lack of research on the effect of salinity on C-PE production (Table 1). Sharma et al. (2014) optimized the salinity level suitable for Spirulina platensis and found that a 0.4 M salt concentration is optimal for increasing the C-PE content [53].

4.4. Nutrients (Nitrogen, Carbon, and Phosphorus)

Nutrient uptake is crucial for cyanobacterial growth and pigment production. Nitrogen is one of the important macronutrients since it is required to synthesize nucleic acids and proteins [58]. Furthermore, nitrogen, which serves as the primary nitrogen store within the cell, is essential for cell viability and can regulate the production of PBP [59]. The nitrogen supply in the growth medium influences metabolite production, as it alters the nutrient assimilation mechanism within the cell [16]. Nitrogen deprivation can impair cellular development rates and contribute to a phenomenon known as chlorosis, which provokes PBP degradation and leads to photosynthesis downregulation [59,60]. The type of nitrogen source and the amounts required are species-dependent [16]. Cyanobacteria have been demonstrated in studies to be capable of assimilating a wide range of nitrogen sources, including nitrate (NO3−), ammonium (NH4+), nitrite (NO2−), and urea (Table 2). A few research studies have compared various nitrogen sources on C-PE synthesis. Most cyanobacteria (including Arthrospira platensis, Fischerella sp., Nodularia sphaerocarpa, and Spirulina maxima) prefer nitrate concentrations ranging from 0.2 to 2.5 g L−1 [52,61,62,63]. Anabaena fertilissima supplemented with nitrite reported the best C-PE production, whereas an ammonium source was reflected in Phormidium sp. and Pseudoscillatoria sp. [48,61]. The addition of an inappropriate nitrogen source (especially ammonium) will result in a significant influx of ammonium ions and a change in pH, which will be toxic to the cell and reduce C-PE production [16,61]. Liotenberg et al. (1996), on the other hand, discovered that ammonium and nitrate sources may alter PBP proportion differentially depending on the cyanobacterial strain. The study revealed that the growth of Calothrix sp. in the presence of ammonium, as compared to nitrate, increased the PC content by 46% but resulted in 35% lower intracellular levels of PE [64]. Given that a PE monomer contains more tetrapyrrole chromophores than the other two phycobilin monomers, it may be assumed that fewer chromophores are required to construct phycobilisomes in ammonium-grown cells than in nitrate-grown cells [64,65]. As phycobilisomes constitute a main cellular investment, with phycobiliproteins accounting for up to 50% of the cell’s soluble protein, a lower nitrogen level, per tetrapyrrole, might be more suitable in nutrient-limited conditions [66]. This statement was consistent with the findings of Hemlata (2009) and Simeunovie et al. (2013) that Anabaena sp. and Nostoc sp. produced higher C-PE when no additional nitrogen source was supplied [49,67]. Further, the nitrogen fixation capabilities of certain cyanobacteria (i.e., Nostoc sp., Oscillatoria sp., and Trichodesmium sp.) can be driven when the inorganic nitrogen source is limited and inhibited when the inorganic nitrogen source is abundant [53,68]. Non-nitrogen-fixing cyanobacteria are normally limited by nitrogen, but nitrogen-fixing cyanobacteria may use ubiquitous atmospheric nitrogen, giving them a competitive advantage in nitrogen-limiting conditions [69]. Yet, although some cyanobacteria can fix atmospheric nitrogen, it is typically preferable to supply the medium with nitrogen [16].
Phosphorus has major structural roles in nucleic acids and is functional in various metabolic processes [59]. A phosphorus deficiency will affect respiration, photosynthesis, and the activities of ATP-dependent enzymes [59,70]. However, phosphorus itself has little effect on growth and pigment content [58]. Thus, only a few studies on the correlation between phosphorus and C-PE productivity have been published, and the main phosphorus source comes from K2HPO4 [62,71]. Instead, the effect of the nitrogen:phosphorus (N:P) ratio has received great attention [59,72,73]. N2-fixation is phosphorus-dependent, as the process needs a high amount of photosynthetically derived energy [74]. Hence, an optimal N:P ratio is crucial. While the N:P ratio can reveal which of these two nutrients is the underlying limiting factor of cyanobacterial growth and pigment synthesis, it is also a better indicator than each nutrient’s absolute concentration [72]. The extreme nutrient-limiting condition is usually encountered when the external N:P is less than 20:1 [73].
Carbon is another essential nutrient that contributes to all organic compounds, and a sufficient carbon source enables cyanobacteria to proliferate and accumulate metabolites of interest [3]. Carbon, like nitrogen, can be utilized by cyanobacteria in both organic and inorganic sources. The flexibility of shifting toward different sources makes the mixotrophic cyanobacteria more independent of light, which does not limit growth as in autotrophic mode, and carbon assimilation can be supplemented by organic compounds, as in the heterotrophic mode. To be more specific, mixotrophic cultured cyanobacteria can use resources more efficiently and produce more biomass by taking advantage of both heterotrophic and photoautotrophic conditions. The higher biomass may cause cells to self-shade, boosting pigment synthesis, such as PE [58]. The dose of carbon sources varied between strains and was highly dependent on light conditions and culture period [16]. For example, upon pretreatment with either light or darkness, Westiellopsis prolifica used exogenous organic carbon more quickly in the light than in the dark incubation [75]. On the contrary, Calothrix elenkenii utilizes glucose more quickly in the dark, especially after prolonged incubation [76]. The culture media of cyanobacteria typically contained NaHCO3 as the carbon source [53,71,77]. Most research concluded that sucrose was the best organic carbon source. Sucrose (5 g L−1)−1 has been shown to enhance C-PE concentration in Anabaena azollae, Anabaena fertilissima, and Nodularia sphaerocarpa (up to 90%) when compared to fructose and glucose [48,52,78]. Sugarcane molasses, as an alternate source of pure sucrose, has been found to be the most promising substrate for producing C-PE in Nostoc sp. [79]. PBP synthesis may increase in the presence of sucrose due to the production of ATP and higher energy-related assimilation [76]. The addition of glucose in the medium has also been shown to enhance growth and C-PE synthesis in Calothrix sp. and Nostoc sp. [76,79,80,81]. Furthermore, glycerol can substitute glucose in enhancing PBP synthesis in Nostoc sp. [80] Lignite, a low-rank carbon byproduct of numerous carbon extractions, has an organic nature and benefits microbial nutrition [16]. Hence, its use at a low concentration (0.06 g L−1) positively affected growth and C-PE synthesis in Spirulina platensis [82].
Table
Table 2. The nutrient sources utilized by cyanobacteria to increase C-PE production. C-PE content is expressed in mg g−1 unless another unit is indicated. The asterisks (*) indicate the value is absorbance value.
Table 2. The nutrient sources utilized by cyanobacteria to increase C-PE production. C-PE content is expressed in mg g−1 unless another unit is indicated. The asterisks (*) indicate the value is absorbance value.
NutrientsCyanobacteriaSourceConcentration Range (g L−1)Optimal Condition (Source; Concentration (g L−1))C-PE ContentReference
NitrogenAnabaena fertilissima (PUPCCC 410.5)KNO3,KNO20.2–0.5KNO2; 0.2193.2[48]
Anabaena sp.NaNO3, urea, NH4+0–0.50TPB: 127.5[49]
Arthrospira platensisNaNO3, KNO3, NH4Cl2.4NaNO3; 2.41.84%DW[61]
Fischerella sp.NaNO3, NH4+0–1.0NaNO3; 0.2126.5[63]
Nodularia sphaerocarpa (PUPCCC 420.1)KNO3, NaNO20.2, 0.5, 1.0, 1.5KNO3; 0.5, NaNO2; 1.0n/a (Increase 40%)[52]
Nostoc sp. (S36)NaNO30, 20~ 160[67]
Phormidium sp.NaNO3, KNO3, NH4Cl1.5NH4Cl; 1.51.67%DW[61]
Pseudoscillatoria sp.NaNO3, KNO3, NH4Cl1.5NH4Cl; 1.51.36%DW[61]
Spirulina maximaNaNO32.5NaNO3, 2.50.02[62]
PhosphorusOscillatoria sp. (OSCI_UFPS001)K2HPO40.04K2HPO4; 0.041.8%DW[71]
Spirulina maximaK2HPO40.5K2HPO4, 0.50.02[62]
CarbonAnabaena azollaeGlucose, jaggery, sucrose5Sucrose; 50.2 (Increase up to 90%)[78]
Anabaena fertilissima (PUPCCC 410.5)Fructose glucose, sucrose5Sucrose; 5197.9[48]
Calothrix elenkeniiGlucose5Glucose; 50.12[76]
Calothrix sp.Glucose1Glucose; 1n/a[81]
Nodularia sphaerocarpa (PUPCCC 420.1)Fructose glucose, sucrose5, 10Sucrose; 5n/a (Increase 40%)[52]
Nostoc sp.Glucose, sugarcane molasses, sucrose,0-5Glucose; 12.633 *[79]
Sucrose; 0.50.84 *
Sugarcane molasses; 11.44 *
Nostoc sp. (2S7B)Glucose, glycerol0–3Glycerol; 3n/a[80]
Oscillatoria sp. (OSCI_UFPS001)NaHCO30.16NaHCO3; 0.161.8%DW[71]
Spirulina platensisNaHCO30, 4.5, 9, 18NaHCO3; 91.97[53]
Lignite0–0.06Lignite; 0.060.60[82]
Trichromus sp.NaHCO30.02NaHCO3; 0.025%DW[77]
n/a represents not available; DW represents the dry weight.

5. Downstream Processing of C-PE

5.1. Extraction of C-PE

C-PE downstream processes are typically multi-step and sophisticated in order to assure quantity, purity, and quality [4]. A proper extraction should allow for the recovery of the greatest amount of C-PE with minimal contamination, which will be valuable for future analyses of its chemical structure and biological activity [4,83].
The selection of the solvent is primordial for efficient C-PE extraction. The choice of extraction buffer must take into account not merely the polarity and solubility of C-PE, but also the economics, availability of the solvent, and simplicity of use [4,16]. The phosphate buffer and tris chloride buffer are the common buffer types reported for the extraction of C-PE (Table 3). These buffers have been widely used for PBP extraction in cyanobacteria and seaweed and have been proven effective [24,84,85]. An acetate buffer at a pH of 5 is occasionally employed as a buffer for C-PE extraction [86]. However, research by Julianti et al. (2019) and Zavrel et al. (2018) demonstrated that the C-PC yield and purity extracted with an acetate buffer are lower than that extracted with a phosphate buffer [87,88]. A comparatively low-cost, double-distilled water was shown to extract the highest quantity of C-PC, with no significant difference in comparison to PC amounts extracted using a phosphate buffer [89]. However, there is no information on utilizing double-distilled water for C-PE extraction. The pH solvent used in most C-PE extractions was between a pH of 7 and a pH of 8. (Table 3). Dilute or low-molarity buffers could provide better extraction than high-molarity buffers due to the salting-in effect [4,90]. Since a pH of 8 is not suitable for other proteins, it can avoid undesirable contamination, making crude extract purification easier [4].
Cyanobacteria cell walls are multi-layered and difficult to disrupt, yet cell rupture is needed to extract the C-PE. The crude extraction can be accomplished using several methods, including enzymatic digestion, high-pressure homogenization, ultrasonication, or continuous freezing and thawing of the biomass [89]. The methods chosen must consider factors such as the physical strength of the cell wall, stability, and the composition of the cyanobacteria [91]. Enzymatic extraction can be completed by using lysozyme to enzymatically digest the biomass. This enzymatic hydrolysis is particularly effective in seaweed, which has a strong cell covering or outer sheath to protect the cells [92]. Nevertheless, as cyanobacterial cells lack special protection, purified enzymes may not be required. Mechanical methods, such as grinding, crushing, homogenization, or sonication, will generate heat transferred to the biomass solvent, causing the C-PE extracts to be denatured even before they are entirely extracted into the buffer [4,24]. Instead, repeated freezing and thawing were recommended, as it is the gentler approach from the protein perspective and no heat was generated throughout the process [4]. Thus, it is the most commonly used technique for C-PE extraction [36,51,84]. Ghosh and Mishra (2020) also concluded that the freezing–thawing method was the best way to achieve the cell disruption of marine cyanobacteria during their screening of several potential C-PE extraction procedures [4].

5.2. Purification of C-PE

Refined purification can increase the commercial value of C-PE extract. Purifying C-PE is time-consuming and labor-intensive since most cyanobacteria produce PC and APC along with the target PE [93]. Although the ultrafiltration process improves C-PE purity, it reduces C-PE concentration due to thermal and mechanical shear forces [36]. Nowadays, PBP is usually purified from crude extract using at least one of the following methods: precipitation, centrifugation, or chromatography (Table 3). According to Kamble et al. (2018), combining different methodologies is more beneficial for increasing the purity of C-PE while retaining a better yield [36,86]. The first step can be completed by ammonium sulfate precipitation [16]. Ammonium sulfate is water soluble at low temperatures, which inhibits bacterial growth and assists in protein concentration and purification [94]. Different amounts and concentrations of ammonium sulfate were utilized as a salting-out agent to exclude unwanted proteins and boost the purity of C-PE [36]. The precipitated C-PE was further purified based on size, color, and polarity using chromatography approaches, including size-exclusion and ion-exchange chromatography. Size-exclusion chromatography, also known as gel-permeation chromatography, commonly uses Sephadex as a gel filtration resin to remove high-molecular-weight proteins [84,85]. Diethylaminoethyl cellulose (DEAE-C), a positively charged resin, was employed in ion-exchange chromatography to eliminate unwanted protein [36,85,86]. A more developed, aqueous, two-phase system has been used effectively as a low-cost alternative to recover C-PC, but no studies have been conducted to assess its efficiency on C-PE [95].

6. Strategies to Improve the Stability of C-PE

Another prerequisite for large-scale production and acceptability in the industry is the stability of C-PE [3]. C-PE is unstable and susceptible from the time it is extracted from the biomass. As C-PE is a protein, pH, temperature, or light changes may modify its molecular structure. For these reasons, C-PE degradation is possible and can be significant during or after the extraction and purification process, resulting in its shorter shelf life [96]. The pH level is the major factor influencing C-PE aggregation and dissociation, either in monomer, trimer, or hexamer forms in the solution. The most stable structure of C-PE, the hexameric form, predominates at a pH of 7 and easily dissociates at a higher or lower pH [97,98]. C-PE extracted in the study completed by Ghosh et al. (2020) was stable within the pH range of 3–8. This broad pH range facilitates its use in the food and beverage industries. As most beverages have an acidic pH, a stable colorant under such conditions is beneficial [4]. Further, it is recommended to keep extracted C-PE at a low temperature. Temperatures exceeding 40 °C reduce the number of alpha helices, resulting in a loss of stability and gradual degradation [99]. It is also not suggested to keep C-PE at temperatures above room temperature since C-PE is more sensitive to deterioration by microorganisms [97]. Sub-zero temperatures are favorable for long-term preservation since they restrict protein water activity to a minimum level [4]. Furthermore, it is preferable to keep C-PE in the dark [3]. Long-term exposure to high light intensity causes C-PE to lose its chromophores, resulting in color degradation and stability loss [99].
Several stabilizing approaches or strategies have been introduced to increase C-PE stability, as outlined by Hsieh-Lo et al. (2019) [3]. The addition of effective preservatives is the most popular and convenient method. Although dithiothreitol (DTT) and sodium azide (NaN3) are commonly employed in C-PE for analytical purposes, both are poisonous and not suitable for use in food [98]. Thus, only edible preservatives can be used to produce food-grade C-PE. According to Mishra et al. (2010), the stability of C-PE treated with citric acid is higher than that treated with sucrose, calcium chloride, or sodium chloride as preservatives [98]. Benzoic acid is another useful edible additive. It has potent antioxidant properties that can preserve and enhance the stability of C-PE while also acting as an antibacterial agent that inhibits bacterial growth [96].
Crosslinking is another non-additive technology used effectively on PE to increase stability [100]. It is a method of reinforcing the folded structure of a protein by covalently joining two chemical groups on its surface using bifunctional reagents, either intermolecularly or intramolecularly [101]. The intramolecular crosslinking of the protein molecule with silver nanoparticles, Ag+, has been shown to prevent protein aggregation and enhance the thermal stability of PE [100]. Other approaches, such as complex formation and microencapsulation, have been established on PC; however, more study is required to determine the efficacy of employing such techniques on PE [102,103,104]. At present, most of the established strategies have focused on enhancing thermal stability. Hence, stability regarding other factors needs to be explored so that extracted C-PE may be widely marketed across various fields.

7. Application of C-PE

7.1. PE in Pharmaceuticals, Nutraceuticals, & Therapeutics

Several bioactive compounds, such as zeaxanthin, α-tocopherol, and caffeic acid, have already been employed in disease prevention and treatment. However, cyanobacterial bilin is more effective than other phytochemicals, attracting the attention of researchers [105]. The study of PBP bioactivities has emerged in recent years, albeit such study is still restricted and mainly focused on PC [16]. Indeed, PE displays a variety of bioactivities that allow it to be used in the nutraceutical, pharmaceutical, and therapeutic industries.
The accumulation of reactive oxygen species (ROS) causes oxidative stress, which stimulates organisms to produce bioactive compounds in response to their defensive mechanisms. An enzymatic or non-enzymatic antioxidant approach can neutralize these ROS. PBP is an example of a non-enzymatic antioxidant mechanism that scavenges ROS and reduces oxidation [16]. Sonani et al. (2014) discovered that C-PE isolated from Lyngbya sp. exhibited higher in vitro, dose-dependent, antioxidant activity than C-PC and C-APC [85]. This might be because C-PE exhibits antioxidant activity through the main pathway, scavenging already produced ROS via a redox reaction, and has lower chelating and reducing ability [106]. Other potential antioxidant properties of C-PE still require further investigation. In addition to its antioxidant capabilities, C-PE from Lyngbya sp. also had anti-Alzheimer’s and anti-aging properties [85,106]. Phycobilins, such as PCB, PEB, PUB, and PVB, have also been potent phytochemical inhibitors of SARS-CoV-2 Mpro and PLpro proteases [107].
Anti-tumor activities and underlying mechanisms were primarily proven in the literature employing C-PC in medications [16,108]. C-PE differs structurally and spectroscopically from C-PC, suggesting that it may potentially have unique anti-tumor properties [13]. R-PE has been incorporated into photodynamic therapy (PDT), a treatment method for lung, stomach, skin, and oral tumors [13,109]. Effective PDT can selectively destroy cancer or tumor cells by producing ROS-mediated damage, vascular damage, and immune system activation while causing no damage to normal cells [13]. R-PE and its subunit had a stronger PDT effect as well as an inhibitory effect on human liver cancer cell SMC 7221 and mouse tumor cell S180. The smaller size of the β-subunit allows it to access the tumor cell easily. Furthermore, it emitted more significant fluorescence, which was employed as a fluorescent marker for detecting binding sites [109]. In this regard, future research could focus on the role and dosage of C-PE in tumor therapy.
Oral treatment with C-PE as a hepatoprotective and neuroprotective drug has ameliorated kidney, redox, hepatobiliary, and hepatocellular biomarkers against CCl4-induced toxicity in rats. It was determined that proteolytic enzymes might break down C-PE in the gastrointestinal system into bilirubin and low-molecular-weight proteins, mediating the pharmacological effects [110]. Furthermore, C-PE was shown to alleviate diabetes complications by lowering oxidative stress and oxidized, low-density, lipoprotein-induced atherogenesis in streptozotocin-induced type 2 diabetic mice. C-PE administration decreased organ weights, food consumption, cholesterol concentrations, and serum concentrations of glucose and raised body weight, bilirubin, total protein, and the ferric-reducing capacity of plasma values [111]. Also, hepatic and renal tissues showed substantial reductions in lipid hydroperoxide, thiobarbituric acid reactive substances (TBARS), and conjugated diene contents, with elevations in superoxide dismutase, catalase, and glutathione peroxidase, and reduced glutathione, vitamin C, and vitamin E levels [13,111]. Although C-PE administration has become widely used, the detailed mode of action of C-PE in many diseases remains unknown. Further study is needed to elucidate the action point of C-PE in different metabolic pathways.

7.2. PE in Cosmetics

C-PE has long been recognized as an excellent quencher of various oxygen derivatives. Due to this capability, phytopigment of Spirulina sp., such as C-PC, PCB, and PEB, are thought to be excellent antioxidant, anti-wrinkle, anti-melanogenic, and anti-aging agents, as well as natural, non-toxic colorants in eye shadows, eyeliners, and lipsticks [14,112,113]. The majority of the PE content in commercial cosmetics and skin care products is in the form of B-PE or R-PE; however, there is no data on PE derived from cyanobacteria [14,15,114]. For example, purified R-PE isolated from Colaconema formosanum demonstrates anti-aging and anti-allergic properties without toxicity on several mammalian cell lines and epidermal tissues, indicating that this compound has potential for cosmetics usage [15]. Various cyanobacteria strains are now commonly utilized in skin care products to treat various skin problems by functioning as anti-wrinkling agents, sunscreens, nourishing moisturizers, whitening agents, or texture enhancers [113,115]. Hence, it is envisaged that C-PE, which has a range of beneficial bioactivities, will be included in cosmetics or skincare products.

7.3. PE in Food and Feed Industries

By appealing to consumers’ growing health awareness, PE with natural colorant properties has become an alternative to the use of synthetic dye [41]. For example, the pink colorant exhibited by PE has been added as a dye into dairy products, such as milkshakes and yogurt, to make processed products more enticing and to provide color to otherwise colorless food [116]. Red phycoerythrin has a unique yellow fluorescence. Transparent lollipops prepared from sugar solution, soft drinks, dried sugar-drop sweets for cake decorating (that fluoresce under UV light), and fluorescent alcoholic beverages that exploit the benefits of these spectral properties are still being researched [117]. Another pivotal reason for using PE as a dye is that it has antioxidant properties that add value to processed food products. Phycoerythrin is famous as an antioxidant and anti-inflammatory drug, with features that are able to avert ROS-related abnormalities, protect against physiological changes caused by oxidative stress, and have anti-aging benefits [16,118]. This substance also has fewer side effects than synthetic additives or chemical drugs. However, C-PE uses in food products need to be explored further and corroborated through toxicity testing. Further, more study on the bioavailability and interaction of diets and C-PE is necessary [116].
Several studies have demonstrated the benefits of using PE as feed. For example, Lee et al. (2021) found that PE can variably increase the immunological response of whiteleg shrimp in vitro and in vivo and that it might be applied as an immunomodulator in shrimp production [119]. Dietary PE supplementation may also modify the gut microbiota to improve intestinal nutrition and disease resistance in animals by increasing the beneficial bacteria and reducing the prevalence of harmful bacteria within the intestine [120].

7.4. PE in Detection, Diagnosis, and Biotechnology

Compared to monomers, the high molar extinction coefficient of PE is ascribed to trimeric and hexameric packing, as well as the presence of open-chain tetrapyrrole chromophores covalently attached to them [41,119]. The unique fluorescent features of PE have broadened its applications as a fluorescent label in immunoassays, flow cytometry, cell biology, and fluorescence microscopy for biomedical research and diagnostics [121,122,123]. Furthermore, the staining of the PE-labelled monoclonal antibody probe was more clear and bright than the PC-labelled probe when detecting invasive amebiasis [124]. PE can also be employed as a marker in electrofocusing or gel electrophoresis [125] and as a photosensitizer in cancer therapy [126].

8. SWOT Analysis of C-PE Derived from Cyanobacteria

The SWOT analysis is employed to assess the importance of the four main components (strengths, weaknesses, opportunities, and threats) in achieving the economic viability of cyanobacteria-derived C-PE (Figure 4). This analysis appraises the current state and forecasts of the worldwide C-PE markets.

8.1. Strengths and Opportunities

C-PE is being pursued due to its natural origin. Cyanobacteria have attracted more interest than eukaryotic microalgae as the sources of bioactive compounds due to their easier culture methods and higher resistance to bacterial contamination [25]. Cyanobacteria are capable of surviving in a wide range of adverse conditions. Their living processes rely on the bare necessities of light, carbon dioxide, water, and inorganic compounds [127]. Furthermore, cyanobacteria may be produced without arable land and they might be the first plants to colonize bare rock and soil habitats [127]. In microalgae, phycobiliprotein, which functions as a nitrogen reserve, will break down under nitrogen deprivation. According to Da Silva et al., the nitrogen-starved cell of Rhodomonas causes a 25-fold reduction in PE content [128]. In contrast, cyanobacteria can satisfy their own nitrogen requirements through nitrogen fixation [129,130]. This peculiarity reduces the cost of the culture media while reducing the problem of contamination by other microorganisms [38]. Since cyanobacteria own a chromatic adaptation mechanism, green light can be applied to enhance phycoerythrin biosynthesis [16]. The co-production of other valuable compounds, such as vitamins and proteins, also corresponds to additional value [131].
Future research potential includes exploring emerging rare C-PE sources and more profound studies of C-PE and its novel technologies and applications. To date, only a minority of cyanobacteria (i.e., Arthrospira sp., Nostoc sp., and Oscillatoria sp.) are well characterized and commercially exploited. The lesser-known potentially PE-producing strains, such as Pseudanabaena sp., can be explored further [36,132]. The expanding applications of C-PE and consumer acceptance of C-PE will position the industry for revenue growth. The development of high-value markets (both niche and commodity) will also drive research. Moreover, the prokaryotic nature of cyanobacteria allows for more superficial genetic manipulation, resulting in hyper-producing strains [133]. There is also a vast market opportunity in developing improved large-scale culture systems (i.e., effective cyanobacterial screening and strain optimization) and economic, efficient, and environmentally friendly C-PE downstream processing techniques with a focus on the circular economy and sustainability. The end-user is expected to shift with a growing interest in lifestyles of health and sustainability (LOHAS) that prioritize the production of natural C-PE from sustainable sources [134].

8.2. Weaknesses and Threats

Natural C-PE production is problematic because it is labor-intensive and costly [14]. The production cost of high-purity PE was around 30 USD per gram [135]. The high production costs limit the commercial value of C-PE and reduce its market share. C-PE-producing cyanobacteria are often reported to exhibit a slower growth rate and low PE production, along with being recalcitrant, making C-PE extraction and purification even more laborious [3]. Low PE content in cyanobacteria can be rectified using cost-effective process optimization methods [3,16]. A high yield of C-PE can also be achieved through strain improvement via genetic engineering [13]. The downstream processes for extracting C-PE from cultures have become significant challenges since they must adhere to as many green chemical principles as possible to last in the future [58]. Several methods, including maceration, ultrasound, microwaving, freeze–thaw extraction, pressurized liquid/solvent extraction, high-pressure homogenization, and sub and supercritical fluid approaches, have been tested in recent years to extract PE from various sources [4,24]. These are intriguing strategies, and more research will be conducted in the coming years to confirm the findings and enhance pigment selectivity. Conventional solvent extraction methods, such as phosphate-buffered saline and sodium or potassium phosphate buffers are the most well-documented. Nowadays, these solvents are generally replaced with double distilled water, a more environmentally friendly solvent with outstanding results and a high extraction yield [89]. Following extraction, another difficult task is purification. Unlike cryptophytes, cyanobacteria produce phycobilisome protein complexes that usually contain allophycocyanin and phycocyanin. This complicates protein purification in C-PE production since distinct PBPs need to be separated from one another [93]. The chromatography approach appears to be the most promising purifying method, as it is based on a solvent system similar to green solvents used for extraction [36]. There are currently fewer commercially acclaimed cyanobacteria strains available. The rivalry from currently existing natural commercial sources of PE, such as rhodophyte, poses hurdles to the acceptance of cyanobacteria-derived PE [1,24].
Extensive research on cyanotoxins is required to completely comprehend their adverse effects on humans and the environment. Despite their numerous benefits, cyanobacteria can trigger algae blooms, which endanger the environment and ecosystem [136]. As a result, deep insight into the biosynthesis pathway and cyanotoxin removal methods is required to pave the way for the safe exploitation and expansion of the applications of cyanobacterial compounds [137,138]. Strict rules and slow-moving legislation may impede the path to market and the development of novel products. Different countries and geographical areas have different safety and regulatory standards. Companies that intend to market products containing microalgae (including cyanobacteria) must follow the legislation in place, which often involves submitting scientific information and health and safety assessments to a region-specific authority for approval [134]. Only a few algae, such as Porphyridium sp. and Arthrospira sp., are regarded as generally recognized as safe (GRAS) for human consumption by the Food and Drug Administration (FDA) [126]. Biotechnology for species modification (genetic engineering) must also consider health standards [139]. Aside from this, there is a lack of efficient, large-scale C-PE production. It is noteworthy that each cyanobacteria species has specific growth requirements that must be studied prior to large-scale production. Also, further research is needed to deal with the contamination issue in open cultivation systems [16].

9. C-PE Market Drive and Commercial Relevance

Despite a growing interest in algal resources, European production lags behind global production. This is due to the scale and competitiveness of Asian algal aquaculture and the scarcity, ambiguity, and fragmentation of information accessible in the sector [134,140]. The existing regulatory framework for the algae industry may need to be optimized to consider market data and construct a database on algal production and potential markets [140]. To commercialize C-PE worldwide, the limitations, needs, and opportunities for supporting sustainable development in the industry must be identified and resolved.
Research on C-PE proved that, as a natural ingredient, this pigment is an eco-friendly alternative to pharmaceuticals, food supplements or additives, and cosmetic compounds [1,3,16]. There is a high market demand for PE added-value products at present. Still, there is a lack of information regarding effective downstream processing and reducing production costs that is needed to spark the market’s interest. Although some cyanobacteria strains are efficacious C-PE producers, the downstream process has yet to be standardized. This offers a massive obstacle for C-PE to be recognized as an economically viable natural ingredient, as it should be produced through a rapid, simple, and cost-effective method. Nowadays, most reported, commercially available PE products are isolated from the well-established source, rhodophyte in R-PE form, rather than from cyanobacteria [3]. Concerns about cyanotoxins have hampered the acceptance of cyanobacteria-derived products [137]. The difficulties of preserving products for long periods of time have also curtailed PE usage, particularly in food products, resulting in a reduction in available commercial PE-containing products. Nonetheless, improvements in research on enhancing PE stability may allow for the development of a new market for C-PE-based products [3].
The potential applications of PE are determined by its purity index (A565nm/A280nm). A purity value of 0.7 is considered food grade, values between 0.7 and 3.9 are classified as reagent grade, and a value greater than 4.0 is regarded as analytical grade [141]. The market price of purified PE ranged between 180 and 250 USD per milligram, depending on the final purity level and intended applications [142,143]. When PE is extracted for food industries, purity is not a concern, and the price can be easily reduced. However, when used for scientific analysis, pharmaceutical applications, or as a fluorescent agent, PE purity must be higher, and the cost can be a hundred times higher. In fact, PE is sometimes more expensive than PC (15 USD per milligram) [16]. The market size for C-PE is challenging to estimate due to a lack of regional statistics and few technology products. However, the market price for PE is expected to reach $6.3 million by 2025 [24]. According to Future Market Insights Inc., the total market in 2022 is estimated to be worth 2.6 billion USD, with the potential to double that figure by 2032. The report also stated that Europe is anticipated to expand the PE market at a lucrative rate throughout the forecast timeline due to an increase in cosmetic product sales in Italy, France, and Germany over the next decade [144].

10. Conclusions

Cyanobacteria are remarkable blue-green factories that produce several immense market-value compounds despite a minority with toxin productions. However, a complete understanding of C-PE remains underexplored. The current work has shed light on the pertinent knowledge gaps regarding C-PE to increase the feasibility of commercial PE production using cyanobacteria. The interest in PE has burgeoned lately due to its plenteous health-promoting properties. C-PE exists in two forms: C-PE-I (freshwater and soil cyanobacteria) and C-PE-II (marine cyanobacteria). Cyanobacteria possess several attractive attributes, such as a short life cycle, higher biomass production, nitrogen-fixing abilities, and photosynthetic efficiency, as well as a high amount of PE, thus suggesting their ability to effectively substitute the current commercial source of PE (red algae). So far, Tolypothrix tenuis has displayed the highest rate of C-PE production at 660 mg g−1. This study revealed higher C-PE content was induced under green/blue light. In addition, repeated freezing–thawing is the prevalent approach in extracting C-PE. The stability of C-PE is one of the crucial aspects required prior to large-scale production; thus, several strategies were included in this study to improve the stability of this pigment. Furthermore, the understanding of C-PE is greatly enhanced via SWOT analysis, which underlined the strengths, opportunities, weaknesses, and threats of C-PE production. The development of holistic, integrated, up- and downstream approaches could substantially affect the feasibility of C-PE production. More intensive research should focus on nanotechnology, innovations in cultivation, methods of optimization and bioengineering techniques, the integration of synergistic extraction strategies, and economic and environmental assessments of C-PE production.

Author Contributions

Conceptualization, H.T.T., Y.S.K. and F.M.Y.; methodology, H.T.T.; software, H.T.T. and Y.S.K.; validation, Y.S.K., M.F.N., N.A.I.N.M., S.A.A., N.A.S., T.K. and F.M.Y.; formal analysis, H.T.T.; investigation, H.T.T.; resources, F.M.Y.; data curation, H.T.T., M.F.N. and N.A.I.N.M.; writing—original draft preparation, H.T.T.; writing—review and editing, Y.S.K., M.F.N., N.A.I.N.M., S.A.A., N.A.S., T.K. and F.M.Y.; visualization, M.F.N., N.A.I.N.M. and F.M.Y.; supervision, S.A.A., N.A.S., T.K. and F.M.Y. project administration, F.M.Y.; funding acquisition, F.M.Y. All authors have read and agreed to the published version of the manuscript.

Funding

This study was supported by the Ministry of Higher Education Malaysia under the HiCOE research grant (grant no. 6369100), and SATREPS-COSMOS (JPMJSA 1509) Japan-Malaysia collaborative project. The funders had no role in the study design, data collection, analyses, the decision to publish, or the preparation of the manuscript.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Acknowledgments

The first author is the recipient of the Excellent Students’ Program (PPC) Ph.D. program scholarship. Technical assistance from staff and students of the Laboratory of Aquatic Animal Health and Therapeutics at the Institute of Bioscience is duly acknowledged.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Ramu Ganesan, A.; Kannan, M.; Karthick Rajan, D.; Pillay, A.A.; Shanmugam, M.; Sathishkumar, P.; Johansen, J.; Tiwari, B.K. Phycoerythrin: A Pink Pigment from Red Sources (Rhodophyta) for a Greener Biorefining Approach to Food Applications. Crit. Rev. Food Sci. Nutr. 2022, 1–19. [Google Scholar] [CrossRef] [PubMed]
  2. Bryant, D.A. Phycoerythrocyanin and Phycoerythrin: Properties and Occurrence in Cyanobacteria. J. Gen. Microbiol. 1982, 128, 835–844. [Google Scholar] [CrossRef] [Green Version]
  3. Hsieh-Lo, M.; Castillo, G.; Ochoa-Becerra, M.A.; Mojica, L. Phycocyanin and Phycoerythrin: Strategies to Improve Production Yield and Chemical Stability. Algal Res. 2019, 42, 101600. [Google Scholar] [CrossRef]
  4. Ghosh, T.; Mishra, S. Studies on Extraction and Stability of C-Phycoerythrin From a Marine Cyanobacterium. Front. Sustain. Food Syst. 2020, 4, 1–12. [Google Scholar] [CrossRef]
  5. Bryant, D.A.; Guglielmi, G.; de Marsac, N.T.; Castets, A.M.; Cohen-Bazire, G. The Structure of Cyanobacterial Phycobilisomes: A Model. Arch. Microbiol. 1979, 123, 113–127. [Google Scholar] [CrossRef]
  6. Onay, M. Enhancing Phycoerythrin and Phycocyanin Production from Porphyridium Cruentum CCALA 415 in Synthetic Wastewater: The Application of Theoretical Methods on Microalgae. Süleyman Demirel Üniversitesi Fen Bilim. Enstitüsü Derg. 2021, 25, 499–512. [Google Scholar] [CrossRef]
  7. Punampalam, R.; Khoo, K.S.; Sit, N.W. Evaluation of Antioxidant Properties of Phycobiliproteins and Phenolic Compounds Extracted from Bangia atropurpurea. Malays. J. Fundam. Appl. Sci. 2018, 14, 289–297. [Google Scholar] [CrossRef]
  8. Pez Jaeschke, D.; Rocha Teixeira, I.; Damasceno Ferreira Marczak, L.; Domeneghini Mercali, G. Phycocyanin from Spirulina: A Review of Extraction Methods and Stability. Food Res. Int. 2021, 143, 110314. [Google Scholar] [CrossRef] [PubMed]
  9. Kuddus, M.; Singh, P.; Thomas, G.; Al-Hazimi, A. Recent Developments in Production and Biotechnological Applications of C-Phycocyanin. Biomed Res. Int. 2013, 2013, 1–9. [Google Scholar] [CrossRef] [Green Version]
  10. Hemlata; Afreen, S.; Fatma, T. Extraction, Purification and Characterization of Phycoerythrin from Michrochaete and Its Biological Activities. Biocatal. Agric. Biotechnol. 2018, 13, 84–89. [Google Scholar] [CrossRef]
  11. Soni, B.; Visavadiya, N.P.; Dalwadi, N.; Madamwar, D.; Winder, C.; Khalil, C. Purified C-Phycoerythrin: Safety Studies in Rats and Protective Role against Permanganate-Mediated Fibroblast-DNA Damage. J. Appl. Toxicol. 2010, 30, 542–550. [Google Scholar] [CrossRef] [PubMed]
  12. Nowruzi, B.; Fahimi, H.; Lorenzi, A.S. Recovery of Pure C-Phycoerythrin from a Limestone Drought Tolerant Cyanobacterium Nostoc sp. and Evaluation of Its Biological Activity. An. Biol. 2020, 13, 115–128. [Google Scholar] [CrossRef]
  13. Chen, H.; Qi, H.; Xiong, P. Phycobiliproteins—A Family of Algae-Derived Biliproteins: Productions, Characterization and Pharmaceutical Potentials. Mar. Drugs 2022, 20, 450. [Google Scholar] [CrossRef]
  14. Román, R.B.; Alvárez-Pez, J.M.; Fernández, F.G.A.; Grima, E.M. Recovery of Pure B-Phycoerythrin from the Microalga Porphyridium cruentum. J. Biotechnol. 2002, 93, 73–85. [Google Scholar] [CrossRef] [PubMed]
  15. Lee, P.T.; Yeh, H.Y.; Lung, W.Q.C.; Huang, J.; Chen, Y.J.; Chen, B.; Nan, F.H.; Lee, M.C. R-Phycoerythrin from Colaconema formosanum (Rhodophyta), an Anti-Allergic and Collagen Promoting Material for Cosmeceuticals. Appl. Sci. 2021, 11, 9425. [Google Scholar] [CrossRef]
  16. Pagels, F.; Guedes, A.C.; Amaro, H.M.; Kijjoa, A.; Vasconcelos, V. Phycobiliproteins from Cyanobacteria: Chemistry and Biotechnological Applications. Biotechnol. Adv. 2019, 37, 422–443. [Google Scholar] [CrossRef]
  17. Yuan, B.; Li, Z.; Shan, H.; Dashnyam, B.; Xu, X.; McClements, D.J.; Zhang, B.; Tan, M.; Wang, Z.; Cao, C. A Review of Recent Strategies to Improve the Physical Stability of Phycocyanin. Curr. Res. Food Sci. 2022, 5, 2329–2337. [Google Scholar] [CrossRef]
  18. Srivastava, A.; Kalwani, M.; Chakdar, H.; Pabbi, S.; Shukla, P. Biosynthesis and Biotechnological Interventions for Commercial Production of Microalgal Pigments: A Review. Bioresour. Technol. 2022, 352, 127071. [Google Scholar] [CrossRef]
  19. Rather, L.J.; Mir, S.S.; Ganie, S.A.; Shahid-ul-Islam; Li, Q. Research Progress, Challenges, and Perspectives in Microbial Pigment Production for Industrial Applications—A Review. Dye. Pigment. 2023, 210, 110989. [Google Scholar] [CrossRef]
  20. Prabha, S.; Vijay, A.K.; Paul, R.R.; George, B. Cyanobacterial Biorefinery: Towards Economic Feasibility through the Maximum Valorization of Biomass. Sci. Total Environ. 2022, 814, 152795. [Google Scholar] [CrossRef]
  21. Dagnino-leone, J.; Pinto, C.; Latorre, M.; Donoso, A.; Vallejos-almirall, A.; Agurto-muñoz, A.; Pavón, J.; Agurto-muñoz, C. Phycobiliproteins: Structural Aspects, Functional Characteristics, and Biotechnological Perspectives. Comput. Struct. Biotechnol. J. 2022, 20, 1506–1527. [Google Scholar] [CrossRef] [PubMed]
  22. Ram, S.; Mitra, M.; Shah, F.; Tirkey, S.R.; Mishra, S. Bacteria as an Alternate Biofactory for Carotenoid Production: A Review of Its Applications, Opportunities and Challenges. J. Funct. Foods 2020, 67, 103867. [Google Scholar] [CrossRef]
  23. Stengel, D.B.; Connan, S. Natural Products from Marine Algae: Methods and Protocols. Nat. Prod. From Mar. Algae Methods Protoc. 2015, 1308, 1–439. [Google Scholar] [CrossRef]
  24. Ardiles, P.; Cerezal-Mezquita, P.; Salinas-Fuentes, F.; Órdenes, D.; Renato, G.; Ruiz-Domínguez, M.C. Biochemical Composition and Phycoerythrin Extraction from Red Microalgae: A Comparative Study Using Green Extraction Technologies. Processes 2020, 8, 1628. [Google Scholar] [CrossRef]
  25. Frances, E.M.; Onate, C.E. Cyanobacteria and Microalgae in the Production of Valuable Bioactive Compounds. Microalgal Biotechnol. 2018, 6, 105–128. [Google Scholar]
  26. Adir, N.; Dines, M.; Klartag, M.; McGregor, A.; Melamed-Frank, M. Assembly and Disassembly of Phycobilisomes. In Complex Intracellular Structures in Prokaryotes; Springer: Berlin/Heidelberg, Germany, 2006; pp. 47–77. ISBN 9783540325246. [Google Scholar]
  27. Stadnichuk, I.N.; Tropin, I.V. Phycobiliproteins: Structure, Functions and Biotechnological Applications. Appl. Biochem. Microbiol. 2017, 53, 1–10. [Google Scholar] [CrossRef]
  28. Sidler, W.A. Phycobilisome and Phycobiliprotein Structures. In The Molecular Biology of Cyanobacteria; Springer: Dordrecht, The Netherlands, 1994; Volume 7002, pp. 140–216. ISBN 978-0-7923-3273-2. [Google Scholar]
  29. Frank, H.A.; Cogdell, R.J. Light Capture in Photosynthesis; Elsevier Ltd.: Amsterdam, The Netherlands, 2012; Volume 8, ISBN 9780080957180. [Google Scholar]
  30. Mimuro, M.; Kikuchi, H. Antenna Systems and Energy Transfer in Cyanophyta and Rhodophyta. In Light-Harvesting Antennas in Photosynthesis; Springer: Dordrecht, The Netherlands, 2003; pp. 281–306. [Google Scholar]
  31. Chakdar, H.; Pabbi, S. Cyanobacterial Phycobilins: Production, Purification, and Regulation. In Frontier Discoveries and Innovations in Interdisciplinary Microbiology; Springer: New Delhi, India, 2016; pp. 45–69. ISBN 9788132226109. [Google Scholar]
  32. Everroad, C.; Six, C.; Partensky, F.; Thomas, J.C.; Holtzendorff, J.; Wood, A.M. Biochemical Bases of Type IV Chromatic Adaptation in Marine Synechococcus spp. J. Bacteriol. 2006, 188, 3345–3356. [Google Scholar] [CrossRef] [Green Version]
  33. Sonani, R.R.; Roszak, A.W.; Ortmann de Percin Northumberland, C.; Madamwar, D.; Cogdell, R.J. An Improved Crystal Structure of C-Phycoerythrin from the Marine Cyanobacterium Phormidium sp. A09DM. Photosynth. Res. 2018, 135, 65–78. [Google Scholar] [CrossRef]
  34. Freitas, M.V.; Pacheco, D.; Cotas, J.; Mouga, T.; Afonso, C.; Pereira, L. Red Seaweed Pigments from a Biotechnological Perspective. Phycology 2021, 2, 1. [Google Scholar] [CrossRef]
  35. Freitas, M.V.; Inácio, L.G.; Martins, M.; Afonso, C.; Pereira, L.; Mouga, T. Primary Composition and Pigments of 11 Red Seaweed Species from the Center of Portugal. J. Mar. Sci. Eng. 2022, 10, 1168. [Google Scholar] [CrossRef]
  36. Mishra, S.K.; Shrivastav, A.; Mishra, S. Preparation of Highly Purified C-Phycoerythrin from Marine Cyanobacterium Pseudanabaena sp. Protein Expr. Purif. 2011, 80, 234–238. [Google Scholar] [CrossRef] [PubMed]
  37. Tan, H.T.; Yusoff, F.M.; Khaw, Y.S.; Nazarudin, M.F.; Noor Mazli, N.A.I.; Ahmad, S.A.; Shaharuddin, N.A.; Toda, T. Characterisation and Selection of Freshwater Cyanobacteria for Phycobiliprotein Contents. Aquac. Int. 2022, 1–31. [Google Scholar] [CrossRef]
  38. Rodríguez, H.; Rivas, J.; Guerrero, M.G.; Losada, M. Enhancement of Phycobiliprotein Production in Nitrogen-Fixing Cyanobacteria. J. Biotechnol. 1991, 20, 263–270. [Google Scholar] [CrossRef]
  39. Keithellakpam, O.S.; Nath, T.O.; Oinam, A.S.; Thingujam, I.; Oinam, G.; Dutt, S.G. Effect of External PH on Cyanobacterial Phycobiliproteins Production and Ammonium Excretion. J. Appl. Biol. Biotechnol. 2015, 3, 38–42. [Google Scholar] [CrossRef] [Green Version]
  40. Fujita, Y.; Hattori, A. Effect of Chromatic Lights on Phycobilin Formation in a Blue-Green Tolypothrix tenuis. Plant Cell Physiol. 1960, 1, 293–303. [Google Scholar]
  41. Sonani, R.R. Recent Advances in Production, Purification and Applications of Phycobiliproteins. World J. Biol. Chem. 2016, 7, 100. [Google Scholar] [CrossRef] [PubMed]
  42. Venkata Mohan, S.; Rohit, M.V.; Chiranjeevi, P.; Chandra, R.; Navaneeth, B. Heterotrophic Microalgae Cultivation to Synergize Biodiesel Production with Waste Remediation: Progress and Perspectives. Bioresour. Technol. 2015, 184, 169–178. [Google Scholar] [CrossRef]
  43. Bachchhav, M.B.; Kulkarni, M.V.; Ingale, A.G. Enhanced Phycocyanin Production from Spirulina platensis Using Light Emitting Diode. J. Inst. Eng. Ser. E 2017, 98, 41–45. [Google Scholar] [CrossRef]
  44. Vonshak, A.; Cheung, S.M.; Chen, F. Mixotrophic Growth Modifies the Response of Spirulina (Arthrospira) Platensis (Cyanobacteria) Cells to Light. J. Phycol. 2000, 36, 675–679. [Google Scholar] [CrossRef] [Green Version]
  45. Stowe, W.C.; Brodie-Kommit, J.; Stowe-Evans, E. Characterization of Complementary Chromatic Adaptation in Gloeotrichia UTEX 583 and Identification of a Transposon-like Insertion in the CpeBA Operon. Plant Cell Physiol. 2011, 52, 553–562. [Google Scholar] [CrossRef] [Green Version]
  46. Mishra, S.K.; Shrivastav, A.; Maurya, R.R.; Patidar, S.K.; Haldar, S.; Mishra, S. Effect of Light Quality on the C-Phycoerythrin Production in Marine Cyanobacteria Pseudanabaena sp. Isolated from Gujarat Coast, India. Protein Expr. Purif. 2012, 81, 5–10. [Google Scholar] [CrossRef] [PubMed]
  47. Mullineaux, C.W.; Emlyn-Jones, D. State Transitions: An Example of Acclimation to Low-Light Stress. J. Exp. Bot. 2005, 56, 389–393. [Google Scholar] [CrossRef] [PubMed]
  48. Khattar, J.I.S.; Kaur, S.; Kaushal, S.; Singh, Y.; Singh, D.P.; Rana, S.; Gulati, A. Hyperproduction of Phycobiliproteins by the Cyanobacterium Anabaena fertilissima PUPCCC 410.5 under Optimized Culture Conditions. Algal Res. 2015, 12, 463–469. [Google Scholar] [CrossRef]
  49. Hemlata; Fatma, T. Screening of Cyanobacteria for Phycobiliproteins and Effect of Different Environmental Stress on Its Yield. Bull. Environ. Contam. Toxicol. 2009, 83, 509–515. [Google Scholar] [CrossRef] [PubMed]
  50. Maurya, S.S.; Maurya, J.N.; Pandey, V.D. Factors Regulating Phycobiliprotein Production in Cyanobacteria. Int. J. Curr. Microbiol. Appl. Sci. 2014, 3, 764–771. [Google Scholar]
  51. Blas-Valdivia, V.; Rojas-Franco, P.; Serrano-Contreras, J.I.; Sfriso, A.A.; Garcia-Hernandez, C.; Franco-Colín, M.; Cano-Europa, E. C-Phycoerythrin from Phormidium Persicinum Prevents Acute Kidney Injury by Attenuating Oxidative and Endoplasmic Reticulum Stress. Mar. Drugs 2021, 19, 589. [Google Scholar] [CrossRef]
  52. Kaushal, S.; Singh, Y.; Khattar, J.I.S.; Singh, D.P. Phycobiliprotein Production by a Novel Cold Desert Cyanobacterium Nodularia sphaerocarpa PUPCCC 420.1. J. Appl. Phycol. 2017, 29, 1819–1827. [Google Scholar] [CrossRef]
  53. Sharma, G.; Kumar, M.; Ali, M.I.; Jasuja, N.D. Effect of Carbon Content, Salinity and PH on Spirulina platensis for Phycocyanin, Allophycocyanin and Phycoerythrin Accumulation. J. Microb. Biochem. Technol. 2014, 6, 202–206. [Google Scholar] [CrossRef] [Green Version]
  54. Johnson, E.M.; Kumar, K.; Das, D. Physicochemical Parameters Optimization, and Purification of Phycobiliproteins from the Isolated Nostoc sp. Bioresour. Technol. 2014, 166, 541–547. [Google Scholar] [CrossRef]
  55. Sudhir, P.R.; Pogoryelov, D.; Kovács, L.; Garab, G.; Murthy, S.D.S. The Effects of Salt Stress on Photosynthetic Electron Transport and Thylakoid Membrane Proteins in the Cyanobacterium Spirulina platensis. J. Biochem. Mol. Biol. 2005, 38, 481–485. [Google Scholar] [CrossRef] [Green Version]
  56. Vogt, J.C.; Abed, R.M.M.; Albach, D.C.; Palinska, K.A. Bacterial and Archaeal Diversity in Hypersaline Cyanobacterial Mats Along a Transect in the Intertidal Flats of the Sultanate of Oman. Microb. Ecol. 2018, 75, 331–347. [Google Scholar] [CrossRef] [PubMed]
  57. Rafiqul, I.M.; Hassan, A.; Sulebele, G.; Orosco, C.A.; Roustaian, P.; Jalal, K.C.A. Salt Stress Culture of Blue-Green Algae Spirulina fusiformis. Pak. J. Biol. Sci. 2003, 6, 648–650. [Google Scholar] [CrossRef] [Green Version]
  58. Pajot, A.; Huynh, G.H.; Picot, L.; Marchal, L.; Nicolau, E. Fucoxanthin from Algae to Human, an Extraordinary Bioresource: Insights and Advances in up and Downstream Processes. Mar. Drugs 2022, 20, 222. [Google Scholar] [CrossRef] [PubMed]
  59. Fernández-Juárez, V.; Bennasar-Figueras, A.; Sureda-Gomila, A.; Ramis-Munar, G.; Agawin, N.S.R. Differential Effects of Varying Concentrations of Phosphorus, Iron, and Nitrogen in N2-Fixing Cyanobacteria. Front. Microbiol. 2020, 11, 541558. [Google Scholar] [CrossRef]
  60. Klotz, A.; Georg, J.; Bučinská, L.; Watanabe, S.; Reimann, V.; Januszewski, W.; Sobotka, R.; Jendrossek, D.; Hess, W.R.; Forchhammer, K. Awakening of a Dormant Cyanobacterium from Nitrogen Chlorosis Reveals a Genetically Determined Program. Curr. Biol. 2016, 26, 2862–2872. [Google Scholar] [CrossRef] [Green Version]
  61. Khazi, M.I.; Demirel, Z.; Dalay, M.C. Evaluation of Growth and Phycobiliprotein Composition of Cyanobacteria Isolates Cultivated in Different Nitrogen Sources. J. Appl. Phycol. 2018, 30, 1513–1523. [Google Scholar] [CrossRef]
  62. Maza, L.D.L.R.; Guevara, M.; Gómez, B.J.; Arredondo-Vega, B.; Cortez, R.; Licet, B. Production of Pigments from Arthrospira maxima Cultivated in Photobioreactors. Rev. Colomb. Biotecnol. 2017, 19, 108–114. [Google Scholar] [CrossRef]
  63. Soltani, N.; Khavarinezhad, R.A.; Tabatabaei, Y.S.; Shokravi, S. Growth and Some Metabolic Features of Cyanobacterium Fischerella sp. FS18 in Different Combined Nitrogen. J. Sci. 2007, 18, 123–128. [Google Scholar]
  64. Liotenberg, S.; Campbell, D.; Rippka, R.; Houmard, J.; Tandeau De Marsac, N. Effect of the Nitrogen Source on Phycobiliprotein Synthesis and Cell Reserves in a Chromatically Adapting Filamentous Cyanobacterium. Microbiology 1996, 142, 611–622. [Google Scholar] [CrossRef] [Green Version]
  65. Glazer, A.N. Phycobilisomes. In Methods in Enzymology; Academic Press: Cambridge, UK, 1988; Volume 167, pp. 304–312. [Google Scholar]
  66. Ting, C.S.; Rocap, G.; King, J.; Chisholm, S.W. Cyanobacterial Photosynthesis in the Oceans: The Origins and Significance of Divergent Light-Harvesting Strategies. Trends Microbiol. 2002, 10, 134–142. [Google Scholar] [CrossRef]
  67. Simeunović, J.; Bešlin, K.; Svirčev, Z.; Kovač, D.; Babić, O. Impact of Nitrogen and Drought on Phycobiliprotein Content in Terrestrial Cyanobacterial Strains. J. Appl. Phycol. 2013, 25, 597–607. [Google Scholar] [CrossRef]
  68. Zehr, J.P. Nitrogen Fixation by Marine Cyanobacteria. Trends Microbiol. 2011, 19, 162–173. [Google Scholar] [CrossRef] [PubMed]
  69. Sohm, J.A.; Webb, E.A.; Capone, D.G. Emerging Patterns of Marine Nitrogen Fixation. Nat. Rev. Microbiol. 2011, 9, 499–508. [Google Scholar] [CrossRef]
  70. Geider, R.J.; La Roche, J. Redfield Revisited: Variability of C:N:P in Marine Microalgae and Its Biochemical Basis. Eur. J. Phycol. 2002, 37, 1–17. [Google Scholar] [CrossRef]
  71. Zuorro, A.; Leal-Jerez, A.G.; Morales-Rivas, L.K.; Mogollón-Londoño, S.O.; Sanchez-Galvis, E.M.; García-Martínez, J.B.; Barajas-Solano, A.F. Enhancement of Phycobiliprotein Accumulation in Thermotolerant Oscillatoria sp. Through Media Optimization. ACS Omega 2021, 6, 10527–10536. [Google Scholar] [CrossRef] [PubMed]
  72. Peng, G.; Wilhelm, S.W.; Lin, S.; Wang, X. Response of Microcystis Aeruginosa FACHB-905 to Different Nutrient Ratios and Changes in Phosphorus Chemistry. J. Oceanol. Limnol. 2018, 36, 1040–1052. [Google Scholar] [CrossRef]
  73. Guildford, S.J.; Hecky, R.E. Total Nitrogen, Total Phosphorus, and Nutrient Limitation in Lakes and Oceans: Is There a Common Relationship? Limnol. Oceanogr. 2000, 45, 1213–1223. [Google Scholar] [CrossRef] [Green Version]
  74. Sañudo-Wilhelmy, S.A.; Kustka, A.B.; Gobler, C.J.; Hutchins, D.A.; Yang, M.; Lwiza, K.; Burns, J.; Capone, D.G.; Raven, J.A.; Carpenter, E.J. Phosphorus Limitation of Nitrogen Fixation by Trichodesmium in the Central Atlantic Ocean. Nature 2001, 411, 66–69. [Google Scholar] [CrossRef]
  75. Adhikary, S.P.; Pattnaik, H. Growth Response of Westiellopsis prolifica Janet to Organic Substrates in Light and Dark. Hydrobiologia 1979, 67, 241–247. [Google Scholar] [CrossRef]
  76. Prasanna, R.; Pabby, A.; Singh, P.K. Effect of Glucose and Light-Dark Environment on Pigmentation Profiles in the Cyanobacterium Calothrix elenkenii. Folia Microbiol. 2004, 49, 26–30. [Google Scholar] [CrossRef]
  77. Haddad, M.F.; Dayioglu, T.; Yaman, M.; Nalbantoglu, B.; Cakmak, T. Long-Term Diazotrophic Cultivation of Trichormus sp. IMU26: Evaluation of Physiological Changes Related to Elevated Phycobiliprotein Content. J. Appl. Phycol. 2020, 32, 881–888. [Google Scholar] [CrossRef]
  78. Venugopal, V.; Prasanna, R.; Sood, A.; Jaiswal, P.; Kaushik, B.D. Stimulation of Pigment Accumulation in Anabaena Azollae Strains: Effect of Light Intensity and Sugars. Folia Microbiol. 2006, 51, 50–56. [Google Scholar] [CrossRef] [PubMed]
  79. Borsari, R.R.J.; Morioka, L.R.I.; Ribeiro, M.L.L.; Buzato, J.B.; Pinotti, M.H.P. Mixotrophic Growth of Nostoc sp. on Glucose, Sucrose and Sugarcane Molasses for Phycobiliprotein Production. Acta Sci.-Biol. Sci. 2007, 29, 9–13. [Google Scholar]
  80. Kovač, D.; Babić, O.; Milovanović, I.; Mišan, A.; Simeunović, J. The Production of Biomass and Phycobiliprotein Pigments in Filamentous Cyanobacteria: The Impact of Light and Carbon Sources. Appl. Biochem. Microbiol. 2017, 53, 539–545. [Google Scholar] [CrossRef]
  81. Lebedeva, N.V.; Boichenko, V.A.; Semenova, L.R.; Pronina, N.A.; Stadnichuk, I.N. Effects of Glucose during Photoheterotrophic Growth of the Cyanobacterium Calothrix sp. PCC 7601 Capable for Chromatic Adaptation. Russ. J. Plant Physiol. 2005, 52, 235–241. [Google Scholar] [CrossRef]
  82. Rivera Gonzalez, M.V.; Gómez Gómez, L.; Cubillos Hinojosa, J.G.; Peralta Castilla, A. Effect of Coal Type Lignite on Growth and Production of Pigments of Arthrospira platensis. Rev. Colomb. Biotecnol. 2016, 18, 73–80. [Google Scholar] [CrossRef]
  83. Denis, C.; Ledorze, C.; Jaouen, P.; Fleurence, J. Comparison of Different Procedures for the Extraction and Partial Purification of R-Phycoerythrin from the Red Macroalga Grateloupia turuturu. Bot. Mar. 2009, 52, 278–281. [Google Scholar] [CrossRef]
  84. Parmar, A.; Singh, N.K.; Kaushal, A.; Sonawala, S.; Madamwar, D. Purification, Characterization and Comparison of Phycoerythrins from Three Different Marine Cyanobacterial Cultures. Bioresour. Technol. 2011, 102, 1795–1802. [Google Scholar] [CrossRef]
  85. Sonani, R.R.; Singh, N.K.; Kumar, J.; Thakar, D.; Madamwar, D. Concurrent Purification and Antioxidant Activity of Phycobiliproteins from Lyngbya sp. A09DM: An Antioxidant and Anti-Aging Potential of Phycoerythrin in Caenorhabditis elegans. Process Biochem. 2014, 49, 1757–1766. [Google Scholar] [CrossRef]
  86. Kamble, S.P.; Vikhe, G.P.; Chamle, D.R. Extraction and Purification of Phycoerythrin-A Natural Colouring Agent from Spirulina platensis. J. Pharm. Chem. Biol. Sci. 2018, 6, 78–84. [Google Scholar]
  87. Zavřel, T.; Chmelík, D.; Sinetova, M.A.; Červený, J. Spectrophotometric Determination of Phycobiliprotein Content in Cyanobacterium Synechocystis. J. Vis. Exp. 2018, 139, 1–9. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  88. Julianti, E.; Susanti; Singgih, M.; Neti Mulyani, L. Optimization of Extraction Method and Characterization of Phycocyanin Pigment from Spirulina platensis. J. Math. Fundam. Sci. 2019, 51, 168–176. [Google Scholar] [CrossRef]
  89. Tan, H.T.; Khong, N.M.H.; Khaw, Y.S.; Ahmad, S.A.; Yuso, F.M. Optimization of the Freezing-Thawing Method for Extracting Phycobiliproteins from Arthrospira sp. Molecules 2020, 25, 3894. [Google Scholar] [CrossRef] [PubMed]
  90. Arakawa, T.; Timasheff, S.N. The Stabilization of Proteins by Osmolytes. Biophys. J. 1985, 47, 411–414. [Google Scholar] [CrossRef]
  91. Pan-utai, W.; Iamtham, S. Physical Extraction and Extrusion Entrapment of C-Phycocyanin from Arthrospira platensis. J. King Saud Univ.-Sci. 2019, 31, 1535–1542. [Google Scholar] [CrossRef]
  92. Dumay, J.; Clément, N.; Morançais, M.; Fleurence, J. Optimization of Hydrolysis Conditions of Palmaria palmata to Enhance R-Phycoerythrin Extraction. Bioresour. Technol. 2013, 131, 21–27. [Google Scholar] [CrossRef]
  93. Mercier, L.; Peltomaa, E.; Ojala, A. Comparative Analysis of Phycoerythrin Production in Cryptophytes. J. Appl. Phycol. 2022, 34, 789–797. [Google Scholar] [CrossRef]
  94. Patel, A.; Mishra, S.; Pawar, R.; Ghosh, P.K. Purification and Characterization of C-Phycocyanin from Cyanobacterial Species of Marine and Freshwater Habitat. Protein Expr. Purif. 2005, 40, 248–255. [Google Scholar] [CrossRef]
  95. Rito-Palomares, M.; Nuez, L.; Amador, D. Practical Application of Aqueous Two-Phase Systems for the Development of a Prototype Process for c-Phycocyanin Recovery from Spirulina maxima. J. Chem. Technol. Biotechnol. 2001, 76, 1273–1280. [Google Scholar] [CrossRef]
  96. Kannaujiya, V.K.; Sinha, R.P. Thermokinetic Stability of Phycocyanin and Phycoerythrin in Food-Grade Preservatives. J. Appl. Phycol. 2016, 28, 1063–1070. [Google Scholar] [CrossRef]
  97. Chaiklahan, R.; Chirasuwan, N.; Bunnag, B. Stability of Phycocyanin Extracted from Spirulina sp.: Influence of Temperature, PH and Preservatives. Process Biochem. 2012, 47, 659–664. [Google Scholar] [CrossRef]
  98. Mishra, S.K.; Shrivastav, A.; Pancha, I.; Jain, D.; Mishra, S. Effect of Preservatives for Food Grade C-Phycoerythrin, Isolated from Marine Cyanobacteria Pseudanabaena sp. Int. J. Biol. Macromol. 2010, 47, 597–602. [Google Scholar] [CrossRef] [PubMed]
  99. Munier, M.; Jubeau, S.; Wijaya, A.; Morançais, M.; Dumay, J.; Marchal, L.; Jaouen, P.; Fleurence, J. Physicochemical Factors Affecting the Stability of Two Pigments: R-Phycoerythrin of Grateloupia turuturu and B-Phycoerythrin of Porphyridium cruentum. Food Chem. 2014, 150, 400–407. [Google Scholar] [CrossRef] [PubMed]
  100. Bekasova, O.D.; Borzova, V.A.; Shubin, V.V.; Kovalyov, L.I.; Stein-Margolina, V.A.; Kurganov, B.I. An Increase in the Resistance of R-Phycoerythrin to Thermal Aggregation by Silver Nanoparticles Synthesized in Nanochannels of the Pigment. Appl. Biochem. Microbiol. 2016, 52, 98–104. [Google Scholar] [CrossRef]
  101. Martelli, G.; Folli, C.; Visai, L.; Daglia, M.; Ferrari, D. Thermal Stability Improvement of Blue Colorant C-Phycocyanin from Spirulina platensis for Food Industry Applications. Process Biochem. 2014, 49, 154–159. [Google Scholar] [CrossRef]
  102. Selig, M.J.; Malchione, N.M.; Gamaleldin, S.; Padilla-Zakour, O.I.; Abbaspourrad, A. Protection of Blue Color in a Spirulina Derived Phycocyanin Extract from Proteolytic and Thermal Degradation via Complexation with Beet-Pectin. Food Hydrocoll. 2018, 74, 46–52. [Google Scholar] [CrossRef]
  103. Yan, M.; Liu, B.; Jiao, X.; Qin, S. Preparation of Phycocyanin Microcapsules and Its Properties. Food Bioprod. Process. 2014, 92, 89–97. [Google Scholar] [CrossRef]
  104. Purnamayati, L.; Dewi, E.; Kurniasih, R.A. Phycocyanin Stability in Microcapsules Processed by Spray Drying Method Using Different Inlet Temperature. In Proceedings of the IOP Conference Series: Earth and Environmental Science, 3rd International Conference on Tropical and Coastal Region Eco Development 2017, Yogyakarta, Indonesia, 2–4 October 2017; p. 012076. [Google Scholar] [CrossRef]
  105. Hirata, T.; Tanaka, M.; Ooike, M.; Tsunomura, T.; Sakaguchi, M. Antioxidant Activities of Phycocyanobilin Prepared from Spirulina platensis. J. Appl. Phycol. 2000, 12, 435–439. [Google Scholar] [CrossRef]
  106. Sonani, R.R. Antioxidant Potential of Phycobiliproteins: Role in Anti-Aging Research. Biochem. Anal. Biochem. 2015, 4, 2161-1009. [Google Scholar] [CrossRef] [Green Version]
  107. Pendyala, B.; Patras, A.; Dash, C. Phycobilins as Potent Food Bioactive Broad-Spectrum Inhibitors Against Proteases of SARS-CoV-2 and Other Coronaviruses: A Preliminary Study. Front. Microbiol. 2021, 12, 645713. [Google Scholar] [CrossRef]
  108. Jiang, L.; Wang, Y.; Liu, G.; Liu, H.; Zhu, F.; Ji, H.; Li, B. C-Phycocyanin Exerts Anti-Cancer Effects via the MAPK Signaling Pathway in MDA-MB-231 Cells. Cancer Cell Int. 2018, 18, 12. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  109. Bei, H.; Guang-Ce, W.; Chen-Kui, Z.; Zhen-Gang, L. The Experimental Research of R-Phycoerythrin Subunits on Cancer Treatment: A New Photosensitizer in PDT. Cancer Biother. Radiopharm. 2002, 17, 35–42. [Google Scholar] [CrossRef] [PubMed]
  110. Soni, B.; Visavadiya, N.P.; Madamwar, D. Ameliorative Action of Cyanobacterial Phycoerythrin on CCl4-Induced Toxicity in Rats. Toxicology 2008, 248, 59–65. [Google Scholar] [CrossRef] [PubMed]
  111. Soni, B.; Visavadiya, N.P.; Madamwar, D. Attenuation of Diabetic Complications by C-Phycoerythrin in Rats: Antioxidant Activity of C-Phycoerythrin Including Copper-Induced Lipoprotein and Serum Oxidation. Br. J. Nutr. 2009, 102, 102–109. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  112. Sathasivam, R.; Radhakrishnan, R.; Hashem, A.; Abd_Allah, E.F. Microalgae Metabolites: A Rich Source for Food and Medicine. Saudi J. Biol. Sci. 2019, 26, 709–722. [Google Scholar] [CrossRef]
  113. Nowruzi, B.; Sarvari, G.; Blanco, S. The Cosmetic Application of Cyanobacterial Secondary Metabolites. Algal Res. 2020, 49, 101959. [Google Scholar] [CrossRef]
  114. D’Agnolo, E.; Rizzo, R.; Paoletti, S.; Murano, E. R-Phycoerythrin from the Red Alga Gracilaria longa. Phytochemistry 1994, 35, 693–696. [Google Scholar] [CrossRef]
  115. Vaibhav, V.; Sahastrabuddhe, S. ‘BLUE’ Is the New ‘GREEN’ for Cosmetic Industry. Int. J. Res. Trends Innov. 2018, 3, 134–144. [Google Scholar]
  116. García, A.B.; Longo, E.; Murillo, M.C.; Bermejo, R. Using a B-Phycoerythrin Extract as a Natural Colorant: Application in Milk-Based Products. Molecules 2021, 26, 297. [Google Scholar] [CrossRef]
  117. Dufossé, L.; Galaup, P.; Yaron, A.; Arad, S.M.; Blanc, P.; Murthy, K.N.C.; Ravishankar, G.A. Microorganisms and Microalgae as Sources of Pigments for Food Use: A Scientific Oddity or an Industrial Reality? Trends Food Sci. Technol. 2005, 16, 389–406. [Google Scholar] [CrossRef]
  118. Patel, S.N.; Sonani, R.R.; Jakharia, K.; Bhastana, B.; Patel, H.M.; Chaubey, M.G.; Singh, N.K.; Madamwar, D. Antioxidant Activity and Associated Structural Attributes of Halomicronema Phycoerythrin. Int. J. Biol. Macromol. 2018, 111, 359–369. [Google Scholar] [CrossRef] [PubMed]
  119. Liu, L.-N.; Su, H.-N.; Yan, S.-G.; Shao, S.-M.; Xie, B.-B.; Chen, X.-L.; Zhang, X.-Y.; Zhou, B.-C.; Zhang, Y.-Z. Probing the PH Sensitivity of R-Phycoerythrin: Investigations of Active Conformational and Functional Variation. Biochim. Biophys. Acta-Bioenerg. 2009, 1787, 939–946. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  120. Qi, H.; Liu, Y.; Qi, X.; Liang, H.; Chen, H.; Jiang, P.; Wang, D. Dietary Recombinant Phycoerythrin Modulates the Gut Microbiota of H22 Tumor-Bearing Mice. Mar. Drugs 2019, 17, 665. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  121. Glazer, A.N. Phycobiliproteins—A Family of Valuable, Widely Used Fluorophores. J. Appl. Phycol. 1994, 6, 105–112. [Google Scholar] [CrossRef]
  122. Bailey, M.P.; Rocks, B.F.; Riley, C. On the Use of Fluorescent Labels in Immunoassay. J. Pharm. Biomed. Anal. 1987, 5, 649–658. [Google Scholar] [CrossRef]
  123. Wilson, M.R.; Crowley, S.; Odgers, G.A.; Shaw, L. Immunofluorescent Labeling Using Covalently Linked Anti-phycoerythrin Antibodies and Phycoerythrin Polymers. Cytometry 1991, 12, 373–377. [Google Scholar] [CrossRef]
  124. Intrarapuk, A.; Awakairt, S.; Thammapalerd, N.; Mahannop, P.; Pattanawong, U.; Suppasiri, T. Comparison between R-Phycocyanin-Labeled and R-Phycoerythrin-Labeled Monoclonal Antibody (MAb) Probes for the Detection of Entamoeba histolytica Trophozoites. Southeast Asian J. Trop. Med. Public Health 2001, 32, 165–171. [Google Scholar]
  125. Aráoz, R.; Lebert, M.; Häder, D.P. Electrophoretic Applications of Phycobiliproteins. Electrophoresis 1998, 19, 215–219. [Google Scholar] [CrossRef]
  126. Li, S.; Ji, L.; Shi, Q.; Wu, H.; Fan, J. Advances in the Production of Bioactive Substances from Marine Unicellular Microalgae Porphyridium spp. Bioresour. Technol. 2019, 292, 122048. [Google Scholar] [CrossRef]
  127. Issa, A.A.; Abd-Alla, M.H.; Ohyama, T. Nitrogen Fixing Cyanobacteria: Future Prospect. In Advances in Biology and Ecology of Nitrogen Fixation; InTech: London, UK, 2014; Volume 2, pp. 24–48. [Google Scholar]
  128. da Silva, A.F.; Lourenço, S.O.; Chaloub, R.M. Effects of Nitrogen Starvation on the Photosynthetic Physiology of a Tropical Marine Microalga Rhodomonas sp. (Cryptophyceae). Aquat. Bot. 2009, 91, 291–297. [Google Scholar] [CrossRef]
  129. Pathak, J.; Rajneesh; Maurya, P.K.; Singh, S.P.; Häder, D.P.; Sinha, R.P. Cyanobacterial Farming for Environment Friendly Sustainable Agriculture Practices: Innovations and Perspectives. Front. Environ. Sci. 2018, 6, 7. [Google Scholar] [CrossRef]
  130. Singh, J.S.; Kumar, A.; Rai, A.N.; Singh, D.P. Cyanobacteria: A Precious Bio-Resource in Agriculture, Ecosystem, and Environmental Sustainability. Front. Microbiol. 2016, 7, 529. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  131. Singh, S.; Kate, B.N.; Banecjee, U.C. Bioactive Compounds from Cyanobacteria and Microalgae: An Overview. Crit. Rev. Biotechnol. 2005, 25, 73–95. [Google Scholar] [CrossRef] [PubMed]
  132. Khan, Z.; Wan Maznah, W.O.; Faradina Merican, M.S.M.; Convey, P.; Najimudin, N.; Alias, S.A. A Comparative Study of Phycobilliprotein Production in Two Strains of Pseudanabaena Isolated from Arctic and Tropical Regions in Relation to Different Light Wavelengths and Photoperiods. Polar Sci. 2019, 20, 3–8. [Google Scholar] [CrossRef]
  133. Berla, B.M.; Saha, R.; Immethun, C.M.; Maranas, C.D.; Moon, T.S.; Pakrasi, H.B. Synthetic Biology of Cyanobacteria: Unique Challenges and Opportunities. Front. Microbiol. 2013, 4, 246. [Google Scholar] [CrossRef] [Green Version]
  134. Novoveská, L.; Ross, M.E.; Stanley, M.S.; Pradelles, R.; Wasiolek, V.; Sassi, J.F. Microalgal Carotenoids: A Review of Production, Current Markets, Regulations, and Future Direction. Mar. Drugs 2019, 17, 640. [Google Scholar] [CrossRef] [Green Version]
  135. Reis, A.; Mendes, A.; Lobo-Fernandes, H.; Empis, J.A.; Novais, J.M. Production, Extraction and Purification of Phycobiliproteins from Nostoc sp. Bioresour. Technol. 1998, 66, 181–187. [Google Scholar] [CrossRef]
  136. Pantelić, D.; Svirčev, Z.; Simeunović, J.; Vidović, M.; Trajković, I. Cyanotoxins: Characteristics, Production and Degradation Routes in Drinking Water Treatment with Reference to the Situation in Serbia. Chemosphere 2013, 91, 421–441. [Google Scholar] [CrossRef]
  137. Abbas, T.; Kajjumba, G.W.; Ejjada, M.; Masrura, S.U.; Marti, E.J.; Khan, E.; Jones-lepp, T.L. Recent Advancements in the Removal of Cyanotoxins from Water Using Conventional and Modified Adsorbents—A Contemporary Review. Water 2020, 12, 2756. [Google Scholar] [CrossRef]
  138. Westrick, J.A.; Szlag, D.C.; Southwell, B.J.; Sinclair, J. A Review of Cyanobacteria and Cyanotoxins Removal/Inactivation in Drinking Water Treatment. Anal. Bioanal. Chem. 2010, 397, 1705–1714. [Google Scholar] [CrossRef]
  139. Moreira, J.B.; Vaz, B.D.S.; Cardias, B.B.; Cruz, C.G.; de Almeida, A.C.A.; Costa, J.A.V.; de Morais, M.G. Microalgae Polysaccharides: An Alternative Source for Food Production and Sustainable Agriculture. Polysaccharides 2022, 3, 441–457. [Google Scholar] [CrossRef]
  140. Araújo, R.; Vázquez Calderón, F.; Sánchez López, J.; Azevedo, I.C.; Bruhn, A.; Fluch, S.; Garcia Tasende, M.; Ghaderiardakani, F.; Ilmjärv, T.; Laurans, M.; et al. Current Status of the Algae Production Industry in Europe: An Emerging Sector of the Blue Bioeconomy. Front. Mar. Sci. 2021, 7, 1247. [Google Scholar] [CrossRef]
  141. Wang, C.; Shen, Z.; Cui, X.; Jiang, Y.; Jiang, X. Response Surface Optimization of Enzyme-Assisted Extraction of R-Phycoerythrin from Dry Pyropia yezoensis. J. Appl. Phycol. 2020, 32, 1429–1440. [Google Scholar] [CrossRef]
  142. Mittal, R.; Raghavarao, K.S.M.S. Extraction of R-Phycoerythrin from Marine Macro-Algae, Gelidium pusillum, Employing Consortia of Enzymes. Algal Res. 2018, 34, 1–11. [Google Scholar] [CrossRef]
  143. Phuong, H.; Massé, A.; Dumay, J.; Vandanjon, L.; Mith, H.; Legrand, J.; Arhaliass, A. Enhanced Liberation of Soluble Sugar, Protein, and R-Phycoerythrin Under Enzyme-Assisted Extraction on Dried and Fresh Gracilaria gracilis Biomass. Front. Chem. Eng. 2022, 4, 718857. [Google Scholar] [CrossRef]
  144. Insights Phycoerythrin Market Overview. 2022. Available online: https://www.futuremarketinsights.com/reports/phycoerythrin-market (accessed on 28 August 2022).
Marinedrugs 21 00028 g001 550
Figure 1. Atomic resolution structure (PDB ID: 5NB4) of C-phycoerythrin from Phormidium sp. (a) α-subunits; (b) β-subunits; (c) three-dimensional structure of phycoerythrobilin; (d) three-dimensional structure of trimer; (e) three-dimensional structure of hexamer (adopted from [33] with permission, 2017, Sonani et al.).
Figure 1. Atomic resolution structure (PDB ID: 5NB4) of C-phycoerythrin from Phormidium sp. (a) α-subunits; (b) β-subunits; (c) three-dimensional structure of phycoerythrobilin; (d) three-dimensional structure of trimer; (e) three-dimensional structure of hexamer (adopted from [33] with permission, 2017, Sonani et al.).
Marinedrugs 21 00028 g001
Marinedrugs 21 00028 g002 550
Figure 2. The UV-visible absorbance spectra (solid line) and fluorescence emission spectra (dotted line) of C-PE extracted from Pseudanabaena sp. (UPMC-A0103). C-PE of Pseudanabaena sp. showed an absorption maxima at 564 nm and fluorescence emission maxima at 574 nm.
Figure 2. The UV-visible absorbance spectra (solid line) and fluorescence emission spectra (dotted line) of C-PE extracted from Pseudanabaena sp. (UPMC-A0103). C-PE of Pseudanabaena sp. showed an absorption maxima at 564 nm and fluorescence emission maxima at 574 nm.
Marinedrugs 21 00028 g002
Marinedrugs 21 00028 g003 550
Figure 3. Potential cyanobacteria strains that produce C-phycoerythrin: (a) Tolypothrix sp.; (b) Pseudanabaena sp.
Figure 3. Potential cyanobacteria strains that produce C-phycoerythrin: (a) Tolypothrix sp.; (b) Pseudanabaena sp.
Marinedrugs 21 00028 g003
Marinedrugs 21 00028 g004 550
Figure 4. Strengths, weaknesses, opportunities, and threats (SWOT) analysis of C-PE.
Figure 4. Strengths, weaknesses, opportunities, and threats (SWOT) analysis of C-PE.
Marinedrugs 21 00028 g004
Table
Table 1. The optimum cultivation conditions of cyanobacteria to achieve high yields of C-phycoerythrin (C-PE). Cultivation parameters include culture media (M), light intensity (I), photoperiod (LP), light color (LC), pH, temperature (T), and salinity (S). Light intensity is expressed in μmol m−2 s−1 unless another unit is indicated. Parameters highlighted in red color indicated the factors optimized in the respective study.
Table 1. The optimum cultivation conditions of cyanobacteria to achieve high yields of C-phycoerythrin (C-PE). Cultivation parameters include culture media (M), light intensity (I), photoperiod (LP), light color (LC), pH, temperature (T), and salinity (S). Light intensity is expressed in μmol m−2 s−1 unless another unit is indicated. Parameters highlighted in red color indicated the factors optimized in the respective study.
CyanobacteriaOptimal Cultivation ParametersCultivation ModeC-PE Content (mg g−1)References
Anabaena sp.M: BG 11 + HEPES; I: 100; LP: 12:12 h; LC: Green; pH: 8; T: 30 °Cautotrophic102[38]
Anabaena fertilissima (PUPCCC 410.5)M: Chu-10 medium; I: 44.5; LP: 14:10 h; LC: white; pH: 8; T: 28 °Cmixotrophic105.8[48]
M: Chu-10 medium; I: 44.5; LP: 14:10 h; LC:blue; pH: 8; T: 28 °C 214.5
Fremyella diplosiphonM: BG 11 + HEPES; I: 15; LP: n/a; LC: Green; pH: 8; T: 27 °Cautotrophic559.69[45]
Gloeotrichia sp.M: BG 11 + HEPES; I: 15; LP: n/a; LC: Green; pH: 8; T: 27 °Cautotrophic414.18[45]
Lyngbya sp. (CCNM 2053)M: ASN-III; I: 60; LP: 12:12 h; LC: n/a; pH: n/a; T: 25 °C; S: n/aautotrophic22.99[4]
Nodularia sphaerocarpa (PUPCCC 420.1)M: Chu-10 medium; I: 44.5; LP: 14:10 h; LC: green; pH: 8; T: 28 °Cmixotrophic283[52]
Nostoc sp. BTA-61M: BG11; I: 54-67; LP: 14:10 h; LC: white; pH: 7.0; T: 28 °Cautotrophic125.11[39]
Phormidium persicinumM: synthetic NM; I: 3000 LUX; LP: 12:12 h; LC: n/a; pH: n/a; T: 21 °C; S: n/amixotrophic32.98[51]
Pseudanabaena sp.M: ASN-III; I: 75-110; LP: 12:12 h; LC: Green/blue; pH: n/a; T: 25 °C; S: n/amixotrophic30[46]
M: BG 11; I: 40; LP: 12:12 h; LC: n/a; pH: 7; T: 25 °Cautotrophic92.57[37]
Spirulina platensisM: Zarrouk’s medium; I: n/a; LP: n/a; LC: n/a; pH: n/a; T: 30 °C; S: 0.4 Mmixotrophic1.61[53]
M: Zarrouk’s medium; I: n/a; LP: n/a; LC: n/a; pH: 7; T: 30 °C; S: n/a 1.44
Tolypothrix tenuisM: N-free mineral medium; I: 200 LUX; LP: 12:12 h; LC: Green; pH: n/a; T: 20–25 °Cmixotrophic660[40]
HEPES, hydroxyethyl piperazineethanesulfonic acid; ASN, artificial seawater nutrient; NM, nutrient media.
Table
Table 3. The extraction and purification methods used to extract C-PE.
Table 3. The extraction and purification methods used to extract C-PE.
CyanobacteriaExtraction MethodSolventPurificationC-PE ContentPurity of C-PEReferences
Halomicronema sp. A27DM.Freezing–thawingTris Cl buffer + sodium azide (3 mM), pH 8.1Ammonium sulfate precipitation + Gel permeation chromatography (Sephadex G-150)5.76 mg mL−1 (66.2%)4.0[84]
Lyngbya sp. (CCNM 2053)Freezing–thawingPhosphate buffer (0.1 M), pH 8n/a22.40 mg g−13.84[4]
Lyngbya sp. (A09 DM)Freezing–thawingPotassium phosphate buffer (20 mm), pH 7.2Ammonium sulfate precipitation (Triton X-100)23.76 mg1.59[85]
Ion exchange chromatography + Gel permeation chromatography (Sephadex G-150)20.92 mg6.75
Lyngbya sp. (A09 DM)Freezing–thawingTris Cl buffer + sodium azide (3 mM), pH 8.1Ammonium sulfate precipitation + Gel permeation chromatography (Sephadex G-150)8.89 mg mL−1 (64.9%)3.7[84]
Phormidium sp. (A27DM)Freezing–thawingTris Cl buffer + sodium azide (3 mM), pH 8.1Ammonium sulfate precipitation + Gel permeation chromatography (Sephadex G-150)9.95 mg mL−1 (62.6%)3.90[84]
Phormidium persicinumFreezing–thawingDistilled waterSize exclusion chromatography5.09 µg mL−12.39[51]
Ammonium sulfate precipitation15.58 µg mL−13.99
Dialysis (phosphate buffer)32.98 µg mL−14.35
Pseudanabaena sp.Sonication + Freezing–thawingSodium phosphate buffer (0.1 mM), pH 7.0n/a0.03 mg mL−1n/a[46]
Pseudanabaena sp.Freezing–thawingPotassium phosphate buffer (100 mm), pH 7.2Ammonium sulfate precipitation5.69 mg mL−12.10[36]
Ammonium sulfate precipitation + Size exclusion chromatography5.77 mg mL−15.32
Ammonium sulfate precipitation + Size exclusion chromatography + Ion exchange chromatography5.91 mg mL−16.86
Spirulina platensisFreezing–thawingAcetate buffer [Sodium chloride (50 mM) + 0.002 M sodium azide (0.002 M)], pH 5.10Ammonium sulfate precipitation 384 µg mL−12.59[86]
Ammonium sulfate precipitation + Dialysis (Acetate buffer)299 µg mL−12.61
Ammonium sulfate precipitation + Ion exchange chromatography374 µg mL−14.57
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Tan, H.T.; Yusoff, F.M.; Khaw, Y.S.; Noor Mazli, N.A.I.; Nazarudin, M.F.; Shaharuddin, N.A.; Katayama, T.; Ahmad, S.A. A Review on a Hidden Gem: Phycoerythrin from Blue-Green Algae. Mar. Drugs 2023, 21, 28. https://doi.org/10.3390/md21010028

AMA Style

Tan HT, Yusoff FM, Khaw YS, Noor Mazli NAI, Nazarudin MF, Shaharuddin NA, Katayama T, Ahmad SA. A Review on a Hidden Gem: Phycoerythrin from Blue-Green Algae. Marine Drugs. 2023; 21(1):28. https://doi.org/10.3390/md21010028

Chicago/Turabian Style

Tan, Hui Teng, Fatimah Md. Yusoff, Yam Sim Khaw, Nur Amirah Izyan Noor Mazli, Muhammad Farhan Nazarudin, Noor Azmi Shaharuddin, Tomoyo Katayama, and Siti Aqlima Ahmad. 2023. "A Review on a Hidden Gem: Phycoerythrin from Blue-Green Algae" Marine Drugs 21, no. 1: 28. https://doi.org/10.3390/md21010028

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

Article Metrics

Back to TopTop