Chlorella vulgaris

Takashi Hasegawa. . Yasunobu Yoshikai, in Studies in Natural Products Chemistry, 2005 Protection against a chemical-induced cancer by Chlorella vulgaris administration Oral administration of Chlorella vulgaris in regard to colorectal carcinomas induced by 1, 2-dimethylhydrazine (DMH) was examined in a rat model chlorella vulgaris 1]. DMH was injected subcutaneously 20 mg/kg body weight once a week for 10 weeks. After 30 weeks, the incidence and total number of colorectal cancers were investigated.

The result showed that Chlorella vulgaris administration was markedly protective against the development of colorectal carcinomas.

In addition, the effect of Chlorella chlorella vulgaris administration on skin papillomas induced by 7, 12-dimethylbenzantracene (DMBA) was examined in mice [ 2]. Topical application of Chlorella vulgaris modulated the percent incidence of mice skin papillomas in the initiation stages of carcinogenesis by DMBA. Alice Ferreira, Luisa Gouveia, in Handbook of Microalgae-Based Processes and Products, 2020 28.4.3 Chlorella vulgaris biorefineries Chlorella vulgaris is one of the most intensively researched microalgae.

Therefore, a lot of work has been done concerning biorefinery from this microalga. Collet et al. (2011) worked on a biorefinery using C. vulgaris with lipid extraction followed by methane production from the remaining biomass.

The authors developed a Life Cycle Assessment (LCA) and demonstrated that the microalgal methane is the worst case, when compared to microalgal biodiesel and diesel, in terms of abiotic depletion, ionizing radiation, human toxicity, and possible global warming.

These negative results are mainly due to a strong demand for electricity. For the land use chlorella vulgaris, algal biodiesel also had a lesser impact than algal methane. However, algal methane is a much better option regarding acidification and eutrophication. Another work by Ehimen et al. (2011) consider the simultaneous production of biodiesel and methane in a biorefinery concept. The authors obtained biodiesel from a direct transesterification process on the Chlorella biomass, and methane through anaerobic digestion of the biomass residues.

Chlorella vulgaris maximum methane concentration obtained was 69% ( v/v), with a specific yield of 0.308 m 3 CH 4/kg VS, at 40°C and a C/N mass ratio of 8.53. The biodiesel yield was not reported ( Ehimen et al., 2011). Gouveia et al. (2014) studied the simultaneous production of bioelectricity and added-value pigments with wastewater treatment.

Fig. 28.4 represents the photosynthetic algal microbial fuel cell (PAMFC), where C. vulgaris is present in the cathode compartment and a bacterial consortium in the anode compartment.

The authors proved that the light intensity chlorella vulgaris the PAMFC power and increases the carotenogenesis process in the cathode compartment. The maximum power produced was 62.7 mW/m 2 with a light intensity chlorella vulgaris 96 μE/(m 2 s).

Fig. 28.4. C. vulgaris biorefinery: photosynthetic algal microbial fuel cell ( Gouveia et al., 2014). Another example of a C. vulgaris biorefinery is a bioethanol-biodiesel-microbial fuel cell as reported by Powell and Hill (2009). This fuel cell consisted of an integration of C. vulgaris (in the cathode) that captures the CO 2 emitted by yeast fermenters (in the anode).

The study demonstrated the possibility of generating electrical power and oil for biodiesel, in a bioethanol production facility. After oil extraction, the remaining biomass could be used for animal feed supplementation ( Powell and Hill, 2009). Eduardo Bittencourt Sydney. . Carlos Ricardo Soccol, in Biofuels from Algae, 2014 4.2.3.1 Chlorella vulgaris The first photosynthetic microbe to be chlorella vulgaris and grown in pure culture was the freshwater microalga Chlorella vulgaris. It is a spherical unicellular eukaryotic green algae that presents a thick cell wall (100–200 nm) as its main characteristic.

This cell wall provides mechanical and chemical protection, and its relation to heavy metals resistance is reported, which explains why C. vulgaris is one of the most used microorganisms for waste treatment. The uptake of carbon by C. vulgaris cells is done through the enzyme carbonic anhydrase, which catalyzes the hydration of CO 2 to form HCO 3 − and a proton.

Hirata and collaborators (1996) studied carbon dioxide fixation by this microalga, which showed important variations comparing cultivation under fluorescent lamps and sunlight. In the first case the chlorella vulgaris rate of carbon dioxide fixation was 865 mg CO 2 L − 1 d − 1; in a sunlight regimen the estimated rate achieved 31.8 mg CO 2 L − 1 d − 1.

Winajarko et al. (2008) achieved a transferred rate of 441.6 g CO 2 L − 1 d − 1 under the same cultivation conditions as Hirata et al. (1996). According to Sydney et al. (2011), in experiments using classic synthetic media and a 12-h light/dark regimen, C. vulgaris biofixation rate of carbon dioxide is near 250 mg L − 1 day − 1. Carbon fixation by Chlorella vulgaris is variable and depends, among other chlorella vulgaris, on the concentration of CO 2 in the gaseous source.

Yun et al (1997) cultivated C. vulgaris in 15% of carbon dioxide and achieved a fixation of 624 mg L − 1 day − 1; Scragg et al. (2002) achieved a fixation of 75 mg L − 1 day − 1 under CO 2 concentration of 0.03%. In the same study, Scragg tested a medium with low nitrogen and the fixation rate was 45 mg L − 1 day − 1, suggesting that nitrogen also influences carbon uptake rate.

Some studies ( Chinassamy et al., 2009; Morais and Costa, 2007) indicate that the best chlorella vulgaris of CO 2 in the gas supplied to C. vulgaris growth is about 6%. Eduardo Bittencourt Sydney. . Carlos Ricardo Soccol, in Biofuels from Algae (Second Edition), 2019 4.3.1 Chlorella vulgaris The first photosynthetic microbe to be isolated and grown in pure culture was the freshwater microalga, Chlorella vulgaris.

It is a spherical unicellular eukaryotic green alga, which presents a thick cell wall (100–200 nm) as the main characteristic. This cell wall provides mechanical and chemical protection and it is reported its relation to heavy metals resistance, which explains why C.

vulgaris is one of the most used microorganisms for waste treatment. The uptake of carbon by C. vulgaris cells is done through the enzyme carbonic anhydrase, which catalyzes the hydration of CO 2 to form HCO 3 − and a proton. Hirata and collaborators [41] studies upon carbon dioxide fixation in 1996 by this microalga showed important variations comparing cultivation under fluorescent lamps and sunlight.

In the first case, the estimated rate of carbon dioxide fixation was 865 mg CO 2 L − 1 d − 1, while in sunlight regimen, the estimated rate achieves 31.8 mg CO 2 L − 1 d − 1. Winajarko et al. [42] achieved a transferred rate of 441.6 g CO 2 L − 1 d − 1 under the same cultivation conditions of Hirata et al. [41].

In experiments using classic synthetic media and 12 h light/dark regimen, C. vulgaris biofixation rate of carbon dioxide is close to 250 mg L − 1 d − 1 [33]. Carbon fixation by Chlorella vulgaris is variable and depends mainly on the concentration of CO 2 in the gaseous source and type of reactor. In 4% CO 2 maximum CO 2 uptake is around 200 mg L − 1 d − 1 [43], while in 5% it reaches 250 mg L − 1 d − 1 [33].

Membrane-photobioreactors is described to allow CO 2 fixation up to 0.275 g L − 1 h − 1, approximately 100% superior chlorella vulgaris comparison to bubble column and airlift systems [27]. Others studies [44,45] indicate that the best concentration of CO 2 fin the gas supplied to C. vulgaris growth is about 6%. The lipid content and productivity of microalgae vary from species to species.

Among the most commonly researched species, Botryococcus braunii and Chlorella vulgaris have the highest lipid contents (% on dry matter basis), and Chlorella sorokiniana and Nannochloropsis oculata have the highest lipid productivity (mass per volume per unit time) ( Table 13.2).

Additionally, the lipid content is also dependent on other factors, such as types of reactors for cultivation (e.g., open pound, and closed photobioreactors), nutrient availability in the medium, salinity, light sources and intensity, pH, temperature, dissolved oxygen levels, etc. ( Abomohra et al., 2012; Chen et al., 2011; Gong and Jiang, 2011; Verma et al., 2010). Microalgae can be cultivated under photoautotrophic, heterotrophic, mixotrophic, and photoheterotrophic conditions.

Different strains of microalgae can grow under one or more cultivation conditions with varied lipid content and biomass productivity. Each of the cultivation conditions has advantages and disadvantages.

Species Cultivation condition Lipid content (%wt d.b.) Lipid productivity (mg/L/day) Biomass productivity (g/L/day) References Chlorophyta/green algae Botryococcus braunii UTEX 572 Phototrophic a 25.0–75.0 5.5 0.03 Yoo et al.

(2010b) Chlorella emersonii CCAP 211/11N Phototrophic b 25.0–34.0 10.3–12.2 0.04 Scragg and Bonnett (2002) Phototrophic a 29.0–63.0 8.1–49.9 0.03–0.05 Illman et al. (2000) Chlorella sp. F&M-M48 Phototrophic a 18.7 42.1 0.23 Huang et al. (2010) Phototrophic a 32.0–34.0 121.3–178.8 0.37–0.53 Chiu et al. (2009) Chlorella protothecoides CCAP 211/8D Phototrophic a 11.0–23.0 0.2–5.4 0.002–0.02 Illman et al.

(2000) Heterotrophic f 43.0–46.0 1881.3–1840.0 4.0–4.4 Yun Cheng (2009) Heterotrophic c 50.3–57.8 1209.6–3701.1 2.2–7.4 Xiong et al. (2007) Heterotrophic c,g 46.1 932.0 2.0 Huang et al.

(2010) Heterotrophic c 43.0–48.7 732.7–932.0 1.7–2.0 Huang et al. (2010) Chlorella vulgaris KCTC AG Phototrophic a 6.6 6.9 0.1 Yoo et al. (2010a) Phototrophic a 33.0–38.0 4.0 chlorella vulgaris Huang et al. (2010) Heterotrophic c,d 23.0–36.0 27.0–35.0 0.08–0.15 Huang et al. (2010) Mixotrophic c,e 21.0–34.0 22.0–54.0 0.09–0.25 Phototrophic b 5.1 7.4 0.18 Gouveia and Oliveira (2009) Dunaliella salina – 6.0–25.0 116.0 0.22–0.34 Mata et al.

(2010) Neochloris oleoabundans UTEX 1185 Phototrophic a,b 15.9–56.0 10.7–38.8 0.03–0.15 Gouveia and Oliveira (2009) Phototrophic b 29.0 26.1 0.09 Phototrophic a 7.0–40.3 38.0–133.0 0.31–0.63 Li et al. (2008) Nannochloropsis oculate NCTU-3 Phototrophic a 22.7–29.7 84.0–142.0 0.37–0.48 Chiu et al.

(2009) Phaeophyta/brown algae Pavlova lutheri CS 182 Phototrophic a 35.5 50.2 0.14 Huang et al. (2010) Pavlova salina CS 49 Phototrophic a 30.9 49.4 0.16 Isochrysis sp. F&M-M37 Phototrophic chlorella vulgaris 27.4 37.8 0.14 Isochrysis sp. (T-ISO) CS 177 Phototrophic a 22.4 37.7 0.17 Rhodophyta/red algae Porphyridium cruentum Phototrophic a 9.5 34.8 0.37 Huang et al.

(2010) Bacillariophyceae/diatoms Phaeodactylum tricornutum Phototrophic a 18.7 44.8 0.24 Huang et al. (2010) Thalassiosira pseudonana Phototrophic a 20.6 17.4 0.08 a CO 2. b Air. c Glucose. d Acetate. e Glycerol. f Jerusalem artichoke hydrolysate (JAH). g Corn powder hydrolysate (CPH). Autotrophic cultivation is the basic type of cultivation for chlorella vulgaris microalgae species. It refers to the condition of CO 2 being the carbon source for cell growth and lipid production ( Chen et al., 2011).

Solar or artificial light provides the energy for photosynthesis, which has been studied extensively on characterizing the susceptible light sources for individual lipid-rich microalgal strains ( Fork, 1979; Kendirlioğlu Şimşek and Cetin, 2017; Napolitano, 1994; Orcutt and Patterson, 1974; Ra et al., 2018; Sung et al., 2018).

Compared to other types of cultivation, autotrophic condition is more robust and has less severe contamination problems. In autotrophic cultivation of Chlorella sp., the highest lipid productivity can reach approximately 179 mg/L/day, sequestrating approximately 2% of CO 2 at 0.25 vvm of air flow ( Chiu et al., 2008, 2009).

Heterotrophic cultivation uses organic carbon as both energy and carbon sources for microalgal growth. In heterotrophic culture, cell growth is significantly influenced by nutrient concentrations in the medium and process factors ( Huang et al., 2010). A high lipid productivity of 1209.6–3701.1 mg/L/day was reported in Chlorella protothecoides cultivation under heterotrophic conditions ( Xiong et al., 2007).

Even under limited light supply, microalgae still can grow with a high yield of biomass as long as favorable carbon sources are provided ( Yoo et al., 2010a). Heterotrophic cultivation could avoid the problems associated with limited light supply in phototrophic cultivation in large-scale photobioreactors. A 40% increase in lipid content was gained in Chlorella protothecoides cultivation by just changing the condition from phototrophic to heterotrophic ( Chen et al., 2011).

Mixotrophic cultivation refers to the condition where microalgae undergo photosynthesis using both organic and inorganic carbon compounds as the carbon source for growth, under either phototrophic or heterotrophic conditions, or both. Compared to phototrophic and heterotrophic, mixotrophic cultivation is rarely used for lipid production because of the high cost of organic carbon and inadequate yield ( Mata et al., 2010).

Photoheterotrophic cultivation is the condition where the microalgae require light as the energy source and organic compounds as the carbon source, while mixotrophic cultivation can use organic compounds to serve this purpose.

Hence, photoheterotrophic cultivation needs external supply of both sugars and light at the same time ( Chojnacka and Marquez-Rocha, 2004). Although frequently used for enhanced production of some light-regulated metabolites ( Ogbonna et al., 2002), photoheterotrophic cultivation is rarely used for producing microalgal lipids. Among these four types of cultivation, heterotrophic is capable of producing a much higher lipid yield than other cultivation conditions.

However, heterotrophic culture is less resistant to bacterial contamination, especially in open-pond systems, in addition to its requirement of organic carbon as the energy source. Phototrophic cultivation can uptake CO 2 in flue gases and is commonly used in lab-scale cultivation.

Compared to the high operation cost in heterotrophic cultivation, phototrophic cultivation is relatively inexpensive ( Brennan and Owende, 2010; Demirbas, 2011), but its lipid productivity is typically low due to the low cell growth rate. Mixotrophic and photoheterotrophic cultivations are also used for lipid production; however, they are restricted by their high contamination risk and special light requirements, which result in higher operating and processing costs ( Amaro et al., 2011; Satyanarayana et al., 2011).

Tomasz Bocheński. . George Stephanopoulos, in Computer Aided Chemical Engineering, 2016 Abstract Microalgae is a promising feedstock chlorella vulgaris the chlorella vulgaris of liquid fuels to be used as replacements of oil-based fuels. In this work, we analyse a Chlorella vulgaris microalgae biorefinery that aims to produce biodiesel and renewable diesel from the lipids contained in the algae. The analysis is carried out by decomposing the biorefinery into a supply of lipids problem and a demand of lipids one.

The results indicate that by just producing fuels out of microalgae, none of the actors involved (suppliers and demanders) make profit. Reconfiguration of the processing pathway by incorporating a cell disruption step together with the addition of a valuable protein stream, displays potential for a feasible biorefinery in which all actors make profit. Species in the genera Chlorella and Scenedesmus are the most cultivated microalgae in wastewater due to their high environmental tolerance, high biomass, and lipid yield, especially Chlorella vulgaris and Scenedesmus obliquus.

The maximum biomass concentration of Chlorella vulgaris (4.25 g/L) is higher than Scenedesmus obliquus (2.3 g/L) when they are cultivated in the AD digestate. As stated in Table 16, the microalgae biomass production and growth rate highly depend on microalgae species, cultivation conditions, and wastewater compositions.

The biomass concentrations of Chlorella vulgaris and Scenedesmus obliquus have a wide range of 0.2–4.25 g/L and 0.24–2.3 g/L in response to different culture conditions, respectively. Biomass productivity is an important parameter for assessing the microalgae growth, which ranged from 29 to 890 mg/L/d for varies microalgae species.

Digestate has been regarded as a suitable medium for biomass production from different microalgae species, although the confrontation of cultivation in different types of wastewater is not straightforward. Because the wide range of cultivation conditions (e.g., cultivation mode, working volume, light intensity, and temperature) affect biomass production ( Kuo et al., 2015). Microalgae species Effluent Cultivation conditions Initial nutrient concentrations (mg/L) Biomass concentration (g/L) Biomass productivity (mg/L/d) References Chlorella vulgaris JSC-6 Swine farm effluent Mixotrophic cultivation, light intensity 200 W/m 2, 0.2 vvm CO 2 (2.5%), 12 d, fivefold dilution 264.8 NH 4 +-N 3.96 Wang et al.

(2015) Chlorella vulgaris Digestate of swine wastewater Light intensity 50, 25 °C, 7 d N/A 0.9 244.8 Deng et al. (2018) Chlorella vulgaris Digestate of swine wastewater Light intensity 300, 25 °C, air: CO 2 (97:3 v/v), 11 d, 5% wastewater 3355 TN, NH 4 +-N 2050, TP = 318.5 1.47 229 Franchino et al.

(2016) Chlorella sp. GD Swine farm effluent Mixotrophic cultivation, light intensity 300, 2% CO 2, 26 °C, 10 d 550 TN, 490 NH 4 +-N, 20 TP 9.5 681 Kuo et al. (2015) Scenedesmus obliquus Digestate of swine wastewater Light intensity 200, 25 °C, 7 d 121 TN, 129 TP (3200, 1600, 1200, 800, and 400 mg/L COD) 1.5 (400)–2.3 (1600) 124.4 (400)–311.3 (1600) Xu et al.

(2015) Giorgos Markou, Florian Monlau, in Biofuels from Algae (Second Edition), 2019 4.2 Anaerobic Digestion and Dark Fermentation Process The first authors to report on the anaerobic digestion (AD) of microalgae biomass were Golueke et al.

[95]. They investigated the anaerobic digestion of Chlorella vulgaris and Scenedesmus grown as part of a wastewater treatment chlorella vulgaris [95]. Since, several other authors have investigated the anaerobic digestion of whole microalgae [96,97] or after lipid-extraction [98–101]. For instance, Mussgnug et al. [97] have investigated the biogas production of six microalgae strains.

They reported the best production for the green alga Chlamydomonas reinhardtii with a methane potential of 387 mL CH 4 g − 1 VS whereas Scenedesmus obliquus led only to 178 mL CH 4 g − 1 VS. Bohutskyi et al. [99] have investigated the methane production of lipid-extracted Auxenochlorella protothecoides in semi-continuous anaerobic digestion (AD). They demonstrated that 0.25 L of methane per g of volatile solids can be recovered from the lipid residue, improving the total energy generation at 30%.

Prajapati and Kumar [88] worked on the closed-loop strategy by using the digestates from the AD of microalgae as nutrient source for further biomass production and reported that microalgae grew best when digestate was added at a portion of 30% either in recycled (exhausted) artificial cultivation medium or in municipal wastewater, demonstrating the partial feasibility of the closed-loop approach.

Likewise, Sforza and Barbera [102] investigated the recycling of digestates from the AD of lipid extracted microalgae and found that liquid digestates obtained after centrifugation and filtration had an insufficient amount of soluble S and P, the latter mainly due to precipitation and its removal in the solid phase. Therefore, in order to obtain an unhindered microalgal growth, the missing nutrients can either be externally supplied or recovered from the solid phase using an extraction method [101].

Dark fermentation (DF) is a simple process that requires low energy and can use various kinds of organic waste [103,104]. Algae biomass and lipid-extracted microalgae, being rich in carbohydrates, chlorella vulgaris great potential as feedstock for biohydrogen production.

In a bibliographic review, Sambusiti et al. [105] have detailed the biohydrogen potentials of various microalgae strains including Chlorella vulgaris sp., D. chlorella vulgaris sp., and Nannochloropsis sp. Among microalgae strains, Chlorella sp. has been one of the most studied with hydrogen potentials varying from 6.1 to 31.2 mL H 2 g − 1 alga [106–108].

Some studies have also investigated the biohydrogen production using lipid-extracted microalgae [109–111]. With regard to algal biomasses after lipid extraction, Yang et al. reported a biohydrogen potential of 21 mL H 2 g − 1 TS for lipid extracted Scenedesmus sp.

microalgae [110]. Additionally, the spent medium after dark fermentation has been demonstrated that could be used to grow microalgae [112]. However, so far limited research is performed in recycling the spent medium of fermented microalgae for the recycling of nutrients for further microalgae cultivation. Nonetheless, when considering the AD and DF of microalgae several limitations should be expected mostly regarding the low digestability/solubilization of biomass constituents.

Microalgae cell wall is mostly composed of organic compounds with slow biodegradability and/or low bioavailability. This resilient chlorella vulgaris wall hinders the methane and/or hydrogen yield [113].

Due to the recalcitrant structure of the microalgae cell wall, a pretreatment step could be recommended prior to biofuels conversion. To overcome these natural barriers, several types of pretreatment technologies commonly used for bioethanol production have been transferred with the purpose of increasing the biohydrogen and methane production from lignocellulosic residues and, more recently, from algal biomass too [103,113].

Pretreatments are generally divided into four categories: (i) thermal; (ii) mechanical; (iii) chemical; and chlorella vulgaris biological methods [103,113,114]. AD of microalgae could face as well ammonia toxicity due to chlorella vulgaris low C/N ratio of the chlorella vulgaris biomass [100,115]. To alleviate these limitations, co-digestion with rich carbon substrates or using carbohydrate-enriched biomass could be used [105,116,117]. G.S. Anisha, R.P.

John, in Advances in Hydrogen Production, Storage and Distribution, 2014 9.2.1 Algal species used for hydrogen production The unicellular green microalga Scenedesmus obliquus was first reported to evolve hydrogen under anaerobic conditions in chlorella vulgaris or light ( Gaffron and Rubin, 1942). Unicellular green algae such as Chlamydomonas reinhardtii ( Pantí et al., 2007), Chlorella vulgaris ( Amutha and Murugesan, 2011), Scenedesmus and nitrogen-fixing cyanobacterium Anabaena cylindrica ( Benemann and Weare, 1974) are examples of algae used for biological production of hydrogen.

Other algal species, such as Chlorococcum littorale and Platymonas subcordiformis have also been experimented on for hydrogen evolution Gloebacter PCC7421, Synechococcus PCC602, and Aphanocapsa montana ( Howarth and Codd, 1985; Serebryakova et al., 1998). We have developed a biorefinery superstructure for the production of biodiesel from the lipid contents of microalgae and the simultaneous conversion of microalgae residue into the useful products.

It includes all the major processing steps/stages for the production of biofuels from Chlorella vulgaris, and at each processing step various potential technological alternatives/options are considered. As shown in Figure 1, each option included in the superstructure is represented by two indices; the first index represents the option number and the subsequent second index represents the processing stage. The list chlorella vulgaris technological options included in the biorefinery superstructure model is given in Table 1.

The empty boxes represent the bypassing of certain processing stages, e.g., to accommodate wet lipid extraction, in-situ transesterification, etc. The detailed description of the problem data and superstructure development can be found in our previous study ( Rizwan et al., 2013). 1,1 Feed 10,5 Empty 1,2 Open pond system 1,6 Base catalyzed transesterification 2,2 Photobioreactor 2,6 Acid catalyzed transesterification 1,3 Flocculation with poly electrolyte 3,6 Enzymatic transesterification 2,3 Flocculation with NaOH 4,6 Alkaline in-situ transesterification 3,3 Flocculation with PGA 5,6 Acidic in-situ transesterification 4,3 Flocculation with chitosan acid solution 6,6 Enzymatic in-situ transesterification 5,3 Bioflocculation + Centrifugation 7,6 Empty 6,3 Centrifugation 1,7 Post transesterification purification 7,3 Auto flocculation (induced by high pH) 2,7 Empty 8,3 Microfiltration + Centrifugation 1,8 Empty 1,4 Grinding in liquid nitrogen 2,8 Empty 2,4 Drying + Ultrasound 3,8 Empty 3,4 Drying+Grinding+Microwave+Ultrasound 4,8 Enzymatic hydrolysis 4,4 Empty 5,8 Empty 5,4 Drying 1,9 Empty 1,5 Grinding assisted lipid extraction 2,9 Empty 2,5 Ultrasound assisted Extraction by [Bmim][MeSO4] 3,9 Fast pyrolysis 3,5 Ultrasound & microwave assisted lipid extraction 4,9 Fermentation 4,5 Wet lipid extraction 5,9 Anaerobic digestion 5,5 Solvent extraction (Bligh & Dyer Method) 1,10 Biodiesel 6,5 Extraction by Mod.

Bligh & Dyer Method 2,10 Glycerol 7,5 Supercritical fluid extraction 3,10 Bio-oil 8,5 Extraction by ionic liquid mixture 4,10 Bioethanol 9,5 Extraction by [Bmim][MeSO4] 5,10 Biogas • About ScienceDirect • Remote access • Shopping cart • Advertise • Contact and support • Terms and conditions • Privacy policy We use cookies to help provide and enhance our chlorella vulgaris and tailor content and ads.

By continuing you agree to the use of cookies. Copyright © 2022 Elsevier B.V. or its licensors or contributors. ScienceDirect ® is a registered trademark of Elsevier B.V. ScienceDirect ® is a registered trademark of Elsevier B.V. Contents • 1 Introduction • 2 Symbiosis • 3 Production • 4 Uses • 4.1 Bioenergy • 4.2 Food • 5 In popular chlorella vulgaris • 6 References Introduction [ edit ] C.

vulgaris is a green eukaryotic microalga in the genus Chlorella, which has been present on earth since the Precambrian period. [3] This unicellular alga was discovered in 1890 by Martinus Willem Beijerinck as the first microalga with a well-defined nucleus. [4] At the beginning of the 1990s, German scientists noticed the high protein content chlorella vulgaris C. vulgaris and began chlorella vulgaris consider it as a new food source. Japan is currently the largest consumer of Chlorella, [3] [5] both for nutritional and therapeutic purposes.

[6] Symbiosis [ edit ] Chlorella vulgaris occurs as a symbiont in tissues of the freshwater flatworms Dalyellia viridis and Typhloplana viridata. [7] Production [ edit ] Chlorella vulgaris world annual production of the various species of Chlorella was 2000 tonnes (dry weight) in 2009, with the main producers being Germany, Japan and Taiwan.

{INSERTKEYS} [3] C. vulgaris constitutes an excellent candidate for production due to its high resistance against rough conditions and invading organisms. In addition, the production of the various organic macromolecules of interest (proteins, lipids, starch) differ depending on the technique used to create biomass and can be therefore targeted. [3] Under more hostile conditions, the biomass decreases but lipids and starch contents increase.

[8] Under favourable conditions, protein content raises along with the biomass. [9] Different growth techniques have been developed. They exploit the autotrophic, heterotrophic or mixotrophic properties of C. vulgaris. Growing C. vulgaris autotrophically is illustrated by the closed photo- bioreactor. Harvesting the biomass is then generally done by centrifugation due to the high process efficiency (95% recovery). Other techniques exist, such as flocculation, filtration [10] and flotation.

[11] Chlorella sp. cultivated in digested and membrane-pretreated swine manure is capable of improving the growth medium performance of microalgae cultivations in terms of final biomass productivity, showing that algal growth depends on the turbidity of liquid digestate streams rather than on their nutrient availability.

[12] Uses [ edit ] Bioenergy [ edit ] C. vulgaris is seen as a promising source of bioenergy. It may be a good alternative to biofuel crops, like soybean, corn or rapeseed, as it is more productive and does not compete with food production. [13] It can produce large amount of lipids, up to 20 times more than crops [14] that have a suitable profile for biodiesel production.

[15] This microalgae also contains high amounts of starch, good for the production of bioethanol. [3] However, microalgal biofuels are far from competitive with fossil fuels, given their high production costs and controversial sustainability. [3] [16] Food [ edit ] The proteins content of C.

vulgaris varies from 42 to 58% of its biomass dry weight. [17] [18] [19] [20] These proteins are considered as having a good nutritional quality compared to the standard profile for human nutrition of the World Health Organization and Food and Agriculture Organization, as the algae synthesizes amino acids. [3] The algae also contains lipids (5–40% of the dry mass), [6] [17] carbohydrates (12–55% dry weight) [21] [22] and pigments with among others chlorophyll, reaching 1–2 % of the dry weight.

[23] [24] C. vulgaris contains minerals and vitamins. [3] C. vulgaris is marketed as dietary supplement, additive, [25] [26] as colourant or food emulsion. [27] They are all in the form of capsules, extracts, tablets or powder. [28] [29] They are consumed in Japan as a medical treatment. [5] [30] However, despite its high protein content, C. vulgaris is not yet widely incorporated in food products. The main reason for this is its dark green colour and its smell, which is close to that of fish.

[31] Vitamin B12, specifically in the form of methylcobalamin, has been identified in Chlorella vulgaris. [32] In popular culture [ edit ] In Alan Dean Foster's short story, "Village of the Chosen", published in The Best of Omni in 1983, a pair of scientists engineer a strain of Chlorella to be in symbiosis with humans, so humans can become photosynthetic.

References [ edit ] • ^ "Chlorella vulgaris". NCBI taxonomy. Bethesda, MD: National Center for Biotechnology Information . Retrieved 5 December 2017. Other names: synonym: Chlorella vulgaris var. viridis Chodat includes: Chlorella vulgaris Beijerink IAM C-27 formerly Chlorella ellipsoidea Gerneck IAM C-27 • ^ Duval B., Margulis L.

(1995). "The microbial community of Ophrydium versatile colonies: endosymbionts, residents, and tenants".

Symbiosis. 18: 181–210. PMID 11539474. • ^ a b c d e f g h Safi, C., Zebib, B., Merah, O., Pontalier, P. Y., & Vaca-Garcia, C. (2014). "Morphology, composition, production, processing and applications of Chlorella vulgaris: A review" (PDF). Renewable and Sustainable Energy Reviews. 35: 265–278.

doi: 10.1016/j.rser.2014.04.007. {{ cite journal}}: CS1 maint: multiple names: authors list ( link) • ^ Beijerinck, M. W. (1890). "Culturversuche mit Zoochlorellen, Lichenengonidien und anderen niederen Algen". Bot. Zeitung. 48: 781–785. • ^ a b Kitada, K., Machmudah, S., Sasaki, M., Goto, M., Nakashima, Y., Kumamoto, S., & Hasegawa, T. (2009). "Supercritical CO 2 extraction of pigment components with pharmaceutical importance from Chlorella vulgaris".

Journal of Chemical Technology and Biotechnology. 84 (5): 657–661. doi: 10.1002/jctb.2096. {{ cite journal}}: CS1 maint: multiple names: authors list ( link) • ^ a b Freitas, Hércules Rezende (2017-08-25). "Chlorella vulgaris as a Source of Essential Fatty Acids and Micronutrients: A Brief Commentary". The Open Plant Science Journal. 10 (1). doi: 10.2174/1874294701710010092.

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Part 1: Preparation and evaluation". Journal of the Science of Food and Agriculture. 90 (10): 1656–1664. doi: 10.1002/jsfa.3999. PMID 20564448. {{ cite journal}}: CS1 maint: multiple names: authors list ( link) • ^ Li, H.-B., Jiang, Y., & Chen, F. (2002). "Isolation and chlorella vulgaris of lutein from the microalga Chlorella vulgaris by extraction after saponification".

Journal of Agricultural chlorella vulgaris Food Chemistry. 50 (5): 1070–1072. doi: 10.1021/jf010220b. PMID 11853482. {{ cite journal}}: CS1 maint: multiple names: authors list ( link) • ^ Fernandes, B., Dragone, G., Abreu, A.

P., Geada, P., Teixeira, J., & Vicente, A. (2012). "Starch determination in Chlorella vulgaris—a comparison between acid and enzymatic methods". Journal of Applied Phycology. 24 (5): 1203–1208. CiteSeerX 10.1.1.1024.1758. doi: 10.1007/s10811-011-9761-5. S2CID 10404393. {{ cite journal}}: CS1 maint: multiple names: authors list ( link) • ^ Liang, S., Liu, X., Chen, F., & Chen, Z. (2004). Ang, Put O (ed.).

Current microalgal health food R & D activities in Chlorella vulgaris. Asian Pacific Phycology in the 21st Century: Prospects and Challenges. pp. 45–48. doi: 10.1007/978-94-007-0944-7. ISBN 978-94-007-0944-7. S2CID 12049767. {{ cite book}}: CS1 maint: multiple names: authors list ( link) • ^ Yamaguchi, K. (1996). "Recent advances in microalgal bioscience in Japan, with special reference to utilization of biomass and metabolites: a review".

Journal of Applied Phycology. 8 (6): 487–502. doi: 10.1007/BF02186327. S2CID 21226338. • ^ Morris, H. J., Carrillo, O. V., Almarales, Á., Bermúdez, R. C., Alonso, M. E., Borges, L., Quintana, M. M., Fontaine, R., Llauradó, G., & Hernández, M. (2009). "Protein hydrolysates from the alga Chlorella vulgaris 87/1 with potentialities in immunonutrition". Biotecnología Aplicada. 26 (2): 162–165. {{ cite journal}}: CS1 maint: multiple names: authors chlorella vulgaris ( link) • ^ Becker, E.

(2007). "Micro-algae as a source of protein". Biotechnology Advances. 25 (2): 207–210. doi: 10.1016/j.biotechadv.2006.11.002. PMID 17196357. • ^ Kumudha A, Selvakumar S, Dilshad P, Vaidyanathan G, Thakur MS, Sarada R. (2015). "Methylcobalamin--a form of vitamin B12 identified and characterised in Chlorella vulgaris". Journal of Food Chemistry. 170: 316–320. doi: 10.1016/j.foodchem.2014.08.035. PMID 25306351. {{ cite journal}}: CS1 maint: multiple names: authors list ( link) • Wikidata: Q309972 • Wikispecies: Chlorella vulgaris • AlgaeBase: 27676 • BioLib: 1259316 • EoL: 921268 • EPPO: CEJVU • GBIF: 5270961 • iNaturalist: 317909 • IRMNG: 10004584 • ITIS: 5815 • NBN: NHMSYS0021059135 • NCBI: 3077 • NZOR: 2ef3a7c6-1688-4b57-80a7-ab0defc5aa8b • uBio: 1967729 • WoRMS: 532029 Edit links • This page was last edited on 25 March 2022, at 14:59 (UTC).

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none Chlorella vulgaris Chlorella vulgaris, a green alga that is widely used as chlorella vulgaris food supplement, has good antioxidant and therapeutic properties.

From: Functional Ingredients from Algae for Foods and Nutraceuticals, 2013 Related terms: • Chlorella • Lipids • Biomass • Pretreatment • Algae • Cell Walls • Proteins • Microalgae • Sewage H.R.B. Raghavendran. . S. Rekha, in Functional Ingredients from Algae for Foods and Nutraceuticals, 2013 12.4.1 In vitro hepatoprotective role of Chlorella vulgaris and Dunaliella bardawil Chlorella vulgaris, a green alga that is widely used as a food supplement, has good antioxidant and therapeutic properties.

C. vulgaris contains a variety of compounds, including antioxidants and a glycoprotein, which may chlorella vulgaris on different pathways of tumour cell growth and survival, triggering an antagonistic effect of Bax and Bcl-2.

Apoptotic signals are generally believed to be mediated through a hierarchy of caspase activation controlled by one of the two distinct pathways that are associated with either mitochondrial caspase-8 or chlorella vulgaris. The initiating caspases then converge on the central effector caspases, caspases-3 and -7. Furthermore, results of previous studies demonstrate that the mitochondrial signaling pathway is involved in C. vulgaris-induced apoptosis of HepG2 cells ( Yusof et al., 2010).

It could be postulated that C. vulgaris does not induce the initiator caspase-8, but might activate the other initiator caspase-9. Among the ten different members of caspases identified in mammalian cells, caspase-3 may serve as a general mediator of apoptosis. When cells are undergoing apoptosis, executioner or effector caspase-3 triggers cellular proteins, such as poly (ADP-ribose) polymerase and DNA fragmentation factor resulting in the characteristic changes chlorella vulgaris apoptosis.

Caspase-3 is synthesized as a 33 kDa inactive proenzyme that requires proteolytic activation. In addition, earlier studies have shown that a high level of proenzyme of caspase-3 was present in untreated tumour cells, and that active caspase-3 gradually increased after C. vulgaris treatment, suggesting that C.

vulgaris could induce apoptosis through a caspase-3-dependent mechanism. It has been postulated that C. vulgaris mediates apoptosis in a p53-dependent manner with increased expression of Bax and decreased expression of Bcl-2 proteins in a time-dependent fashion. The activation of p53 and related family members can either enforce cell cycle arrest or induce apoptosis.

The p53-dependent responses are directed against the damaged cell to protect the organism. The rules that govern the choice between growth, arrest, and apoptosis are likely to be enforced by other proteins that can antagonize or synergize with p53 to regulate apoptosis.

Bax expression in HepG2 cells after C. vulgaris supplementation increased p53 accumulation ( Mohd Azamai et al., 2009). It has been hypothesized that C.vulgaris protects liver from CCl 4 and cadmium induced toxicity ( Shim et al., 2008; Li et al., 2011). C. vulgaris has the ability to modulate the hepatic levels of drug metabolizing enzymes and lipid peroxidation. C. vulgaris induced a significant increase in the hepatic levels of glutathione S-transferase (GST) and sulfhydryl (-SH) in foetal and neonatal systems after treatment of gestating or lactating mice.

It also modulated inhibition of cytochrome b5 (Cyt. b5), cytochrome P450 (Cyt. P450), and malondialdehyde (MDA) level in the hepatic tissue following the transplacental or translactational exposure. The observed modulation in the levels of hepatic drug metabolizing enzymes and chlorella vulgaris peroxidation suggest the chemopreventive potential of C. vulgaris via perinatal passage of active constituents and/or metabolites ( Singh et al., 1998).

Despite chlorella vulgaris importance of macroalgae in healthcare and medicine, Chlorella, a green microalga, has gained more attention in the pharmaceutical industry because of its ability to enhance the nutritional content of conventional food preparations and positively affect the health of humans and animals.

Chlorella is rich in protein, glycerol, sugars, and bases esterified to saturated or unsaturated fatty acids. Some vital fatty acids, such as omega-3 and omega-6, are abundant in Chlorella, and a high amount chlorella vulgaris PUFAs, equivalent to that found chlorella vulgaris fish chlorella vulgaris, has also been observed.

Among liver disorders, obstructive jaundice of the liver could be due to poor appetite and decreased food intake. An chlorella vulgaris study has reported the role of microalgae in intestinal barrier function and oxidative stress in obstructive jaundice, which is characterized by the absence of intestinal bile flow and impaired functioning of the reticuloendothelial system as well as immune defense systems.

Administration of Chlorella has been found to be effective in inhibiting lipid peroxidation and oxidation of GSH in bile duct-ligated rats. Both decreased malondialdehyde (MDA) and oxidized GSH are associated with lower endotoxin levels. It has been found that the primary site for de novo GSH synthesis is the liver, which supplies approximately 90% of the circulating plasma GSH.

Oxidation of hepatic GSH was observed to occur during biliary obstruction because of the significant increase in the concentration of oxidized glutathione. Although Chlorella could prevent lipid peroxidation induced by biliary obstruction, it could not protect the liver from cholestatic injury, as evaluated by conventional biochemical markers of liver damage. It is possible that a homeostatic response of the liver cells could have upregulated the synthesis of GSH, preventing further damage in bile duct-ligated rats.

The results from pathological scoring also showed that pathological changes were improved in rats after Chlorella treatment. Chlorella is capable of improving the ability of the body to remove oxygen-free radicals, mitigate membrane lipid peroxidation, and thereby effectively protect intestinal epithelial cells ( Bedirli et al., 2009). Similarly, another carotenoid-producing alga Dunaliella bardawil is being considered for use in nutraceuticals. An earlier study showed moderate but statistically significant mean serum enzyme values following D.

bardawil treatment, when compared with higher mean chlorella vulgaris in animals that received CCl 4 alone ( Vanitha et al., 2007). Likewise, lipid peroxidation, measured by thiobarbituric acid reactive substances (TBARS) activity, was slightly less elevated following algae treatment.

The study also demonstrated the protective effect of the alga against DNA strand breaks in hepatocytes, as measured by single cell gel electrophoresis.

Further, liver histopathology was less severe following 14-day D. bardawil treatment, thus supporting its apparent protective effect on hepatic oxidative injury. Horacio G. Pontis, in Methods for Analysis of Carbohydrate Metabolism in Photosynthetic Organisms, 2017 2.3.3 Preparation of Extracts from Photosynthetic Microorganisms Representative procedures for protein extraction from model unicellular algae ( Chlorella vulgaris, Chlamydomonas reinhardtii, and Ostreococcus tauri), unicellular cyanobacteria ( Synechocystis sp.

PCC 6803, Synechococcus sp. PCC 7002, Microsystis aeruginosa PCC 7806), filamentous cyanobacteria ( Anabaena ( Nostoc) sp. PCC 7120) cyanobacteria, and a single-celled flagellate photosynhtetic protist ( Euglena gracilis) will be described below. 2.3.3.1 Chlorella vulgaris Materials Chlorella vulgaris (Beijerinck strain 11468) cultures (500 mL) Washing buffer: 25 mM Tris-HCl (pH between 7.0 to 8.0) containing 1 mM EDTA and 5 mM β-mercaptoethanol Extraction buffer: 100 mM Hepes-NaOH (pH 7.5) containing 20 mM β-mercaptoethanol, 2 mM EDTA, 2% ethylenglycol, and 0.5 mM phenylmethylsulfonyl fluoride Glass powder Procedure Carry out all operations at 2–4°C.

Collect the cells by centrifugation at 3000×g for 5 min and wash twice with the washing buffer. Resuspend packed cells in five to eight times their volume with extraction buffer.

Cells can be broken either by sonication (c.0.1–1 g of fresh weight) at 40–100 W (eg, 3 pulses of 10 min) in the presence of glass powder (keeping refrigerated at 10–12°C), or by passage through a French press (c.3–15 g of fresh weight) at 25,000 psi.

Check disintegration of cells by light microscope observation. After removing cell debris by centrifugation (30,000×g for 30 min) and desalting, the extract can be used for enzyme activity determination.

Eventually, proteins can be concentrated with solid ammonium sulfate bringing the supernatant to 70% saturation, while the pH is kept at 7.0 by addition of ammonium hydroxide, or with an ultrafiltration concentration system. Desalt the protein extract before enzyme activity determination. Comments Although sonication and French press disruption are capable of disintegrating all cells, usually higher enzyme activities can be obtained when c.80% of total protein is released to the medium.

A similar procedure was used in the study of the sugar metabolism enzymes of Scenedesmus obliquus (strain 11457) and Prototheca zopfii. chlorella vulgaris Chlamydomonas reinhardtii Materials Chlamydomonas reinhardtii (strain CC124) cultures (1 L) Washing buffer: 10 mM Tris-HCl (pH 8.0) containing 1 mM EDTA chlorella vulgaris 8.0) Extraction buffer: 100 mM Chlorella vulgaris (pH 7.5) containing 20 mM β-mercaptoethanol, 2 mM EDTA (pH 7.0), 20 mM MgCl 2, 0.5 mM phenylmethylsulfonyl fluoride, 20% glycerol, and 2% ethylenglycol Glass powder Procedure Carry out all operations at 2–4°C.

Collect the cells by centrifugation at 3000×g chlorella vulgaris 5 min, wash the cell pellet with washing buffer, precipitate the cells by centrifugation at 3000×g for 5 min, and resuspend them in 20 mL of extraction buffer. Disruption of cells can be performed by two cycles of slow freezing to −80°C followed by thawing to room temperature, or with a French press at 700 psi (high).

Centrifuge the homogenate at 30,000×g for 20 min. Measure enzyme activity in the supernatant and/or proceed to further purification. Comments The Chlamydomonas cell structure can be also disrupted by brief exposure to sonication (a total of 30–60 s at 4°C at 30–40 W).

Acetone powders for enzyme extraction can also be easily prepared with Chlamydomonas cells. A procedure similar to those described for unicellular algae can be used to prepare protein extracts from Euglena gracilis (a single-celled flagellate protist). The extraction buffer consists of 50 mM Hepes-NaOH (pH 7.5) containing 20 mM β-mercaptoethanol, MgCl 2 20 mM, 2 mM EDTA, 2% ethylenglycol, 0.5 mM phenylmethylsulfonyl fluoride, and glycerol 20%, and breaking cells by sonication (0.1–1 g fresh weight of cells, 3 pulses of 40 W for 10 s) or with a French press (3–15 g fresh weight of cells at 3000 psi).

2.3.3.3 Ostreococcus tauri Materials Ostrococcus tauri (strain 0TTH0595) cultures (600 mL) Washing buffer: 25 mM Tris-HCl (pH 7.5) containing 1 mM EDTA and 5 mM β-mercaptoethanol Extraction buffer: 50 mM Tris-HCl (pH 7.5) containing 1 mM EDTA, 100 mM NaCl, 20 mM β-mercaptoethanol, and 1 mM phenylmethylsulfonyl fluoride Glass powder Procedure Carry out all operations at 2–4°C. Collect the cells by centrifugation at 3000×g for 5 min and wash twice with the washing buffer.

Resuspend packed cells in five volumes of extraction buffer. Conduct three cycles of freezing in liquid nitrogen/thawing. Then, break the cells in the presence of glass powder either with a Potter–Elvehjem homogenizer with conic-finished glass stick with Teflon tip or by sonication (using a 2 mm-tip probe, 10 cycles of 10 s at 40 W, with 20 s pauses).

Centrifuge 30,000×g for 30 min to remove cell debris and desalt the supernatant through Sephadex G-50 equilibrated chlorella vulgaris the extraction buffer. 2.3.3.4 Cyanobacteria Materials Anabaena (also named Nostoc) sp. PCC 7120 cultures Washing buffer: 25 mM Hepes–NaOH (pH 7.5) containing chlorella vulgaris mM EDTA (pH 7.0) and 5 mM β-mercaptoethanol Extraction buffer: 100 mM Hepes–NaOH (pH 7.5) containing 2 mM EDTA (pH 7.0), 2% (v/v) ethylene glycol, 20% (v/v) glycerol, 20 mM MgCl 2, 20 mM β-mercaptoethanol, and 1 mM phenylmethylsulfonyl fluoride Small glass beads (<200 μm in diameter) or glass powder Procedure A Collect cells (from c.150–300 mL) by centrifugation at 3000×g for 10 min.

Wash the pellet twice by addition of washing buffer and centrifuging each time at 3000×g for 10 min. Resuspend the packaged cells in the extraction buffer (2 mL of buffer per gram of fresh weight) and distribute in five eppendorf tubes. Add glass beads (or glass powder) to the cell paste and freeze in liquid nitrogen.

Submit the cell paste contained in each tube to five cycles of freezing and disintegration with a Teflon tipped glass rod (or with a conic-finished frosted glass stick), precooled with liquid nitrogen, placed in a vertical laboratory stirrer for 30 s each cycle. Centrifuge the extract 30,000×g for 30 min to remove cell debris. Take the supernatant with a Pasteur pipette and place the liquid in a cooled tube.

Desalt the clarified protein extract through a Sephadex G-50 bed (see Section 2.4). Procedure B Collect cells by centrifugation and add two times their volume of washing buffer. Centrifuge and weigh the pellet (c.3–8 g of cells). Homogenize the cell paste in the presence of glass beads and extraction buffer (2 mL of buffer per gram of fresh weight) in a −20°C-precooled mortar using a pestle, under liquid nitrogen. Sonicate the extract for four cycles of 30 s at 4°C, at 40 W.

Centrifuge at 30,000×g for 30 min to remove debris and then, centrifuge the supernatant at 100,000×g for 1 h. Desalt the supernatant through a disposable (or reusable) desalting column or as described in Section 2.4 before the enzyme activity assay. Comments Alternatively, protein extraction can be carried out from −80°C-stored cells that have been harvested, washed, and weighed before storage. Usually, enzyme activities are lower when the extract is prepared from frozen cells than from fresh cells.

Similar chlorella vulgaris procedures can be applied for protein extraction from Syechocystis sp. PCC 6803, Synechococcus sp. PCC 7002, or Microsyctis aeruginosa PCC 7806. A conductometric biosensor using immobilized Chlorella vulgaris microalgae as bioreceptors was used as a bi-enzymatic biosensor in ( Chouteau et al., 2004, 2005).

The use of microorganisms for multi-detection can be a good alternative, each living cell containing a large number of enzymes. Local conductivity variations caused by algal alkaline phosphatase and acetylcholinesterase activities could be detected using the following enzymatic reactions ( Fig.

6.22): [6.25] These two enzymes are known to be inhibited by distinct families of toxic compounds: heavy metals for alkaline phosphatase, carbamates and organophosphorus pesticides for acetylcholinesterase. The bi-enzymatic conductometric biosensors were tested to study the influence of heavy metal ions and pesticides on the corresponding enzymes. It appears that these biosensors are quite sensitive to Cd 2+ and Zn 2+ (limit of detection (LOD) = 10 ppb for a 30 min exposure) while Pb 2+ gives no significant inhibition, as this ion seems to adsorb preferably on albumin.

For pesticides, first experiments showed that paraoxon-methyl inhibits Chlorella vulgaris AChE, contrary to parathion-methyl and carbofuran. Biosensors were then exposed to different mixtures (Cd 2+/Zn 2+, Cd 2+/paraoxon-methyl) but no synergetic or antagonistic effect could be observed. A good repeatability could be achieved with biosensors, chlorella vulgaris the relative standard deviation did not exceed 8 %, while the response time was 5–7 min. Monica Gallo, in Encyclopedia of Food Security and Sustainability, 2019 Protein Content The first studies on cultivation of the chlorophyte Chlorella vulgaris showed a high nutritional value in proteins of this species, that could represent a major source of food for the growing human population.

However, this goal has not yet been reached due to the very high cultivation costs, which chlorella vulgaris to a non-competitive price compared to other biomass protein sources, such as soy.

The pioneering studies trace back to 1950, carried chlorella vulgaris with the eucaryote microalgae Chlorella and Scenedesmus, and later, Spirulina. These organisms chlorella vulgaris contain up to a maximum of 78% protein dry weight, which is a value much higher than that of most cultivated plants ( Angell et al., 2016). The nutritional value of microalgae and cyanobacteria has been well documented. However, it has also been shown that the digestibility and the global nutritional value of these sources are not only due to the genetic traits of an individual strain, but also depend on the technological process used for biomass production.

The protein content of algal biomass depends on the availability of nutrients, such as potassium, sodium, and nitrogen. Furthermore, the protein content is determined by the growth phase as well as by the quality of light ( Markou et al., 2014).

Because proteins are considered the most important component of algal biomass, many efforts have been directed towards determining the metabolic control of protein synthesis.

I.S. Chronakis, M. Madsen, in Handbook of Food Proteins, 2011 14.5.2 Chlorella vulgaris Evaluations have been made of the capacity chlorella vulgaris the biomass of the microalga Chlorella vulgaris as a fat mimetic and its ability as an emulsifier. Pea protein emulsions with an addition of C. vulgaris (green, 60% protein, and orange–carotenogenic, 6% protein) chlorella vulgaris prepared at different protein and oil contents ( Raymundo et al., 2005).

The addition of C. vulgaris proved to be beneficial in terms of enabling lesser oil contents in the emulsions without disturbing their structural and textural properties. Although the microalgal biomass (Cv green) has a high protein content, it cannot be used as the only emulsifier in these types of emulsion systems.

Possible interactions between pea protein and microalgal chlorella vulgaris can also contribute to the reinforcement chlorella vulgaris the emulsion structure via the formation of physical entanglements.

This effect was more significant for Cv green, which must be related to its higher chlorella vulgaris content (60% for Cv green vs.

6% for Cv orange). Chlorella vulgaris total oil content can be reduced in this case, yielding emulsions with the same rheological and sensory properties.

For this reason, it was considered that the biomass acted as a fat mimetic with a mechanism that likens that of xanthan gum. The rheological properties of the chlorella vulgaris food emulsions were also chlorella vulgaris in terms of the viscoelastic properties and steady state flow behaviour and texture properties ( Raymundo et al., 2005). The effect of addition of oil on the viscoelastic properties of the 3% pea emulsions with 2% C.

vulgaris (green and orange) can be seen in Fig. 14.10. These emulsions present mechanical spectra typical of protein-stabilised emulsions in which an elastic network develops owing to the occurrence of an extensive bridging flocculation process.

It can be observed from the dynamic measurements that, for a certain protein and microalgae concentration, a higher oil content induces a reinforcement of the emulsion structure. However, texture does not differ between Cv green and Cv orange performance as a fat mimetic; in both cases, the exponential increase of firmness observed with oil contents was not significantly different.

Overall, the above results support the potential benefit of using the Chlorella vulgaris microalgae to act as a fat mimetic, in addition to the possible advantages as a colouring and antioxidant agent. Norman P.A. Huner. . Gunnar Öquist, in Cell and Molecular Response to Chlorella vulgaris, 2002 6.2 Cold acclimation As predicted, photosynthetic adjustment during cold acclimation of the unicellular green algae Chlorella vulgaris and Dunaliella salina by growth at low temperature and moderate irradiance 5°C/150 μmol m –2 s –1 (5/150) mimics photoacclimation of these algal species grown at high light and moderate temperatures (27/2200) ( Huner et al., 1998).

Cells grown at 5/150 are indistinguishable from those grown at 27/2200 with respect to photosynthetic efficiency, photosynthetic capacity, pigmentation, Lhcb content and sensitivity to photoinhibition. These results are explained on the basis that cultures grown at either 5/150 or 27/2200 indeed are exposed to comparable excitation pressure measured as 1– qP ( Huner et al., 1998).

Similar conclusions regarding the role of excitation pressure have been reported for thermal and photoacclimation of Laminaria saccharina ( Machalek et al., 1996) and cold acclimation of the filamentous cyanobacterium, Plectonema boryanum ( Miskiewicz et al., 2000).

These results are consistent with the thesis that exposure to low temperature creates a similar imbalance in energy budget as exposure to high light. These green algal and cyanobacterial species are unable to up-regulate carbon metabolism and thus are unable to adjust the capacity of electron-consuming sinks during growth and development at low temperature ( Savitch et al., 1996; Miskiewicz et al., 2000).

As a consequence, these organisms are unable to adjust n · τ –1 significantly with respect to changes in growth temperature. Thus, to attain photostasis, these organisms adjust σ PSII · I through a reduction in the size of PSII light-harvesting complex coupled with an increased capacity for NPQ which result in a decrease in σ PSII.

The redox state of the PQ pool acts as a chloroplastic sensor regulating the expression of the nuclear chlorella vulgaris Lhcb genes whereas the trans-thylakoid ∆pH acts as a sensor regulating xanthophyll cycle activity and hence NPQ ( Wilson and Huner, 2000). In contrast, the redox sensor for Plectonema boryanum appears to be located downstream of the PQ pool and Cyt b 6/ f complex ( Miskiewicz et al., 2000).

In addition, Plectonema boryanum modulates I by accumulating the carotenoid, myxoxanthophyll, in the cell membrane. This carotenoid is a nonphotosynthetic pigment and acts as a natural sunscreen to protect the chlorella vulgaris apparatus from excess light ( Miskiewicz et al., 2000).

Cold temperate conifers such as Lodgepole pine ( Pinus contorta L.) and herbaceous cereals such winter wheat ( Triticum aestivum L.) and winter rye ( Secale cereale L.) are representative chlorella vulgaris some of the most cold tolerant plants that retain their foliage during the autumn and winter ( Levitt, 1980). This capacity to cold acclimate is an essential requirement for the development of maximum freezing tolerance, which allows them to survive the freezing temperatures chlorella vulgaris the winter.

However, these two groups of plants exhibit quite different strategies for the utilization of light energy during growth and cold acclimation (Öquist et al., 2001; Savitch chlorella vulgaris al., 2002). Cold acclimation of conifers induces the cessation of primary growth in contrast to winter cereals, which require continued growth and development during the cold acclimation period to attain maximum freezing tolerance ( Fowler and Carles, 1979).

In the context of these different growth strategies, the requirement for photosynthetic assimilates also differs considerably. Conifers exhibit a decreased requirement for photosynthetic assimilates upon the induction of dormancy and cold acclimation.

In contrast, overwintering cereals maintain a high demand for photoassimilates during cold acclimation. As a consequence of the decreased sink demand for photoassimilates, that is, a decrease in n · τ –1, conifers exhibit feedback inhibition of CO 2 assimilation ( Savitch et al., 2002). To attain photostasis under these conditions, conifers adjust their capacity and efficiency to absorb light by decreasing the content of PSII reaction centers as well as the content of Lhcb polypeptides and their associated pigments.

In addition, conifers increase their capacity for NPQ through the up-regulation of PsbS and the xanthophyll cycle with the concomitant aggregation of the major light harvesting pigment-protein complexes ( Ottander et al., 1995; Savitch et chlorella vulgaris, 2002).

Energetically, this results in a highly quenched state. Since conifers chlorella vulgaris the capacity to recover fully from this quenched state with the onset of spring ( Ottander et al., 1995), this capacity to down-regulate photosynthesis during cold acclimation is an important mechanism for the successful establishment of evergreen conifers in cold temperate and subarctic climates.

In contrast, winter cereals maintain maximum efficiency and capacity for light absorption through the light harvesting complexes and reaction chlorella vulgaris of PSII and PSI with a minimum investment in the expression of PsbS and the capacity for nonphotochemical quenching of absorbed light energy ( Huner et al., 1998; Savitch et al., 2002).

This maximizes σ PSII · I which should lead to maximum excitation pressure at low temperature. However, excitation pressure is moderated due to an increased capacity for CO 2 assimilation through the up-regulation of transcription and translation of genes coding for Rubisco, the rate-limiting enzyme for photosynthetic CO 2 fixation, as well as regulatory enzymes of cytosolic sucrose and vacuolar fructan biosynthesis ( Hurry et al., 1996). This reprogramming of carbon metabolism to match the continued absorption of light energy has a dual function: it not only maximizes the chemical energy and carbon pool necessary for the renewed growth in the spring but the accumulation of photosynthetic end-products such as sucrose provides cryoprotectants to stabilize the cells during freezing events during the winter ( Hurry et al., 1996).

The response of cereals to excitation pressure extends beyond photosynthesis to include the regulation of plant morphology and freezing tolerance ( Huner et al., 1998). Ali Bahadar. . K. Jalwana, in Handbook of Marine Microalgae, 2015 Abstract Supercritical carbon dioxide fluid extraction was carried out to extract oil from microalgae ( Chlorella vulgaris) for biofuel production.

The extraction was performed at temperature ranges of 40–80 °C and pressures of 4000–9000 psi to investigate the optimum process parameters for microalgae biomass. Static/dynamic supercritical fluid extraction (SCFE) was used and total extraction time was held at 3 h for all experiments.

Four operating process parameters (pressure, temperature, extraction time, and solvent flow rate) were optimized by response surface methodology to obtain a response (oil yield) with a central composite design. The results obtained were successfully fitted in a second-order polynomial (quadratic) model. The predicted and actual oil yields for C. vulgaris were in close agreement. The optimum yield for microalgae was 17.7 wt% (based on total oil content present in the biomass), which was in close agreement with the time-consuming n-hexane based solvent extraction.

In addition to the proper design of processes, the advances and applications of kinetic models for describing SCFE from various solid matrices are presented in this chapter. The theoretical models reviewed here include linear driving force, broken and intact cells, and shrinking cores. Komal V. Mahindrakar, Virendra K. Rathod, in Innovative and Emerging Technologies in the Bio-marine Food Sector, 2022 5.5.4.1 Ultrasonic assisted extraction of phenolic compounds from microalgae Optimized extraction by ultrasonic assistance yielded 133.7 mg GAE/g polyphenolic compounds by Folin assay from Chlorella vulgaris in 146 min at 72°C temperature when 71% ethanol as an extractant and 1:62 solid to solvent ratio was used.

Multiresponse surface methodology coupled with Central composite design was used successfully with 95% confidence level. However, ultrasonic extraction recovered enhanced yield when compared with the solvent extraction ( Mohd Nasir et al., 2017). Ultrasonic extraction of phenolics Arthrospira platensis got lesser yield than microwave and high pressure temperature extraction techniques.

This may have happened as the ultrasonic parameters were not optimized ( da Silva et al., 2017). B. Klamczynska, W.D. Mooney, in Sustainable Protein Sources, 2017 20.5.2 Safety Chlorella has been in the human food supply for centuries, and it is recognized as safe.

Chlorella vulgaris and Chlorella pyrenoidosa are considered not novel in the EU (Regulation (EC) No 258/97). Recently, Chlorella protothecoides was recognized as Generally Regarded as Safe and a “no questions” letter was received from the Food and Drug Administration (FDA) in the United States ( FDA, 2014).

The whole-cell C. protothecoides was evaluated for dietary safety in a 13-week feeding trial in rodents, as well as evaluated for food allergy potential. No adverse effects related to the whole algae protein were reported ( Szabo, Matulka, & Chan, 2013).

In another study, an enzymatic protein hydrolysate from C. vulgaris was tested in the nutritional recovery of malnourished mice. The study showed no negative effects from C. vulgaris ( Morris et al., 2011). Suphi S. Oncel. . Giuseppe Torzillo, in Handbook of Marine Microalgae, 2015 3.3 Biogas The first laboratory study of anaerobic digestion using algae biomass as a substrate was done by Golueka using Scenesmus sp.

and Chlorella vulgaris for waste treatment processes ( Golueka and Oswald, 1959). In later studies, open ponds were used to treat wastewater with microalgae, and microalgal biomass has been used for anaerobic digestion ( Oswald and Golueka, 1960; Golueka and Oswald, 1965). This work can be considered as an important milestone for the development of algal studies in terms of integrating bioprocesses and also for using the chlorella vulgaris cultivation concept.

Anaerobic digestion of microalgae was more like a literature knowledge until the 1970 fuel criris. However similar to other alternative fuels, biogas began to be considered again as a possibility, with a special emphasis on the biorefinery concept ( Ward et al., 2014). In the case of biogas, microalgal biomass is itself the main substrate ( Samson and LeDuy, 1982). Thus the cellular composition of biomass directly affects the yield of anaerobic digestion ( Droop, 1983; Sialve et al., 2009).

The complex structure of algae made it hard to estimate substrate ratios for digestion, so basic equations were adapted from conventional strategies ( Symon and Buswells, 1933; Harris and Adams, 1979; Sialve et al., 2009). Currently, rather than being a separate process, biogas production from microalgae is considered as a step in the biorefinery chlorella vulgaris ( Mussgnug et al., 2010; Oncel, 2013).

The disrupted and oil-extracted waste biomass still contains high concentrations of proteins, carbohydrates, and other compounds. Utilization of waste for biogas production could be a more feasible pathway for the waste treatment policy, too.

Until now, diverse research on microalgal biogas production has been conducted. Commercially important strains such as Spirulina sp., Chlorella sp., Tetraselmis, Scenesmus, Dunaliella salina, Arthrospira and their biogas production capacities have been investigated.

A brief description of these results appears in the latest review related to anaerobic digestion of microalgal biomass ( Ward et chlorella vulgaris, 2014). • About ScienceDirect • Remote access • Shopping cart • Advertise • Contact and support • Terms and conditions • Privacy policy We use cookies to help provide and enhance our service and tailor content and ads.

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Contents • 1 Classification • 1.1 Species • 2 Description and Significance • 2.1 Genome Structure • 3 Cell Structure, Metabolism and Life Cycle • 4 Ecology and Pathogenesis • 5 References Classification NCBI: Taxonomy http://www.ncbi.nlm.nih.gov/Taxonomy/Browser/wwwtax.cgi?mode=Info&id=3077&lvl=3&lin=f&keep=1&srchmode=1&unlock Species Eukaryota, Chlorophyta, Trebouxiophyceae, Chlorellales, Chlorellaceae, Chlorella Description and Significance "Chlorella vulgaris" is a eukaryotic, unicellular green algae.

"C. vulgaris" is estimated to have been on Earth for more than 2.5 billion years. During that time, it has needed to evolve for survival, resulting in many of the useful functions we use today and in the future(Liebke). Most of the important features deals with its ability to rapidly grow. Common practice normally involves growing populations in photobioreactors(Sacasa 2013). These chambers are consistently shaken and used to control certain aspects of metabolism in "C. vulgaris".

Variables such as media, carbonation, and light have been researched heavily to understand the best means of optimal growth. Yuvraj et al. (2016) demonstrated that photoautotrophic growth of C. vulgaris is generally limited by depletion of nutrients (especially nitrogen), light attenuation, change in pH, carbon limitation, and accumulation of photosynthetic oxygen.

Several uses of "C. vulgaris" have been researched. First, due to its high mineral and protein levels, it is used in vitamins and has even thought to be a viable food when dehydrated(Belasco 1997). It has powerful effects in boosting human health(Liebke). Secondly, many algae produce lipids through photosynthesis. This makes these organisms a viable source for biofuel. "C. vulgaris" is lipid content per biomass is approximately 42%. This is more than soybeans, sugarcane, and corn; making it a viable alternative for biodiesel(Yujie 2011).

With current technology, it can match oil prices of $63.97 per barrel. This is not even mentioning the potential to make money back through waste water treatment(Yujie 2011). Waste water is treated even in textile production. Research shows that "C. vulgaris" reduced the color dye by 41.8%, Ammonium by 44%, Phosphate by 33%, and Carbon dioxide by 33-62%(Lim 2010). "C. vulgaris" ability has also been considered for reducing emissions from power plant.

This mostly deals with the ability for rapid growth and the variety of uses. Despite the range of benefits, a negative aspect is the cost to grow "C. vulgaris". Vast areas would need to be used to make much of an impact. CO2 is a limiting resource for large quantities of C. vulgaris to grow rapidly, in exception to a coal burning power plant. Photobioreactors chlorella vulgaris often carbonated with brings an extremely high cost in energy to the equation. Genome Structure C.

vulgaris is a small, spherical algae that has a size of 5-10µm. It contains 16 chromosones consisting of a range between 0.98 Mb to 4 Mb("Chlorella vulgaris" C-169). This range is rather large due to different geographic location and being a free-living algae. Full sequencing of the chloroplast was found to contain 150,613 bp.

The total genome does not chlorella vulgaris large sections that repeat. This missing inverted repeat is found in most alga. A genomic section found in "Escherichia coli" that are responsible for cell division was found in "C. vulgaris", indicating that chloroplast division in "C.

vulgaris" resembles the division taking place in bacteria. Red and brown algae contribute no homologous genomic sections in its chloroplast which means that "C. vulgaris" is surprising more similar to land vegetation(Wakasugi 1997). When chlorella vulgaris at life history, plants are thought to be evolved from green algae.

This correlates with gene segments in "C. vulgaris" that exist in many plants that have been sequenced to date. Currently, no Mitochondrial sequencing has been performed. Cell Structure, Metabolism and Life Cycle Chlorella vulgaris of the cell wall is unique for "C. vulgaris" compared to most related green algae. It possess and enzyme-digestible cell wall which is unlike other green algae. C. vulgaris is somewhat versatile with fixing carbon.

"C. vulgaris" is a photolithoautotroph. Depending on the environment (media) it exists in, changes the byproducts resulting from metabolic processes. C. vulgaris is similar to most phototrophs because light is absorbed via the chloroplast. Green algae then fixes the CO2 into fatty acids within the cell. C. vulgaris fatty acid biomass changes with different amounts of carbohydrates present in the media.

The presence of carbohydrate causes the formation of intercellular fatty acids to have long chains. In situations with little or no carbohydrates, this green algae forms linolenic acid.

Similar to most green and red algae, C. vulgaris doesn’t not make unsaturated fatty acids(harris). This is important for the high lipid amounts found in green algae biomass. Also rather important, when C. vulgaris is grown on inorganic media, it contains more linolenic acid. This relates to the vast research about using Chlorella as a food source.

Some green algae can be high in protein and is believed to be a healthy substance for human consumption(Warren 1997). Once the appropriate fatty acids are formed oxygen is then respirated and the CO2 is stored. Ecology and Pathogenesis Similar to most green algae, C. vulgaris is a freshwater micro-algae. All Chlorella species combine attribute to the largest source of chlorophyll(Liebke). During its many years on earth, it has developed important functions that we value in regards to human health.

C. vulgaris is known to survive under certain stressors such as viruses, bacteria, fungi, and many types of pollutants.

The reason for these attributes stems from its ability to rapidly repair its DNA(JGI).

When a break occurs, C. vulgaris is able to mutate and assimilate rapidly. Because of this, researchers are trying to further understand this process to maybe benefit human health. Furthermore, when consumed it acts similar to an antibiotic.

Chlorellan(substance produced by Chlorella) also can have properties of antitumor, antiviral, and even chlorella vulgaris benefits(Liebke). Patients in need of detoxification can sometimes be treated with Chlorella(mix of species),making it a valuable tool for healthcare.

References Sacasa Castellanos, Claudia, "Batch and Continuous Studies of Chlorella Vulgaris in Photo-Bioreactors" (2013). University of Western Ontario - Electronic Thesis and Dissertation Repository.

Paper 1113. http://ir.lib.uwo.ca/etd/1113 Belasco, Warren (July 1997). "Algae Burgers for a Hungry World? The Rise and Fall of Chlorella Cuisine". Technology and Culture 38 (3): 608–34.doi:10.2307/3106856.JSTOR 3106856. Feng, Y., Chlorella vulgaris, C., & Zhang, D. (2011). Lipid production of Chlorella vulgaris cultured in artificial wastewater medium. Bioresource Technology, 102(1), 101-105.

Harris, R., Harris, P., & James, A. (1965). The fatty acid metabolism of Chlorella vulgaris. Biochimica et Biophysica Acta (BBA) - Lipids and Lipid Metabolism, 106(3), 466-473. Home - Chlorella vulgaris C-169. (n.d.). Home - Chlorella vulgaris C-169. Retrieved April 30, 2014, from http://genome.jgi-psf.org/Chlvu1/Chlvu1.home.html Liebke, F. (n.d.).

Chlorella Vulgaris - Medicinal Food. Klinghart Academy. Retrieved April 28, 2014, from http://www.klinghardtacademy.com/Articles/Chlorella-Vulgaris-Medicinal-Food.html Lim, S., Chu, W., & Phang, S.

(2010). Use of Chlorella vulgaris for bioremediation of textile wastewater. Bioresource Technology, 101(19), 7314-7322.

Nichols, B. (1965). Light induced changes in the lipids of Chlorella vulgaris. Biochimica et Biophysica Acta (BBA) - Lipids and Lipid Metabolism, 106(2), 274-279. Wakasugi, T. (1997). Complete nucleotide sequence of the chloroplast genome from the green chlorella vulgaris Chlorella vulgaris: The existence of genes possibly involved in chloroplast division.

Proceedings of the National Academy of Sciences, 94(11), 5967-5972.

Yuvraj, Vidyarthi, A.S., & Singh, J. (2016). Enhancement of Chlorella vulgaris cell density: Shake flask and bench-top photobioreactor studies to identify and control limiting factors. Korean Journal of Chemical Engineering, 33(8), 2396-2405. Page authored by David Wells, student of Prof. Jay Lennon at IndianaUniversity.
This article is about the genus of algae.

For the bacterial infection, see Cholera and Vibrio cholerae. Chlorella Chlorella vulgaris Scientific classification (unranked): Viridiplantae Division: Chlorophyta Class: Trebouxiophyceae Order: Chlorellales Family: Chlorellaceae Genus: Chlorella M.Beijerinck, chlorella vulgaris Species • Chlorella autotrophica • Chlorella coloniales • Chlorella lewinii • Chlorella minutissima • Chlorella pituita • Chlorella pulchelloides • Chlorella pyrenoidosa • Chlorella rotunda • Chlorella singularis • Chlorella sorokiniana • Chlorella variabilis • Chlorella volutis • Chlorella vulgaris Chlorella is a genus of about thirteen species of single- celled green algae belonging to the division Chlorophyta.

The cells are spherical in shape, about 2 to 10 μm in diameter, and are without flagella. Their chloroplasts contain the green photosynthetic pigments chlorophyll-a and -b. In ideal conditions cells of Chlorella multiply rapidly, requiring only carbon dioxide, water, sunlight, and a small amount of minerals to reproduce.

[1] The name Chlorella is taken from the Greek χλώρος, chlōros/ khlōros, meaning green, and the Latin diminutive suffix ella, meaning small. German biochemist and cell physiologist Otto Heinrich Warburg, awarded with the Nobel Prize in Physiology or Medicine in 1931 for his research on cell respiration, also studied photosynthesis in Chlorella. In 1961, Melvin Calvin of the University of California received the Nobel Prize in Chemistry for his research on the pathways of carbon dioxide assimilation in plants using Chlorella.

Chlorella has been considered as a source of food and energy because its photosynthetic efficiency can reach 8%, [2] which exceeds that of other highly efficient crops such as sugar cane. Contents • 1 As a food source • 1.1 History • 1.2 Current status • 1.2.1 Production difficulties • 2 Use in carbon dioxide reduction and oxygen production • 3 Dietary supplement • 4 Health concerns • 5 Aquaria • 6 See also • 7 References As a food source [ edit ] Chlorella is a food source because it is high in protein and other essential nutrients; when dried, it is about 45% protein, 20% fat, 20% carbohydrate, 5% fiber, and 10% minerals and vitamins.

Mass-production methods are now being used to cultivate it in large man-made circular ponds. It is commonly used as a superfood and chlorella vulgaris be found as an ingredient in certain liquid-based cocktails. When first harvested, Chlorella was suggested as an inexpensive protein supplement to the human diet. Advocates sometimes focus on other supposed health benefits of the algae, such as claims of weight control, cancer prevention, and immune system support.

[3] According to the American Cancer Society, "available scientific studies do not support its effectiveness for preventing or treating cancer or any other disease in humans". [4] Under certain growing chlorella vulgaris, Chlorella yields oils that are high in polyunsaturated fats— Chlorella minutissima has yielded eicosapentaenoic acid at 39.9% of total lipids.

[5] Some companies producing Chlorella commercially as a human food include TerraVia (formerly Solazyme) and Allma. [6] History [ edit ] Following global fears of an uncontrollable human population boom during the late 1940s and the early 1950s, Chlorella was seen as a new and promising primary food source and as a possible solution to the then-current world hunger crisis.

Many people during this time thought hunger would be an overwhelming problem and saw Chlorella as a way to end this crisis by providing large amounts of high-quality food for a relatively low cost.

[3] Many institutions began to research the algae, including the Carnegie Institution, the Rockefeller Foundation, the NIH, UC Berkeley, the Atomic Energy Commission, and Stanford University.

Following World War II, many Europeans were starving, and many Malthusians attributed this not only to the war, but also to the inability of the world to produce enough food to support the increasing population.

According to a 1946 FAO report, the world would need to produce 25 to 35% more food in 1960 than in 1939 to keep up with the increasing population, while health improvements would require a 90 to 100% increase. [3] Because meat was costly and energy-intensive to produce, protein shortages were also an issue. Increasing cultivated area alone would go only so far in providing adequate nutrition to the population. The USDA calculated that, to feed the U.S.

population by 1975, it would have to add 200 million acres (800,000 km 2) of land, but only 45 million were chlorella vulgaris. One way to combat national food shortages was chlorella vulgaris increase the land available for farmers, yet the American frontier and farm land had long since been extinguished in trade for expansion and urban life.

Hopes rested solely on new agricultural techniques and technologies. Because of these circumstances, chlorella vulgaris alternative solution was needed. To cope with the upcoming postwar population boom in the United States and elsewhere, researchers decided to tap into the chlorella vulgaris sea resources.

Initial testing by the Stanford Research Institute showed Chlorella (when growing in warm, sunny, shallow conditions) could convert 20% of solar energy into a plant that, when dried, contains 50% protein. [3] In addition, Chlorella contains fat and vitamins. The plant's photosynthetic efficiency allows it to yield more protein per unit area than any plant—one scientist predicted 10,000 tons of protein a year could be produced with just 20 workers staffing a 1000-acre chlorella vulgaris 2) Chlorella farm.

[3] The pilot research performed at Stanford and elsewhere led to immense press from journalists and newspapers, yet did not lead to large-scale algae production.

Chlorella seemed like a viable option because of the technological advances in agriculture at the time and the widespread acclaim it got from experts and scientists who studied it. Algae researchers had even hoped to add a neutralized Chlorella powder to conventional food products, as a way to fortify them with chlorella vulgaris and minerals. [3] When the preliminary laboratory results were published, the scientific community at first backed the possibilities of Chlorella. Science News Letter praised the optimistic results in an article entitled "Algae to Feed the Starving".

John Burlew, the editor of the Carnegie Institution of Washington book Algal Culture-from Laboratory to Pilot Plant, stated, "the algae culture may fill a very real need," [7] which Science News Letter turned into "future populations of the world will be kept from starving by the production of improved or educated algae related to the green chlorella vulgaris on ponds." The cover of the magazine also featured Arthur D.

Little's Cambridge laboratory, which was a supposed future food factory. A few years later, the magazine published an article entitled "Tomorrow's Dinner", which stated, "There is no doubt in the mind of scientists that the farms of the future will actually be factories." Science Digest also reported, "common pond scum would soon become the world's most important agricultural crop." However, in the decades since those claims were made, algae have not been cultivated on that large of a scale.

Current status [ edit ] Since the growing world food problem of the 1940s was solved by better crop efficiency and other advances in traditional agriculture, Chlorella has not seen the kind of public and scientific interest that it had in the 1940s. Chlorella has chlorella vulgaris a niche market for companies promoting chlorella vulgaris as a dietary supplement. [3] Production difficulties [ edit ] Chlorella culture, L'Eclosarium, Houat. The experimental research was carried out in laboratories, rather than in the field, and scientists discovered that Chlorella would be much more difficult to produce than previously thought.

To be practical, the algae grown would have to be placed either in artificial light or in shade to produce at its maximum photosynthetic efficiency. Chlorella vulgaris, for the Chlorella to be as productive as the world would require, it would have to be grown in carbonated water, which would have added millions to the production cost.

A sophisticated process, and additional cost, was required to harvest the crop, and, for Chlorella to be a viable food source, its cell walls would have to be pulverized. The plant could reach its nutritional potential only in highly modified artificial situations. Another problem was developing sufficiently palatable food products from Chlorella. [8] Although the production of Chlorella vulgaris looked promising and involved creative technology, it has not to date been cultivated on the scale some had predicted.

It has not been sold on the scale of Spirulina, soybean products, or whole grains. Costs have remained high, and Chlorella has for the most part been sold as a health food, for cosmetics, or as animal feed. [8] After a decade of experimentation, studies showed that following exposure to sunlight, Chlorella captured just 2.5% of the solar energy, not much better than conventional crops.

[3] Chlorella, too, was found by scientists in the 1960s to be impossible for humans and other animals to digest in its natural state due to the tough cell walls encapsulating the nutrients, which presented further problems for its use in American food production. [3] Use in carbon dioxide reduction and oxygen production [ edit ] See also: Carbon sequestration In 1965, the Russian CELSS experiment BIOS-3 determined that 8 m 2 of exposed Chlorella could remove carbon dioxide and replace oxygen within the sealed environment for a single human.

The algae were grown in vats underneath artificial light. [9] Dietary supplement [ edit ] Chlorella in pill form. Chlorella vulgaris is consumed as a dietary supplement. Manufacturers of Chlorella products falsely assert that it has purported health effects, [10] including an ability to treat cancer, [11] for which the American Cancer Society stated "available scientific studies do not support its effectiveness for preventing or treating cancer or any other disease in humans".

[11] The United States Food and Drug Administration has issued warning letters to supplement companies for falsely advertising health benefits of consuming chlorella products, such chlorella vulgaris one company in October chlorella vulgaris.

[12] Health concerns [ edit ] A 2002 study showed that Chlorella cell walls contain lipopolysaccharides, endotoxins found in Gram-negative bacteria that affect the immune system and may cause inflammation.

[13] chlorella vulgaris [15] However, more recent studies have found that the lipopolysaccharides in organisms other than Gram-negative bacteria, for example in cyanobacteria, are considerably different from the lipopolysaccharides in Gram-negative bacteria. [16] Aquaria [ edit ] Chlorella can be a nuisance organism in freshwater aquaria. [ citation needed] See also [ edit ] Wikimedia Commons has media related to Chlorella.

• Calvin cycle • List of ineffective cancer treatments • Quorn (food product): made from mycoprotein • Soyuz 28, a 1978 space mission which included experiments on Chlorella • Spirulina (dietary supplement) • Chlorellosis, a disease caused by the infection of Chlorella.

References [ edit ] • ^ Scheffler, John (3 September 2007). "Underwater Habitats". Illumin. 9 (4). • ^ Zelitch, I. (1971). Photosynthesis, Photorespiration and Plant Productivity. Academic Press. p.

275. • ^ a b c d e f g h i Belasco, Warren (July 1997). "Algae Burgers for a Hungry World? The Rise and Fall of Chlorella Cuisine". Technology and Culture. 38 (3): 608–34. doi: 10.2307/3106856. JSTOR 3106856. • ^ "Chlorella". American Cancer Society. 29 April 2011. Archived from the original on 5 September 2013. Retrieved 23 August 2013. • ^ Yongmanitchai, W; Ward, OP (1991). "Growth of and omega-3 fatty acid production by Phaeodactylum tricornutum under different culture conditions". Applied and Environmental Microbiology.

57 (2): 419–25. Bibcode: 1991ApEnM.57.419Y. doi: 10.1128/AEM.57.2.419-425.1991. PMC 182726. PMID 2014989. • ^ Rack, Jessie (11 August 2015). "Protein Goes Chlorella vulgaris Can Algae Become The Next Soy?". NPR. Retrieved 15 January 2021. • ^ Burlew, John, ed. (1953). Algal Culture-from Laboratory to Pilot Plant. Carnegie Institution of Washington. p. 6. ISBN 978-0-87279-611-9. • ^ a b Becker, E.W. (2007). "Micro-algae as a source of protein". Biotechnology Advances.

25 (2): 207–10. doi: 10.1016/j.biotechadv.2006.11.002. PMID 17196357. • ^ "Russian CELSS Studies". Space Colonies. Permanent. Retrieved 3 November 2012. • ^ Sun Chlorella, Going Green from the Inside Out – LA Sentinel • ^ a b "Chlorella". American Cancer Society. 29 April 2011. Archived from the original on 5 September 2013. Retrieved 13 September 2013. • ^ William A. Correll (20 October 2020). "FDA Warning Letter to ForYou Inc". Inspections, Compliance, Enforcement, and Criminal Investigations, US Food and Drug Administration.

Retrieved 9 March 2021. • ^ Sasik, Roman (19 January 2012). "Trojan horses of Chlorella 'superfood' ". Robb Wolf. • ^ Armstrong, PB; Armstrong, MT; Pardy, RL; Child, A; Wainwright, N (2002). "Immunohistochemical demonstration of a lipopolysaccharide in the cell wall of a chlorella vulgaris, the green alga, Chlorella".

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