Enzyme assays were carried at 60 DAS, which corresponded to 10 days pre-tasseling. Any kind of environmental stresses like nutrient or moisture deficiency at this late vegetative growth stage greatly affects the crop performance and therefore this time was chosen to assess any stress response due to the different nutrient management treatments (McWilliams et al., 1999). For carrying out enzyme assays, leaf tissues (0.1 g fresh weight) were collected at 60 DAS and homogenized using a mortar and pestle with liquid nitrogen. The tissue was then extracted with 50 mM Tris extraction buffer (pH 7.5) containing 0.2% (w/v) Triton X-100, 0.1 mM EDTA, 1 mM polymethylsulphonylfluoride, and 2 mM dithiothreitol for glutathione reductase (GR) activity. For ascorbate peroxidase (APX) assay, instead of Tris, 50 mM potassium phosphate buffer (pH 7.0) amended with 2 mM ascorbate was used for extraction. Both the homogenates were vortexed and centrifuged at 20,000 × g for 30 min at 4◦C. The supernatants were collected and stored at −80◦C until further analysis. Protein content was estimated according to Bradford method (1976) using bovine albumin as a standard and measured using UV–Vis spectrophotometer. The APX (EC <IP_ADDRESS>) and GR (EC <IP_ADDRESS>) activity were measured according to Nakano and Asada (1981) and Edwards et al. (1990), respectively. The concentration of oxidized ascorbate was calculated using an extinction coefficient of 2.8 mM−1 cm−1 , while GR activity was calculated using an extinction coefficient of 6.22 M−1 cm−1 for NADPH. One unit of APX is defined as 1 mmol ascorbate oxidized per min per ml and expressed in units per mg protein. One unit of GR is defined as 1µmol NADPH oxidized per min per ml at 25◦C and expressed in units per mg protein.

#### Analytical Methods

The total nitrogen content in the grain was estimated by standard semi-micro-Kjeldahl method (Thimmaiah, 1999) and expressed in terms of crude protein using a conversion factor of 6.25 (Jones, 1941), while Na and K content was determined using flame photometry (Flame photometer 128, Systronics, India) according to the method prescribed by Jackson (1973). Phosphorus (P) content in grains was determined by the Vanado-Molybdate yellow method (Jackson, 1973) following wet digestion with HNO3–HClO4 (10:4) di-acid mixture. Carbohydrate in grains was estimated by the method of Dubois et al. (1956). Fat content in grains was determined using petroleum ether (at 70◦C) in a Soxhlet apparatus (AACC, 1987)

The available N in soil was analyzed by alkaline permanganate method (Subbiah and Asija, 1956). The available K in soil was extracted by neutral normal ammonium acetate method (Hanway and Heidel, 1952) and available P by sodium bicarbonate as per Olsen et al. (1954) and estimated by flame photometer (Flame photometer 128, Systronics, India) and spectrophotometer (Varian Cary-50 Bio, Varian Inc., USA), respectively. Organic carbon was analyzed according to the method of Walkley and Black (1934).

#### Statistical Analysis

The treatments were subjected to statistical analysis by analysis of variance (ANOVA) using InfoStat statistical software V 2012 (Di Rienzo et al., 2011). Post-hoc comparison of means was carried out using Fisher's Least Significant Difference (LSD) at the probability level of 5% and presented in the tables.

#### RESULTS

#### Effects on Growth Parameters of Maize

The plant heights recorded in different treatments at 20, 40, 60, and 80 DAS are presented in **Table 2**. The lowest plant height was recorded at all the growth stages under the absolute control treatment where no fertilizer was applied (T1). Compared to RRF, the treatment receiving N nutrition equally through LMBR from Chlorella and chemical source (T5) gave an initial fillip to the plant height as early as 20 DAS that was maintained significantly till 80 DAS (P < 0.001). However, all the treatments employing LMBR from Lyngbya were at par with RRF with respect to plant height at all the days of observations, except T8 at 60 DAS; however, there was no change in the number of nodes due to any of the treatments. Interestingly, the length of the tassels formed by applying either of the LMBRs at any of the substitution rates was markedly higher than that formed under RRF (P < 0.05), except that in the treatments employing the lowest level of LMBR from Chlorella and the highest level of that from Lyngbya, where were at par.

Leaf chlorophyll index and net photosynthetic rates recorded at different growth stages are tabulated in **Table 3**. The chlorophyll index of LMBR treated plants was significantly higher than that of RRF only at 60 DAS (P < 0.001) and that also in the T4 and T5 treatments in which 75 and 50% chemical N substitution was done through LMBR from Chlorella and in T10 which replaced 25% of N. All these three treatments were at par. The treatment T10 maintained this advantage even at 100 DAS. T5 and T10, which recorded the highest chlorophyll index at 60 DAS using Chlorella and Lyngbya LMBR, respectively, also recorded the significantly highest net photosynthetic rate (P < 0.001) compared to RRF for the respective LMBR at the corresponding time of observation. Both these treatments recorded higher rate of photosynthesis at the early growth stage (30 DAS) as well, however, they were at par with T4 and T6.

The DMA in stem, leaves, and roots and their total at harvest are presented in **Table 4**. Supplying 75% of N through LMBR from Chlorella (T4) resulted in the maximum DMA in stem and roots which were significantly higher over RRF (P < 0.001). The leaf dry biomass in this treatment was nevertheless at par with RRF. However, this treatment recorded the significantly highest (P < 0.001) total dry matter in the vegetative parts than that in any of the other treatments. There was a marked decline in the DMA in stem and root in all the treatments receiving Lyngbya LMBR, although interestingly, none of them except T10 showed decline in leaf dry weight compared to RRF. This, however, resulted in overall significant decline in the total vegetative DMA, compared to RRF. Further, all the Lyngbya substitution treatments were found at par with respect to total DMA.

*Values followed by different alphabets in columns are significantly different at p* < *0.05 by LSD.*

*Values followed by different alphabets in columns are significantly different at p* < *0.05 by LSD.*

TABLE 4 | Effect of different LMBR treatments on dry matter accumulation in above-ground vegetative parts and roots of maize plant.

T10 28.66 ± 2.78cd 41.80 ± 4.22a 45.58 ± 4.30a 40.26 ± 2.75b 38.52 ± 3.58a 37.65 ± 2.82bc 37.83 ± 0.91a

*Values followed by different alphabets in columns are significantly different at p < 0.05 by LSD.*

#### Stress Enzymes

Preliminary studies on LMBR treated plants also revealed positive results with respect to the stress enzymes studied which was carried out with a view to assess whether the application of the LMBRs on maize plants elicit any stress response. Two key enzymes, namely, ascorbate peroxidase (APX) and glutathione reductase (GR) activities were assayed (**Figure 1**). It was found that the treatment receiving no external nutrition recorded the highest APX activity, which was significantly higher than that in RRF (P = 0.011). Higher APX is suggestive of greater oxidative stress. All the other LMBR treatments had significantly lower activity of this enzyme (P ≤ 0.027) than RRF except in T3, which received N nutrition solely through LMBR of Chlorella. Interestingly, the APX activity decreased with decreasing dose of Chlorella LMBR. However, all the treatments employing LMBR of Lyngbya recorded significantly lower activity (P < 0.001) over RRF, which were at par with each other and thus was suggestive of decreased oxidative stress. GR activity was lowest in T1, which was, however, at par to RRF. Among the Chlorella treatments, only T5 recorded significantly higher values of GR activity over RRF (P = 0.015), while the rest were at par. All the Lyngbya treatments except that utilizing LMBR solely as N source recorded significantly higher GR activity over RRF, the highest being in T9, which was superior to all the other nine treatments (P < 0.001).

#### Effect on Yield Attributes and Grain Yield

The length, width, fresh weight as well as the dry weight of the maize cobs with or without the outer husk remained statistically similar in all the LMBR treatments comparable to that in the RRF. Similarly, the same trend was found in number of kernel rows per cob and number of kernels (grains) per row, which determine the total number of grains formed (**Table 5**). Notably, T8 treatment that substituted 75% of chemical N by Lyngbya LMBR had significantly heavier grains compared to RRF (P = 0.008), which however was at par with all other LMBR treatments except in T7 where 100% of N was supplied by LMBR from Lyngbya. The highest grain weight (65.16 g plant−1 ) was measured in T10 where 25% of the total N was supplied by Lyngbya derived LMBR and rest by urea, which was closely followed by that (63.48 g plant−1 ) measured in T5 treatment where N was supplied to the plants in equal proportion through Chlorella LMBR and urea. However, both these treatments were found at par with other levels of substitutions with LMBRs from either of the algae and were also equivalent to RRF. The yield and its attributes were found the lowest in the absolute control treatment where no external nutrition was supplied.

#### Effect on Nutritional Quality of Maize

The carbohydrate, crude protein, and crude fat in maize grains were determined and are presented in **Table 6**. The carbohydrate content was the maximum where 75% of N was supplied through LMBR of Chlorella. This treatment was at par to all the other combinations of Chlorella LMBR, but was significantly superior to RRF (P = 0.018) as well as all other Lyngbya treatments (P ≤ 0.021). However, all the Lyngbya treated plants were at par with RRF treated ones. There was no change in crude protein content in grains due to either of the two LMBR treatments as

compared to that in RRF. All the algal biomass treated plants were equivalent with respect to crude fat content, except T3 where 100% N was supplied through Chlorella derived LMBR. With respect to nutritional quality, it was found that there was no change in nitrogen content of the grains due to any of the LMBR treatments when compared to RRF. Compared to RRF, T3, and T10 had the significantly (P < 0.05) highest phosphorus content in the Chlorella and Lyngbya derived LMBR treatments, respectively. However, all the LMBR treatments were at par with each other with respect to P content in grains. Compared to RRF, there was significantly (P < 0.001) higher potassium uptake in grains in all the LMBR treatments except where N was entirely supplied through Chlorella derived LMBR. There was a marked increase in sodium as well as potassium content of grains in T10 treatment when compared to that in RRF.

# Effects on Physico-chemical Properties of Soil at Harvest

The changes in physico-chemical properties brought out due to the various treatments are presented in **Table 7**. It was found that there was no significant difference in pH, EC, organic carbon, and available N content in the LMBR treated soil when compared to RRF treated ones. Other than the treatments supplying 100% (T3) and 75% (T4) N through Chlorella derived LMBR, all other treatments were at par to RRF with respect to available P content. The two aforesaid treatments were significantly (P ≤ 0. 006) better than all other treatments, but at par with each other. T4 and T5 in case of Chlorella based LMBR and T8 and T10 in case of Lyngbya based LMBR treatments recorded significantly lower available K in soil at harvest, while all other treatments were at par with RRF.

# DISCUSSION

Among the LMBR from the two algal species, Chlorella derived one apparently was found to be more responsive in improving the plant height as no response was found through LMBR from Lyngbya when compared to RRF (**Table 2**). Nitrogen nutrition

TABLE 5 | Effect of different LMBR treatments on yield attributes and grain yield of maize.

*Values followed by different alphabets in columns are significantly different at p < 0.05 by LSD.*

*Values followed by different alphabets in columns are significantly different at p* < *0.05 by LSD.*

TABLE 7 | Effect of different LMBR treatments on physico-chemical properties of soil at harvest.

*Values followed by different alphabets in columns are significantly different at p* < *0.05 by LSD.*

derived equally from both organic and chemical sources seemed to be the best balanced dose that sustained higher plant height relative to RRF starting from as early as 20 DAS till 80 DAS. It was observed that substitution of 100–50% of total N (T3–T5) through Chlorella LMBR treatments maintained the plant height at par with RRF or was even higher until 60 DAS. However, lowest rate of substitution in T6 (25%) brought about a diminution suggesting the role of the algal biomass toward sustained supply of nutrients or other active factors to the plants.

With respect to total DMA in the above ground vegetative parts and roots, contrasting results were found between LMBR from Chlorella and Lyngbya (**Table 4**). Whereas, 100 to 50% N substitutions through Chlorella biomass resulted in higher or equivalent DMA, similar levels of substitution by Lyngbya resulted in significant decrease when compared to that in RRF, suggesting the presence of one or more factor in the latter that are responsible for diminution of growth. Substitution of 25% N through either of the LMBR invariably brought a significant decrease in the aforesaid total DMA, connoting the requirement of Chlorella in higher amounts and confirming the retardant activity of LMBR from Lyngbya even at lower doses of substitution. The differential response between the two algal LMBRs might be on the account of their composition, which thus needs to be fully unraveled. This may indicate the presence of some toxic ingredients in Lyngbya species that negatively affect the plant growth. There are reports of toxicity of the microalgal extracts affecting the growth of plants (Romanowska-Duda and Tarczyñska, 2002) thus validating the argument (Grzesik and Romanowska-Duda, 2015) that untested algal residues should not be employed for treatment on plants.

Interestingly, there was no decrease in DMA in leaf due to the Lyngbya treatments except that in T10 where least level of substitution (25%) was done, which was also equivalent to all the other Lyngbya treatments. T4 fared the best among all the treatments for total DMA which could be primarily attributed to increase in stem and root biomass, while T5 formed the highest leaf biomass, which was although similar to that in RRF. The highest plant height and leaf biomass found in the LMBR treatment using biomass of Chlorella in equal proportion as chemical N (T5) at 60 DAS can be well-corroborated by the leaf chlorophyll index and net photosynthetic rate recorded during this time (**Table 3**), which were found to be the highest among all the Chlorella treatments and significantly superior to RRF. Notably, in spite of lower DMA in the vegetative biomass in all the Lyngbya treatments, compared to RRF, there was no significant decrease in net photosynthetic rate at the leaf level, as evident from its values recorded at 30 and 60 DAS (**Table 3**). Rather, it was significantly higher than all other treatments at 60 DAS. This treatment also showed the best chlorophyll index among all the Lyngbya LMBR treatments at all the times and was significantly higher than RRF before (60 DAS) as well as after tasseling stage (100 DAS) of the plant. Grzesik and Romanowska-Duda (2015) also similarly reported intensification of several metabolic processes such as net photosynthesis and other gas exchange parameters, stability of cytomembranes which eventually led to enhanced shoot and root dry weight of willow plants by application of sonicated biomass of two species of Cyanobacteria (Microcystis sp. and Anabaena sp.) and one green algae (Chlorella sp.).

Thus, the observations recorded on plant height, leaf chlorophyll index, net photosynthetic rate and above ground vegetative DMA in the treatments using LMBR from Chlorella suggest that any level of substitution resulted in either maintaining the growth parameters or improving it in some of the treatments. This possibly explained for the equivalent grain yield obtained from all the Chlorella derived LMBR treatments, compared to RRF. Besides, microalgae, which itself is photosynthetic, have all the plant essential elements in its biomass (**Table 1**) which are left in the residue after oil extraction and upon decomposition are available to the plants fertilized with it. Even though there was a detrimental effect on the total DMA by using LMBR from Lyngbya (**Table 4**), the maintenance of similar or higher photosynthetic rate leading to attainment of similar leaf biomass might have resulted in better partitioning of photosynthates to the grains, thus explaining for the similar grain yields despite the mentioned limitations. Evidently, even though the grain yield of maize was found similar to RRF in all the LMBR treatments, irrespective of the species of algae it was derived from, there were differences in the quality of grains due to the different treatments (**Table 6**). Whereas, there was no change in the protein content, T4 was nutritionally superior in terms of carbohydrate content, while T3 was superior with respect to fat and phosphorus content when compared over RRF. T10 was superior in terms of P, K as well as Na content when compared to RRF.

Strikingly, it was found that all the LMBR treatments except T3 had marked enhancement in the potassium content. Potassium has a prominent role in imparting stress tolerance in plants (Zörb et al., 2014). Another notable observation was that all the LMBR treatments exhibited a significantly longer tassel compared to that in RRF except that in T6 and T7, which were also numerically superior (**Table 1**). A longer tassel has been reported to be desirable under stress environment to ensure sufficient and extended pollen availability (Sofi, 2007). Even though the plants in the present experiment were not subjected to any specific stress, nevertheless, plants do experience biotic and abiotic stress, even at different times of the day during its growth and reproductive phase when grown under uncontrolled conditions (Layek et al., 2015). Such stress may be due to temperature extremities during day or night, incipient wilting during mid-day, variation in soil moisture, pest attack, etc., all of which could induce instantaneous production of reactive oxygen species (ROS). The results obtained from our studies on ROS scavengers like APX suggests that LMBRs may also be important for imparting tolerance under environmental stress and may be explored in the future on this aspect. The presence of bioelicitors in the LMBRs may also be explored which can perhaps trigger plant defense pathways in response to biotic and abiotic stresses. Such elicitors have been reported in other green algae like Ulva lactuca (Grzesik and Romanowska-Duda, 2015).

Evidently, no detrimental effect was found in the pH and EC of the soil due to the various LMBR treatments (**Table 7**). Although, there was no change in organic carbon content of soil, continuous application of the carbon rich LMBR may have a significant beneficial effect in the long run. The treatments employing 100% (T3) and 75% (T4) N through Chlorella-derived LMBR had a striking effect in improving available phosphorus in soil which may be ascribed to the improved microbial and associated biochemical activity of such carbonaceous materials (Anand et al., 2015). Interestingly, T3 also had the highest P content in the grains, all of which connote toward greater mineralization of soil phosphorus and also the higher mobilization of phosphorus to the grains of soil, which needs to be investigated in future. Grzesik and Romanowska-Duda (2014) recently reported that application of microalgal residues to corn plants significantly enhanced the activity of phosphatase enzyme, which is responsible for distribution of P in plants. The use of LMBR may also confer another advantage by preventing the loss of soil nutrients by way of leaching or volatilization, thus being a sustainable release nutrient source. Mulbry et al. (2005) reported that only 3% of algal N was present as plant available N in soil at the time of application of dried algal biomass (day 0). Similar to our studies, they also found that dried algal biomass was equivalent to fertilizer in supplying N and P to cucumber and corn. It was further conjectured that NH3 volatilization would not occur by use of dried algal biomass as manure in contrast to the use of chemical fertilizers.

Chemical solvent extraction is the most common method to effectively extract lipids from algae cells, but use of solvents like n-hexane and electric power, makes the cost of oil production high. In our case, LMBR was produced on a smaller scale for the purpose of conducting pot experiments. Other methods like wet extraction (Reddy et al., 2014), supercritical fluid technologies using CO2 (Halim et al., 2011), milking extraction methods (Kleinegris et al., 2010) are emerging as newer technologies for lipid extraction that would considerably economize on the cost and energy requirement. Even though efforts to extract the microalgal lipids and recover the solvent used has been recently demonstrated by using solar driven solvent extractor (Ghosh et al., 2013), nevertheless, it is extremely important that the algal residue obtained after extraction of the main product (oil) should be used in a biorefinery approach, which was the primary objective of the study. We found that the LMBRs deserved proper exploitation as fertilizer substitute as they are rich in plant nutrients. Considering the state of the art of algal production and post-harvest technology, the current production of biofuel from microalgae is technically feasible, but its economic feasibility is still a major challenge (Chiaramonti et al., 2015). The results of the present experiment applying the biorefinery concept to valorize the LMBR as a plant nutrient source is one of the ways to achieve economic sustainability of algal biofuel.

In view of promising results and due to the limited information available in the literature on this aspect of worldwide interest, further studies are required to evaluate the effect of the use of LMBRs on crop plants over the long term and understand the mechanism of action as well. The effects of LMBRs on soil microbial and biochemical properties, which have bearing on the nutrient cycle, also need to be studied in the future. Other active ingredients or anti-nutrients present in LMBRs also need to be characterized.

#### CONCLUSION

In the present study, nitrogen-rich LMBRs of two microalgal species, namely, Chlorella and Lyngbya were utilized to substitute for the chemical nitrogen fertilizer requirement of maize plants in different proportions viz., 100, 75, 50, and 25%. The grain yield obtained using either of the LMBRs were equivalent to that under treatment employing RRF solely through chemical fertilizers. Even though LMBR of Lyngbya resulted in diminution of the stem and root biomass, there was no reduction observed in the formation of leaf biomass and photosynthetic rate as a result of which the translocation of assimilates from the source to sink was maintained and similar grain yields to that in RRF were obtained using it. Substitution of 75% of the total chemical N through Chlorella LMBR (T4) resulted in the highest DMA in above-ground vegetative biomass and roots and this treatment also recorded the highest grain carbohydrate content, all of which were significantly superior to that in RRF. Among the Lyngbya treatments, substitution at 25% level (T10) was found to be the best in terms of grain yield production as well as phosphorus

#### REFERENCES

and potassium content in grains, while carbohydrate, protein as well as fat content were found at par with RRF. Decreased APX enzyme activity, indicative of lower plant stress was also apparent in most of the LMBR treatments. There was also no detrimental effect of the various LMBR treatments on soil properties. Based on the results, it is concluded that both of the LMBRs can substitute, wholly or partially, the chemical nitrogen fertilizer without affecting the yield and quality of the maize crop, thereby reducing the usage of chemical fertilizers in agriculture and simultaneously making gainful use of the LMBRs generated during biodiesel preparation. The results obtained from the present study thus helps to valorize the LMBR in the commodity markets, thereby advancing toward the economic sustainability of algae production for biofuels.

#### ACKNOWLEDGMENTS

This manuscript bears the number CSIR-CSMCRI – 129/2015. We acknowledge CSIR for providing the support for this work through MoES-NMITLI (TLP 0096). We gratefully thank Dr. Ruma Pal and Mr. Gour Gopal Satpati for providing Lyngbya sp. biomass. The authors would like to acknowledge Mr. Chhaganbhai Jambucha for help in pot experiment. The authors also acknowledge Dr. Parimal Paul, DC, ADCIF, CSIR-CSMCRI, Bhavnagar for providing the instrumentation facility. RM, KC, TG, KT, IP, and DK would like to acknowledge CSIR, CSC 0203 and CSC 0105 for awarding fellowship and AcSIR for Ph.D. enrollment. We also thank Dr. P. K. Ghosh, Dr. Arvind Kumar and Dr. P. K. Agarwal for their generous support and helpful advice.

Spirodela oligorrhiza. Environ. Toxicol. 17, 434–440. doi: 10.1002/tox. 10076

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Maurya, Chokshi, Ghosh, Trivedi, Pancha, Kubavat, Mishra and Ghosh. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Astaxanthin-Producing Green Microalga *Haematococcus pluvialis*: From Single Cell to High Value Commercial Products

#### Md. Mahfuzur R. Shah1 , Yuanmei Liang1 , Jay J. Cheng1, 2 and Maurycy Daroch1 \*

1 School of Environment and Energy, Peking University, Shenzhen Graduate School, Shenzhen, China, 2 Department of Biological and Agricultural Engineering, North Carolina State University, Raleigh, NC, USA

Many species of microalgae have been used as source of nutrient rich food, feed, and health promoting compounds. Among the commercially important microalgae, Haematococcus pluvialis is the richest source of natural astaxanthin which is considered as "super anti-oxidant." Natural astaxanthin produced by H. pluvialis has significantly greater antioxidant capacity than the synthetic one. Astaxanthin has important applications in the nutraceuticals, cosmetics, food, and aquaculture industries. It is now evident that, astaxanthin can significantly reduce free radicals and oxidative stress and help human body maintain a healthy state. With extraordinary potency and increase in demand, astaxanthin is one of the high-value microalgal products of the future.This comprehensive review summarizes the most important aspects of the biology, biochemical composition, biosynthesis, and astaxanthin accumulation in the cells of H. pluvialis and its wide range of applications for humans and animals. In this paper, important and recent developments ranging from cultivation, harvest and postharvest bio-processing technologies to metabolic control and genetic engineering are reviewed in detail, focusing on biomass and astaxanthin production from this biotechnologically important microalga. Simultaneously, critical bottlenecks and major challenges in commercial scale production; current and prospective global market of H. pluvialis derived astaxanthin are also presented in a critical manner. A new biorefinery concept for H. pluvialis has been also suggested to guide toward economically sustainable approach for microalgae cultivation and processing. This report could serve as a useful guide to present current status of knowledge in the field and highlight key areas for future development of H. pluvialis astaxanthin technology and its large scale commercial implementation.

Keywords: *Haematoccoccus pluvialis*, astaxanthin, nutraceuticals, algae cultivation and processing, biorefinery

# INTRODUCTION

"Green microalgae" comprise more than 7000 species growing in a variety of habitats. Haematococcus pluvialis (Chlorophyceae, Volvocales) is unicellular fresh water microalga distributed in many habitats worldwide. It is considered as the best natural source of astaxanthin and the main producing organism of this commercial product

*Edited by:*

Flavia Vischi Winck, University of São Paulo, Brazil

#### *Reviewed by:*

Rosana Goldbeck, Universidade Estadual de Campinas, Brazil Xianhai Zeng, Xiamen University, China

> *\*Correspondence:* Maurycy Daroch<EMAIL_ADDRESS>
#### *Specialty section:*

This article was submitted to Plant Biotechnology, a section of the journal Frontiers in Plant Science

*Received:* 31 October 2015 *Accepted:* 04 April 2016 *Published:* 28 April 2016

#### *Citation:*

Shah MMR, Liang Y, Cheng JJ and Daroch M (2016) Astaxanthin-Producing Green Microalga Haematococcus pluvialis: From Single Cell to High Value Commercial Products. Front. Plant Sci. 7:531. doi: 10.3389/fpls.2016.00531 (Lorenz, 1999; Ranga Rao et al., 2010). Astaxanthin (3,3′ dihydroxy-ß-carotene-4,4′ -dione) is a bright red secondary carotenoid from the same family as lycopene, lutein, and β-carotene, synthesized de novo by some microalgae, plants, yeast, bacteria, and present in many of our favored seafood including salmon, trout, red sea bream, shrimp, lobster, and fish eggs (Higuera-Ciapara et al., 2006; Ranga Rao et al., 2014). Astaxanthin contains two chiral centers and can exist in three different stereoizomers, (3S, 3′ S); (3R, 3′ S), and (3R, 3 ′R). The ratio of 1:2:1 of these isomers is obtained during chemical synthesis of this compound. H. pluvialis biosynthesizes predominantly the 3S, 3′ S stereoisomer, the most valuable one (Yang et al., 2013; Al-Bulishi et al., 2015). Astaxanthin synthesis in H. pluvialis is directly correlated in space and time with deposition of cellular reserves in lipid droplets under conditions of cellular stress (Solovchenko, 2015). From biochemical perspective astaxanthin is synthesized through carotenoid pathway from glyceraldehyde-3-phosphate and pyruvate. Both of these compounds are products of photosynthesis and/or glycolysis depending on cultivation conditions. These two key metabolic intermediates then enter non-mevalonate (MEP) pathway to generate IPP—key intermediate for the synthesis of all carotenoids including astaxanthin. Astaxanthin has a wide range of applications in the food, feed, cosmetic, aquaculture, nutraceutical, and pharmaceutical industries because of its free radical scavenging capacity. In terms of antioxidant activity astaxanthin is 65 times more powerful than vitamin C, 54 times stronger than β-carotene, 10 times more potent than β-carotene, canthaxantin, zeaxanthin, and lutein; and 100 times more effective than α-tocopherol (Miki, 1991; Borowitzka, 2013; Koller et al., 2014; Pérez-López et al., 2014; Cyanotech, 2015). Currently, over 95% of the astaxanthin available in the market is produced synthetically; while H. pluvialis derived natural astaxanthin corresponds to <1% of the commercialized quantity (Koller et al., 2014). Synthetic astaxanthin, synthesized from asta-C15 -triarylphosphonium salt and the C10 -dialdehyde in a Wittig reaction (Krause et al., 1997), has 20 times lower antioxidant capacity than its natural counterpart and to date has not been approved for human consumption (Lorenz and Cysewski, 2000; Koller et al., 2014). Moreover, there are concerns about the safety of using synthetic astaxanthin for direct human consumption due to both different stereochemistry and potential carryover of synthesis intermediates. These concerns make natural astaxanthin from H. pluvialis a preferred choice for highend markets (Li et al., 2011). In addition, H. pluvialis has been already approved as a color additive in salmon feeds and as a dietary-supplement ingredient for human consumption in the USA, Japan, and several European countries (Yuan et al., 2011). Nevertheless, there is no EFSA (European Food Safety Authority) approval for the therapeutic application so far. Supercritical CO2 extracts from H. pluvialis have been granted "novel food" Status by the UK Food Standards Agency (FSA), whilst US FDA (Food and Drug Administration) granted astaxanthin from H. pluvialis "GRAS" status (Generally Recognized As Safe) (Grewe and Griehl, 2012; Capelli and Cysewski, 2013).

The increasing demand for natural astaxanthin and its high price raises interest in efficient systems to produce astaxanthin from H. pluvialis. Various cultivation and astaxanthin production methods in photoautotrophic, heterotrophic, and mixotrophic growth conditions, both indoors; in open raceway ponds or closed photobioreactors using batch or fed-batch approach have been reported (Kang et al., 2005, 2010; Kaewpintong et al., 2007; Ranjbar et al., 2008; García-Malea et al., 2009; Issarapayup et al., 2009; Zhang et al., 2009; Li et al., 2011; Han et al., 2013; Wang et al., 2013a,b). The most recent advances in H. pluvialis cultivation for astaxanthin production include a two-stage mixotrophic culture system (Park et al., 2014) and an "attached cultivation" system using the immobilized biofilm (Zhang et al., 2014). Along with the cultivation process, the induction of carotenoid synthesis in H. pluvialis has a direct correlation with the astaxanthin content of cells and total astaxanthin productivity. The accumulation of astaxanthin is affected by both environmental factors such as light (Saha et al., 2013; Park et al., 2014); temperature (Yoo et al., 2012); pH (Hata et al., 2001); salt concentration (Kobayashi et al., 1993); and nutritional stresses (Boussiba et al., 1999; Chekanov et al., 2014), as well as various plant hormones and their derivatives (Yu et al., 2015). Attempts were made to genetically enhance the capacity of H. pluvialis to produce astaxanthin using both classical mutagenesis (Hu et al., 2008), and more recently genetic engineering (Sharon-Gojman et al., 2015). Once biomass is successfully grown and achieved high cell density, efficient harvesting, cell disruption, dehydration, and recovery/extraction of astaxanthin from H. pluvialis biomass are important factors. Harvesting have been so far achieved through a combination of passive settling and subsequent centrifugation (Han et al., 2013; Pérez-López et al., 2014), or flotation and centrifugation (Panis, 2015). Mechanical processes such as expeller pressing and bead milling are commonly used cell disruption methods to enhance recovery of astaxanthin from H. pluvialis (Razon and Tan, 2011). For the dehydration of H. pluvialis biomass, spray drying (Li et al., 2011; Panis, 2015), freeze drying, lyophilization, and cryodesiccation (Milledge, 2013) methods have been utilized.  Although many studies have explored various methods of extraction and downstream processing of astaxanthin from H. pluvialis biomass, more comprehensive investigation is required to take an advantage of the biological potential of this microalga and its highly valuable product. Since astaxanthin has a great potential in the global market (280 mt, \$447 million in 2014 for both synthetic and natural astaxanthin) and high market value (\$2500–7000/kg); (Koller et al., 2014; Pérez-López et al., 2014; Industry Experts, 2015), in depth investigation of H. pluvialis biology, physiology, efficient culture techniques, downstream bioprocessing, and product formation are highly desired for further development of this sector. Even though currently several commercial companies (Cyanotech Corporation, Mera Pharmaceuticals Inc, AIgatechnologies, Fuji Chemical Industry Co. Ltd etc.) are involved in large scale production of H. pluvialis

and astaxanthin, the production capacity is far beyond the global demand of natural astaxanthin.

This review summarizes both classical knowledge and most recent advances in the cell biology, physiological, and biochemical characteristics, responses to environmental stresses, and their effect on astaxanthin accumulation, genetic engineering, growth conditions, and different cultivation techniques, harvesting, and post harvest downstream bioprocessing of H. pluvialis. The biorefinery concept, global market potential, challenges, and future direction for development of H. pluvialis and astaxanthin production in commercial scale also are discussed.

# BIOLOGY OF *H. PLUVIALIS*

### Taxonomy and Occurrence

The freshwater unicellular biflagellate green microalgae H. pluvialis Flotow belongs to the class Chlorophyceae, order Volvocales and family Haematococcaseae (Bold and Wynne, 1985; Eom et al., 2006). It is also known as Haematococcus lacustris or Sphaerella lacustris. Haematococcus was first described by J. Von Flotow in 1844 and later in 1899 Tracy Elliot Hazen extensively presented its biology and life cycle (Hazen, 1899; Leonardi et al., 2011). H. pluvialis is common in small transient freshwater bodies and widely distributed in many habitats worldwide. It occurs primarily in temporary water bodies like ephemeral rain pools, artificial pools, natural and man-made ponds, and birdbaths (Czygan, 1970; Burchardt et al., 2006). This microalga can be usually found in temperate regions around the world and has been isolated from Europe, Africa, North America, and Himachal Pradeslv India (Pringsheim, 1966; Suseela and Toppo, 2006). It has been also found across diverse environmental and climate conditions: in brackish water on the rocks on the seashore (Chekanov et al., 2014); freshwater basin in the rock filled with melted snow on Blomstrandhalvøya Island (Norway) (Klochkova et al., 2013); dried fountain near Rozhen, Blagoevgrad in Bulgaria Gacheva et al., 2015, freshwater fishpond in Bihor, Romania (Dragos et al., 2010); rooftop surface of a building of KIOST in Seoul Korea (Kim et al., 2015). It is well suited for survival under conditions of expeditious and extreme in light, temperature, and salt concentration that would be deleterious to many other microalgae, due to its ability to encyst (become enclosed by thick membrane) in a rapid manner (Proctor, 1957).

#### Cellular Morphology and Life Cycle

Cellular structure of H. pluvialis is similar to most of other members of volvocalean unicellular green algae. The life cycle of H. pluvialis consists of four types of distinguishable cellular morphologies: macrozooids (zoospores), microzooids, palmella, and hematocysts (aplanospores) (Hazen, 1899; Elliot, 1934). Macrozooids (zoospores), microzooids, and palmella stages are usually called "green vegetative phase" (**Figures 1A,B**). Hematocysts (aplanospores) are referred as "red nonmotile astaxanthin accumulated encysted phase" of the H. pluvialis life cycle (**Figures 1C,D**). Macrozooids (zoospores) are spherical, ellipsoidal, or pear-shaped cells with two flagella of equal length emerging from anterior end, and a cup-shaped chloroplast with numerous, scattered pyrenoids (**Figure 1A**). The macrozooid cells are between 8 and 20 µm long with a distinct gelatinous extracellular matrix of variable thickness. Numerous contractile vacuoles are irregularly distributed near the protoplast surface of the cell (Hagen et al., 2002). These flagellated fast-growing vegetative cells predominate under favorable culture conditions in the early vegetative growth stage (**Figure 1A**) Macrozooids may divide into 2–32 daughter cells by mitosis (Wayama et al., 2013) (**Figures 2A,B**). Under unfavorable environmental or culture conditions, macrozooids start losing flagella, and expand their cell size. They form an amorphous multilayered structure in the inner regions of the extracellular matrix or the primary cell wall as they develop into non-motile "palmella" and become resting vegetative cells (Hagen et al., 2002) (**Figure 1B**). With the continued environmental stress (i.e., nutrient deprivation, high light irradiance, high salinity) and cessation of cell division, "palmella" transform into the asexual "aplanospores" (**Figures 1C,D**). At this stage, cells contain two distinct structures, a thick and rigid trilaminar sheath, and secondary cell wall of acetolysis-resistant material. Such cells become resistant to prevailing extreme environmental conditions (Santos and Mesquita, 1984; Boussiba and Vonshak, 1991). Mature aplanospores; accumulate large amounts of secondary carotenoids, particularly astaxanthin, in lipid droplets deposited in the cytoplasm, which results in a characteristic bright red color of these cells (Hagen et al., 2002). Some H. pluvialis strains are reported to be capable of accumulating astaxanthin without forming aplanospores (Brinda et al., 2004). Once environmental or culture conditions return to optimal, red aplanospores germinate to form flagellated zoospores to initiate a new vegetative growth cycle (**Figure 1A**). In some cases, gametogenesis may occur in aplanospores (**Figures 3A–D**). Such process requires an exposure to extreme adverse conditions (freezing, desiccation, or nutrient starvation) followed by return to favorable culture conditions. During gametogenesis, aplanospore cells can produce up to 64 gametes which are known as microzooids. The microzooids are smaller in size (<10 µm) compared to the zoospores (20–50 µm), and exhibit high-speed motility after their release from gametocysts. Sexual reproduction is rarely observed in H. pluvialis, and has been largely replaced by vegetative reproduction (Triki et al., 1997).

## Ultrastructural Changes of *H. pluvialis* during the Life Cycle

In H. pluvialis cells, large amount of ultrastructural changes occurs during their life cycle which is frequently associated with responses to stress conditions in the environment. In the green vegetative palmella cells, a thick cell wall surrounds the cell, and two layers of extracellular matrix present near the cell wall (**Figure 4A**). Nucleus is located in the center of the cell, and highly developed chloroplasts are located at the periphery (**Figure 4A**). Few astaxanthin granules are present which located around the nucleus (**Figure 4A**). Conspicuous pyrenoids with electron-dense matrix can be found in the stroma (**Figures 4A,B**) (Wayama et al., 2013). During the starting of encystment process,

FIGURE 1 | Light microscopic images of *H. pluvialis* cells in life cycle. (A) Green vegetative motile cell; (B) Green vegetative palmella cell; (C) Astaxanthin accumulating palmella cell in transition to aplanospore; (D) Astaxanthin accumulated aplanospore cell. Scale bar: 10 µm.

Haematococcus turned into greenish-orange cells (with some astaxanthin accumulation) which can be referred as intermediate stage cells. In this stage, conspicuous pyrenoids with electrondense matrix located in the stroma, remain surrounded by thick starch capsules and many starch grains are located around the pyrenoids (**Figure 5**). Round oil droplets with various sizes, containing astaxanthin located around the nucleus (**Figure 5**). At this stage, thylakoids become partial degraded (Wayama et al.,

FIGURE 3 | Gametogenesis in *H. pluvialis*. (A) C, cyst; G, gametocyst; S, a sporocyst; (B) vegetative zoospore, F, flagella (indicated by arrow); (C): gametocyst before releasing gametes; (D): release of gametes from gametocyst. Reproduced with permission from Triki et al., 1997, Phycologia, Allen Press Publishing Services. Copyright (1997) Allen Press Publishing Services.

2013). With the gradual accumulation of astaxanthin chloroplast reduce in volume but photosynthetic activity remains. In the aplanosopre or cyst stage, astaxanthin accumulates and cells form cysts. Oil droplets and astaxanthin accumulation patterns may differ among cyst cells. For example, electron-dense astaxanthin granules in oil droplets (**Figures 6A,B**) occurred in some cells. In other cells, relatively large oil droplets occurred throughout the cell (**Figures 6C,D**). Chloroplasts are degenerated and localized in the interspace between oil droplets (Wayama et al., 2013).

# BIOCHEMICAL COMPOSITION OF *H. PLUVIALIS*

Because of the unique life cycle of H. pluvialis, cellular composition of this microalga varies tremendously between its "green" and "red" stages of cultivation (**Table 1**).

#### Protein

In green stage, during favorable growth conditions most H. pluvialis strains are rich in protein (29–45) (**Table 1**), lower protein content (23.6%) have been however observed in a Bulgarian strain Haematococcus cf. pluvialis Rozhen-12 during green stage cultivation (Gacheva et al., 2015). It was estimated that proteins contribute to 21 (Kim et al., 2015) and 23% (Lorenz, 1999) of cellular content during red stage cultivation of H. pluvialis. Amino acid composition of proteins in the red stage indicated that proteins were mainly composed of aspartic acid, glutamic acid, alanine, and leucine with total amino acid content of 10.02/100 mg, 46.0% of which belonged to essential amino acids (Lorenz, 1999; Kim et al., 2015).

#### Carbohydrates

In green stage, carbohydrate content approximates to 15–17%, about a half of the red stage (**Table 1**). In the red stage, under conditions of stress (e.g., nutrient starvation, light stress, high acidity, temperature variations etc.), H. pluvialis accumulates

FIGURE 4 | Transmission electron micrographs of green vegetative cells of *H. pluvialis*. (A) General ultrastructure. The cell wall is surrounded by extracellular matrix (arrowheads). Arrows indicate astaxanthin granules. (B) Chloroplast and pyrenoid. C, chloroplast; CW, cell wall; N, nucleus; P, pyrenoid. Scale bars in (A,B): 5 and 1 µm, respectively. Figure reproduced from Wayama et al. (2013) distributed under the terms of the Creative Commons Attribution License.

higher content of carbohydrates (starch), for example 38 (Lorenz, 1999), 60 (Recht et al., 2012), and 74% (Boussiba and Vonshak, 1991). Under prolonged stress conditions starch is consumed in the cell.

### Lipid

In green stage, total lipid content varies from 20 to 25%, with approximately 10% lipids composed predominantly of short (C16, C18) polyunsaturated fatty acids deposited in the chloroplasts. Neutral lipids are predominant lipid class in both green and red cells (**Table 1**). In red stage, prolonged stress conditions direct larger flux toward the synthesis of neutral lipids—triacylglycerols (TAG). Red stage cells can accumulate up to 40% of their cell weight as cytoplasmic lipid droplets (LD), and considerable amount of secondary metabolites including up to 4% of the ketocarotenoid astaxanthin (Boussiba et al., 1992, 1999; Saha et al., 2013). The phospholipid content does not change compared to the green stage, while the glycolipid fraction nearly doubles in red cells when compared with green vegetative cells (Damiani et al., 2010). The total fatty acid profile of H. pluvialis is relatively complex. Palmitic (16:0), linoleic (18:2), and linolenic (18:3) acids are predominant components of the profile with highly polyunsaturated species also present in considerable amounts (**Table 2**). Based on the comparative studies on fatty acids profile of two different H. pluvialis, it was revealed that both strains varied in composition, especially of palmitic (16:0), oleic (18:1), linoleic (18:2), and linolenic (18:3) acids. This variation might be associated with several factors such as culture environment, stress conditions, culture parameters, variation of strain origin, nutrient etc. Higher lipid content of H. pluvialis grown under nutrient starvation and the suitable profile of its fatty acids indicate a possibility of biodiesel production from this microalga (Damiani et al., 2010; Saha et al., 2013). The massive astaxanthin accumulation in H. pluvialis is a cellular response to stress conditions and is accompanied by the enhanced biosynthesis of triacylglycerols (TAG) (Zhekisheva et al., 2002, 2005; Cerón et al., 2007), and the reduction in photosynthetic activity of PSII, loss of cytochrome f, and subsequent reduction in electron transport, and increased respiration rate (Boussiba, 2000). During transition from green vegetative cells to red aplanospores after exposure to stress conditions astaxanthin start to accumulate as fatty acid mono- or diesters in cytoplasmic lipid droplets (LD) (Aflalo et al., 2007). As cells undergo transition to red stage, both chlorophyll and protein contents drop.

# Carotenoid

The carotenoid fraction of green vegetative cells consists of mostly lutein (75–80%), β-carotene (10–20%) and others, including chlorophyll a and b, primary carotenoids, violaxanthin, neoxanthin, lactucaxanthin, and zeaxanthin (Renstrøm et al., 1981; Harker et al., 1996a). In the red stage, the total carotenoid content is markedly enhanced, and the characteristic primary carotenoid pattern of vegetative stage is replaced by secondary carotenoids, mainly astaxanthin (80–99% of total carotenoids) (Harker et al., 1996a; Dragos et al., 2010). The ratio of carotenoids to chlorophylls is about 0.2 in the green stage and increases in the red stage by an order of magnitude and reaches about 2–9. The majority of astaxanthin is not deposited in its free form but it exists within the cell as fatty acid esters of astaxanthin, usually mono- or diesters of palmitic (16:0), oleic (18:1), or linoleic (18:2) acids. This type of modification is required for the deposition of this highly polar molecule within non-polar matrix of lipid droplets. Approximately 70% monoesters, 25% diesters, and only 5% of the free ketocarotenoid is present in the mature "red" cells of H. pluvialis (Zhekisheva et al., 2002; Solovchenko, 2015). Under certain conditions of stress H. pluvialis has been shown to accumulate up to 3–5% DW of astaxanthin (Han et al., 2013; Chekanov et al., 2014).

# *H. PLUVIALIS*-DERIVED ASTAXANTHIN

# *H. pluvialis* as a Major Source of Astaxanthin

H. pluvialis can accumulate up to 5% DW of astaxanthin and is considered as the best natural source of this highvalue carotenoid pigment (Wayama et al., 2013). Dietary supplements containing Haematococcus astaxanthin has proved to be safe to humans and widely used for over 15 years as a nutraceutical supplement with no adverse side-effects of its supplementation (Capelli and Cysewski, 2013; Yang et al., 2013). Natural astaxanthin from H. pluvialis or krill oil is available in the market as a dietary supplement in dosages from 3.8 to 7.6 mg per day due to potential health benefits (Yang et al., 2013). As societies nowadays are looking toward "green" solutions, natural astaxanthin form H. pluvialis seems to be more favorable than its synthetic counterpart due to structure, function, application, and

security (Choubert and Heinrich, 1993; Capelli and Cysewski, 2013; Pérez-López et al., 2014).

### Biosynthesis of Astaxanthin in *H. pluvialis*

Biosynthesis of astaxanthin in H. pluvialis is a complex process that is highly up-regulated in conditions of stress and which coincides with the accumulation of triacylglycerols (TAGs). Both compounds are deposited in the cytosolic lipid bodies during the "red" stage of H. pluvialis cultivation. Astaxanthin belongs to carotenoids, a C40 tetraterpenes, synthesized from isoprene units. Isopentenyl pyrophosphate (IPP) is a key intermediate of carotenoid synthesis. In principle, IPP can originate from two dissimilar pathways: mevalonate pathway (MVA) located in cytosol and non-mevalonate (MEP) located in the chloroplast (Lichtenthaler et al., 1997; Lichtenthaler, 1999; Eisenreich et al., 2001). Alternative name for MEP is DOXP, due to the formation of 1-deoxy-D-xylulose-5-phosphate in the first stage of the pathway. Comparative transcriptomic analysis of astaxanthin biosynthesis in H pluvialis have shown that the key intermediate-IPP is most likely synthesized using the DOXP pathway. H. pluvialis lacks three key enzymes of the mevalonate pathway (MVA) catalyzing the formation of isopentenyl pyrophosphate (IPP) from acetoacetyl-CoA (Gwak et al., 2014). There has been numerous evidence of the full set of enzymes required for the conversion of photosynthesisderived products i.e., pyruvate and glyceraldehyde-3-phosphate into isopentenyl pyrophosphate through DOXP pathway inside H. pluvialis chloroplasts (Gwak et al., 2014). It makes it the most likely source of IPP in H. pluvialis cells. The process of astaxanthin biosynthesis is presented on **Figure 7**. IPP derived from DOXP pathway is an initial building block of astaxanthin synthesis. In the subsequent step the IPP undergoes isomerization to dimethylallyl diphosphate (DMAPP). It has been long assumed that this conversion was catalyzed exclusively by isopentenyl pyrophosphate isomerase (IPI) (Sun et al., 1998; Lichtenthaler, 1999). However, recent transcriptomic studies suggest that neither of the two ipi genes of H pluvialis (ipi1 and ipi2IPI2) are up-regulated during cellular accumulation of astaxanthin (Gwak et al., 2014). Suggestions have been made that another enzyme of similar activity, 4-hydroxy-3-methylbut-2-enyl diphosphate reductase (HDR) was more likely to be responsible for catalyzing interconversion between IPP and

TABLE 1 | Typical composition of *H. pluvialis* biomass in green and red cultivation stages.

Adapted from Grewe and Griehl (2012). n.d., no data.

DMAPP (Hoeffler et al., 2002; Rohdich et al., 2002; Gwak et al., 2014). Further studies are required to assess the contribution of these three enzymes to this key biosynthetic step of astaxanthin formation. Elongation of the isoprenoid chain is initiated with a molecule of DMAPP and a subsequent linear additions of three molecules of IPP catalyzed by an enzyme geranylgeranyl pyrophosphate synthase (GGPS) (Britton, 1993; Cunningham and Gantt, 1998). The final step of this process is the formation of a C20 compound, geranylgeranyl pyrophosphate (GGPP), a shared precursor with other isoprenoids. The first committed step of carotenoid synthesis is catalyzed by phytoene synthase (PSY) and results in a head-to-tail condensation of two GGPP molecules to form a C40 compound—phytoene that serves as a precursor for astaxanthin and other carotenoids (Cunningham and Gantt, 1998). The expression of the phytoene synthase gene (psy) was up-regulated in Haematococcus cells stressed with high light and undergoing transformation from "green" to "red" stage (Steinbrenner and Linden, 2001; Vidhyavathi et al., 2008; Gwak et al., 2014). The subsequent formation of highly unsaturated compound—lycopene proceeds through four desaturation steps catalyzed by two phytoene desaturases (PDS) and a ζ-carotene desaturase (ZDS) with two plastid terminal oxidase (PTOX 1, PTOX 2) acting as co-factors for electron transfer between C40 carotenoid intermediates, plastoquinone and final electron acceptor—oxygen (Li et al., 2010; Nawrocki et al., 2015). Of the two, PTOX 1 was found to be co-regulated with astaxanthin synthesis in H. pluvialis (Wang et al., 2009; Nawrocki et al., 2015). Desaturation reactions increase the number of conjugated carbon-carbon double bonds that form the chromophore in carotenoids and convert a colorless molecule of ζ-carotene to a pink colored lycopene (Cunningham and Gantt, 1998). Both termini of lycopene undergo cyclization catalyzed by lycopene

cyclases (LCY-e and LCY-b). Cyclization is a branching point of the carotenoid biosynthesis in most organisms, yielding αcarotene (precursor of lutein) and β-carotene (precursor of other carotenoids including astaxanthin). In H. pluvialis vast majority of the carbon flux is directed into the latter (Gwak et al., 2014), and high level of LCY-b transcripts have been observed under stress conditions (Lorenz and Cysewski, 2000; Gwak et al., 2014). Final two oxygenation steps catalyzed by β-carotene ketolase (BKT) and β-carotene hydroxylase (CrtRb) are rate limiting steps of astaxanthin synthesis (Linden, 1999; Steinbrenner and Linden, 2001; Vidhyavathi et al., 2008). Although in principle the reactions catalyzed by these two enzymes can proceed in any order, higher substrate specificity of BKT toward β-carotene than zeaxanthin favors initial addition of keto group before enantio-selective hydroxylation of canthaxanthin to astaxanthin is catalyzed by CrtR-b (Lotan and Hirschberg, 1995). Enantioselectivity of astaxanthin synthesis is of primary importance for the nutraceuticals market and the major advantage of H. pluvialis astaxanthin over its synthetic counterpart. Since astaxanthin has two identical chiral centers at the positions of 3 and 3′ it can exist in four different configurations which yield three different isomers: (3R, 3′ S); (3R, 3′R); (3S, 3′ S) depending on the spatial orientation of the hydroxyl (OH) groups in chiral carbon. During chemical synthesis these isomers are present in the ratio of 2:1:1, respectively, yielding only 25% of the naturally occurring (3S, 3 ′ S) isoform. H. pluvialis synthesizes the (3S, 3′ S) stereoisomer of astaxanthin and is therefore a much sought-off product in the nutraceutical market.

### Effect of Small Molecules on Astaxanthin Synthesis

Astaxanthin is a secondary metabolite, a carotenoid synthesized by H. pluvialis in a response to stress conditions such as high light, salinity, or carbon to nitrogen ratio (Gao et al., 2012a). Regulation of these pathways can be affected by numerous small molecules like plant hormones or their analogs. An array of such molecules has been explored to modulate astaxanthin accumulation by H. pluvialis. Plant hormones that are usually associated with stress response mechanisms e.g., abscisic acid (ABA), jasmonic acid (JA), methyl jasmonate (MJ) or growth regulators like gibberellic acid (GA3), salicylic acid (SA) or brassinosteroid—2, 4-epibrassinolide (EBR) were found particularly promising in increasing astaxanthin accumulation in H. pluvialis (Kobayashi et al., 1997b; Gao et al., 2012a,b, 2013a,b; Yu et al., 2015). It was found that these compounds affect numerous genes involved in astaxanthin biosynthesis and result in their even six to 10 fold up-regulation. All of these compounds were tested in various concentrations and the highest improvement of astaxanthin accumulation was achieved with salicylic acid. At relatively low concentrations of the hormone 50 mg L−1 and low light 25 µmol photons m−2 s −1 the content of astaxanthin raised seven fold from 0.391 mg L−1 to 2.74 mg L−1 . Higher levels of these hormones had however deleterious effect on both growth and astaxanthin accumulation (Gao et al., 2012a). Correlation between mRNA transcript levels of five key enzymes of astaxanthin synthesis pathway (ipi, psy, pds, crtO, and crtR-b encoding respectively isopentenyl-diphosphate δ-isomerase (IPI), phytoene synthase (PSY), phytoene desaturase (PDS), β-carotene oxygenase (CrtO), and β-carotene hydroxylase (CrtR-b) with alga growth and astaxanthin production suggested a complex, multiple regulatory mechanisms at transcriptional, translational, and post-translational levels controlling entire process of carotenoid synthesis in H. pluvialis (Li et al., 2010). Small molecules can have multiple effects on the regulation of each of these genes and more detailed investigation of the molecular responses to their application is required for both understanding gene regulation in H. pluvialis and enhancing its capacity as commercial astaxanthin producer.

# Genetic Improvement of *H. pluvialis* for Astaxanthin Production

Green eukaryotic microalgae are among the organisms that are notoriously difficult to genetically engineer. In principle, genetic engineering of microalgae has been reported for over 30 strains of various species, including Haematococcus (Rosenberg et al., 2008; Radakovits et al., 2010; Forján et al., 2015). In majority of these reports however, only a transient transgene expression has been achieved and a desired, stable, hereditary, and efficient genetic modification existed only for model species such as Chlamydomonas reinhardtii and Volvox carteri. Due to these constrains genetic improvement of H. pluvialis strains have been long limited to classical mutagenesis. Combination of mutagenic treatment and various screening methods resulted in the development of numerous interesting mutants of H. pluvialis including those of higher astaxanthin accumulation capacity (Tjahjono et al., 1994a; Chumpolkulwong et al., 1997; Tripathi et al., 2001; Chen et al., 2003; Hu et al., 2008; Hong et al., 2012; Gomez et al., 2013). Various mutagenic treatments have been tested, and can be broadly divided into UV and chemical mutagenesis. Chemical mutagenesis has been generally found to be more suitable for H. pluvialis due to alga's intrinsic capacity to limit the damage from light. Typical chemicals used for mutagenesis include ethyl methanesulphonate (EMS) and N-methyl-N-nitro-N-nitrosoguanidine (MNNG). In all studies their concentrations were adjusted to target 85–95% cell mortality. Successful mutants are usually screened using a combination of factors that promote identification of mutants capable of high astaxanthin accumulation.  Screening relies on identifying colonies that are capable of surviving and/or growing well in the presence of inhibitory concentrations of these compounds and high illumination. Surviving strains should in principle exhibit better capacity to synthesize carotenoids and divert larger, or more efficient carbon flux toward these compounds. A number of successful mutants have been isolated this way and typical improvement of astaxanthin accumulation ranges from several percent (Gomez et al., 2013) to two or three fold improvement (Hu et al., 2008). In former case the mutated strain was tested in commercial scale cultivation system (120,000 L) and retained the improved capacity for astaxanthin production. An alternative approach to strain improvement relying on selection of photosensitive mutants was recently attempted (Hong et al., 2012). Photosensitive mutant with an ability to grow under hetero or mixotrophic conditions should be in principle advantageous over wild type strains due to faster growth rates and more efficient stress trigger. Screening for successful mutants was performed in a three stage process. Since photosensitivity is connected with impaired photosynthesis, these impaired mutants were selected in the first screening. Secondary screening tested for the ability of heterotrophic growth of these photosensitive isolates. Tertiary screening involved mixotrophic conditions with moderate illumination to obtain mixotrophic photosensitive strain that accumulated 4.7% (w/w) of astaxanthin under much shorter induction time (Hong et al., 2012). The mutated strain was stable for at least 1.5 year and is an interesting example of using classical mutagenesis for improvement of H. pluvialis. Mutagenic strain improvement can be expanded by breeding or creating hybrid strains from previous genetic improvement efforts. Technique of protoplast

fusion has been successfully applied to H. pluvialis (Tjahjono et al., 1994a). Two mutagenized strains, norflurazon-resistant and nicotine-resistant have been fused to create a hybrid containing genetic material of two initial strains and showed 30% higher astaxanthin accumulation than the initial wild type strain, when neither of the fused strains showed such characteristics (Tjahjono et al., 1994a). Till very recently H. pluvialis was one of the organisms in which genetic engineering of its nuclear genome was considered difficult due to lack of suitable shuttle vectors and satisfactory transformation efficiencies (Sharon-Gojman et al., 2015). A number of unsuccessful approaches have been tried that included various transformation methods (particle bombardment, electroporation, Agrobacterium), vectors, promoters, and strains (Sharon-Gojman et al., 2015). To address these limitations and open a new array of possibilities in H. pluvialis and astaxanthin biology and technology new developments were required. In recent years these developments emerged and stable transformations of H. pluvialis chloroplast (Gutiérrez et al., 2012) and nuclear genomes (Sharon-Gojman et al., 2015) were achieved. Most recent nuclear transformation vectors are capable to transform one or two transgenes into the nuclear genome either 5′ or 3′ of the endogenous dominant selection marker, in the absence of any additional antibiotic resistance genes. The selection marker used in this system is a phytoene desaturase (pds) variant that confers resistance to a herbicide norflurazon due to a single point mutation (L504A). Successful transformation of H. pluvialis was obtained with particle bombardment and numerous constructs based on pds selection marker were delivered and incorporated to the genome showing stability of integration for over 16 months of subculturing (Sharon-Gojman et al., 2015). Genetic engineering of chloroplast genome of H. pluvialis have been also achieved relatively recently (Gutiérrez et al., 2012). So far these studies are limited to expressing exogenous antibiotic resistance gene (aadA

cassette) between Internal Transcribed Spacer region and 16S gene of H. pluvialis chloroplast genome, but this technique may in the near future have significant impact on protein production in H. pluvialis (Gutiérrez et al., 2012) as higher protein yields are generally obtained during chloroplast expression of transgenes in other microalgal strains (Li et al., 2015). These new developments in genetic engineering of H. pluvialis can open a new chapter for the development of this organism as both industrial astaxanthin producer and an interesting model for carotenoid synthesis and accumulation studies.

#### APPLICATIONS OF *H. PLUVIALIS* ASTAXANTHIN

#### Astaxanthin in Human Health and as Nutraceutical

Astaxanthin possesses various human health benefits and nutraceutical applications and plenty of published information available with evidences, mainly from in vitro and animal models (Guerin et al., 2003; Chew et al., 2004; Higuera-Ciapara et al., 2006; Palozza et al., 2009; Yuan et al., 2011). The effect of Haematococcus derived astaxanthin on various physiological systems in animal and human subject is presented in **Table 3**. Astaxanthin is considered as "super anti-oxidant" which possesses one of the strongest known antioxidant effects. Its unique structure allows it to span biological membranes and act as an antioxidant by reducing and stabilizing free radicals (Hussein et al., 2006; Liu and Osawa, 2007; Ranga Rao et al., 2010). It is very good at protecting membrane phospholipids and other lipids against peroxidation (Naguib, 2000). There are several studies which showed high antioxidant activity of astaxanthin from H. pluvialis in rats supplemented with diet (Kamath et al., 2008; Ranga Rao et al., 2010, 2013; Augusti et al., 2012). Astaxanthin can terminate the induction of inflammation in biological systems. It can help to fight symptoms of ulcer disease from Helicobacter pylori (Liu and Lee, 2003); protect against gastric lesions (ulcers), improve gastrointestinal health (Nishikawa et al., 2005; Kamath et al., 2008); and treat gastrointestinal discomfort (Andersen et al., 2007; Kupcinskas et al., 2008). Astaxanthin offers protection against free radical damage to preserve immune-system defenses. The immunomodulating capacity of astaxanthin has been found to be superior to that of β-carotene and canthaxanthin (Chew and Park, 2004). Astaxanthin has shown significant effect on immune function in a number of in vitro and in vivo assays using both animal models (Chew et al., 2004) and humans (Park et al., 2010). Astaxanthin is a potential therapeutic agent against atherosclerotic cardiovascular disease (Fassett and Combes, 2011). Astaxanthin supplementation can be beneficial for people with enhanced risk for heart attacks. It is carried by VLDL, LDL, and HDL (high-density lipoprotein) in human blood and protects LDL-cholesterol against oxidation (Iwamoto et al., 2000); has a role in the reduction of blood plasma level (Karppi et al., 2007); and increases basal arterial blood flow (Miyawaki et al., 2008). Oxidative stress is a causative or at least ancillary factor in the pathogenesis of major neurodegenerative diseases (Alzheimer's, Huntington's, Parkinson's, and amyotrophic lateral sclerosis-ALS). Diets high in antioxidants offer the potential to lower the associated risks (Ferrante et al., 1997). Natural astaxanthin can cross the blood-brain barrier in mammals and can extend its antioxidant benefits beyond that barrier. Therefore, astaxanthin can help to alleviate the effects of Alzheimer's disease and other neurological diseases. Astaxanthin can improve respiratory and sympathetic nervous system activities (Nagata et al., 2006), inhibit the growth of fibrosarcoma, breast, and prostate cancer cells and embryonic fibroblasts (Palozza et al., 2009); cell death, cell proliferation and mammary tumors (Nakao et al., 2010). Astaxanthin supplementation can help to protect against UVinduced photooxidation; as an oral sun-protectant; can prevent skin thickening and reduce collagen reduction against UV induced skin damage (Ranga Rao et al., 2013) and can improve skin condition across its layers i.e., corneocyte, epidermis, basal, and dermis by combining oral supplementation and topical treatment (Seki et al., 2001; Yamashita, 2002; Tominaga et al., 2012). Results have shown that semen quality, pregnancy rate and sperm velocity in human subject can be improved (Elgarem et al., 2002; Comhaire et al., 2005) whereas idiopathic infertility can be decreased by astaxanthin (Andrisani et al., 2015).

# Astaxanthin in Aquaculture and Poultry Industry

During last 20 years, synthetic astaxanthin has been widely used for pigmentation of fish. Haematococcus astaxanthin has great potential in aquaculture industry, due to increasing consumer demands for natural products and ability of Haematococcus astaxanthin to provide necessary supplementation for adequate growth and reproduction of commercially valuable fishes (Salmonid, Red sea bream), rainbow trouts, and shrimps. Microalgae- derived astaxanthin has been demonstrated as safe and effective compound for flesh pigmentation of these fish (Torrissen and Naevdal, 1984; Tolasa et al., 2005). Utilization of H. pluvialis meal for pigmenting has resulted in significant astaxanthin deposition in flesh and skin, flesh coloration enhancement, enhanced antioxidant system, fish egg quality, better growth and survival of fry of Salmonid, sea bream, and rainbow trout (Arai et al., 1987; Sommer et al., 1991; Choubert and Heinrich, 1993; Sheikhzadeh et al., 2012a,b), ornamental fishes (Ako and Tamaru, 1999), and shrimp (Arai et al., 1987; Parisenti et al., 2011). A recent study indicated that diets supplemented with H. pluvialis can improve large yellow croaker fish growth more than diets supplemented with synthetic astaxanthin (Li et al., 2014). H. pluvialis-derived natural astaxanthin has shown to be efficient in pigmentation of egg yolks, egg production (Elwinger et al., 1997) in hen and breast muscle tissue improvement and higher feed efficiency in broiler chicken (Inborr and Lignell, 1997; Inbbor, 1998). It has also been proved to improve health and fertility of chicken and to decrease their mortality (Lignell and Inborr, 1999, 2000).

#### TABLE 3 | Effect of *H. pluvialis-*derived astaxanthin on various physiological systems in human and animal subjects.

# CULTIVATION AND PROCESSING OF *H. PLUVIALIS* FOR ASTAXANTHIN PRODUCTION

# Culture Conditions and Requirements for Cell Growth and Astaxanthin Formation

Optimization of the various culture parameters, such as growth medium composition, light, pH, temperature etc. is necessary to achieve high biomass and astaxanthin production. Most of these parameters have different optima for biomass accumulation and astaxanthin production. For carotenogenesis induction, the stronger exposure to stress conditions, the higher astaxanthin accumulation. The origins of this stress can be diverse and successful astaxanthin accumulation has been induced with both, high levels of one stressor, or from a combination of multiple stress factors.  Various types of growth media are used for cultivation of H. pluvialis. The most commonly used media are BG- 11 (Rippka et al., 1979), BBM (Bischoff and Bold, 1963), OHM (Fábregas et al., 2000); KM1-basal medium with organic carbon sources in the form of sodium acetate (Kobayashi et al., 1993), and their modifications. An ideal composition of the medium to achieve high growth rate and biomass accumulation is different from ideal composition for high accumulation of astaxanthin. Sodium nitrate was found to be the most optimal inorganic nitrogen source (Sarada et al., 2002a), alternatively an organic source such as urea can be used. When culture is subjected to nutrient deficiency, it leads to accumulation of astaxanthin within the cells (Saha et al., 2013). Nitrogen limitation leads to approximately twice the rate of astaxanthin production than the limitation of phosphorus. It can be due to higher cellular damage resulting from a lack of nitrogen, which manifests in significant degradation of chlorophyll, compared to the phosphorus starvation (Boussiba et al., 1999). Micronutrients such as selenium and chromium result in an increased biomass and astaxanthin production (Tripathi et al., 1999; Fábregas et al., 2000; Domínguez-Bocanegra et al., 2004). Formation of astaxanthin can also be induced by adding NaCl (0.25–0.5% w/v) to the media. Also, when NaCl is added together with 2.2 mM sodium acetate, astaxanthin accumulation can be increased (Sarada et al., 2002b). Addition of 0.45 mM Fe2+ in the form of ferrous sulfate may significantly increase the biosynthesis of carotenoids in cysts due to formation of hydroxyl radicals. (Kobayashi et al., 1997b). This effect may be enhanced by combining Fe2+ treatment with an addition of sodium acetate and high temperature exposure (Kobayashi et al., 1993; Tjahjono et al., 1994b). According to most studies, the suitable temperature for the growth and astaxanthin accumulation of H. pluvialis is between 20 and 28◦C (Fan et al., 1994; Hata et al., 2001; Lababpour et al., 2005; Kang et al., 2010; Yoo et al., 2012; Wan et al., 2014a). However, temperature above 30◦C induces a transition from green vegetative stage to red stage and formation of red cysts can be observed within 2 days. This transition is combined with a significant slowdown in growth, while astaxanthin accumulation is 2–3 times higher than at 20◦C. The increased temperature is likely to affect the synthesis of astaxanthin through stimulation of oxygen radicals formation and their higher reactivity (Tjahjono et al., 1994b). It is preferred that the temperature change takes place gradually, allowing better acclimation to the new conditions (Hata et al., 2001). pH can also significantly affect the cell growth and synthesis of chlorophyll and carotenoids in H. pluvialis. In terms of biomass and astaxanthin production optimal pH is within the range of 7.00–7.85 (Hata et al., 2001; Sarada et al., 2002a). The typical irradiation for H. pluvialis cultivation ranges between 40 and 50 µmol photons m−2 s −1 (Hata et al., 2001; Chekanov et al., 2014; Park et al., 2014). Optimal irradiation to achieve a high growth rates tend to be higher, namely 70 (Zhang et al., 2014), 80 (Saha et al., 2013), 90 (Fan et al., 1994), or even up to 177 µmol photons m−2 s −1 (Domínguez-Bocanegra et al., 2004). These different optimal values may be caused by other cultivation parameters such as media composition, temperature, or the strain of H. pluvialis. During vegetative stage cultivation of H. pluvialis, the regular cycles of alternating light and dark 12:12 or 16: 8 h are often used (Saha et al., 2013; Park et al., 2014), but the higher density cultures are achieved with continuous illumination (Domínguez-Bocanegra et al., 2004). The best practice to date appears to be white or blue LED lighting (Saha et al., 2013) or the mixture of both at the ratio of 3:1 at 7000 lx (∼95 µmol photons m−2 s −1 ). These conditions promote morphologic changes from green vegetative cells to red cyst cells (Sun et al., 2015). Carotenogenesis is induced in cells upon exposure to higher light intensity than the corresponding light saturation point (LSP). However, specific optimum value of LSP differ between studies. The lowest intensities that have been reported utilized irradiation of around 100–150 µmol photons m−2 s −1 (Zhang et al., 2014) followed by 240 (Saha et al., 2013), 345 (Domínguez-Bocanegra et al., 2004), and 480 µmol photons m−2 s −1 (Chekanov et al., 2014). Lower optimal irradiation was found to be influenced by other stress conditions, such as deficiency of nutrients (Saha et al., 2013; Zhang et al., 2014) or elevated temperature (Tjahjono et al., 1994b). It proves that for effective induction of carotenogenesis excessive irradiation may not be necessary if other stressors are present. With the reduced requirements of light for cultivation in photobioreactors, the costs of cultivation can be minimized which is essential for astaxanthin production in an industrial scale. Regarding the type of illumination, the highest carotenoid content was obtained by using a continuous PAR lighting (Saha et al., 2013). An interesting alternative to an immediate change in the radiation intensity to induce the transition from the vegetative phase to carotenoid production is gradually increasing the level of lighting. Gradual increase of light intensity can result in gradual transformation of cells to cysts and can also contribute to better accumulation of astaxanthin, because the cells are capable to cope with increasing higher levels of stress (Park et al., 2014).

#### Culture Systems

Astaxanthin-producing H. pluvialis is capable of growing in photoautotrophic, heterotrophic, or mixotrophic growth conditions in indoors, open raceway ponds or closed photobioreactors in batch, fed batch, or continuous modes.

#### Photoautotrophic Culture

Photoautotrophic culture of H. pluvialis is mainly carried out in open raceway ponds or closed photobioreactors. Typical photobioreactors used for its cultivation include tubular, bubble column and airlift photobioreactors. As the culture conditions for maximum growth and maximum astaxanthin content are mutually exclusive, a two-step cultivation strategy is commonly adopted for the commercial production. The first step, green vegetative growth phase ("green stage") is to promote algal growth under favorable culture conditions (e.g., low light and nitrogen-replete) (Boussiba, 2000; Aflalo et al., 2007; Del Rio et al., 2007). When high cell density is reached, the culture enters into the second step, reddish inductive production phase ("red stage"), where algal cells are subjected to stress conditions such as high light intensity and nitrogen depletion, excess acetate addition, pH or salt stress, phosphate deficiency, or the addition of specific cell division inhibitors. These stress factors (either one or combination of more) induce the astaxanthin production in H. pluvialis (Fábregas et al., 2001; Torzillo et al., 2003; Orosa et al., 2005; He et al., 2007; Hu et al., 2008; Li et al., 2010; Choi et al., 2011). Therefore, carotenoid induction method has a direct correlation with both the astaxanthin content and total astaxanthin productivity. The optimal environmental and nutritional conditions for each stage are quite different (Del Rio et al., 2007). The reported biomass productivities in green stage and red stage ranged from 0.01 to 0.5 g L−1 d −1 and 0.01 to 4.8 g L−1 d −1 , respectively. Astaxanthin productivity and astaxanthin content ranged from 0.44 to 21 mg L−1 d −1 and 0.8 to 4.8% of DW, respectively (**Table 4**). Astaxanthin can be also produced efficiently by H. pluvialis using a simpler "one-step strategy." This strategy involves the administration of nitrate starvation and specific average irradiance in the culture medium, resulting in simultaneous algal cell growth and astaxanthin accumulation (Del Río et al., 2005; Del Rio et al., 2007; Del Río et al., 2008; García-Malea et al., 2009). At the laboratory scale and under continuous illumination, mean astaxanthin productivity of 20.8 mg L−1 d −1 has been reported for the one-step method (Del Río et al., 2008). The technical feasibility of this approach at a pilot scale was demonstrated in an outdoor tubular photobioreactor, which resulted in biomass and astaxanthin productivities of 0.7 g L−1 d −1 and 8 mg L−1 d −1 , respectively (García-Malea et al., 2009). One-stage cultivation seems attractive, since it is less complicated than the twostage process and the production of astaxanthin takes place in a continuous mode. It has however two serious drawbacks. First, the actual astaxanthin production is significantly lower compared to the two-stage approach. Second, this cultivation is unsuitable for outdoor setting, since it requires incessant illumination during night as well what makes the process too expensive (Aflalo et al., 2007). An "attached cultivation" approach was successfully applied in the induction of H. pluvialis for astaxanthin production. In this method green cells are cultured in the conventional water column and then deposited on the membrane to increase light stress surface area in the second phase of cultivation. Under the optimized conditions, biomass, and astaxanthin productivities in the attached cultivation system were 2.8 (3.7 g m−2 d −1 ) and 2.4-fold (65.8 mg m−2 d −1 ) higher than those of the suspended bioreactor, respectively (Wan et al., 2014b). Other studies that used similar approach have reported higher astaxanthin productivities of 124 mg m−2 d −1 (Yin et al., 2015) and 164.5 mg m−2 d −1 (Zhang et al., 2014). Attached cultivation approach is superior to suspended induction in several aspects such as, lower water consumption and smaller risk of contamination. This indicates that attached induction approach can provide a promising way to boost economic benefits and considerably reduce production cost of astaxanthin from H. pluvialis (Zhang et al., 2014; Wan et al., 2014b). Recently, Park et al. (2014) established a two-stage "perfusion culture" system for H. pluvialis combining it with stepwise increase of light irradiance. Approach is based on repeated replacement of the growth medium. Cells are grown in a photobioreactor and are periodically retained in the cell settling chamber whilst growth medium is being replaced in the photobioreactor. Cells are later recycled to the bioreactor and can efficiently utilize fresh growth medium which is free of inhibitory metabolic by-products. Perfusion culture can provide high biomass productivities of 0.18 g L−1 d −1 . Under stepwise increased light irradiance (150– 450 µE/m2 /s), cellular density of 12.3 g L−1 of have been obtained which is 3.09 and 1.67 times higher than batch and fed-batch processes, respectively whilst the productivity of astaxanthin reached 602 mg L−1 (Park et al., 2014).

#### Heterotrophic and Mixotrophic Culture

High light irradiance is often employed for enhancing astaxanthin formation in H. pluvialis cultures. However, light absorption and scattering caused by mutual shading of cells in large-scale cultures severely affects the productivity and quality of algal biomass and products. The high cost of illumination is another problem hindering the commercialization of Haematococcus products. To overcome this drawback, heterotrophic culture approach may be considered. Under heterotrophic growth conditions light is not needed as organic substrates serve as carbon and energy sources for growth and synthesis of secondary metabolites. Also, since lipid accumulation and astaxanthin biosynthesis are connected in space and time the effect of carbon source on lipid accumulation can have significant effect on overall productivity. It has been shown in Haematococcus and other microalgae lipid content and lipid profiles of microalgae are dependent on the cultivation conditions with various stress factors such as starvation or salt stress are efficient triggers of lipid accumulation, and can result in the alteration of fatty acid profiles due to cellular adjustment to particular stressor (Damiani et al., 2010; Lei et al., 2012; Saha et al., 2013; Chen et al., 2015). Various types of organic carbon sources have been used for heterotrophic cultivation and induction using acetate has been found effective for Haematococcus encystment and initiation of astaxanthin production (Kobayashi et al., 1991; Kakizono et al., 1992; Orosa et al., 2000; Hata et al., 2001; Kang et al., 2005). However, unlike many microalgae in which oversupply of easily accessible carbon in combination with nitrogen limitation yields diversion of

Shah et al. Astaxanthin Production from H. pluvialis

 4 | Summary of various methods of *H. pluvialis* biomass cultivation and corresponding astaxanthin productivities.

TABLE

aObtained from one-step culture process.

performed outdoors;

\*Detail medium composition

dObtained from a two-step process in which astaxanthin

 can be found in relevant references.

bObtained from two-step culture process. Productivity

 value was calculated based on total time required by the "green stage" and "red stage" of cultivation.

> productivity

> was calculated based on time spent on the "red stage" only.

cInduction of astaxanthin

 was

the carbon flux toward lipid accumulation (Miao and Wu, 2006; Jia et al., 2014), H. pluvialis grows at a relatively low rate (0.22 d −1 ) and accumulates negligible amount of astaxanthin, too low to be considered for commercial scale production if single step cultivation is considered (Kobayashi et al., 1992, 1997a; Moya et al., 1997). Additionally, heterotrophic cultivation of Haematococcus increases the risk of bacterial or fungal contamination (Hata et al., 2001; Olguín et al., 2012). H. pluvialis can be also produced indoors mixotrophically employing an organic acid (e.g., acetate) or carbohydrates as an additional carbon and energy source (Kobayashi et al., 1993). Studies have shown that both growth and astaxanthin production can be enhanced under mixotrophic culture conditions. A final cell density of 0.9–2.65 g L−1 and a maximum astaxanthin content of 1–2% DW were obtained from mixotrophic cultures of H. pluvialis (Chen et al., 1997; Zhang et al., 1999; Wang et al., 2003). A sequential, heterophotric-photoautotrophic culture mode was also explored. Heterotrophic culture was used in the green stage to produce algal biomass, while astaxanthin production was induced in photoautotrophic culture conditions. The induction of astaxanthin accumulation was performed under nitrogen deprivation conditions and whilst using bicarbonate or CO2 as carbon sources. As a result, a very high cellular astaxanthin content of 7% (DW) was achieved, 3.4-fold higher than heterotrophic induction whilst astaxanthin productivity of 6.25 mg L−1 d −1 was obtained (Kang et al., 2005). Results indicate that photoautotrophic induction of astaxanthin production in H. pluvialis is more effective than heterotrophic one. Based upon the information obtained thus far, heterotrophic and mixotrophic culture modes are less cost-effective than the photoautotrophic one for Haematococcus mass culture.

# Microbial Contamination and Possible Control Measures

Since interest in commercial microalgae cultivation is increasing, microbial contaminants that hamper production by resulting in reduced biomass yield and quality received great attention recently. Mass culture of H. pluvialis is reported to be contaminated by fungal parasites and zooplanktonic predators (e.g., amoebas, ciliates, and rotifers), as well as other microalgae and cyanobacteria (Han et al., 2013). A parasitic chytrid/blastoclad fungus Paraphysoderma sedebokerenses is found to be responsible for reduced astaxanthin productivity and frequent culture collapses in commercial Haematococcus cultivation facilities (Hoffman et al., 2008; Strittmatter et al., 2015). Detection of contaminants is prerequisite for preventing and controlling of microbial contamination in mass microalgal culture. Methods of detection usually include microscopy and staining, flow cytometry, molecular based detection and monitoring. In order to cope with microalgae culture contamination, the techniques that are generally used include abiotic stresses such as NO− 3 limitation, pH stress, temperature stress, light stress, toxic substances, and shear forces. There are some other techniques for parasite removal including salvage harvest, chemical agents (abscisic acid, copper sulfate), physical methods, biological methods (selective breeding and biological agents) (Carney and Lane, 2014). Recently, several patent applications relating to control of fungus P. sedebokerenses have been developed in the USA and China to protect production losses in commercial Haematococcus culture facilities across the world (McBride et al., 2013; Zhang et al., 2013; Carney and Sorensen, 2015).

#### Harvesting

Harvesting remains one of the most challenging issues and a limiting factor for commercial algal biomass production. Harvesting of H. pluvialis refers to the selection of appropriate techniques to recover the "red" biomass, after the accumulation of astaxanthin in the cells and also that can facilitate costefficient astaxanthin extraction in the extraction phase. For the large scale harvesting of H. pluvialis centrifugation is the most common method and combined with other processes. Usually haematocysts are separated from the water through passive settling and subsequently concentrated with centrifugation (Lorenz and Cysewski, 2000; Olaizola, 2000; Li et al., 2011; Han et al., 2013; Pérez-López et al., 2014). Through the combination of these processes total suspended solid of 13.5% in the algal cake is achieved (Li et al., 2011). Flotation and disk-stack centrifugation have been also reported as another alternative for H. pluvialis harvest. Both showed more than 95% biomass recovery efficiency (Panis, 2015).

# Cell Disruption

Different techniques have been developed in order to disrupt the algal cell and recover the intracellular metabolites. The most appropriate cell disruption methods to enhance recovery of astaxanthin from H. pluvialis at a commercial scale involve mechanical processes and more specifically expeller pressing and bead milling (Lorenz and Cysewski, 2000; Olaizola, 2003; Mercer and Armenta, 2011; Razon and Tan, 2011). During pressing (pulverization) microalgae cells are squeezed under high pressure in order to rupture the thick sporopollenin wall. Main advantage of expeller pressing is simple operation and minimization of contamination from external sources. Algal oil recovery efficiency of 75% can be achieved in a single step. Bead milling utilizes vessels filled with tiny glass, ceramic or steel beads that are agitated at high speeds. The dried biomass is fed in these vessels, where continuous exposure of biomass to the grinding media (beads) leads to cell-wall rupture, and subsequent release of intracellular compounds. This method is most effective when biomass concentration in the algal cake after harvesting is between 100 and 200 g/l (Greenwell et al., 2010). Both methods are reliable and widely applied for the H. pluvialis cells disruption at a commercial scale.

### Dehydration

In commercial scale astaxanthin production, dehydration (drying) ensures the quality of the pigment and leads to the formulation of the final product (Mata et al., 2010; Li et al., 2011). After algal cell walls have been disrupted, biomass must be processed rapidly within few hours to avoid spoilage. Thus, dehydration is a process applied prior to recovery of the desired metabolite, in order to extend the shelf-life of the algal biomass (Mata et al., 2010). The most known dehydration techniques that have been employed on microalgae are solar drying, spray drying, and freeze drying (Molina Grima et al., 2003; Brennan and Owende, 2010; Milledge, 2013). Spray drying has been considered as the most appropriate method to dry high-value microalgal products including H. pluvialis astaxanthin (Leach et al., 1998; Brennan and Owende, 2010; Li et al., 2011; Han et al., 2013; Milledge, 2013; Panis, 2015). The recovery efficiency of dry biomass (in powder) using this method exceeds 95% and in some occasions may approach 100% (Leach et al., 1998). After spray drying, the moisture content in "red" biomass is lowered to about 5% (Pérez-López et al., 2014). The main drawbacks of spray drying include high operational costs and the risk of microalgae pigments deterioration (Molina Grima et al., 2003).

### Recovery of Astaxanthin

Once the cell wall is disrupted and the biomass is fully dried, the recovery of the desired product is possible. Astaxanthin is a lipophilic compound and can be dissolved in solvents and oils. There is an abundance of astaxanthin extraction methods from H. pluvialis utilizing solvents, acids, edible oils, supercritical carbon dioxide (SC-CO2) as well as microwave-assisted and enzyme-assisted approaches. Among the recovery methods used solvent extraction and supercritical carbon dioxide (SC-CO2) extraction are considered as the most efficient, compatible, and widely used methods for astaxanthin extraction from H. pluvialis. The summary of various extraction methods of astaxanthin from H. pluvialis with recent updates is presented in **Table 5** Supercritical carbon dioxide (SC- CO2) extraction has been widely used for industrial applications due to its many processing advantages. Due to low critical temperature of carbon dioxide, the SC- CO2 system can be operated at moderate temperatures, preventing the degradation of valuable substances (Machmudah et al., 2006). Several studies have reported experiments on supercritical CO2 extraction for the recovery of astaxanthin from H. pluvialis. Considering astaxanthin quality as the most important criterion, supercritical CO2 extraction is the most favorable option. Supercritical CO2 provides shorter extraction time and limits the use of toxic organic solvents. By contrast to most solvents, CO2 is relatively cheap, chemically inert, non-toxic, and stable (Guedes et al., 2011). Supercritical fluid extraction has also been tested with Haematococcus, aiming at improving the extraction efficiency. For instance, supercritical carbon dioxide (SC- CO2) coupled with ethanol or vegetable oil as a co-solvent can further increase the extraction efficiency of astaxanthin (80–90%) (Nobre et al., 2006; Krichnavaruk et al., 2008). There is an array of alternative approaches that can assist astaxanthin extraction from H. pluvialis such as solvents, acids, edible oils, enzymes, or pressurized liquids (Sarada et al., 2006; Kang and Sim, 2008; In, 2009; Jaime et al., 2010; Zou et al., 2013; Dong et al., 2014) Pressurized liquid extraction has several advantages over traditional solvent extraction. PLE requires shorter time, can be automated, uses less solvent, and retains the sample in an oxygen-free and light-free environment in contrast to traditional organic solvent extraction (Jaime et al., 2010). Recently, a simple method for the direct extraction of lipids from high moisture H. pluvialis microalgae was successfully achieved using liquefied dimethyl ether (Boonnoun et al., 2014).

# BIOREFINERY APPROACH FOR *H. PLUVIALIS*

Microalgae have been often proposed as third generation feedstock for biofuel production that does not compete for freshwater or land resources (Daroch et al., 2013a). However, despite significant advances in recent years it becomes apparent that cultivation of microalgae for the sole purpose of biofuel production is unlikely to be possible unless a major low-energy breakthrough technologies in algae cultivation, dewatering, and harvesting are developed (Li et al., 2015). In the meantime, microalgae are extremely important producers of many high-value nutraceutical compounds such as polyunsaturated fatty acids or astaxanthin that can justify high cost of microalgae cultivation and processing technologies. Integration of simultaneous production of numerous compounds within one system maximizing the benefits and limiting the costs is called biorefining (Li et al., 2015). Taking into consideration these findings H. pluvialis emerges as a very useful organism for the development of a dedicated microalgal biorefinery. It fits numerous requirements of for the development of first microalgal biorefineries especially the "high value product first" principle (Li et al., 2015). First, H. pluvialis is the best-known producer of astaxanthin-high value product worth in excess of several thousand US \$ per kilogram. This product itself can easily justify expensive cultivation systems required for this organism. Second, H. pluvialis grown under nutrient starvation conditions induces both carotenogenesis (astaxanthin formation) and deposition of storage materials (triglycerides). It has been shown that these two responses are closely related and coincide in both space and time and triglycerides are essential for deposition of astaxanthin inside lipid bodies to confer its protective function (Solovchenko, 2015). In traditional approaches of microalgae to biofuels starvation-induced lipid accumulation is considered as significant challenge for commercialization of these systems as the overall lipid productivity of culture can drop significantly due to impaired growth rates under starvation conditions (Daroch et al., 2013b). In case of high value product like astaxanthin this drop becomes much less of the burden as the high value of the main product will compensate for this delay in final product formation. Due to the coexistence of astaxanthin and triglycerides in space and time it is possible to simultaneously obtain high value product (astaxanthin) and a biofuel feedstock (triglycerides) from a single algal feedstock. Since fatty acid content in the astaxanthin-containing 'red' cells can be as high as 30–60% of algae dry weight (Solovchenko, 2015) making H. pluvialis a very good candidate for biorefining strain. The fatty acid profiles of the alga have been evaluated by several studies and are summarized in **Table 2**, indicating that fatty acid profiles of the algae are suitable for biodiesel production

#### TABLE 5 | Summary of astaxanthin extraction methods from *H. pluvialis*.

SC, supercritical; EA, ethyl acetate.

(Damiani et al., 2010). Third, H. pluvialis have been found to be a mixotrophic alga what is highly advantageous for development of microalgae biorefineries. H. pluvialis is capable of utilizing carbon dioxide, carbonates, and carbohydrates as carbon sources, this opens a possibilities of lowering cultivation costs and/or speeding up the cultivation of the strain through using various waste streams like flue gasses or waste streams containing carbon and nutrient compounds (Wu et al., 2013). Auto-, hetero-, and mixo-trophic cultivation modes require energy and nutrients, both of which can be to an extent recycled from anaerobic digestion process. Carbon sources vary depending on cultivation mode. Photoautotrophic cultivation requires CO2 that can be recycled from energy production at anaerobic digestion stage. Heterotrophic cultivation requires reduced carbon source—such as carbohydrates or acetate which need to be supplied from alternative source. These compounds can also originate from waste streams. For example, food industry is rich in carbohydrate-rich waste streams that can be used in heterotrophic cultivation of H. pluvialis (Wang, 2014). Mixotrophic cultivation can take advantage of both sources of carbon. After simultaneous extraction of both high value product-astaxanthin and biofuel product-triglycerides algal cake composed of residual biomass can be utilized as a supplementary feedstock for biogas production using anaerobic digestion that would further assist in extraction of residual energy from this integrated bioprocess. These three features of H. pluvialis make it a suitable strain for the development of algal biorefineries producing high value product (astaxanthin) and biofuel molecule (biodiesel and/or biogas). The proposed biorefinery scheme is presented on **Figure 8** and employs a classical two stage cultivation of H. pluvialis in green and red stage.

# CURRENT GLOBAL MARKET AND MARKET PLAYERS OF *H. PLUVIALIS* ASTAXANTHIN

Synthetic astaxanthin dominates current commercial market, of the total value exceeding \$200 million, corresponding to 130 metric tons of product per year (Li et al., 2011). According to recent reports, microalgae-derived astaxanthin corresponds to less than 1% of the commercialized quantity due to much lower price of synthetic astaxanthin and technological problems associated with large-scale algae cultivation (Koller et al., 2014; Pérez-López et al., 2014). In recent years, there has been a growing trend toward using natural ingredients in food, nutraceutical, and cosmetic markets, resulting from increasing concerns for consumer safety and regulatory issues over the introduction of synthetic chemicals into the human food chain. The demand for natural astaxanthin derived from H. pluvialis in the global market has been "sky-rocketing" in recent years owing to increasing consumer awareness of its health benefits. Global market for both synthetic and natural source astaxanthin in aquaculture feed, nutraceuticals, cosmetics, and food and

beverages is estimated at 280 metric tons valued at \$447 million in 2014. It is further projected to reach 670 metric tons valued at \$1.1 billion by 2020 (Industry Experts, 2015; Panis, 2015). Synthetic astaxanthin, astaxanthin rich Phaffia yeast, and Paracoccus bacteria are predominantly used in the aquaculture sector, while the astaxanthin derived from H. pluvialis is the main source for human applications such as dietary supplements, cosmetics, and food and beverages. Nowadays, the estimated market value of astaxanthin depending on products' purity varies from \$2500–7000/kg to about \$15,000/kg pigment from H. pluvialis in some cases (Borowitzka, 2013; Koller et al., 2014; Pérez-López et al., 2014; Industry Experts, 2015), while the production cost is estimated at about \$1000 per kg of astaxanthin from H. pluvialis (Li et al., 2011). Natural astaxanthin is three to four times more valuable than the synthetic alternative in nutraceutical and pharmaceutical markets (Han et al., 2013). Since there is growing market demand for natural astaxanthin for specific commercial applications (e.g., the nutraceuticals market) to replace the synthetic astaxanthin, mass cultivation of H. pluvialis in industrial scale has great potential and attractive business opportunity. However, current market demand for natural astaxanthin is not met. It is expected that in the foreseeable future after the optimization of the production technology, the production costs of the natural astaxanthin from H. pluvialis should be more competitive to these of the synthetic alternative (Pérez-López et al., 2014). Since the mid 1990's, several leading companies are successfully producing H. pluvialis at commercial scale and marketing natural astaxanthin from H. pluvialis worldwide. The list of the leading companies with their products is presented in **Table 6**. The size of the nutraceutical astaxanthin market is growing day by day and this market is very attractive to Haematococcus astaxanthin producers since the price of these products is significantly higher than those of feed applications. Haematococcus producers need to invest their attention for increasing astaxanthin production capacity to meet the global demand. It is worthy to mention that several manufacturers already have doubled the cultivation capacities in recent years. Apart from an increase in existing players' capacities, new producers, such as BGG in China, have entered the market with significant production capacities.

# MAJOR CHALLENGES FOR THE IMPROVEMENT OF *H. PLUVIALIS* BIOMASS AND ASTAXANTHIN PRODUCTION

There are many challenges and problems for the development of large scale production of biomass and astaxanthin from H. pluvialis. Due to these obstacles the productivity can be hampered and in some cases a failure of the production system can make the production process economically unsustainable. Following issues are considered as most important challenges for the development of H. pluvialis astaxanthin production process:

• Lack of effective solution to prevent or treat microbial contaminations of mass cultures in a commercial scale.

#### TABLE 6 | Leading commercial companies and their *H. pluvialis-*derived astaxanthin and related products in the world market.

#### CONCLUSION AND PERSPECTIVES

This review provides an insight about the latest scientific and technological advancements in various aspects of astaxanthin-producing microalga H. pluvialis such as cell biology, reproduction, biosynthesis pathway, stress mechanism, biomass production, and downstream processing. It also contemplates a broader image including potential benefits, global market opportunities and integration of astaxanthin production into biorefining. In recent years there is an increased interest for natural astaxanthin from green microalga H. pluvialis. Wide ranges of scientific improvements have been achieved during the last decade in terms of productivity and bioprocessing in order to obtain a refined astaxanthin product. Yet its commercial production, especially for low-end markets is too expensive for mass adoption of natural astaxanthin over its synthetic counterpart. H. pluvialis has been shown to be cultured in photoautotrophic, heterotrophic or mixotrophic growth conditions in various culture systems. Research have been conducted on the optimization of the various culture parameters, such as growth medium composition, light, pH, temperature etc. to achieve high biomass and astaxanthin production. Most of these parameters have been optimized and found different for biomass accumulation and astaxanthin production. Little can be done to address this limitatiation as it is funadamentally connected with the life cycle of this microalgae. We believe there exist three key areas where further improvements are required and interesting novel approaches have been recently developed: cultivation efficiency and cost; good cultivation practice and predator control; and astaxanthin isolation and purification.

First, due to complex life-cycle of H. pluvialis it is important to maximase cell densities of alga at "green stage" of cultivation to maximize astaxanthin yield from the "red stage." We think that a number of recent developments can make significant impact in maximizing cell densities. Especially attached cultivation approach and a two-stage "perfusion culture" system can be considered most promising due to the capability of producing several fold higher biomass and astaxanthin productivity and some other benefits such as lower water consumption and smaller risk of contamination. These improvements may boost economic benefits and reduce production cost of astaxanthin from H. pluvialis (Park et al., 2014; Wan et al., 2014b; Zhang et al., 2014). Alternatively, utilization of supplementary carbon source and adoption a two-stage sequential heterotrophic-photoautotrophic approach could improve biomass and astaxanthin production. Especially utilization of waste carbon and nutrient sources in biorefinery setup could help to decrease cultivation costs. Unfortunately, these researches are still in laboratory stage and need to be tested in large-scale commercial production for further validation.

Second, control of contaminants, parasites, and predators remains to be primary concern for Haematococcus growers and major issue in culture stability and astaxanthin productivity. Since there is very little that can be done once contamination takes place it is important to limit the possibility of such disruption and identify it as soon as possible, and avoid spreading to other parts of culture. Traditional detection methods such as microscopy and staining can be used to visualize algal parasites, however this technique may be too labor intensive to perform on a routine basis for most commercial operations. For routine detection, more automated systems such as flow cytometry would be ideal. Alternatively, molecular-based techniques that are considered as the most informative and sensitive for the detection and identification of parasites. Following techniques are worth further exploring DNA sequencing (Sanger, shotgun, or next generation) and then monitoring for these specifically using qPCR or phylochip technology. Decreasing costs of next generation DNA sequencing can make DNA sequencing for culture diagnostic purposes more accessible in the near future.

Third, combination of low cell densities and robust trilayer cell walls of astaxanthin-containing aplanospores make isolation of astaxanthin difficlut and expensive. Currentlly harvesting by centrifugation, cell wall disruption by expeller pressing and bead milling are the most common described methods for commercial scale astaxanthin production from H. pluvialis. After cell walls disruption, biomass is usually processed by spray drying or freeze drying. A number of astaxanthin extraction methods such as (solvents, acids, edible oils, supercritical carbon dioxide, microwave-assisted, and enzyme-assisted approaches have been reported for H. pluvialis and supercritical carbon dioxide (SC- CO2) extraction has been widely used for industrial applications. Two recently developed methods allow efficient extraction of astaxanthin-containing lipids from wet biomass at yields comparable to conventional drying-solvent extraction method. Efficient extraction of astaxanthin from wet H. pluvialis biomass was achieved with liquefied dimethyl ether (Boonnoun et al., 2014) and also cell germination process in conjunction with ionic liquids treatment (Praveenkumar et al., 2015).

Despite significant advances in research and development of H. pluvialis astaxanthin production is still in laboratory stage and often faces difficulties to become implemented in large-scale commercial production. There is a number of other areas of improvement that will contribute to the expansion of Haematococcus production capacity, lowering the production cost, and increasing market penetration at low end applications. These include: next-generation culture systems along with advanced management practices; better understanding of astaxanthin biosynthesis, metabolic pathways and their regulation, genetic engineering, and omics-scale understanding of astaxanthin accumulation; development of genetic manipulation toolbox; exploration of integration of H. pluvialis cultivation with other processes. Yet, we firmly believe that three key areas of focus should be: cultivation efficiency and cost; good cultivation practice and predator control; and astaxanthin isolation and purification. Further developments in these fields can have a profound effect on the commercial deployment of H. pluvialis astaxanthin products and can act as a catalyst for the development of an entire microalgae industry in the near future.

### AUTHOR CONTRIBUTIONS

MS, collected data, participated in preparation of draft manuscript, participated in assembly and editing of the final manuscript; YML, collected data, participated in preparation of draft manuscript; JJC, participated in assembly and editing of the final manuscript; MD, collected data, participated in preparation of draft manuscript, participated in assembly and editing of the final manuscript.

# FUNDING

Authors would like to acknowledge the support of National Natural Science Foundation of China for Young International Scientists Grant no.<PHONE_NUMBER>4 and<PHONE_NUMBER>7, Shenzhen Municipal Government for Special Innovation Fund for Shenzhen Overseas High-level Personnel KQCX20140521150255300 and Shenzhen Knowledge and Innovation Basic Research Grant JCYJ20150626110855791, State Ocean Administration Grant 201305022, and National Thousand People Plan Grant of Jay Jiayang Cheng.

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pluvialis and Nannochloropsis. Appl. Energ. 88, 3507–3514. doi: 10.1016/j.apenergy.2010.12.052

astaxanthin with attached cultivation. Bioresour. Technol. 163, 26–32. doi: 10.1016/j.biortech.2014.04.017

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2016 Shah, Liang, Cheng and Daroch. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

*Yibo Xiao† , Yue Lu† , Junbiao Dai\* and Qingyu Wu\**

*Ministry of Education Key Laboratory of Bioinformatics, Center for Synthetic and Systems Biology, School of Life Sciences, Tsinghua University, Beijing, China*

#### *Edited by:*

*Diego Mauricio Riaño-Pachón, Centro Nacional de Pesquisa em Energia e Materiais, Brazil*

#### *Reviewed by:*

*S. Venkata Mohan, CSIR-Indian Institute of Chemical Technology, India Jaciane Lutz Ienczak, Centro Nacional de Pesquisa em Energia e Materiais, Brazil*

#### *\*Correspondence:*

*Junbiao Dai<EMAIL_ADDRESS>Qingyu Wu<EMAIL_ADDRESS>
*† Yibo Xiao and Yue Lu have contributed equally to this work.*

#### *Specialty section:*

*This article was submitted to Plant Biotechnology, a section of the journal Frontiers in Bioengineering and Biotechnology*

*Received: 30 July 2015 Accepted: 02 October 2015 Published: 19 October 2015*

#### *Citation:*

*Xiao Y, Lu Y, Dai J and Wu Q (2015) Industrial fermentation of Auxenochlorella protothecoides for production of biodiesel and its application in vehicle diesel engines. Front. Bioeng. Biotechnol. 3:164. doi: 10.3389/fbioe.2015.00164*

Microalgae-derived biodiesel has been regarded as a promising alternative for fossil diesel. However, the commercial production of microalgal biodiesel was halted due to its high cost. Here, we presented a pilot study on the industrial production of algal biodiesel. We began with the heterotrophic cultivation of *Auxenochlorella protothecoides* in a 60-m3 fermentor that produced biomass at 3.81 g L−1 day−1 with a neutral lipid content at 51%. Next, we developed plate-frame filter, natural drying, and ball milling methods to harvest, dry, and extract oil from the cells at low cost. Additionally, algal biodiesel was produced for a vehicle engine test, which indicated that the microalgal biodiesel was comparable to fossil diesel but resulted in fewer emissions of particulate matter, carbon monoxide, and hydrocarbon. Altogether, our data suggested that the heterotrophic fermentation of *A. protothecoides* could have the potential for the future industrial production of biodiesel.

Keywords: microalgae biodiesel, *Chlorella*, heterotrophic cultivation, cost assessment, diesel engine test

# INTRODUCTION

We will eventually have to face the time when all fossil fuels, i.e., petroleum, natural gas, and coal, run out. Even before that time, climate change might destroy the entire planet if we continue to rely heavily on fossil fuels as we have, and we are now already experiencing warming of over 1°C (http://news.yahoo.com/not-run-fossil-fuels-op-ed-231243174.html). Recent advances in drilling and hydraulic fracturing technology to explore shale gas (Weber and Clavin, 2012) are exciting and may relieve concerns about fossil fuels for some time. However, even this resource may eventually be exhausted, as it is still not infinite. Meanwhile, concerns are rising due to its potential damage to the environment, especially land and habitat fragmentation, and its impact on air and water quality (Entrekin et al., 2011; Olmstead et al., 2013; Vidic et al., 2013). Therefore, there continues to be interest in developing new technologies to utilize other renewable and sustainable energy sources, such as solar, wind, geothermal, and biomass (Hidy, 2012).

Due to their faster growth rate, higher lipid content and less impact on food security than agriculture-based feedstocks, algae have attracted increasing amounts of attention for biofuel production in recent years (Brennan and Owende, 2010; Weyer et al., 2010). However, projections have largely depended upon small-scale experimental data. Most open pond systems and photobioreactors have been used to culture microalgae for high-value products but not for biodiesel due to the limitations of the lipid content under such cultivation conditions and the high cost for biomass (Rawat et al., 2013). To simultaneously achieve fast cell growth and high lipid content, the heterotrophic cultivation of *Auxenochlorella protothecoides* has been developed (Miao and Wu, 2006; Xiong et al., 2008; Ceron-Garcia et al., 2013). After optimization, the lipid productivity of *A. protothecoides* could reach 11.8 g L−1 day−1 (Xiong et al., 2010), much higher than that of photoautotrophic algae [0.2 g L−1 day−1 as reported by Rodolfi et al. (2009)]. It could, therefore, potentially be applicable for the commercial production of microalgal biodiesel. In addition, a previous study to evaluate microalgal biodiesel prepared from an 11-ton bioreactor suggested that it was comparable to conventional fossil fuels and complied with the US biodiesel standard (Li et al., 2007). Lately, many improvements at the lab scale for biodiesel preparation from algal lipids or directly from dry and wet cells using chemical or enzymatic approaches have been made to increase the efficiency of transesterification and to obtain high quality final products (Miao and Wu, 2004; Johnson and Wen, 2009; Levine et al., 2010). However, due to the difficulty in obtaining enough biomass for industrial production of microalgal biodiesel, only a few studies in recent years have been able to apply algal biodiesel to original or unmodified vehicle engines (Haik et al., 2011; Al-Lwayzy and Yusaf, 2013).

With the aim to produce microalgal biodiesel for commercial application, we report here the first large-scale trial of an algal biodiesel manufacturing and evaluation process, including the step-wise enlargement of biomass production, cell harvesting and drying, and biodiesel preparation by transesterification followed by quality assessment. After generating enough algal biodiesel for application tests, combustion experiments in a four-cylinder vehicle diesel engine were conducted, and several main features, including engine torque and emission, were monitored.

#### MATERIALS AND METHODS

#### Microalga Strain and Basal Culture Medium

The microalga *A. protothecoides* strain was originally obtained from the Culture Collection of Algae at University of Texas (Austin, TX, USA) and screened in the Laboratory of Microalgae Fermentation and Bioenergy at Tsinghua University, Beijing, China. The basic culture media were the same as reported previously (Xiong et al., 2008).

### Production and Harvest of Microalga Biomass

The production of *A. protothecoides* biomass was carried out at North China Pharmaceutical Huasheng Co., Ltd., and the scalingup fermentation was divided into three steps (**Figure 1**). First, the seed cultivation was performed in three 700 L stirred tank bioreactors containing 300 L basal medium with turbine impellers. Two liters of *A. protothecoides* cells at the exponential stage was inoculated into each fermentor, for which the temperature, pH, aeration rate, and agitation speed were initially set at 28 ± 0.5°C, 6.3 ± 0.1, 25 m3 h−1 (1:1 vvm), and 360 rpm, respectively. The pressure of the inner fermentor was kept at 0.04 Mpa except for the 60-m3 fermentor, which was kept at 0.2 Mpa. After 10 days of cultivation, the cells were partly inoculated into an 11-m3 fermentor containing 5 m3 of medium and kept at the same conditions

6, with 40 m3 medium) industrial fermentors. S1–S4 are supplemental tanks for feeding glucose, nitrogen, NaOH, and antifoam, respectively.

as those of the 700 L fermentors. Finally, cells at the exponential stage from the 11-m3 fermentor were completely inoculated into a 60-m3 fermentor with 40 m3 of medium. We followed the optimized concentration of nitrogen and carbon source according to our previous published paper (Xiong et al., 2008). During fermentation, the cells were sampled every 6 h to determine their glucose concentration by the dinitrosalicylic acid (DNS) method (Miller, 1959) and the organic nitrogen concentration by the Nessler's reagent method (Thompson and Morrison, 1951). Concentrated glucose solution (40%), corn steep liquor (0.3%), and antifoam solution (10%) were fed depending upon the substrate consumption. Five molar KOH was injected continuously to keep the pH at ~6.0.

#### Cell Growth Estimation

To monitor the cell growth, 10 mL cell suspensions were collected. After centrifugation followed by twice washing with distilled water, the cells were oven dried until constant weight to determine the cell density (dry cell weight g L−1 ).

At the end of the fermentation, the cells were harvested by filtration at 0.05–0.12 MPa using plate-frame vertical pressure filters with 32 pieces of cloth for 12 h and were then spread on the open ground under sunlight for 2 days to dry. The water fraction of the algal biomass, represented by the moisture content (MC), was calculated as

$$\text{MC} = \left(\left(\mathcal{W}\_{\text{w}} - \mathcal{W}\_{\text{d}}\right) / \,\mathcal{W}\_{\text{w}} \times 100\,\%\right.\tag{1}$$

where *Wd* is the weight of the completely dried cells using a frozen vacuum dryer and *Ww* is the weight of the biomass (*Ww* − *Wd* was the water left in the biomass).

#### Oil Content Determination and Extraction

For the oil content (OC) determination, 10 g freeze-dried cells was hand-ground with silica in a mortar. The oil was extracted with *n*-hexane by the Soxhlet extraction method for 72 h. The *n*-hexane was subsequently evaporated, and the OC (weight of oil/weight of biomass × 100%) was calculated. For large-scale oil extraction, 30 kg dry cells, 30 kg steel grinding balls (3–5 mm in diameter) and 30 L isopropyl alcohol (IPA) were loaded into a 100-L steel ball mill machine (an industrial piece of equipment used in the food, oil and mining industries, XQM-100, Tianchuang Powder Equipment Co., Ltd., China) and ground at 200 rpm for 1 h. After recovering the milling balls using a stainless steel screen with a mesh size of 2 mm, the solution was filtered by a plate and frame filter. The oil-IPA miscella was vacuum distilled to recover the IPA. The crude oil was refined, and the oil extraction efficiency (OEE) was calculated as follows:

$$\text{OEE} = \text{OW}\_x \mid \text{(BW}\_x \times \text{OC}\text{)} \times 100\text{\%} \tag{2}$$

where OWx is the weight of the oil extracted with the ball milling process and BWx is the weight of the dried biomass.

To investigate the effects of the ball mill machine on cell breakage and OEE, the microstructure of the broken cells (after oil extraction) was examined by scanning electron microscopy (Oberkochen, Germany). Both intact and broken cells were dehydrated sequentially with 50, 75, 90, and 100% ethanol (volume fraction) for 1 h each. After natural volatilization, the samples were spread on conductive tape, coated with gold powder and subjected to microscope examination.

#### Preparation of Microalgal Biodiesel

The transesterification of the microalga oil was catalyzed by liquid lipase (12,000 U g−1 , Sichuan Habio Bioengineering Co., Ltd., China) with the volume fraction of catalyzer:methanol (batch addition):oil at 1:10:100. The reaction was carried out in a 50-L bioreactor under constant stirring at 180 rpm and a temperature of 50°C for 8 h. The processes of methanol feeding and product separation were conducted according to a previous report (Li et al., 2007). After washing and distilling the remaining methanol, catalyst, and glycerol, the crude biodiesel at the top layer was rectified to obtain high quality biodiesel in a 100-L distillation tower with a maximum temperature at 220°C and vacuum degree at 0.1 Pa.

# Components and Quality Analysis of Algal Biodiesel

The biodiesel components were analyzed by gas chromatography–mass spectroscopy (GC–MS) (Thermo, USA). A dual-stage quadrupoles GC apparatus was equipped with a Varian VF-5 ms column (30 m × 0.25 mm ID DF = 0.25 μm), and the GC was manipulated with a flow rate at 10 mL min−1 . The physical and chemical properties of the biodiesel were determined according to ASTM D6751 standards at the Institute Analysis Center, SINOPEC CORP. The petroleum diesel (PD) on the market was used for comparison.

#### Vehicle Diesel Engine Test