The ability to alter gene expression, and as such, metabolic pathways within green microalgae, is of a crucial importance in optimizing those algae for biofuel pro­duction. Achieving such alterations and rational modifications of expression relies primarily on the identification of transcription factors (TFs), genes, and enzymes implicated in metabolic pathways driving the synthesis of oil and hydrogenated metabolites that can potentially be used as biofuels. Post-identification, via struc­tural and functional annotations of genomes, the establishment of a library of cloned, ready to transform, open reading frames (ORFs) and TFs makes the optimization process one step closer (Fig. 9.3). Attempts to clone the metabolic ORFeome of model green algae, C. reinhardtti, have been made and published by Ghamsari et al. (2011) and Chang et al. (2011). The structural annotation of the ORFs was based on models generated by the Joint Genome Institute (JGI) (Merchant et al. 2007) and AUGUSTUS gene prediction algorithm (http://augustus. gobics. de/predictions/chlamydomonas/). The outcome leads to the generation of Gateway-compatible clones that can easily be moved to a plethora of available expression vectors.

Another attempt, as part of an ongoing work, is the genome-wide cloning of TFs in C. reinhardtti to add a regulatory dimension to the available clone resources (unpublished data). The completion of such an endeavor will result in a library of TF clones that are Gateway compatible and readily available for transformation to

alter and modify the expression of target genes. Clones can also be obtained from the Chlamydomonas Resource Center (http://www. chlamy. org/). The center hosts a multitude of C. reinhardtti strains, plasmids, molecular kits, and cDNA libraries.

The importance and relevance of cloning has been underlined as a major initial step in the process of algal optimization for biofuel production by making available a library of metabolic and transcription factor clones that can be used to modify the genetic information and expression patterns and, subsequently, directionally alter the metabolism in algal cells toward a higher yield of hydrogenated metabolites suitable for biofuel use.

A method that is similar to the use of metal salts is electro-coagulation-flocculation or ECF. In ECF, metals are electrolytically released from a sacrificial anode. At the cathode, hydrogen gas and hydroxyl anions are produced. The metal cations react with the hydroxyl anions and form metal hydroxides that can cause flocculation in a similar way to metal salts. Due to the high conductivity of seawater, energy demand is much lower when ECF is used in seawater than in freshwater, which makes this method attractive for harvesting marine microalgae (Vandamme et al. 2011). The overall efficiency of the process may be improved by changing the polarity of the electrodes during operation (Kim et al. 2012). Although ECF may be promising for harvesting marine microalgae, contamination of the biomass with metals remains an issue. Recently, an electrolytic method was proposed for flocculating microalgae that uses inert carbon electrodes (Misra et al. 2014).

There is no universal solution to cost effectively harvesting low-density algal cul­tures (pond or photobioreactor) on the scale necessary for commercial-scale algal biofuel production. Traditional methods of centrifugation and drying are too costly to be applicable to biofuel production—therefore, continued innovation is necessary in this area. The diversity of the algal biofuel production processes, ranging from extraction of high lipid biomass of a unialgal culture and conversion of *21 % solid biomass derived from a mixed culture of native grown algal strains by hydrothermal liquefaction to harvesting a secreted product either directly or indi­rectly, make finding a single universal harvesting method elusive. Table 14.3 summarizes the advantages and disadvantages of the various systems described in this review.

Innovative approaches are being tested both in academic and in industrial lab­oratories, and the expectation is that economical harvesting methods, either new or based on adaptations of existing procedures, will be developed that spur the

deployment and commercial success of algal biofuels and associated co-products. All of the recent R&D focused on this area and discussed in this review indicates that innovation is ongoing, and the authors are confident that economical harvesting of dilute algal systems will be achievable in the near future.

14.2 Conclusion

Microalgal harvesting methods are quite varied, and the choice of a particular method is dictated by the ultimate use of the harvested biomass. As companies try to reduce the cost of microalgal-based biofuels by the production of high-value coproducts from the biomass, additional constraints are placed on the harvesting technology to maintain the quality of both the primary biofuel and the coproduct. A careful balance needs to be maintained to reduce the costs of harvesting dilute algal cultures to maintain a positive impact on the overall production costs of the port­folio of products. The continuing research to reduce the costs of algal harvesting will make a positive contribution to reduce the commercial threshold of biobased fuels and bioproducts derived from microalgae.

Acknowledgments The authors gratefully acknowledge financial support from the US DOE CCS Program (Grant No. DE-FE0001888 to Phycal) that funded research presented for filtration of microalgae in this review.

Navid R. Moheimani, David Parlevliet, Mark P. McHenry, Parisa A. Bahri and Karne de Boer

Abstract Microalgae cultivation is a promising methodology for solving some of the future problems of biomass production (i. e. renewable food, feed and bioenergy production). There is no doubt that in conjunction with conventional growth sys­tems, novel technologies must be developed in order to produce the large-scale sustainable microalgae products. Here, we review some of the most promising existing microalgae biomass growth technologies and summarise some of the novel methodologies for sustainable microalgae production.

1.1 Introduction

There has recently been extensive research focus on biology, physiology, engi­neering and their integration for microalgae cultivation to produce sustainable products such as biofuel, food, feed and high-value products. Algae belong to many different and unrelated taxonomic groups that all contain chlorophyll a and are able to utilise solar energy and fix CO2 to produce organic compounds (Borowitzka 2012). More than a dozen algal species have been mentioned in the literature as potential candidates for large-scale cultivation. However, conclusive information obtained through commercial trials is not yet available to assess suitability of most of these species. The ideal microalga must be able to grow very well even under high biomass concentration and varying environmental conditions. It must be able to produce high concentration of product of interest (i. e. high-value products, lipids and hydrocarbons). However, it is unknown how many species of algae exist, with

N. R. Moheimani (H)

Algae R&D Center, School of Veterinary and Life Sciences, Murdoch University, Murdoch, WA 6150, Australia e-mail: n. moheimani@murdoch. edu. au

D. Parlevliet • M. P. McHenry • P. A. Bahri • K. de Boer

School of Engineering and Information Technology, Murdoch University,

Murdoch, WA 6150, Australia © Springer International Publishing Switzerland 2015

N. R. Moheimani et al. (eds.), Biomass and Biofuels from Microalgae,

Biofuel and Biorefinery Technologies 2, DOI 10.1007/978-3-319-16640-7_1

estimates ranging between several hundred thousand and several million different species—with new types identified all of the time (Guiry 2012). Only a small portion of microalgal species (several thousand) can be kept alive in culture, and only a handful of them have been successfully grown commercially. Table 1.1 summarises the main microalgae species tested for medium — to large-scale pro­duction (especially for feed, high-value products and biofuel).

However, to date, only a few of these species were successfully grown in large scale. Commercial large-scale production of microalgae for bioproducts began in early 1960s and 1970s with Chlorella and Spirulina and followed in the 1980s with production of P-carotene from Dunaliella salina (Borowitzka 2013a). All three species were successfully grown in mixed or unmixed open ponds (Craggs et al. 2013). The ability to grow at highly selective environments is the main reason for the successful growth of these species (Spirulina = high pH and high HCO3-, D. salina = high salinity and Chlorella = high nutrients) (Craggs et al. 2013). Moheimani and Borowitzka (2006) also showed that Chrysotila carterae reliable long-term culture in raceway pond is successful due to the ability of this alga to grow at very high pH. Other species that do not have this selective advantage may need to be grown in closed photobioreactors. The selection of growth technologies or production systems for microalgae will need to be based to a large extent on the microalga of choice and cultivation system.

Many strains of microalgae are able to simultaneously produce oil and many other useful and expensive metabolites, such as antioxidants (Fujita et al. 2008; Ogbonna

2009) and bioactive compounds. Such metabolites can be separated and purified, thereby making the process profitable. After oil extraction, residual biomass can be converted to biofuels, such as bioethanol, biohydrogen, and biomethane via ther­mochemical and biochemical methods (Golueke et al. 1957; Gunaseelan 1997). As an example, the anaerobic digestion of microalgae produces methane, which can then be used to generate electricity. The generated electricity can be used for microalgae cultivation, dewatering, extraction, and transesterification process. Harun et al. (2011) estimated that the electricity generated from methane reduces the cost of biodiesel production by as much as 33 %. It can also be used to produce bioplastic materials (Chiellini et al. 2008), or used as a fertilizer, soil amendment, or feed for fish or livestock (Mulbry and Wilkie 2001; Mulbry et al. 2005; Roeselers et al. 2008).

3.8 Conclusions

Although microalgae biodiesel has many advantages over plant biodiesel in terms of effects on food security and sustainability, the present cost of microalgae oil is still much higher than that of vegetable oils. This is mainly due to low cell growth rate and low final cell concentrations in photoautotrophic cultures. Cell growth rates, final cell concentrations, and in some cases intracellular oil contents are higher in heterotrophic and mixotrophic cultures than photoautotrophic cultures. These lead to increase in oil productivity and reduction in the cost of production. The qualities of oil produced under these culture conditions have also been reported to be more suitable for biodiesel oil production than photoautotrophic microalgae oils. Heterotrophic and mixotrophic culture systems should therefore be exploited for large-scale biodiesel oil production.

Various microalgae species accumulate biomass under different CO2 concentrations. Two experiments on S. obliquus supplied with 12 % CO2 resulted in 1.14 and 1.81 g L 1 as a maximum dry weight biomass yield, whereas a higher biomass concentration (3.5 g L-1) was obtained with Spirulina sp. when using a higher light intensity at the same 12 % CO2 input gas (Table 7.2) (De Morais and Costa 2007a, b). As shown in Table 7.2, the highest biomass concentration of Chlorococcum littorale in three runs under 20 % CO2 was 14.4 g L 1 in a small PBR (Kurano et al.

1995) . There has been several research publications evaluating the biomass pro­ductivity of different strains of microalgae under different CO2 concentrations (De Morais and Costa 2007a; Radmann et al. 2011; Yoo et al. 2010). Table 7.3 shows the biomass productivity of 0.077 g L-1 d-1 for Botryococcus braunii under

5.5 % CO2 (Yoo et al. 2010). A biomass productivity of 0.09 g L-1 d-1 for C. vulgaris was achieved (under 12 % CO2) (Radmann et al. 2011). Similarly, Spirulina sp. was used under 6 % CO2 and biomass productivity of 0.18 g L-1 d-1, signifi­cantly higher than most other reports, was obtained (De Morais and Costa 2007a). This higher biomass productivity might be due to lower CO2 concentration (6 %) used for Spirulina sp. in comparison with 12 and 20 % CO2 supplied for species in other reports. Similarly, microalgal species have shown differing capabilities for CO2 fixation at different concentrations of CO2 (10-20 % is a common range of CO2 for microalgae production systems with enhanced CO2 delivery (Ho et al. 2011)). The CO2 fixation rate of 0.17 g L-1 d-1 was obtained (Eberly and Ely 2012) for Thermosynechococcus elongates at 20 % CO2. Similar input CO2 concentration (20 %) (Tang et al. 2011) was supplied for S. obliquus SJTU-3 and C. pyrenoidosa SJTU-2, fixing 0.244 and 0.223 g L 1 d 1 CO2, respectively. However, the researchers also cultured the same strains under 10 % CO2 and obtained higher fixation rates, such as 0.288 g L-1 d-1 for S. obliquus SJTU-3 and 0.260 for C. pyrenoidosa SJTU-2, when providing high levels of CO2 into culture mediums that leads to acidification and lowering fixation rates of S. obliquus SJTU-3 and C. pyrenoidosa SJTU-2 at 20 % CO2. Research (Kurano et al. 1995) with C. littorale under 20 % CO2 assessed removing CO2 by 4, 0.65, and 0.85 g L-1 d-1 for selected culture volumes of 10 mL, 4 L and 20 L, respectively. The research indicated that C. littorale may achieve a better CO2 fixation rate than T. elongates, S. obliquus SJTU-3, and C. pyrenoidosa SJTU-2, although the differences were not significant. The higher CO2 fixation rate may be either due to engineering issues in scaling up or due to higher light intensity used for C. littorale (15650 lux) compared to other

investigations (11,200 lux) (Table 7.4). Light intensity controls photosynthetic growth in any microalgal system, CO2 removal rates, biomass concentrations, and overall growth rates. While increasing light intensity is usually accompanied by increasing CO2 removal rates in microalgal systems, any photosynthetic system has a saturation point where further increasing light intensity will either produce no benefit or may decrease productivity.

Several integrated toolsets are available for constraint-based analysis. These include Pathway Tools, COBRA Toolbox, SNA, and MetaFluxNet (Table 10.1). The most widely used tools are Pathway Tools and COBRA Toolbox, with COBRA offering

the largest number of tools. A few of these tools (mostly available in COBRA) are described below (Fig. 10.5).

The requisite pH jump for autoflocculation by Mg(OH)2 can be achieved through photosynthesis under CO2-starved conditions, through lime addition, or both. For example, sea water in contact with atmospheric CO2 at PCo2 = 10-35 atmospheres
has a pH of * 8.2 and a total inorganic carbon content of *2 mM. Conversion of this carbon to biomass raises the pH to above 10 and lowers the PCO2 by several orders of magnitude; part of the carbon uptake is by formation of calcite. Such alkaline swings are routinely observed in sewage treatment ponds when photo­synthetic activity is high [for example see (Ayoub et al. 1986; Sukenik and Shelef 1984)]. pH boosting with slaked lime (calcium hydroxide) is relatively inexpensive, and calcium in flocculated microalgae is considered a plus if the spent microalgae are to be used as cattle feed (e. g. Schlesinger et al. 2012). Schlesinger et al. (2012) showed that lime usage could be cut by having an earlier, CO2-starvation step.

Figure 13.1 shows calculated changes in sea water chemistry as lime is added: as pH rises, first calcite forms, and then near pH 10.3 Mg(OH)2 forms. Ca levels rise as lime addition proceeds; Mg levels decrease. The curves in Fig. 13.1 were cal­culated using the chemical speciation code PHREEQC (Parkhurst et al. 1999) and the llnl. dat thermodynamic database. The solubility product of freshly precipitated Mg(OH)2 was set to 10-92 (after Gjaldbsk 1924). Using the latter value predicts that sea water should precipitate Mg(OH)2 at pH > 10.4. Smith and Davis (2012) saw a precipitate forming above pH 10.4 from BG-11 solution containing

9.6 mM Mg. But maximum microalgae flocculation began at pH 10.2, suggesting that flocculation was driven by charge-reversing Mg+2 sorption to microalgae, formation of metastable surface precipitates of Mg(OH)2, or both (Smith and Davis 2012). In sum, maximum autoflocculation occurs 0.2-0.3 pH units below the pH where Mg(OH)2 is predicted to form. Additional formation of Mg(OH)2 does not increase flocculation (Smith and Davis 2012). It therefore seems reasonable to simply set Mg and pH autocoagulation targets to 0.2 pH units below the pH where Mg(OH)2 is predicted to form. The latter will depend on the amount of Mg and to a lesser extent the amount of calcium in solution.

Smith and Davis (2012) estimate Mg requirements of autoflocculation as 1.5-2.4 % of the microalgae’s ash-free mass. For a 2 g/L microalgae culture, 2.4 % is * 2 mM. There are only roughly 0.4 mM of microalgae surface sites in a 2 g/L

culture, suggesting that the bulk of the Mg forms surface precipitates of Mg(OH)2 on the microalgae through the reaction:

Mg(OH)2$ Mg+2 + 2OH — K = {Mg+2}{OH-}2

Given a known microalgal solution Mg level, the target minimum pH, that is a flocculation critical pH—pHc (see Sukenik and Shelef 1984)—that must be achieved, either through lime addition or CO2-starvation, can be calculated for maximum autoflocculation to occur:

pHc = 9.4 — 0.2-0.5log{Mg+2}

where {Mg+2} is the thermodynamic activity of dissolved Mg. High Mg fluids such as sea water will require a lower pHc than a low Mg brackish water. This is shown in Fig. 13.2 which plots the pHc for sea water (Mg = 0.052 mol/L; pHc = 10.06) and the pHc for successive dilutions of sea water having lower Mg levels. High levels of EOM or bicarbonate that form aqueous complexes with Mg+2 will increase the autoflocculation target pH. To account for activity coefficient and solution speci — ation effects, the calculation is most accurately undertaken using a chemical spe — ciation code.

A small consideration must be made regarding sea water’s pH buffering capacity when it comes to high volume open ponds with sea water medium. The buffering capacity can be attributed to the reaction: 2HCO3- $ CO2 + H2O + CO3-2, in which the acid or base added is consumed by the formation of HCO3 . Then, it will be a function of the total dissolved CO2 [T(CO2)]; that is to say (CO2) + (HCO3) + ( HCO3-) + ( CO32-). Natural sea water representative of the oceans has a T(CO2) of around 2 mM and an average buffer intensity around

0. 3 meq/L (Pytkowicz and Atlas 1975). This is a small amount when considering the cost-effectiveness of inducing autoflocculation, as it has reasonably higher values only for pH close to pKa values (bicarbonate has two pKa’s, because it can

gain a proton to become carbonic acid or lose a proton to become carbonate). According to Pytkowicz and Atlas (1975), the buffer intensity of sea water reaches a maximum at pH around 6.1 and 9.1. A starvation step prior to raising pH could solve the issue of the extra base needed to overcome the pH buffering capacity of sea water. Nevertheless, the cost of autoflocculation is very low even if a small amount of extra base is required to increase pH (Vandamme et al. 2013).

There are several key benefits to a system that can coproduce electricity and chemical energy from the solar spectrum. Our proposed system would filter the light before it is provided to the microalgae ponds. This will reduce the total amount of energy provided to the algae in the parts of the spectrum where it is not required for photosynthesis. This in turn will reduce the heating of the ponds and subsequent evaporation of the water. It is to be noted that most places with high light irradiance, that are suitable for microalgae cultivation, also have a high evaporation rates and limited supplies of freshwater. By reducing evaporation, the use of freshwater in the production facility can be reduced. This also reduces the salinity of the microalgae ponds, allowing alga with a lower salt tolerance to be grown for a longer period of time.

Our proposed system would also generate electrical energy. This can be used to aid the production by powering motors and other electrical items at the production facility. This is advantageous in remote areas where grid connection and stable electricity supplies can be an issue. Introducing a method of cogeneration of electrical energy has benefits in the remote areas that microalgae cultivation takes place, such as northern and central Australia. In these areas, the cogeneration of electricity would reduce the reliance on grid-supplied electricity and diesel gener­ators. By generating some of its own electricity, rather than purchasing electricity (or diesel fuel), the costs associated with production, dewatering, and extraction of oil from microalgae can be reduced which leads to more cost-effective production of biomass.

Alternatively, the electricity can also be used augment the light received by the algae to aid their growth. As shown in the model, we have described that this additional illumination could more than double the amount of energy absorbed by the microalgae. This would result in an increase in productivity and growth.

15.6 Conclusion

While there are a number of factors that influence biomass productivity, photo­synthesis places upper limits on how effectively solar energy can be transformed into chemical energy (in the form of carbohydrate, lipid, and proteins). As ulti­mately light via photosynthesis is the main limit to the growth and a key component of the productivity of microalgae, it is understandable that an increase in the amount light provided for photosynthesis will result in more photosynthesis and thus more productivity. Solar panels are another established mechanism for utilizing the solar spectrum. In this case, they convert sunlight into electricity. While these two methods of energy production would normally be competing for the same resource, we have shown in the model described here that they can complement each other. By allowing the portions of the solar spectrum not required by the microalgae to be diverted to highly efficient solar panels, we can generate both electrical and chemical energy from a single facility.

There are several advantages to using a filter system to divert portions of the solar spectrum to different tasks. These include a reduction in heating and evapo­ration, co-production of electricity, and a subsequent boost in the productivity of the microalgae. This allows for the cheaper and more efficient/sustainable production of biofuel or value-added crops in remote locations which are located away from sources of electrical power.

The growth of algae, irrespective of cultivation system, requires large volumes of water. Almost all areas with high solar energy also have a high evaporation rate. Therefore, it is logical to use sea water for large-scale algae biomass production. As highlighted previously, it is also critical to recycle the culture medium to reduce the nutrient use. Sea water must also be used to replace evaporative loss. This means that the salt concentration in the pond will gradually increase over the time. For instance, in conditions with evaporations rate of 2 m year-1, productivity of 20 g m 2 day 1 and 80 % medium recycling, the medium salinity will rise from

3.5 % NaCl to 25 % NaCl in 490 days. Salinity is usually growth-limiting at the extremes of salt tolerance in some microalgal species, and every microalga has an optimum salinity range (Borowitzka and Moheimani 2013a). The effect of salinity on microalgal growth relates to osmoregulation, which in microalgae is achieved through diverse strategies. Osmoregulatory metabolites are organic substances produced by microalgae that, when the latter are exposed to water stress conditions, respond appropriately to the changes in extracellular water activity. Microalgae main osmoregulators (function as intracellular osmotic regulators) are as follows: (a) polyhydric alcohols (i. e. glycerol, mannitol or sorbitol), (b) variety of glycosides (i. e. galactosyl glycerides, floridoside and isofloridoside) and (c) amino acids (i. e. glutamic acid and proline).

Freshwater algae grow between 0 and 1-2 % NaCl; hypotonic algae grow between 3.0 and 5-5.5 % NaCl; halotolerant algae grow between 6-7 and 14-15 % NaCl; and halophylic algae can grow above 15-16 % NaCl. The majority of microalgae can grow in freshwater and hypotonic conditions. Some microalgae (i. e. diatom, chlorophyta and cyanobacteria) are capable of growth under halotol — erant conditions. However, only a few species of microalgae are hypersaline (i. e. D. salina). There is no single strain of algae capable of optimal growth in the whole range of salinity from sea water to saturation. Interestingly, almost all companies interested in large-scale algae production focus on growing either freshwater or hypotonic algae which will not be sustainable (Moheimani et al. 2013c). Halotolerant algae can normally grow under optimal condition in a wider range of salinities (Fon Sing 2010).

An alternative method of cultivation is to mix microalgae while salinity is being increased. In such a method, a new species can be introduced to the culture of a mono-species while the salinity is rising. That is, a halotolerant microalga species will be introduced to the culture of hypotonic algae when salinity is above the optimum growth condition of the hypotonic algae. If the halotolerant alga can flourish while salinity is increased, the medium use can be maximised. The same technique can be applied for mixed cultures of halotolerant and halophylic mic­roalgae. Such a mixed cultivation technique is yet to be tested at the laboratory or outdoor conditions. One very important advantage of mixed microalgae cultivation is avoiding unnecessary water and nutrient discharge. Considering the species change throughout the cultivation, less negative effect of autoinhibitors is also expected. When the salinity of the culture becomes very high, one option is to have large evaporation ponds for the hypersaline wastewater. Alternatively, this hyper­saline water can also be used in salt gradient solar ponds for generating additional energy.