Environmental Stewardship by Microalgae: Air and Water Juhyon Kang Iowa State University

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2015 Environmental stewardship by microalgae: air and water Juhyon Kang Iowa State University

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Environmental stewardship by microalgae - air and water

Juhyon Kang

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Co-majors: Food Science and Technology; Biorenewable Resources & Technology

Program of Study Committee: Zhiyou Wen, Major Professor Tong Wang Byron F. Brehm-Stecher Hongwei Xin Jacek A. Koziel

Iowa State University

Ames, Iowa

2015

TABLE OF CONTENTS

Page ACKNOWLDEGEMENTS ...... v ABSTRACT ...... vii CHAPTER 1. GENERAL INTRODUCTION ...... 1 CHAPTER 2. LITERATURE REVIEW ...... 2 1. Microalgae ...... 2 1.1. Introduction of microalgae ...... 2 1.2. Application of microalgae ...... 3

2. Effect of CO 2 and nitrogen (ammonia) to microalgae cultivation ...... 4

2.1. Role of CO 2 in microalgae cultivation ...... 4 2.2. Ammonia as a nitrogen source ...... 10 3. Astaxanthin production from microalgae ...... 12 3.1. Chemistry of astaxanthin: Origin and effectiveness ...... 12 3.2. Biosynthesis of astaxanthin ...... 16 3.3. Production of astaxanthin ...... 20 4. Wastewater treatment ...... 24 4.1. Overall wastewater treatment procedure ...... 24 4.2. Current phosphorus removal methods ...... 27 References ...... 31

CHAPTER 3. USE OF MICROALGAE FOR MITIGATING AMMONIA AND CO 2 EMISSIONS FROM ANIMAL PRODUCTION OPERATIONS – EVALUATION OF GAS REMOVAL EFFICIENCY AND ALGAL BIOMASS COMPOSITION ...... 41 ABSTRACT ...... 41 1. Introduction ...... 42 2. Materials and Methods ...... 44 2.1 Cell species and growth conditions ...... 44 2.2 Experimental set up ...... 45

2.3 Evaluation of ammonia and CO 2 mitigation efficiencies by algal cultures ...... 46 2.4. Microalgae cell composition analysis ...... 47 2.5. Statistical analysis...... 47

iii

3. Results and Discussion ...... 48 3.1 Cell growth performance ...... 48 3.2 Ammonia gas removal by the algal culture system ...... 50

3.3 CO 2 gas removal by the algal culture system ...... 51 3.4 Biomass chemical composition ...... 52 Conclusion ...... 56 References ...... 57 CHAPTER 4. ASTAXANTHIN PRODUCTION FROM HAEMATOCOCCUS PLUVIALIS USING AMMONIA GAS FROM ANIMAL PRODUCTION OPERATIONS ...... 70 ABSTRACT ...... 70 1. Introduction ...... 71 2. Materials and Methods ...... 73 2.1. Cell seed and culture medium ...... 73 2.2. Photobioreactor set up ...... 74 2.3. Ammonia feeding strategies ...... 75 2.4. Induction of astaxanthin accumulation ...... 75 2.5. Analyses...... 77 2.6. Statistical analysis...... 78 3. Results and discussion ...... 78 3.1. Effect of initial pH on the cell growth and astaxanthin production ...... 78 3.2. Cell growth and astaxanthin production of H. pluvialis under different N sources ...... 79 3.3. Cell growth and astaxanthin production using ammonia gas as nitrogen source ...... 80 3.4. Cell growth and astaxanthin production under different ammonia feeding strategies ... 83 3.5. Induction of astaxanthin from vegetative cells grown in ammonia gas culture ...... 84 Conclusion ...... 85 References ...... 86 CHAPTER 5. PHOSPHORUS REMOVAL DYNAMICS IN REVOLVING MICROALGAL BIOFILM SYSTEMS TREATING WASTEWATER ...... 97 ABSTRACT ...... 97 1. Introduction ...... 98 2. Materials and methods ...... 100

2.1. Algae cell strain and preparation ...... 100 2.2. RAB configuration and operation...... 101 2.3. Analytical methods ...... 103 2.4. Statistical analysis...... 104 3. Result and discussions ...... 104 3.1. Characterization of the different wastewater streams ...... 104 3.2. Cell growth of different wastewater streams ...... 105 3.3. Nutrient removal of different wastewater streams ...... 106 3.4. Dynamics of luxury P uptake ...... 108 3.5. P content in biomass ...... 111 3.6. P mass balance ...... 111 Conclusion ...... 114 References ...... 115 CHAPTER 6. GENERAL CONCLUSION ...... 130 Future Research Recommendations ...... 131

ACKNOWLDEGEMENTS

First and foremost, I am very grateful to my major professor, Dr. Zhiyou Wen, for his persistent help in every aspect of graduate school and family life. His support, guidance, and understanding have been imperative for finishing my Master’s and doctoral degree- and with a new family. I also want to thank my POS committee members, Dr. Tong Wang, Dr. Byron F.

Brehm-Stecher, Dr. Hongwei Xin, and Dr. Jacek A. Koziel. Their insightful preliminary exam questions were very helpful for building my research philosophy, and their encouragements were truly uplifting and carried me through my graduate years.

I also need to thank my lab mates, Martin Gross, Xuefei Zhao, Zhixia Ji, Clayton Michael,

Yi Liang, Yanwen Shen, as well as my department colleagues. They always reminded me that I am not alone in this journey. My third project received many assistance from an undergraduate student, Monica Primacella, such as maintaining reactors and cleaning lab wares. She definitely lightened my lab work burden.

My appreciation also extends to Food Science and Human Nutrition department. The coursework, department seminars, and group meetings were very educational and kept me updated with current science. I was greatly impressed by Drs. Ruth MacDonald, Lester Wilson,

Stephanie Clark, and Jay-Lin Jane, for their genuine kindness and brilliant scholarly work. I am also grateful for the Writing and Media Center at ISU, for their tremendous help in improving my writing skills. While in ISU, I got several scholarships and especially CCAMPIS and CAP were very helpful to afford for childcare service fee.

My life in Ames has been blessed through St. Thomas Aquinas church in many ways.

Their generosity and hospitality have made me feel at home. Their prayer was truly holy, and I

vi met many whom are genuine examples of a good Christian, such as Bob and Nannet Horton,

Glenn and Pam Sibbel, Peter and Patti Orazem, John and Barb Stafford, Rob and Donna Evans,

Shari Reilly, and so many others that I cannot all list here. From the parish educational programs,

I learned many beauties of the Church and was inspired by other parishioners’ practices. I was fortunate to meet Mary and Steve Stritzel through Teams of Our Lady, as they have been our extended family in Ames.

I have the deepest appreciation for my husband, Younjun Kim, who continually and convincingly kept his trust in me. I began graduate school at ISU since we met and had our first born son, Sunkyu James Kim, during my last semester of M.S. and our second born daughter,

Sunjae Regina Kim in my last semester of Ph.D. It has been God’s grace to meet him and walk together down this long pilgrimage. My life with him has been a blessing and a joy.

vii

ABSTRACT

This body of research aimed to prove microalgal potential to utilize environmental pollutants in an effective and economic way. Ammonia gas, carbon dioxide, and wastewater were targeted as pollutants to remove in three different technical papers.

The first work, an extension of the previous paper, studied carbon dioxide effect on the microalgal ammonia scrubber system. Freshwater green algae Scenedesmus dimorphus, the same species as the previous study, were grown in a flat panel photobioreactor aerated with ammonia- and CO 2-laden air. The ammonia mass loading was fixed at an optimal level found in the previous study and CO 2 mass loading rates was varied in a range found in typical concentrated animal feeding operations. The data showed high ammonia and CO 2 removal rate and removal efficiency and the biomass contained amino acid profiles close to the ideal protein profiles for typical animal feeds, with a high ratio of essential amino acids. Therefore, microalgae effectively mitigate ammonia and CO 2 emissions, with a potential for use as an animal feed additive.

The second work was still about microalgal ammonia scrubber system using different species, Haematococcus pluvialis, known as the most productive astaxanthin producers, to enhance its economic feasibility. Currently, most H. pluvialis cultivation systems use two stage mode: optimal growth mode and stress mode to induce astaxanthin. In this study, it was hypothesized that ammonia gas can serve as both growth nutrient and stress to induce astaxanthin. Tapered bubble columns were used to avoid cell settlement because H. pluvialis cells gets bigger when they enter carotenogenesis. Before testing ammonia gas, pH and different nitrogen sources were tested because ammonia/ammonium form depends highly on pHs to compare growth performances between nitrogen sources. After confirming a good growth

viii performance at pH 4, pH 5, and ammonium, ammonia gas was fed directly to the reactor at pH 4 and 5. Further efforts to increase astaxanthin content were made by feeding ammonia gas with different schedule or adding extra chemicals. Among all trials, only C/N ratio 1 was effective.

The third work was about treating phosphorus in wastewater (synthetic BBM medium, wastewater streams A, B, and C) using Revolving Algal Biofilm (RAB) systems. The two keys of microalgal phosphorus removal are efficient cultivation system and efficient operation design.

As to prove the efficiency of RAB cultivation system, cell growth and liquid nutrient removal performances in different growth media were compared with other cultivation systems. As to design the optimal operation of the systems, phosphorus uptake kinetics and mass balance were demonstrated by analyzing bound-P, acid-soluble P (ASP), and acid-insoluble P (AISP). In continuous culture, it was shown that harvest frequency and wastewater replacement rate can be optimized according to the cellular activities.

CHAPTER 1. GENERAL INTRODUCTION

Microalgae has gained its huge attention in recent decades due to its fast growth, low cultivation cost, capability to assimilate wastes, and unique cell composition. In an era of climate change and welfare, employing microalgae is appealing because it is a natural way to remedy environmental problems. However, the concept of utilizing microalgae to convert environmental pollutants into valuable products should be verified by actual demonstration. In this dissertation, microalgae were grown with air-born and water-born pollutants for environmental stewardship and the produced biomass was evaluated for further utilization.

In Chapter 2, principles of chemistry and biology of microalgae relates to the following chapters were covered. In Chapter 3, ammonia gas and carbon dioxide, main wastes from concentrated animal feeding operations, were fed to microalgae. The gas removal efficiency and cell growth performance were evaluated and the produced algae biomass was surveyed for its potential use for animal feed. In Chapter 4, Haematococcus pluvialis , known as productive astaxanthin producers, were grown with ammonia gas. The growth condition and astaxanthin induction methods were tested for cell density and astaxanthin content. In Chapter 5, microalgae was cultivated in Revolving Algal Biofilm (RAB) systems to treat phosphorus in wastewater with batch and continuous mode. The phosphorus uptake mechanism and phosphorus mass balance were analyzed to understand cellular metabolic activities and phosphorus fate in the system. In Chapter 6, general conclusion was reached and future work was suggested.

CHAPTER 2. LITERATURE REVIEW

1. Microalgae

1.1. Introduction of microalgae

Microalgae are highly diverse group of photoautotrophic microorganisms. The diversity of microalgae lies on cell size, cellular structure, chemical compositions, morphology, pigments for photosynthesis, and ecology and habitats (Barsanti and Gualtieri, 2014). Microalgae can be prokaryotic and eukaryotic. The prokaryotic cyanobacteria are often included in microalgae divisions (so called ‘blue-green algae’) as they can do photosynthesis and the origin of microalgal chloroplast is believed to be a symbiotic cyanobacterium (Kaplan and Reinhold,

1999). Except cyanobacteria, majority of microalgae species are eukaryotic. Microalgae and plants are similar in that they both do photosynthesis and use similar mechanical and chemical defense strategies against predators and parasites (Spiteller, 2008). But they are obviously different in morphology, reproduction system, and life cycle (Barsanti and Gualtieri, 2014).

Microalgal morphology can be unicellular, colonial, or filamentous and can be motile or nonmotile depending on their taxonomy or growth stage (Stevenson, 1996). Microalgae are found in almost everywhere from water to air and play an important role in ecosystem. They serve as the primary energy source, generate oxygen via photosynthesis, modulate chemicals by converting inorganic nutrients to organic forms, and stabilize sediments by overgrowing on it

(Stevenson, 1996; Barsanti and Gualtieri, 2014).

Microalgae reproduce either asexually such as cell division and fragmentation in optimal conditions; or sexually such as fusion of gametes and formation of zygospores in stressful conditions (Agrawal, 2012). Asexual reproduction is a fast and efficient way of increasing cell

3 number but restricts genetic diversity while sexual reproduction is less efficient but allows genotype variability (Barsanti and Gualtieri, 2014). The microalgal life cycle (growth, reproduction, spore germination, and re-growth) is usually affected by various environmental factors such as nutrients, light, temperature, pH, medium alkalinity, and physical shock (Agrawal,

2012).

1.2. Application of microalgae

The microalgal biomass market produces about 5,000 ton (dry matter) per year and generates a turnover of U.S. $1.25×10 9 per year (Pulz and Gross, 2004). The microalgae biomass can be used for various purposes, such as ethanol (Hirayama et al., 1998), hydrogen (Ueno et al.,

1998), bio-oil (Miao and Wu, 2004), biogas (Mussgnug et al., 2010), biodiesel (Chisti, 2007), human food, animal feed, fertilizer, pharmaceuticals, nutraceuticals, and cosmetics (Oswald,

2003).

The microalgae biomass industry can be more promising by culturing microalgae with various wastes (greenhouse gases, heavy metals, wastewater, etc.) as a nutrient source.

Depending on the growth condition, microalgae alter their essential biomass composition

(protein, lipid and carbohydrate) as an acclimation mechanism. Some microalgae species can also produce unique “niche” products such as carotenoids, phycocyanin, phenolics, sulpholipids, and sulphated polysaccharides (Baky and El-Baroty, 2013). Therefore, optimizing growth condition and selecting microalgae species that can utilize wastes and produce valuable byproduct are the key streams of current microalgae cultivation research.

There are several advantages of utilizing microalgae biomass. First, microalgae obtain energy via photosynthesis and do not require organic substrates to grow (Goldman et al., 1972).

Second, microalgae can grow in non-arable land and some microalgae can grow in saline water and therefore, do not compete with plants for water and land. Third, microalgae have short doubling time and high biomass productivity. Although these traits make microalgal cultivation an appealing process, the cost for current algal production, particularly at large scale, is still not economic due to various reasons.

2. Effect of CO 2 and nitrogen (ammonia) to microalgae cultivation

2.1. Role of CO 2 in microalgae cultivation

CO 2 is the carbon source in the microalgal photosynthesis process. The simple from of photosynthesis reaction can be expressed as:

6 H 2O + 6 CO 2 → C 6H12 O6 + 6 O 2 (1)

2.1.1. Chemistry of CO 2 in water

- 2- In algal culture medium, CO 2 exists as CO 2-HCO 3 -CO 3 equilibrium depending on the

1 pH and alkalinity of the water, which is the acid neutralizing capacity at the CO 2 end point

(Morel and Hering, 1993). The total carbonate in liquid medium can be expressed as

(parentheses indicate moles/liter):

1 CO 2 end point: when no strong acid or base exist in the solution at the end point of the titration (Morel and Hering, 1993)

(2) ∗ = + + (3) ∗ = + . . ≫ Ionization of water:

(4) ⇆ + ; ℎ = = 10 25℃

The equilibrium of CO 2 dissolution in water can be expressed as (Goldman et al., 1972):

(5) ⇆ . (6) . 10 . = , = 25℃ , = 10 Hydration of CO 2can be expressed as (Goldman et al., 1972):

(7) . + ⇆ ; First ionization of carbonic acid ( ) (Goldman et al., 1972): ∗

(8) ∗ ⇆ + (9) = ∗ = 4.45 × 10 25℃ Second ionization of bicarbonic acid ( ) (Goldman et al., 1972):

(10) ⇆ +

(11) = = 4.6 × 10 25℃ We can know the relative proportions of all carbon species as a function of pH from the total alkalinity of water (in equivalent liter -1):

(12) = + 2 + − Then each carbon species can be expressed as following:

(13)

∗ − + = 2 1 + (14) − + = 2 1 + (15)

− + = 2 1 +

Based on the above various equilibrium, the relative proportions of all carbon species as a function of pH can be expressed as follows (Figure 1).

Figure 1. pH effect on inorganic carbon species distribution in water (adapted from Goldman et al., 1972)

2.1.2. CO 2 utilization by microalgae

Microalgae play an important role in recycling carbon into potentially valuable biomass through photosynthesis. They can fix several folds more CO 2 per unit area than trees or crops

(Salih, 2011). There are four main CO 2 sources that microalgae can obtain from: (1) diffusion from atmosphere; (2) respiration of heterotrophs; (3) anaerobic fermentation; and (4) bicarbonate alkalinity (Goldman et al., 1972).

Three key questions regarding microalgal CO 2 utilization need to be addressed, i.e., (1) which CO 2 species do microalgae most prefer? (2) How microalgae acclimatize to different CO 2 levels? (3) What CO 2 level can microalgae endure or is the optimal for growth?

(1) Which CO 2 species do microalgae utilize most?

- It has been assumed that either one of the CT, , or HCO 3 can be utilized ∗ HCO (Goldman et al., 1972; Goldman, 1973; Shapiro, 1973). With advancement of biochemical tools,

- it became evident that different algae species has different substrate specificity but HCO 3 is the substrate that is readily utilized for inorganic carbon fixation (Kaplan and Reinhold, 1999). The inorganic carbon uptake by microalgae is mediated by a membrane transport mechanisms: CO 2

- is passively diffused across the cytoplasmic membrane and then is converted into HCO 3 , while

- HCO 3 is actively transported through plasma membrane and chloroplast envelope through carrier proteins (Kaplan and Reinhold, 1999). Interchange between carbon species happens in periplasmic space and is catalyzed by carbonic anhydrase (CA) (Kaplan and Reinhold, 1999).

(2) How microalgae acclimatize to different CO 2 levels?

Microalgae adapt to a wide range of CO 2 concentrations by changing their cellular responses and compositions. The most well-documented acclimation response is CO 2 concentrating mechanisms (CCMs). As the most rate limiting reaction in photosynthetic CO 2 fixation process occurs under the reactions of Rubisco, an enzyme that catalyzes the entry of CO 2 into Calvin cycle, (Badger et al., 1998), microalgae have developed inducible CCMs to ensure that Rubisco, is supplied with enough CO 2 substrate (Kaplan and Reinhold, 1999). CCMs include membrane carbon transport system, specialized localization for both internal and external

CA, and localization of Rubisco within the chloroplast where CO 2 can be elevated (Badger et al.,

1998). In high CO 2 concentrations cells repress the membrane transport system while in low CO 2 concentrations the cells induce the transport system (Matsuda and Colman, 1995).

In addition to CCMs, CO 2 acclimation responses include pH homeostasis and cellular composition change. These capabilities are crucial for algal cells to tolerate to extreme CO 2 levels (>20%) (Solovchenko and Khozin-Goldberg, 2013). The intra- and extracellular pH homeostasis is important to maintain enzyme activities. At high CO 2 level, microalgae regulate their intracellular pH by actively pumping H + from cytoplasm to vacuoles and their extracellular pH by uptaking nitrate in the media to excrete alkaline ammonia (Solovchenko and Khozin-

Goldberg, 2013). The change of CO 2 level also changes lipid and starch content and fatty acid composition (Tsuzuki et al., 1990). The alternation of lipid metabolism provides buffer for stress conditions and the fatty acid composition change enables membrane rearrangement to adapt to the environmental conditions. (Solovchenko and Khozin-Goldberg, 2013).

(3) What CO 2 level can microalgae endure or is the optimal level for growth?

This question has been studied for a long time by many researchers (Goldman et al., 1972) and is now a popular topic under the context of CO 2 sequestration. Researches have shifted from optimizing CO 2 level in earlier days to exploring microalgae species or growth conditions that can mitigate a high level of CO 2 (summarized in Salih, 2011; Solovchenko and Khozin-Goldberg,

2013). In general, the optimal CO 2 level for algal growth depends on the specific algae species and other growth factors. For example, the denser algal culture the more CO 2 the algae need, at the same time, light penetration becomes a severe problem. The general conclusion is that higher

CO 2 level increases photosynthesis activities but when CO 2 exceeds a certain concentration, cell growth becomes inhibitory. The inhibition caused by high level of CO 2 is due to acidification of stromal compartment of the chloroplast and cytoplasm and consequent inactivation of key enzymes of Calvin cycle (Kraus and Weis, 1991).

2.2. Ammonia as a nitrogen source

Microalgae uptake nitrogen mainly in the form of inorganic nitrogen, such as ammonium or nitrate. Ammonium is the reduced nitrogen form and needs less energy to be assimilated by the cells (Fernández et al., 2009). While nitrate assimilation involves two steps (nitrate to nitrite in cytosol, nitrite to ammonium in chloroplast), ammonium is directly incorporated into carbon skeletons in chloroplast by glutamine synthetase/glutamate synthase pathway (Fernández and

Galván, 2007).

2.2.1. Ammonia dissolution in water

Unlike CO 2, ammonia is much more soluble in water (Morel and Hering, 1993). The acid-base reactions of ammonia in water can be summarized as follows.

(16) ⇄ (17)

= = 57 .5 25℃ (18) ⇄ + , (19) . = = 10 25℃

The logC versus pH diagram can be drawn from the above equations (Figure 2).

Figure 2. pH effect on inorganic ammonium species distribution in water

2.2.2. Toxicity of ammonia to microalgae

For higher plants, most species grow well with nitrate but develop severe toxicity symptoms with ammonium as the sole nitrogen source (Britto et al., 2001). The response of microalgae to ammonia, however, is species specific (Kratz and Myers, 1955; Thacker and Syrett,

1972). For example, in the presence of ammonium in the culture Chlamydomonas reinhardii , the genes involved in nitrate/nitrite assimilation will be repressed and the cells can use ammonium as the nitrogen sources but do not accumulate nitrogen to inhibitory level within intracellular compartments (Fernández et al., 2009). For many other species, however, ammonia is inferior to nitrate as a nitrogen source due to its toxicity, which is caused by elevated intracellular pH

(Przytocka-Jusiak, 1976; Matusiak et al., 1976; Azov and Goldman, 1982; Tam and Wong,

1996). The toxicity of ammonia depends on media pH and temperature as these factor influence the balance between unionized ammonia (toxic to algae) and ionized ammonium (less toxic)

(Azov and Goldman, 1982; Källqvist and Svenson, 2003).

3. Astaxanthin production from microalgae

3.1. Chemistry of astaxanthin: Origin and effectiveness

3.1.1. Astaxanthin chemistry

Astaxanthin ((3S-3’S)-dihydroxy-β,β-carotene-4,4’-dione) is an orange-red and lipid- soluble pigment and is a member of keto-carotenoids (Figure 3). Carotenoids is a group of more than 700 natural pigments, with the colors derived from the long chain of conjugated double bonds (polyene chain) in the middle of the molecules. Based on the existence of the oxygen in its molecules, carotenoids can be classified into two sub-groups: xanthophylls which contain oxygen; and carotenes which are purely hydrocarbons. Since astaxanthin contains ketone group

(C=O) and hydroxyl group (OH-), it belongs to a group of xanthophyll. Astaxanthin is not synthesized by most animals de novo due to lack of enzymes required to convert carotenoids into astaxanthin, but is present in crustacean (i.e., shrimp, crab) and marine animals (i.e. salmon, trout, and ornamental fish) through digesting microorganisms that can synthesize astaxanthin (Lorenz and Cysewski, 2000).

Figure 3. Astaxanthin structure (Lorenze and Cysewski, 2000)

3.1.2. Effectiveness of astaxanthin

Astaxanthin has important applications in the nutraceutical, cosmetic, food and feed, and pharmaceutical industries. It can be used as antioxidant, colorant and nutrient, and clinical reagents. .

3.1.2.1 High antioxidant activity

Astaxanthin has been recognized as an effective antioxidant and its antioxidant properties surpass those of b -carotene, zeaxanthin, canthaxanthin, vitamin C, vitamin E, or a -tocopherol

(Guerin et al., 2003). Astaxanthin places its orientation perpendicular to the plane of membrane lipid bilayer, due to its hydrophilic ends (ketone groups (C=O) and hydroxyl groups (OH-)) and hydrophobic center (the polyene chain), which make the membranes rigid and limit the permeation lipid oxidation promoters such as Fe (II) ions and H 2O2 (Barros et al., 2011).

Astaxanthin is able to protect cells against oxidative stress by maintaining mitochondrial redox state (Wolf et al., 2010) and scavenging free radicals and destroying peroxide chain reactions

(Lorenz and Cysewski, 2000).

3.1.2.2 Colorant and nutrient

Astaxanthin has been used in aquaculture as a pigmentation reagent due to consumer appeal. Astaxanthin is widely preferred over other carotenoids for example, b- carotene and canthaxanthin, due to its higher colorant effect and higher muscle deposition efficiency due to a better bio-absorption (Supamattaya et al., 2005).

More importantly, it also serves as nutritional component and improves growth and reproduction (Higuera-Ciapara et al., 2006). For example, research has also shown that astaxanthin supplement increases fishery reproduction, such as rate of fertilization and egg survival (Ahmadi et al., 2006) and survival rate by enhancing resistance to stresses (Chien and

Shiau, 2005).

3.1.2.3 Clinical effect

The clinical effects of astaxanthin are related to its high antioxidant activity. Due to its amphiphilicity, astaxanthin can readily cross the blood-retinal-brain barrier and protect the retina against photo-oxidation and prevent the loss of photoreceptor cells (Guerin et al., 2003). The clinical effects of astaxanthin include skin protection from UV light, improvement of age-related macular degeneration, protection against chemically induced cancers, increased high-density lipoproteins (HDL) and prevention of cardiovascular diseases, reduction of coronary heart disease, prevention of exercise induced muscle damage, and enhanced immune system (Lorenze and Cysewski, 2000; Higuera-Ciapara et al., 2006; Djordjevic et al., 2012).

3.1.3. Astaxanthin-producing microorganisms

A limited number of microorganisms, such as algae, bacteria, and fungi are capable of synthesizing astaxanthin. The astaxanthin-producing microorganisms are summarized in Table 1.

Among these microorganisms, H. pluvialis possesses the highest astaxanthin productivity and widely utilized as a commercial producer.

Table 1. Examples of astaxanthin occurrence in microorganisms

Content (%, Microorganism Examples Reference w/w dry wt.)

Haematococcus pluvialis 4 Jaime et al., 2010

Sivathanu and Palaniswamy, Chlorococcum humicola 3.7 2012

Neochloris wimmeri 0.6 Orosa et al., 2001 Green algae Nannochloropsis gaditana <0.3 Lubián et al., 2000

Chlorella vulgaris 0.2 Gouveia et al., 1996

Spirulina 0.02 Mansoreh et al., 2010

Chlorella zofingiensis <0.01 Ip and Chen, 2005

Agrobacterium auratim 0.01 Yokoyama and Miki, 1995

Bacteria Mycobacterium lacticola, n/a Neils and de Leenheer, 1991 Brevibacterium 103,

Xanthophyllomyces 0.47 Rodríguez-Sáiz et al., 2010 dendrorhous Fungi Candida utilis 0.04 Miura et al., 1998

Peniophora sp. n/a Neils and de Leenheer, 1991

3.2. Biosynthesis of astaxanthin

3.2.1. Life cycle of H. pluvialis

The life cycle of H. pluvialis can be divided into four different stages: vegetative cell growth (green flagellate); encystment (brown palmelloid); maturation (red cyst cells); and germination (cyst to vegetative cells). The life cycle of H. pluvialis is shown in Figure 4.The ellipsoidal flagellate contains high levels of chlorophyll and protein with low levels of carotenoids. Under stress conditions, the cells become spherical immotile cyst cells with rigid cell walls, increased lipid accumulation, enhanced carotenoid biosynthesis, and degraded chlorophyll and protein contents. The cysts are two to three times larger than the flagellates. The cysts are in resting stage and show extremely low activity in photosynthesis and oxygen uptake due to the low metabolism. The stiff cell wall provides several benefits to survive in stressful environment: hampers digestion by predators; conveys resistance to high light and UV irradiation; and restricts mechanical breakage (Aflalo et al., 2007).

When cyst cells are transferred to a fresh medium, they germinate again with increased chlorophyll and protein syntheses and carotenoid degradation. The astaxanthin in cells can provide resistance against stresses (Kobayashi et al., 1991; Kobayashi et al., 1992; Klochkova et al., 2013; Hagen et al., 1993).

Figure 4. Life cycle of Haematococcus pluvialis (concepts and images adapted from Kobayashi et al., 1997a; Klochkova et al., 2013).

3.2.2. Biological mechanism of astaxanthin synthesis

Astaxanthin is built from eight C 5 isoprenoid primers, yielding in sequence of C 10 , C 15 , and C 20 compounds. Dimerization of C 20 produces C 40 , phytoene, the first C 40 carotenoid. From phytoene, several carotenoids are produced by enzymatic polymerization. The astaxanthin production scheme is depicted in Figure 5.

The produced carotenoids along the pathway are essential in the reactions of photosystems with these functions: i) light-harvesting processes during photosynthesis in the antenna complexes of the chloroplast, ii) protecting cells from free radical oxidation stress by strong light exposure (Miki, 1991), and iii) protecting protein complexes against singlet O 2

(Zhang et al., 1999).

Figure 5. Biosynthetic pathway of astaxanthin (adapted from Barros et al., 2011; Cardozo et al., 2007).

3.2.3. Reasons for astaxanthin accumulation

It is known that reactive oxygen species (ROSs) mediate the astaxanthin synthesis under stressful conditions, where there is an excess of reducing power for photosynthesis. The stress

19 conditions employed to induce astaxanthin in literature include high irradiance, nutrient deficiency (high C/N ratio), high salinity, high temperature (30-33°C), and aging (Kakizono et al., 1992; Sarada et al., 2002).

However, the role of astaxanthin biosynthesis has been explained in different ways in the literature. First, Yong and Lee (1991) proposed a role of physical screening photoprotection function. The authors observed that astaxanthin disperses towards the periphery of H. lacustris cells under light induction and moves back towards the center of the cell when the light was discontinued. The amount of astaxanthin was not different before and after the discontinued illumination and it was only the relocation of pigment.

Second, Hagen et al. (1993) proposed a role of physicochemical barrier for photo-damage protection. By testing flagellates with different amounts of secondary carotenoids (including astaxanthin), the authors found a higher resistance of red flagellates to photo-oxidative stress

(Hagen et al., 1993). Therefore, they hypothesized that the ketocarotenoids, including astaxanthin, might act as a physicochemical barrier for genomes from free radical-mediated damage by accumulated in lipid vacuoles around the nucleus (Hagen et al., 1993). Wang et al.

(2003) proved this hypothesis in molecular level. When H. pluvialis was under a high irradiance stress, the D1 protein of PSII decreased significantly while the photosynthesis rate was still high.

When the astaxanthin was formed and deposited under continuous light exposure, the D1 protein did not decrease anymore thus the cells enabled to maintain PSII function and structural integrity.

Therefore, it was concluded that astaxanthin acts as a physicochemical barrier to protect the replicating DNA from ROSs photo-oxidation (Wang et al., 2003).

Third, Kobayashi et al. (1997b) proposed a role as an antioxidant against ROSs. By

•- adding superoxide anion radicals (O 2 ) to mature (astaxanthin-rich) and immature cysts of H. pluvialis , Kobayashi et al. (1997b) found a higher antioxidant activity in mature cells. When astaxanthin was extracted from the mature cells, no antioxidant activity was found. The results suggested that astaxanthin functions as an antioxidant against excessive oxidative stress

(Kobayashi et al., 1997b). The defense mechanism against oxidative stress by astaxanthin biosynthesis was also proved at genetic level by Li et al. (2008) using real-time reverse transcription PCR. In their study, the astaxanthin biosynthesis consumed as much as 9.94% of reactive oxygen species (ROS), and it was done through two ways: “(1) extensive oxygen dependent processes, and (2) conversion of molecular oxygen into water using electrons derived from carotenogenic desaturation steps to plastid terminal oxidase via the photosynthetic plastoquinone pool” (Li et al., 2008).

3.3. Production of astaxanthin

3.3.1. H. pluvialis cultivation processes

In open pond culture, H. pluvialis is easily contaminated by many other undesirable algal species because of its relative slow growth. Therefore, H. pluvialis culture prefers closed systems such as tubular photobioreactor, which increases the cultivation cost at large scale (Dufossé et al.,

2005).

Another challenge in cultivation of H. pluvialis is the long life cycle of this species. Since

H. pluvialis starts to accumulate astaxanthin by encystment under stress conditions, there are two different optimum conditions for growth and astaxanthin induction (“two-stage”); therefore a

21 long cultivation time is required for astaxanthin production with high cultivation cost (Harker et al., 1996; Choi et al., 2011).

3.3.2. Autotrophic vs. heterotrophic vs. mixotrophic production

Microalgae cultivation method can be classified into three major types depending on the energy and carbon sources: autotrophic, heterotrophic, and mixotrophic. In autotrophic culture, light is used as an energy source and CO 2 as a carbon source. Either open ponds or photobioreactor systems can be used for algal culture. In these systems, light penetration, CO 2 concentration, and control of contamination are the key factors of successful cell cultivation

(Chen, 1996).

In heterotrophic culture, organic carbon substances are used as sole energy and carbon sources. For this reason, cell growth does not require light source and contamination can be easily controlled by employing enclosed vessel and sterilized medium. The higher cost of the medium compared to autotrophic culture can be balanced by adopting carbon source from downstream processing. However, few microalgae species can grow heterotrophically and the solubility of organic carbon source for optimal growth should be carefully maintained (Chen,

1996).

In mixotrophic culture, microalgae cells are exposed to both light and organic carbon source and therefore, it holds advantages and drawbacks from both sides.

The majority of H. pluvialis cultivation studies use autotrophic system. Those studies usually focus on growth factors (medium composition, reactor configuration, initial biomass

22 density, etc.) or astaxanthin induction methods (different stress conditions). Research also have been made for heterotrophic or mixotrophic cultivation of Haematococcus sp. (Kobayashi et al.,

1991; Kakizono et al., 1992; Kobayashi et al., 1992; Chen et al., 1997; Hata et al., 2001; Kang et al., 2005). Several studies showed that autotrophic culture has higher astaxanthin productivity than heterotrophic cultivation but it requires excessively longer periods of time for carotenogenesis (within several weeks) compared to heterotrophic cultivation (within several days) (Kobayashi et al., 1991; Kang et al., 2005). Therefore, there were efforts to combine heterotrophic and autotrophic systems, such as growing vegetative cells heterotrophically to obtain high cell density and then inducing astaxanthin autotrophically (Hata et al., 2001). There were also attempts to apply mixotrophic cultivation (Chen et al., 1997) based on the findings of

Kobayashi et al. (1992) that H. pluvialis can do photosynthesis and organic carbon oxidative assimilation simultaneously in mixotrophic culture and the specific growth rate in mixotrophic culture is the sum of those in heterotrophic and autotrophic conditions. Ip and Chen (2005) stimulated astaxanthin accumulation under heterotrophic condition by adding reactive oxygen species.

3.3.3. One-stage production vs. two-stage production

Most astaxanthin production is carried out in a two-step process, based on the fact that the condition for green flagellate growth is different from that for the astaxanthin accumulation by red cysts. In two-stage cultivation process, the green vegetative cells are grown under optimal growth condition to achieve the maximum biomass, and then adverse environmental condition is exposed to induce astaxanthin.

One group (Del Río et al., 2005; Del Rio et al., 2007; García-Malea et al., 2009) also examined astaxanthin production in one-step (continuous) process in both lab scale and outdoor scale and proved the possibility under moderate nitrogen limitation and irradiance. The ground reason for these studies was that actively growing cells also have ability to accumulate astaxanthin so significant astaxanthin productivity can be achieved by optimizing cell productivity and astaxanthin content. However, this one stage approach was challenged by

Aflalo et al. (2007) for several reasons: the one-step process gives lower astaxanthin productivity; red cells from one-step process are enriched with other carotenoids and require separation step to purify astaxanthin from those carotenoids; most cells in one-step culture are either in flagellated or palemlloid forms with soft cell wall, which are less protective to predators and mechanical breakage in upscale; and therefore, two-step system is more suitable for commercial large scale implementation.

Nonetheless, comparison of the one-step versus two-step H. pluvialis culture for astaxanthin production is worth to further study particularly optimizing various cultivation parameters for the one-step system. As noted in Choi et al. (2011), the two-step system has intrinsic problem of determining the right timing to transfer to the second stage. Additionally, whether or not the two-stage system gives higher astaxanthin productivity for a short amount of time is inconclusive in current knowledge. Moreover, one-step system can eliminate transferring effort to another reactor in the middle of cultivation.

4. Wastewater treatment

4.1. Overall wastewater treatment procedure

The wastewater treatment system generally can be summarized as five steps: preliminary treatment, primary treatment, secondary treatment, disinfection, and sludge treatment.

4.1.1. Preliminary treatment

The purpose of this step is to remove large inorganic materials to protect the equipment, such as pumps and pipes (NYC, 2015). The mechanism behind this treatment is physical separation by size. For this reason, mechanically cleaned screens consisting of upright bars spaced one to three inches apart are used (NYC, 2015). The separated materials are transported to landfills. After the preliminary treatment, the wastewater is pumped to the sedimentation tank where the primary treatment occurs.

4.1.2. Primary treatment

The purpose of this step is to remove organic and inorganic materials by sedimentation and flotation (Sonune and Ghate, 2004). When the flow of water is slowed down, heavier solids settle down to the bottom of the tank and lighter materials (grease and small plastic material) float to the surface of water so that damage to pumps and clogging of pipes can be prevented

(NYC, 2015). The floatable trashes are screened and the settled solids, called primary sludge, are centrifuged and pumped through cyclone degritters. The grits are sent to landfills and the degritted primary sludge is pumped to the anaerobic digesters (NYC, 2015). Primary treatment takes for one to two hours and can reduce BOD by 25-50 % and total suspended solids by 50-70 %

(Sonune and Ghate, 2004). After primary treatment, the primary effluent moves to the secondary treatment system.

4.1.3. Secondary treatment

The purpose of secondary treatment is to oxidize dissolved organic matter and remove fine suspended using microbes that are remaining after primary treatment (Sonune and Ghate,

2004). There are several variations of secondary treatment, such as activated sludge, trickling filtration, rotating biological contactors (RBC), and lagoons and ponds (World bank, 2015). The activated sludge, lagoons and ponds use suspended microorganisms while the trickling filtration and RBC use attached microbes. Aerobic bacteria degrade most of the fine suspended and soluble organic matters and produce heavier sludges that will settle later in the secondary clarifier process (NYC, 2015). The secondary treatment takes about three to six hours and can reduce BOD and suspended solids by 85-95% (World bank, 2015). The sludges are pumped to anaerobic digesters or re-circulated back to the aeration tank to combine with the activated sludge.

4.1.4. Tertiary treatment (advanced wastewater treatment)

Tertiary treatment is an additional and optional treatment that is needed to remove suspended and dissolved substances remaining after secondary treatment. Although primary and secondary treatment removes the majority of BOD and suspended solids, the treated water is still not sufficient to protect the receiving waters (Sonune and Ghate, 2004). Examples of tertiary treatment includes, ion exchange, coagulations, membrane processes, filtration, activated carbon adsorption, and electrodialysis depending on the targeted pollutant (Sonune and Ghate, 2004;

Mahmoud and Hoadley, 2012; Verlicchi et al., 2012). Tertiary treatment is costly as compared to primary and secondary treatment methods but can remove more than 99 percent of all the impurities from wastewater (World bank, 2015). The pollutants can be organic chemicals,

26 suspended solids, color, nutrients (P and N), heavy metals, and pharmaceutical products (Sonune and Ghate, 2004; Mahmoud and Hoadley, 2012; Verlicchi et al., 2012).

4.1.5. Disinfection

Before the final effluent is released into the receiving waters, it is disinfected for 15-20 minutes to reduce the remaining pathogenic microorganisms (NYC, 2015). The most common reagent used for this purpose is chlorine gas or a chlorine-based disinfectant such as sodium hypochlorite (NYC, 2015). Other disinfection options include UV light, ozone, and peracetic acid (Kitis, 2004).

4.1.6. Sludge treatment

Sludge is generated from all the treatment steps and can be managed in four steps: thickening, dewatering, stabilization, and main treatment (Suh and Rousseaux, 2002). The thickening and dewatering are the steps that reduce the liquid volume up to 90% of the incoming sludge using gravity, pressurization, or spinning (Suh and Rousseaux, 2002; NYC, 2015). The separated water is sent back to the aeration tank for additional treatment (NYC, 2015).

The sludge is then stabilized to minimize the mobility of pollutants by any means or combinations of lime-stabilization, composting, anaerobic digestion, and incineration and melting (Suh and Rousseaux, 2002; Hong et al., 2009). Lime-stabilization is simply mixing limes to the sludge so that the high pH values destruct microbial communities and reduce heavy metal bioavailability (Samaras et al., 2008). In anaerobic digester, the anaerobic bacteria decompose the organic material at least 35 °C and stabilize the thickened sludge by converting into water and biogas, such as CO 2 and methane gas. The produced methane gas can be used as an energy source for the local area (NYC, 2015). The treated sludge, known as biosolids, is then applied to

27 agricultural field to increase soil nutrients (Akrivos et al., 2000). Although its simplicity, composting has limitation in use due to the pathogen and heavy metal issues (Hong et al., 2009).

Incineration and melting methods burn the sludge at high temperature (800-900 °C and

1300-1800 °C, respectively). However, those processes generate significant amount of air-borne pollutants and the high temperature increases the implementation cost (Hong et al., 2009).

4.2. Current phosphorus removal methods

4.2.1. Physical treatment

Particulate phosphorus can be removed by using screen, filter press, or centrifuge.

However, filtration alone removes phosphorus very little (5-30 %) without adding chemicals

(Bowers, 2011). The most advanced physical phosphorus treatment method is membrane process.

Depending upon the material used for membrane and separation mechanism, the membrane process can be classified into microfiltration (MF), ultrafiltration (UF), electrodialysis

(ED), nanofiltration (NF), and reverse osmosis (RO) (Li et al., 2007). MF membranes have the largest pore size and separate large particles and microorganisms. UF membranes have smaller pore size than MF and can exclude bacteria and soluble macromolecules like proteins (Schrotter and Bozkaya Schrotter, 2010). The mechanism of MF and UF is sieving. RO membranes are ‐ effectively non-porous and reject small particles and micromolecules such as salt ions and organics. NF membranes are sometimes called loose RO membranes. NF mechanism is sieving, solution, diffusion, and exclusion while RO mechanism is solution, diffusion and exclusion. MF,

UF, NF, and RO use pressure to transport water across the membrane. MF removes particulate matter while RO removes many solutes. The cost of membrane process increases as the size of

28 solute decreases. Membrane process can work at ambient temperature without phase change without product accumulation inside membrane (unlike ion exchange resins, which needs replacement) and there is no need of chemical additives for separation (Schrotter and Bozkaya ‐ Schrotter, 2010).

4.2.2. Chemical precipitation

The chemicals can be added before a primary clarifier or other solids separation device.

The removal efficiency depends on the chemical dose, pH, temperature, mixing intensity, and age of the precipitant (Neethling et al., 2008).

Conventionally, simple chemicals were added to react with the soluble phosphorus species and produce solid precipitants.

+3 -3 Al + PO 4 → AlPO 4 (20)

+3 -3 Fe + PO 4 → FePO 4 (21)

After the precipitants flocculate, remove the particulate phosphorus by adsorption or filtration.

However, the conventional method is expensive and causes an increase of sludge volume by up to 40 %.

Another chemical method to remove phosphorus is struvite (magnesium ammonium phosphate). By adding magnesium and ammonium, phosphate in wastewater forms a solid, called struvite, which can be easily transported.

+2 + -3 Mg + NH 4 + PO 4 → MgNH 4PO 4 (struvite) (22)

Increasing magnesium concentration and pH enhances struvite formation. The produced struvite can be used as a fertilizer (Barak and Stafford, 2006). Although struvite formation is relatively fast, separating struvite from a mixture of suspended solids is problematic because the specific gravity of struvite (1.6) is similar to other suspended organic matters (Barak and Stafford, 2006).

4.2.3. Biological treatment

Phosphorus is required for microbes for cell growth and reproduction and it is stored for subsequent use. Growth of microorganisms in wetland (Martin et al., 2013), waste stabilization ponds (Powell et al., 2009), lagoons and microalgae ponds are the examples of advanced wastewater treatment for phosphorus removal. Among the advanced biological treatment, the most widely used method is Enhanced Biological Phosphorus Removal (EBPR) (Mino et al.,

1998).

EBPR is a modification of the activated sludge process. It utilizes unique microbes called polyphosphate accumulating organisms (PAOs) that possess a capability to store large quantities of phosphate (luxury P uptake) and hydrolyze them under certain conditions. PAOs are normally present in all aerobic suspended growth cultures, but only display EBPR when they are cycled between aerobic or anoxic and anaerobic zones (Mino et al., 1998).

In anaerobic environment, the organic matter in wastewater undergoes fermentation and produce acetic acid and short-chain volatile fatty acids. The acetic acid is transported across the

PAOs’ cell membrane and polymerized to form a long chain poly-β-hydroxy butyrate (PHB). As the acetic acid metabolism requires ATP, polyphosphate stored in the cell is hydrolyzed to replenish the ATP and excess inorganic phosphorus is released from the cell into the water.

Therefore, PAOs store carbon sources from sludge in the form of PHB, degrade polyphosphate, and release orthophosphate (Mino et al., 1998).

In aerobic environment, PAOs grow up aerobically by taking up external carbon sources and inorganic phosphate, and store polyphosphate. When the external carbon source is depleted,

PAOs use its PHB reserves to continue the aerobic growth and keep taking up the orthophosphate and storing polyphosphate (Mino et al., 1998).

In anoxic environment, when no oxygen is present but nitrate is available, PAOs utilize nitrate as an electron acceptor and performs the same function as in the aerobic condition.

Although the cell yield and phosphorus uptake rates are lower than that of the aerobic condition, the phosphorus removal performance of anaerobic-anoxic is as good as the anaerobic-aerobic system (Mino et al., 1998).

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CHAPTER 3. USE OF MICROALGAE FOR MITIGATING AMMONIA AND CO 2

EMISSIONS FROM ANIMAL PRODUCTION OPERATIONS – EVALUATION OF GAS

REMOVAL EFFICIENCY AND ALGAL BIOMASS COMPOSITION

A manuscript accepted by Algal Research

Juhyon Kang and Zhiyou Wen

Abstract

The emissions of ammonia and CO 2 from animal housing operations are a major challenge for concentrated animal feeding operations (CAFOs). A microalgae culture process was developed as a bio-scrubber to remove these gases. The green algae Scenedesmus dimorphus was grown in a flat panel photobioreactor aerated with ammonia- and CO 2-laden air, with ammonia mass loading being fixed at 42.4 mg/L day and CO 2 mass loading rates ranging from

0.64 to 5.49 g/L day. Within the range of the CO 2 mass loading rate investigated, the ammonia gas removal rate was kept constant (41‒42 mg/L day) with a high removal efficiency (> 95%) from the inlet gas. The majority (80‒90%) of the ammonia was removed by algal cell assimilation, with a small portion (10‒12%) being removed via liquid dissolution. The CO 2 removal rate and removal efficiency, however, highly depended on the CO 2 mass loading rate used. The highest CO 2 removal rate (0.6 g/L day) was achieved at CO 2 mass loading rate of 1.18 g/L day, but the highest CO 2 removal efficiency (78%) was achieved at the 0.64 g/L day CO 2 mass loading rate. The cell biomass obtained contained 7‒8% lipid, 55‒60% protein, and 21‒28%

42 of carbohydrates. The amino acid profiles of the algal biomass are close to the ideal protein profiles for typical animal feeds, with a high ratio of essential amino acids. Collectively, the study shows that microalgae culture is an effective method for mitigation of ammonia and CO 2 emissions from CAFOs, with the potential for use as an animal feed additive.

Keywords: ammonia, CO 2, microalgae, nitrogen mass balance, biomass composition

1. Introduction

Modern livestock production occurs primarily in confined buildings to increase production efficiency and protect animals from the wild outside environment (EPA, 2012). A common feature of large-scale animal houses is accumulation of manure in a pit or storage area underneath the floor for an extended period (six months to a year) before land application. Under certain circumstances, such as prolonged contact of manure with bacterial enzymes, high manure moisture, adequate pH, and elevated temperature, the organic nitrogen contained in the manure is conducive to degradation into ammonia (Groot Koerkamp, 1994; EPA, 2012).

Ammonia volatilization is one of the main concerns in animal house operations because ammonia gas causes serious health problems for both workers and animals (Donham et al., 1989).

Releasing ammonia gas into the atmosphere also causes the eutrophication of surface water where phosphorus concentration is high, and deteriorates air quality by forming particulate matter (e.g., PM 2.5 ). That said, the US Environmental Protection Agency (EPA) is about to start enforcing the farm regulation of 100 lbs NH 3/day/facility (EPA, 2014). Developing a stable and sustainable ammonia removal system is therefore an urgent demand.

Mitigating ammonia gas through “end-of-pipe” methods such as an acid scrubber and a bacteria-based biofilter has been studied (Philippe et al., 2011). Those methods can have very high ammonia removal efficiencies (Choi et al., 2003); however, the high capital and maintenance costs of those systems hinder their implementations at commercial scale. Also, the acid scrubber system eventually causes acid stress in the environment (Ndegwa et al., 2008).

Recent research has demonstrated that microalgae cultivation can be a novel approach for mitigating ammonia gas emissions from animal production operations (Kang et al., 2014). In a laboratory flat panel photobioreactor setting, the microalga Scenedesmus dimorphus can assimilate up to 98.6% of ammonia from the inlet gas, with an ammonia removal rate 51 mg/L day (Kang et al., 2014).

In addition to ammonia, greenhouse gases such as CO2 also accumulate to a high level in large animal houses. For example, in a southeast US broiler operation, CO 2, methane, and nitrous oxide emissions can reach to 4.64 Mg, 3.41 kg, and 1.72 kg, respectively, for each 1,000 birds produced annually (Burns et al., 2008). In another study, Wang et al. (2010) reported CO 2 emissions in two laying henhouses (92,000 and 95,000 birds, respectively) were around 60‒100 g/bird/day. With US meat and egg consumption increasing in recent decades, and the trend of fewer, but larger size farm, ammonia and CO 2 emissions will become more a challenging and pressing issue facing the US livestock industry. Therefore, developing an environmentally sound approach for mitigation of those gas emissions is urgently needed.

In our previous study, microalgae-based ammonia gas mitigation was achieved by blending the ammonia gas with air. The CO 2 in the blend gas was at atmospheric level (~395 ppm) (NOAA, 2013). For typical poultry layer houses, however, the indoor CO 2 concentration can reach 600‒1,000 ppm in summer and 1,500‒3,000 ppm in winter (Hayes et al., 2013). On the

other hand, CO 2 plays an important role in algal cultivation by serving as a carbon substrate for microalgae photosynthesis. Therefore, it most likely creates a synergism for microalgal cells to assimilate these two gas species simultaneously. To the best of our knowledge, however, using microalgae for simultaneously removing ammonia gas and CO 2 and their interaction on algal culture have not been attempted. The aim of this work was to investigate the compound effect of ammonia gas and CO 2 when using microalgal cultivation as a bioscrubber for mitigation of these two gas species. The chemical composition of the algal biomass was also characterized to evaluate its potential use as animal feed, biofuel feedstock, or fertilizer.

2. Materials and Methods

2.1 Cell species and growth conditions

Scenedesmus dimorphus (UTEX 1237) was used as a continuation of our previous study

(Kang et al., 2014). The strain was chosen due to its strong ability to utilize ammonia and high tolerance to CO 2 (Hanagata et al., 1992). The strain was kept on agar slant at 4°C and transferred to 250-mL Erlenmeyer flasks containing 50 mL of modified Bold’s Basal Medium (Anderson,

2005). The medium was autoclaved at 121°C for 15 minutes. The seed flask cultures were incubated at 25°C in an orbital shaker (200 rpm) with continuous (24 hours/day) illumination at

110‒120 µmol s -1 m-2 using florescent light bulbs. The cultures were incubated for 5 days and then stepwise transferred to a bubble column reactor (300 mL working volume) and then to a flat panel photobioreactor (5-L working volume).

2.2 Experimental set up

A flat panel photobioreactor was used for the investigation of algae-based ammonia and

CO 2 mitigation. The detailed set up and operation of the reactor were described previously (Kang et al., 2014). The reactor was operated in continuous (semi-batch) mode by daily exchanging 10% of liquid volume which led to a 0.1 day -1 dilution rate. Nitrogen-free Bold’s Basal Medium was used for growing cells so the ammonia gas in the inlet served as the nitrogen source for the algal culture. The pH of the medium was maintained at 7.0 throughout the culture. The above operational conditions were selected at the levels that have been proved optimal in our previous report (Kang et al., 2014)

Compressed ammonia gas, CO 2, and air were mixed and bubbled to the reactor at 2.77

L/min (0.56 vvm). No mechanical agitation was used for the liquid mixing. The volumetric flow rates of these gas streams were each regulated by digital flow meter (Alicat Scientific, Tucson,

AZ). Our previous research has shown that ammonia gas at a loading rate of 42.4 mg/L day (60 · ppm in inlet gas) resulted in the best cell growth (Kang et al., 2014). As the 60 ppm ammonia gas concentration is a common concentration for the animal house air (10-80 ppm), the loading rate were thus a practical level achievable for animal house operation. The CO 2 gas loading rate was varied by adjusting the flow rate of the compressed CO 2 into the reactor. The ratio of the compressed CO 2 to the ammonia-laden air was in the range of 0‒3000 ppm. Considering air contains 395 ppm of CO 2 (NOAA, 2013), the absolute amount of CO 2 added to the reactor was in the range of 395 to 3395 ppm. This concentration range represents the typical CO 2 level in a henhouse operation (Hayes et al., 2013). Table 1 lists the CO 2 mass loading rates that correspond to each concentration of CO 2 in the inlet gas. The continuous algal culture was operated at a certain inlet CO 2 level until reaching equilibrium and then switched to another CO 2 level. Each

46 operational condition was considered to have reached steady state after at least five consecutive daily samples having less than 5% variation in cell density.

2.3 Evaluation of ammonia and CO 2 mitigation efficiencies by algal cultures

Ammonia gas mitigation was evaluated based on the volumetric ammonia removal rate

(R NH3 , mg NH 3/L·day) and the ammonia gas removal efficiency (E NH3 , %) as follows:

(1) × × =

× (2) = 100 where F is the volumetric flow rate of the aerated air (L/day), X NH3(in) and X NH3(out) are ammonia concentration (ppm) in the inlet and outlet gas, respectively; r NH3 is the ammonia gas density at the operating condition (0.696 g/L); and V is the reactor working volume (L).

CO 2 mitigation was evaluated by the CO 2 removal rate (R CO2 , g CO 2/L·day) and CO 2 removal efficiency (E CO2 , %). The CO 2 removal rate was approximated as the CO 2 fixation rate by the algal cells, assuming the dissolved CO 2 in the liquid was minimal compared to the CO 2 consumed by the cells (Chiu et al., 2008) and was calculated as follows (Ho et al., 2010):

(3) ∙ = / = . where P is the biomass productivity (g/L day), C cell is the biomass density (g/L), D is the dilution

-1 rate (day ), Y cell/CO2 is the biomass yield based on CO 2 utilization. Based on the literature, the

algal biomass can be approximately represented as CO 0.48 H1.83 N0.11 P0.01 (Chisti, 2007), and the value of Y cell/CO2 can then be derived as 0.532.

The CO 2 removal efficiency (E CO2 , %) was evaluated by as follows.

(4) = [ ] × 100 where [CO 2] is the CO 2 mass loading rate (g/L day).

2.4. Microalgae cell composition analysis

Fatty acid was analyzed based on the total fatty acid content. Fatty acid methyl esters

(FAME) were prepared from the algal biomass (Pyle et al., 2008) and quantified by GC (Liang et al., 2011). Total nitrogen content of the biomass was determined by a combustion method

(AOAC 990.03) using a Vario MAX Carbon Nitrogen analyzer (Elementar Analysensysteme

GmbH, Hanau, Germany). Protein content was estimated by multiplying the total nitrogen content by 5.95 (Lopez et al., 2010). Ash was determined by a combustion method, burning at

550˚C overnight. Carbohydrate was estimated by subtracting the protein and lipid contents from the ash free biomass. The amino acid profile was determined based on AOAC Official method

982.30 E (a, b, c) Ch. 45.3.05.

2.5. Statistical analysis

Differences in the data reported as significant were compared at the 95% confidence level by one-way analysis of variance (ANOVA) using PROC GLM and LSMEANS, with the statistical software package SAS (version 9.3 (32) SAS Institute, Inc., Cary, NC, USA).

3. Results and Discussion

3.1 Cell growth performance

Our previous research showed that when ammonia gas was blended with air (without additional CO 2), the cell density and productivity reached the maximum level at 42.4 mg/L day of ammonia day loading rate (60 ppm concentration in the inlet gas). The ammonia loading rate was fixed at this level throughout this study with CO 2 loading rate varying.

As shown in Figure 1, the CO 2 mass loading rate significantly affected S. dimorphus biomass yield and productivity. The highest biomass yield (3.18±0.051 g/L) and productivity

(0.318±0.005 g/L day) reached the highest levels at the 1.18 g/L day CO 2 mass loading rate and · then decreased as the CO 2 mass loading rate increased (p<0.05).

It is well known that CO 2 plays an important role in microalgal photosynthesis by adjusting the carboxylating and oxygenating activity of Rubisco, and facilitating photosynthetic electron transport between PSII and PSI, leading to an increased photon yield and energy for

CO 2 fixation (Gordillo et al., 1999; Dubinsky et al., 1986). However, a CO 2 level that is too high has adverse effects on microalgal photosynthesis due to lower affinity for external CO 2, lower intracellular carbonic anhydrase (CAint) activity, lower CO 2 concentrating capacity, and higher

O2 photosynthetic sensitivity (Berry et al., 1976; Badger et al.,1980; Moroney et al., 1985).

Various studies have been performed to evaluate the algal growth under different CO 2 levels. For example, de Morais and Costa (2007) used 0, 6, and 12% CO 2 to feed Scenedesmus obliquus culture and reached maximum biomass productivity and highest specific growth rate at

12% CO 2. Chiu et al. (2008) studied 2‒15% CO 2 to feed Chlorella sp. and obtained the highest

biomass yield at 2% CO 2. In another study with S. obliquus , Ho et al. (2010) tested 5‒70% CO 2 and achieved maximum biomass productivity and highest specific growth rate at 10% CO 2.

However, it should be noted that reporting CO 2 effects on algal growth based on CO 2 concentration in the inlet gas does not provide a holistic view of CO 2 supply scenarios as the flow rate of inlet gas also determines the CO 2 supply levels. Therefore, we used the CO 2 mass loading rate as a more precise baseline parameter to evaluate the CO 2 loading level to the algal culture system. We therefore further calculated the CO 2 mass flow rate in the above studies. The mass loading rates used by de Morais and Costa (2007) were in the range of 0‒91.8 g/L day, the · CO 2 mass loading rates used by Chiu et al. (2008) were in the range of 12.9‒96.9 g/L day. Those · values are much higher than the mass loading rate used in this study. However, the mass loading rate in the Ho et al. (2010) study was 0.385‒5.39 g/L day CO 2 loading rate, which is similar to · the range reported here.

In general, unlike nitrate or ammonium salts which serve as a cation source to capture excess CO 2 and result in higher CO 2 utilization, ammonia cannot serve as the cation source and thus, cannot capture excessive CO 2 (Tew et al., 1962). This may be the reason why the overall cell growth did not increase at the higher CO 2 levels. The interference of the physical dissolution of CO 2 and ammonia in a CO 2-NH 3-water system also affects the utilization of these two gas species. Both the experimental data and the thermodynamics model showed that the CO 2-holding efficiency of the ammonia-laden water system decreased compared to the ammonia-free system.

Similarly, ammonia partial pressure decreased as the CO 2 partial pressure increased. As a result, the availability of ammonia for the cells decreased under high CO 2 concentration and vice versa

(Darde et al., 2010). Therefore, the co-existence of ammonia and CO 2 in the inlet gas resulted in a negative effect for cell growth.

3.2 Ammonia gas removal by the algal culture system

Figure 2 shows the ammonia removal performance by algal culture under different CO 2 loading rates. Throughout the range of CO 2 loading rates investigated, the volumetric ammonia removal rate (R NH3 ) and the ammonia gas removal efficiency (E NH3 ) were maintained at a relatively constant level without significant difference ( p>0.05). Such high and consistent ammonia removal rates and efficiencies were probably due to the high adsorption of ammonia by algal cells and high solubility of ammonia in the liquid (pK a = 9.24).

A mass balance of nitrogen was performed to delineate the fate of the nitrogen in the inlet gas, liquid medium, algal biomass generated, and exhaust gas, as follows:

Ninlet = N cell + N liquid + N outlet + ε (5)

where N inlet , N cell , N liquid , and N outlet , respectively, represent the mass flow of nitrogen (g/day) contained in the inlet gas, assimilated by cells, dissolved in the liquid, and contained in the exhaust gas. The value of ε represents the experimental error in order to balance the mass. The calculations for each parameter in Equation (5) followed the description in our previous report

(Kang et al., 2014).

As shown in Table 2, the majority of nitrogen (ammonia) in the inlet gas was assimilated into algal cells. A portion of the ammonia was also trapped in the liquid due to its high solubility.

The nitrogen in the ventilation (exhaust) stream was minimal, indicating superior ammonia removal efficiency. The highest cellular nitrogen assimilation (93.1 ± 1.16%) appeared at the

1.18 g/L day CO 2 mass loading rate and then decreased as the mass loading rate further increased.

This trend agreed with the cellular activity and biomass productivity as reported in our previous

51 study (Kang et al., 2014). The amount of ammonia physically trapped in the liquid was relatively consistent among the CO 2 loading rate range tested because the ammonia gas feeding rate and medium pH were maintained constant.

Collectively, the above results show that high ammonia removal efficiency can be achieved with the algal culture system and the majority of the removed ammonia was due to the biological adsorption by the algal cells. It should be noted that although the physical adsorption of ammonia by the algal culture liquid was not affected by the CO 2 mass loading (Table 2), the higher CO 2 loading did help maintain an acidic pH level. As a matter of fact, the external base used to maintain the constant pH was reduced when CO 2 mass loading was higher. Therefore, although increasing the CO 2 loading rate slightly reduces cell growth performance as discussed in Section 3.1, the high CO 2 mass loading rate was still preferred from an ammonia removal point of view.

3.3 CO 2 gas removal by the algal culture system

CO 2 fixation performance was evaluated based on the CO2 removal rate (R CO2 ) and the removal efficiency (E CO2 ) as defined in Equations (3) and (4). As shown in Figure 3A, S. dimorphus had the highest R CO2 (0.60 g/L day) at the CO 2 mass loading rate of 1.18 g/L day. · · However, higher CO 2 removal efficiency (78.3%) was achieved at the CO 2 mass loading rate of

0.64 g/L day. The drop in CO 2 removal efficiency at the higher CO 2 mass loading load rate was · due to the increased CO 2 input and relatively constant CO 2 removal (Figure 3).

The values of R CO2 obtained in this study were higher than previous reports, in the range of 0.057‒0.55 g/L day (Ho et al., 2010; de Morais and Costa 2007; Basu et al., 2013). However,

CO 2 removal efficiency (E CO2 ) was in the range of 7.8‒78.0% (Figure 3B), while a range of

0.12‒58.00% of E CO2 was reported previously in the literature (Chiu et al., 2008; Cheng et al.,

2006; de Morais and Costa, 2007; Vidyashankar et al., 2013; Basu et al., 2013). The descending

ECO2 with increased CO 2 mass loading implies that S. dimorphus has a limit for CO 2 consumption and the CO 2 loading rate should be carefully monitored to effectively remove CO 2.

In order to increase E CO2 by algal cultures, several factors to consider are algal species

(Scenedesmus sp.: de Morais and Costa, 2007, Ho et al., 2010, Basu et al., 2013 and

Vidyashankar et al., 2013; Chlorella sp.: Cheng et al., 2006 and Chiu et al., 2008; and Spirulina sp.: de Morais and Costa, 2007), CO 2 concentration/mass loading rate (Cheng et al., 2006; de

Morais and Costa, 2007; Chiu et al., 2008; Ho et al., 2010; Vidyashankar et al., 2013), CO 2 holding time (Cheng et al., 2006; Chiu et al., 2008), and temperature (Basu et al., 2013). Among these factors, CO 2 mass loading rate needs to be carefully controlled in order to achieve higher removal efficiency. Indeed, our study shows that the value of E CO2 is highly dependent on the

CO 2 mass loading rate. Also, the CO 2 mass loading rates used in previous studies varied widely; thus, further research is needed to evaluate the CO2 removal efficiency relative to the CO 2 mass loading rate.

3.4 Biomass chemical composition

Table 3 shows the proximate analysis of the algal biomass grown under different CO 2 loading rates. At each loading rate, the nitrogen concentration in the medium was maintained at a relative constant level (37-45 mg/L). The biomass contained about 7‒10% ash content. The lipid content of the biomass was relatively constant (7‒9 %). The protein content was also relative

stable, except it increased to 61% at the highest CO 2 mass loading rate. Similarly, the carbohydrate content was also maintained at a relatively constant level except the value decreased at the highest CO 2 mass loading rate. The increase in protein content may be due to the increased carbon building blocks available for amino acid production, increased levels of enzymes such as extracellular carbonic anhydrase (Nimer et al., 1998), and the inorganic carbon concentrating mechanism (CCM) for cells to adapt to a high CO 2 concentration (Kaplan and

Reinhold, 1999). On the other hand, the decreased carbohydrate content at the highest CO 2 loading rate may be due to the decreased intracellular carbonic anhydrase (CAint) activity resulting from excessive CO 2 accumulation (Walsh and Henry, 1991; Henry, 1996).

Fatty acid composition is important when algal biomass is being considered for animal feed. The fatty acid profiles of the biomass are shown in Table 4. Within the range of CO 2 loading rates investigated, the fatty acid compositions were relatively stable. C16:0, C18:0,

C18:1, and C18:2 accounted for more than 90% of TFA, which were in agreement with previous report (Ho et al., 2010). The saturated-, monounsaturated-, and polyunsaturated-fatty acids portion of the algal biomass were also monitored. Saturated fatty acids tended to decrease while unsaturated fatty acids increase with the increasing of CO 2 loading rate. The trends of saturated vs. unsaturated fatty acids were probably due to the decreased intracellular carbonic anhydrase

(CAint) activity and enhanced unsaturation of cell membrane fatty acyl chain at high CO 2 levels

(Walsh and Henry, 1991; Henry, 1996).

Amino acid profile is another important parameter for evaluating the nutritional value of the algal biomass. As shown in Table 5, the individual amino acid composition (% total amino acid) was not significantly different overall except for hydroxyproline, threonine, proline, glycine, cysteine, methionine, isoleucine, lysine, and arginine ( p<0.05). The total amino acid

contents (% DW) obtained at the lower CO 2 mass loading rates (0.64 to 1.71 g/L-day) were significantly different from those obtained at the higher CO 2 mass loading rates (2.26 to 5.49 g/L-day) ( p<0.05). The trend of amino acid contents with the CO 2 loading rate can be explained by the localization of amino acid biosynthesis pathway at plastid (Reyes-Prieto and Moustafa,

2012); the extracellular carbonic anhydrase (CAext) (Nimer et al., 1998), and inorganic carbon concentrating mechanism (Kaplan and Reinhold, 1999) under different CO 2 availability levels.

Table 5 also shows the ratio of essential amino acids (EAA) to non-EAA of the biomass was roughly 1:1. The ratio of EAA to non-EAA has been an important parameter to minimize N excretion from farm animals (Boisen et al., 2000), and it is suggested a minimum portion of

EAA to be 0.45 for pigs (Wang and Fuller, 1989) in order to balance the surplus nitrogen from non-EAA with EAA. Therefore, the 1:1 ratio of EAA to Non-EAA obtained in this work exceeds this minimal level, indicating the algal biomass satisfies the recommendation.

The above results show that animal biomass obtained in this work can be a good protein source for animals. However, whether the algal protein can be fed to animal directly or blended with other animal feeds depends on the compositions of feed typically used for different animals.

Here, the value of biomass as a potential animal feed was evaluated by comparing the algal amino acid profile to the “ideal” amino acid profile for different animals such as broilers, pigs, and cows. The “ideal” amino acid profile is defined as the composition that can be efficiently consumed by the animal with minimal ammonia excretion and no loss of performance (Boisen et al., 2000). As shown in Table 6, the algal protein has roughly the same amino acid composition as the “ideal” amino acid profiles for the three animals, although some specific amino acids may be over supplied (e.g., phenylalanine and phenylalanine+tyrosine for broilers; methionine and phenylalanine for pigs; and methionine+cysteine and phenylalanine+tyrosine for cows). These

55 differences can be adjusted by blending the algal protein with other protein sources to reach the final desirable ratios.

The above results show that the microalga culture can be potentially used as a bio- scrubber for removing ammonia gas and CO 2 from animal house operations. However, several issues need to be addressed in order to implement such a system to a commercial animal farm.

First, the collection of ammonia- and CO 2-rich air from the animal house and delivery to the algal reactor need to be carefully designed. For a commercial animal feeding operation, the algal reactor can be installed indoor or outdoor of the animal house. The indoor design facilitates the gas delivery and exhaust of the reactor, but the reactor may be light limited. The outdoor design takes advantage of using natural sunlight; however, the gas delivery system needs to be sealed to avoid any leaking. The exhaust gas may also need to be recycled back to the animal house to reduce the loss of the gas out of the animal house. Second, the composition of “real” gas from the feeding house will be complicated compared to the artificial ammonia- and CO 2-blended gas as used in this work. The trace amount of volatile compounds such as NO x, methane, H 2S can affect the algal growth performance. As it is difficult to change air composition by modifying the animal feeding operation, the algal culture system needs to be robust and flexible to the gas composition variations. Lastly, future studies on pilot scale algal culture are needed in order to estimate the cost of removing ammonia and CO 2 under the context of different sizes of the commercial feeding operations.

Conclusion

In summary, this work reported an algal culture as an effective process for removal of ammonia gas and CO 2 from inlet gas. Within the wide range of the CO 2 mass loading rates studied, the ammonia and CO 2 removal efficiency can reach as high as 97% and 78%, respectively. This process can be potentially be used as a bio-scrubber for mitigating ammonia and CO 2 emissions from animal production operations. In addition, the algal biomass contained high protein content with an amino acid profile similar to the ideal protein compositions for different animal feeds.

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Table 1 . The CO 2 content in the inlet gas and the corresponding CO 2 mass load rate for the algal culture reactor.

CO 2 content in the CO 2 mass loading rate

inlet gas (ppm) (g/L day)

395 a 0.64

728 1.18

1055 1.71

1394 2.26

2608 4.22

3395 5.49 a No external CO 2 was introduced into the reactor. The CO 2 content in this case was based on the atmospheric CO 2 in the inlet air, which was estimated as 395 ppm based on NOAA (2013).

a Table 2 . Nitrogen mass balance for different CO 2 loading rates.

Nitrogen CO 2 mass loading rate (g/L-day) distribution 0.64 1.18 1.71 2.26 4.22 5.49 (%)

NInlet 100 100 100 100 100 100

NCell 91.5±1.35 93.1±1.16 90.8±1.27 88.0±0.50 86.3±0.99 82.8±2.40

NLiquid 11.0±0.16 12.5±0.78 12.8±0.24 10.7±0.30 10.6±0.49 12.7±0.22

NOutlet 3.27±0.03 2.50±0.01 3.30±0.02 3.08±0.10 2.81±1.28 2.80±0.13

error -5.71 -8.07 -6.90 -1.73 0.27 1.66 a Data are expressed as mean ± standard deviation of five consecutive samples at each steady state of the continuous algal culture.

Table 3. Proximate analysis of the algal biomass. a

Residual Algal biomass composition CO 2 mass nitrogen loading rate concentration Ash Lipid c Protein Carbohydrate (g/L day) (mg/L) (%) (%) (%) (%) 0.64 38.5 ± 0.47 7.16 ± 0.57 8.31 ± 1.02 55.78 ± 0.73 28.8 ± 1.34 1.18 45.6 ± 2.91 7.57 ± 0.28 8.68 ± 0.22 54.98 ± 0.28 28.8 ± 0.46 1.71 45.6 ± 0.66 7.88 ± 0.21 7.36 ± 1.01 55.04 ± 0.51 29.7 ± 0.83 2.26 39.2 ± 0.78 7.99 ± 0.14 8.16 ± 0.24 55.24 ± 0.28 28.6 ± 0.10 4.22 37.2 ± 1.75 8.82 ± 0.19 7.85 ± 0.03 57.22 ± 0.80 26.1 ± 0.59 5.49 44.7 ± 0.75 10.05 ± 0.37 7.38 ± 0.47 61.03 ± 0.77 21.5 ± 0.54 a The liquid sample for residual nitrogen and biomass samples for composition analyses were all collected at the steady state of each condition. b Data are expressed as mean ± standard deviation of five consecutive samples collected at each steady state. c The total fatty acid content was used to approximate as the lipid content.

a Table 4 . Fatty acid profile of the algal biomass obtained at different CO 2 mass loading rates.

CO mass loading rate (g/L day) Fatty acid composition 2 5.49 (% TFA) 0.64 1.18 1.71 2.26 4.22 12:0 4.66±0.06 4.95±0.01 4.60±0.04 4.37±0.02 5.32±0.00 5.34±0.07 12:1 2.64±0.03 2.77±0.00 2.67±0.02 2.57±0.02 3.23±0.00 3.24±0.03 16:0 13.6±0.10 13.6±0.03 15.5±0.05 14.3±0.11 13.0±0.02 12.1±0.10 16:1 4.34±0.03 4.08±0.04 3.31±0.01 2.82±0.02 3.85±0.03 5.56±0.15 18:0 11.3±0.06 11.6±0.02 12.9±0.08 11.7±0.08 10.7±0.01 11.0±0.11 18:1 28.5±0.23 29.6±0.07 34.3±0.17 31.6±0.29 29.4±0.01 31.4±0.34 18:2 31.3±0.34 29.8±0.05 22.6±1.32 29.2±0.24 31.4±0.02 28.5±0.77 18:3 3.14±0.03 3.14±0.01 3.38±0.02 2.82±0.03 2.65±0.01 2.60±1.30 20:0 0.51±0.00 0.49±0.00 0.70±0.00 0.54±0.00 0.38±0.00 0.38±0.09 32.5 30.7 33.7 31.0 29.4 28.8 Saturates (%TFA) 38.3 36.4 40.3 37.0 36.5 40.2 Monosaturates (%TFA) 29.2 32.9 26.0 32.0 34.1 31.1 Polyunsaturates (%TFA) a Data are expressed as mean ± standard deviation of five consecutive samples at each steady state of the continuous algal culture.

a Table 5 . Amino acid (AA) profiles of the algal biomass at different CO 2 mass loading rates.

Amino acid CO 2 mass loading rate (g/L-day) composition b 5.49 0.64 1.18 1.71 2.26 4.22 (% total AA) Taurine 0.06±0.01 0.06±0.01 0.06±0.00 0.06±0.00 0.05±0.01 0.04±0.01 Hydroxyproline 0.18±0.10 0.21±0.08 0.41±0.00 0.15±0.00 0.21±0.06 0.17±0.03 Aspartic Acid 9.65±0.03 9.64±0.03 9.62±0.06 9.60±0.01 9.58±0.03 9.61±0.03 Threonine 5.24±0.13 5.31±0.13 5.29±0.04 5.22±0.02 5.05±0.08 5.00±0.04 Serine 3.91±0.18 4.00±0.17 4.13±0.26 3.85±0.06 3.81±0.07 3.78±0.10 Glutamic Acid 10.68±0.12 10.62±0.12 10.84±0.25 10.74±0.01 10.78±0.05 10.95±0.05 Proline 5.20±0.11 5.21±0.11 5.23±0.01 5.12±0.01 5.11±0.05 4.96±0.02 Lanthionine 0.00±0.00 0.00±0.00 0.00±0.00 0.00±0.00 0.00±0.00 0.00±0.00 Glycine 6.40±0.07 6.42±0.07 6.42±0.02 6.47±0.03 6.58±0.06 6.50±0.03 Alanine 8.63±0.05 8.65±0.05 8.71±0.06 8.67±0.02 8.62±0.04 8.61±0.04 Cysteine 1.28±0.06 1.31±0.06 1.33±0.03 1.33±0.03 1.23±0.02 1.21±0.01 Valine 6.06±0.05 5.92±0.05 5.96±0.01 6.01±0.07 6.01±0.05 6.04±0.06 Methionine 2.43±0.02 2.42±0.02 2.41±0.01 2.43±0.01 2.44±0.01 2.45±0.00 Isoleucine 4.26±0.07 4.16±0.07 4.16±0.02 4.23±0.05 4.27±0.02 4.33±0.05 Leucine 9.61±0.04 9.57±0.04 9.49±0.04 9.58±0.04 9.52±0.03 9.51±0.02 Tyrosine 4.24±0.07 4.24±0.07 4.10±0.13 4.21±0.01 4.21±0.02 4.22±0.03 Phenylalanine 6.28±0.11 6.21±0.11 5.98±0.22 6.20±0.03 6.14±0.02 6.11±0.03 Hydroxylysine 0.28±0.08 0.27±0.08 0.14±0.14 0.25±0.00 0.25±0.01 0.25±0.00 Ornithine 0.24±0.08 0.33±0.06 0.18±0.08 0.29±0.06 0.30±0.05 0.31±0.03 Lysine 6.66±0.11 6.58±0.12 6.55±0.04 6.65±0.02 6.68±0.01 6.85±0.02 Histidine 2.27±0.03 2.26±0.03 2.24±0.02 2.28±0.01 2.30±0.02 2.30±0.01 Arginine 5.68±0.13 5.68±0.13 5.69±0.01 5.74±0.01 5.91±0.01 6.01±0.03 Tryptophan 0.78±0.13 0.94±0.11 1.07±0.06 0.92±0.10 0.96±0.06 0.79±0.13 Total AA 50.2 48.8 49.3 48.2 50.2 52.1 (% DW) EAA: non-EAA 49.3:50.7 49.1:50.9 48.8:51.2 49.3:50.8 49.3:50.7 49.4:50.6 a Data are expressed as mean ± standard deviation of five consecutive samples at each steady state of the continuous algal culture. b The amino acid in bold are essential amino acids.

Table 6 . The essential amino acid profile of algal biomass compared to the ideal amino acid profiles for different animals. a

Amino acid Ideal protein compositions for different animals composition Microalgae (as of % lysine) Broilers Growing pigs Dairy cows Lysine 100 100 100 100 Methionine 36‒37 36‒48 n/a 31‒37 Methionine+Cysteine 54‒57 72‒81 50‒63 n/a Threonine 73‒81 65‒74 60‒75 47‒75 Arginine 85‒88 105‒110 31‒42 43‒67 Valine 88‒91 50‒85 64‒75 73‒85 Isoleucine 63‒64 48‒75 50‒61 61‒74 Leucine 139‒145 80‒144 72‒110 110‒131 Tryptophan 12‒16 16‒19 15‒19 15 Histidine 34 32‒35 26‒36 33‒42 Phenylalanine 89‒94 n/a n/a 62‒76 Phenylalanine+Tyrosine 151‒159 n/a 88‒122 n/a Reference This study Salehifar et al., 2012 Boisen, 2000 Haque et al., 2012 a Lysine is the limiting amino acid for most protein sources and is used as a benchmark; all the other amino acids are represented as a relative percentage to lysine.

3.5 0.35 Cell density Biomass productivity 3 0.3

2.5 0.25

2 0.2

1.5 0.15

BIomass (g/L) BIomass yield 1 0.1 Biomass productivity (g/L-day) Biomass productivity 0.5 0.05

0 0 0.64 1.18 1.71 2.26 4.22 5.49

CO 2 loading rate (g/L day) ···

Figure 1 . Biomass yield and productivity of S. dimorphus under different CO 2 mass loading rates (the corresponding CO 2 concentration in the inlet gas is shown in Table 1). Dilution rate, medium pH, and ammonia mass loading rate were kept at 0.1 day -1, 7.0, and 42.4 mg/L day, respectively. Data are means of five consecutive samples at equilibrium stage of the continuous· culture and error bars represent standard deviations.

0.05

0.04

/L-day) 0.03 3

(g NH (g 0.02 NH3 R 0.01

0 0.64 1.18 1.71 2.26 4.22 5.49

CO 2 loading rate (g/L day) ···

100

60 (%) NH3

E 40

0 0.64 1.18 1.71 2.26 4.22 5.49

CO 2 loading rate (g/L day) ···

Figure 2. Ammonia gas removal rate (R NH3 ) and removal efficiency (E NH3 ) by the continuous culture of S. dimorphus under different CO 2 mass loading rates. Dilution rate, medium pH, and ammonia loading rate were kept at 0.1 day -1, 7, and 42.4 mg/L day, respectively. Data are means of five consecutive samples at equilibrium stage of the continuous· culture and error bars represent standard deviations.

0.70 (A) 0.60

0.50

0.40 /L-day) 2 0.30 (g CO (g 0.20 CO2 R 0.10

0.00 0.64 1.18 1.71 2.26 4.22 5.49

CO 2 loading rate (g/L day)

··· 100 (B)

60 (%) CO2

E 40

0 0.64 1.18 1.71 2.26 4.22 5.49

CO 2 loading rate (g/L day)

···

Figure 3. CO 2 removal rate (R CO2 ) and removal efficiency (E CO2 ) by the continuous culture of S. dimorphus under different CO 2 mass loading rates. Dilution rate, medium pH, and ammonia loading rate were kept at 0.1 day -1, 7, and 42.4 mg/L day, respectively. Data are means of five consecutive samples at equilibrium stage of the continuous· culture and error bars represent standard deviations.

CHAPTER 4. ASTAXANTHIN PRODUCTION FROM HAEMATOCOCCUS PLUVIALIS

USING AMMONIA GAS FROM ANIMAL PRODUCTION OPERATIONS

Juhyon Kang and Zhiyou Wen

Abstract

Ammonia gas from concentrated animal feeding operations (CAFOs) causes detrimental effects on environment and health. The increasing demand for clean air and reinforced regulations urge to develop an effective and environmentally sound ammonia gas removal system. In our previous studies, microalgae showed excellent ammonia gas removal performance and ammonia removal rate. Microalgal ammonia scrubber system can enhance its feasibility by co-producing high value byproducts from the microalgae biomass. In this study, Haematococcus pluvialis was grown in the ammonia scrubber system to produce astaxanthin. Astaxanthin is a valuable ketocarotenoid used in aquaculture, cosmetic, and pharmaceutical industries due to its pigmentation and antioxidant effects. In our study, we used 1L tapered bubble column to achieve both high cell density and astaxanthin induction in one reactor. The growth condition was tested at different pH levels (4, 5, 6, 7, and 8) and different nitrogen sources (ammonium, nitrate, and urea). H. pluvialis showed highest cell density at pH 5-7 but produced the highest astaxanthin at pH 4. After confirming that H. pluvialis can grow with ammonium as sole nitrogen source, the ammonia gas was fed at different concentrations at pH 4 and 5. In order to increase astaxanthin content, different fed-batch strategies and different chemicals to generate stressful condition

71 were examined. From the data, it was concluded that H. pluvialis can utilize ammonia gas as a nitrogen source but further astaxanthin induction was only effective at C/N ratio 1.

Keywords: ammonia gas mitigation, Haematococcus pluvialis, astaxanthin

1. Introduction

Ammonia emission has been a culprit for a number of environmental and human health problems. With presence of acidic species such as NO x or SO x, ammonia gas readily reacts and forms particulate matter (PM 2.5 ), causing smog and respiratory diseases. When ammonia reaches the surface water or soil it causes eutrophication, soil acidification, affecting the ecosystem

(Breemen et al., 1982; Sommer 2001). Currently, the majority of ammonia emissions in the

United States is from concentrated animal feeding operations (CAFOs) (EPA, 2015), from either the direct volatilization from the liquid or degradation of organic nitrogen from manure.

Various methods have been attempted for minimizing ammonia generation and emission from concentrated animal feeding operation. Those efforts are based on either animal production such as changing dietary feed composition, chemical treatment, physical adsorption, or “end-of- pipe” approaches such as acid scrubbing, or bacteria-based biofilters. While those methods are overall effective in term of the ammonia reduction, the implementation is usually prohibitively expensive (Groot Koerkamp, 1994; Philippe et al., 2011; Xin, 2011) or causing additional environmental problems such as acid stress (Ndegwa et al., 2008).

Microalgae cultivation has been proved as an efficient method for reducing ammonia emission. For example, in a laboratory study using model reagent grade ammonia gas, we have

72 achieved >95% ammonia removal efficiency using green alga Scenedesmus dimorphus , with majority ammonia gas assimilated into algal biomass (Kang et al., 2014). In another study, ammonia gas was blended with different concentration of CO 2 to mimic the “real ventilated air” in animal poultry house. A similar high ammonia removal efficiency (>95%) and moderate level of CO 2 removal efficiency (78%) was achieved (Kang and Wen, 2015). These studies show the great potential of using microalgae as ammonia mitigation solution in animal production operations.

One key to make the algal-based ammonia mitigation economically feasible is to explore the value of the biomass, which can compensate the cultivation cost. In the previous study, for example, the alga S. dimorphus contained an appreciable amount of proteins with an amino acid makeup close to the ideal protein profiles for typical animal feeds (Kang and Wen, 2015). In order to further improve the economic value of the algal based ammonia mitigation, however, it is desirable to produce an algal biomass with higher value than animal proteins. To address this requirement, we hereby explore the feasibility of using the alga Haematococcus pluvialis as a candidate strain for producing a high-value astaxanthin using ammonia gas.

Astaxanthin as an antioxidant is effective for quenching singlet oxygen and deactivate free radicals through its conjugated double bond and ketone (C=O) and hydroxyl (OH-) group

(Rodriguez-Amaya, 2001). It can also enhance human immune system and reduce tumor growth and inflammation (Higuera-Ciapara et al. 2006). In aquaculture, astaxanthin is responsible for reddish hue on the shell of crustacean and on the skin or flesh of marine animals such as salmon, trout, and ornamental fish (Del Campo et al. 2007). However, human and animals cannot synthesize astaxanthin de novo , but rather obtain it by digesting microalgae that is the primary astaxanthin producer (Lorenze and Cysewski, 2000).

Astaxanthin is predominantly produced through chemical synthesis. However, there is an increasing interest in naturally occurring astaxanthin (Higuera-Ciapara et al. 2006; Margalith,

1999). Among various astaxanthin-producing microorganism, H. pluvialis has been recognized as the most promising producers (Orosa et al., 2001; Domínguez-Bocanegra et al. 2004; Lorenze and Cysewski, 2000). The aim of this research is to demonstrate the feasibility of using H. pluvialis as a potential ammonia scrubber for concentrated animal house operations while producing a high value astaxanthin.

2. Materials and Methods

2.1. Cell seed and culture medium

Haematococcus pluvialis (UTEX 2505) was obtained from the University of Texas Cell

Collection. The strain was stored on agar slant at 4°C. To prepare the seed culture, the cells in the agar slant were transferred to 250 mL Erlenmeyer flasks each containing 100 mL medium, and incubated in an orbital shaker (150 rpm) at 22°C with continuous illumination at 90-100

2 µmol/m ∙s. The MES-volvox medium used for subculture contained 0.5 mM Ca(NO 3)2∙4H 2O,

0.16 mM MgSO 4∙7H 2O, 0.16 mM Na 2glycerophosphate∙5H 2O, 0.67 mM KCl, 10 mM MES, 0.5 mM NH 4Cl, 12 µM Na 2EDTA∙2H 2O, 2.16 µM FeCl 3∙6H 2O, 1.26 µM MnCl 2∙4H 2O, 0.22 µM

ZnCl 2, 0.05 µM CoCl 2∙6H 2O, 0.102 µM Na 2MoO 4∙2H 2O, 0.135 mg/L vitamin B 12 , 24 mg/L

HEPES buffer pH 7.8, and 0.025 mg/L biotin. The medium was pH 4 before autoclaved at 121°C for 15 min. The seed flask culture was inoculated into 1-L bubble column photobioreactors for investigating various culture conditions. The inoculation size was 10% and the average initial cell density was 3.8 · 10 4 cells/ml.

2.2. Photobioreactor set up

Five tapered glass bubble column photobioreactors (6.5 cm diameter × 50 cm height) with 1-L working volume were used for growing H. pluvialis under different experimental conditions. Air was bubbled from the bottom to the reactor. Each glass column was sealed with a rubber stopper with three ports for aeration tube, gas exhaustion, and sample collection, respectively. The aeration rate was maintained at 800 mL/min (0.8 vvm) to prevent cells from settlement. The reactors were operated at batch mode. The reactors were maintained at 22°C with continuous light illumination at 230 µmol/m 2∙s.

In the study of the pH effect, the initial medium pH was adjusted to a range of pH 4-8 using 4 M NaOH, the medium composition was the same as MES-volvox medium used in the seed culture. In order to test the effects of nitrogen source, Ca(NO 3)2∙4H 2O and NH 4Cl in the

MES-volvox medium were replaced with Ca(NO 3)2∙4H 2O, NH 4Cl, or urea as single nitrogen source at the equivalent nitrogen level (1.5 mM). In the study of using ammonia gas for algal culture, ammonia-laden air was bubbled through the reactors, while the original nitrogen sources in MES-volvox medium was eliminated.

Compressed air and ammonia gas were pre-mixed at a certain ratio of flow rate to achieve the ammonia concentration in the inlet gas. The flow rate of compressed air was maintained at 800 ml/min, while the volumetric flow rates of the ammonia gas were varied to realize the ammonia concentration in the inlet gas as 20, 41, 82, and 164 ppm. Correspondingly, the mass flow rate readings (i.e., the standard cubic centimeter per minute, SCCM) of the ammonia gas were 0.032, 0.041, 0.066, and to 0.133 mL/min. The flow rate of ammonia gas and air were respectively regulated by a digital flow meter (MCS series for ammonia gas; MC series

for air, Alicat Scientific, Tucson, AZ). The ammonia gas loading rate (Q, unit: gNH 3/L-min) was calculated based on the following equation,

(1) [ ]×[] = Where [NH 3] is ammonia concentration in inlet gas (SCCM), r NH3 is the ammonia density

(0.704 g/L at operational conditions), and V is the reactor working volume (1 L).

2.3. Ammonia feeding strategies

Ammonia gas was initially fed to the photobioreactors on a constant way with a certain flow rate and concentration in the inlet gas, which resulted in a fixed mass loading rate (Q). To balance the cell growth and the astaxanthin formation in the cells, different ammonia feeding strategies were used for feeding the photobioreactors. As shown in Table 1, the feeding tests were divided into three groups, with each group of the culture was fed the ammonia gas at certain concentration (20, 41, and 82 ppm, corresponding 32.4, 41.6, and 67.3 mg/L/day loading rate). Within each group, the reactors were fed with either ammonia-containing gas or ammonia- free gas based on different schedules (Table 1). This “on and off” scheme of ammonia feeding enable the cells experienced an alternative of the nitrogen-suffice and nitrogen-starving status, which can alternatively support cell growth and induce the astaxanthin formation. MES-volvox-

N medium was used for the cell culture. Each reactor was inoculated with the same amount of seed (10 %).

2.4. Induction of astaxanthin accumulation

Astaxanthin induction of ammonia-gas based algal culture was performed by applying different chemicals (sodium acetate, H 2O2, and FeCl 36H 2O) to the vegetative cells of H.

76 pluvialis culture. The chemical species and concentrations were chosen based on the literature. i.e., sodium acetate was used to adjust different C/N ratios of the medium (Kakizono et al., 1992);

H2O2 was used to generate free radicals (Ip and Chen, 2005); and iron (Borowitzka et al., 1991) was used as an electron acceptor.

The cells from the ammonia feeding strategy test (67.3 mg/L/day loading rate columns 2 and 4 at day 11 culture, Table 1) were used as a seed and inoculated into 250-mL Erlenmeyer flasks (50-mL working volume) containing MES-volvox-N medium and various inducing chemical at different concentration. The flasks containing no inducing reagent were used as control. The initial ammonia concentration in the column liquid (~ 1 mg/L) was measured and sodium acetate trihydroxide was added accordingly to make the C/N ratios of the medium at 1, 2,

2+ 5, and 10. In case of H 2O2, 0.05, 0.1, and 0.2 mM were added with 18µM Fe to generate oxidative species by Fenton chemistry as Ip and Chen (2005) achieved the highest astaxanthin content at 0.1 mM H2O2.

2+ 3+ - • Fe + H 2O2 → Fe + OH + OH (2)

FeCl 36H 2O was tested for 0.1, 0.2, 0.5, and 1 mg/L concentrations as Borowitzka et al. (1991) tested up to 0.24 mg/L and induced little astaxanthin.

The flasks were autoclaved at 121°C for 12 min before inoculation of the seed. The flasks were placed at 22°C with continuous illumination 230 µmol/m 2∙s. The cells were harvested after eight days of incubation. The dry biomass weight and astaxanthin content were measured after freeze drying the cells.

2.5. Analyses

Cell growth of H. pluvialis was evaluated by maximum cell density and the specific growth rate. The cell density was based on counting the cell number with a haemocytometer.

The specific growth rate was measured as:

(3) = Where X is cell density (cells/ml) and t is time (day).

In the astaxanthin induction test (Section 3.5), the dry cell weight concentration was used instead of the cell number for evaluating the cell growth.

To determine the astaxanthin content, the cell biomass in the broth was harvested by centrifuging the broth at 10,000 g for 2 min. The cell pellet was then freeze-dried for astaxanthin analysis. The supernatant was used for ammonia concentration analysis which was determined using an Orion high performance ammonia electrode (model 9512 HPBNWP Thermo Scientific,

Beverly, MA) according to EPA-approved standard methods 4500-NH 3 E.

The extraction of the astaxanthin from the biomass was based on the method reported by

Mendes-Pinto et al. (2001). In summary, the 40 mg lyophilized biomass was submerged with acetone (2 ml, HPLC grade). After being left static for 16 hours at room temperature, the upper phase of the mixture was collected, and the cell pellet was re-extracted using the same procedure for two to three times until the cell debris became colorless. The whole procedure was performed in dim light environment. The collected astaxanthin-containing solution was then condensate to 1 ml by nitrogen gas evaporation, and analyzed using LC-APCI-MS (liquid chromatography- atmospheric pressure chemical ionization-mass spectrometry). The APCI was in the positive ion mode using reverse phase XDB-C18 column (50×4.6 mm, 1.8µm). The MS analysis was on the

Q-TOF system. The astaxanthin-containing condensate was eluted using a mobile phase of

78 isocratic solvent system consisting of methanol:water (90:10, v/v) for 15 minutes at a flow rate of 0.8ml/min and injection volume was 1.0 uL. The LC column was re-equilibrated with acetone prior to injection of the next sample. The APCI vaporizer temperature was maintained at 350 oC and the corona discharge needle current was set at 4.0 uA in APCI interface. Free astaxanthin and its esterified forms were identified by comparing retention time and quasimolecular ions [M-

+ + + + H2O] , [M+H] , [ME+H] , and [ME-H2O] with authentic astaxanthin standard (Adipogen Co.,

San Diego, USA) and astaxanthin esters identified by Holtin et al. (2009).

2.6. Statistical analysis

The significant difference between data was analyzed by One-way analysis of variance

(ANOVA) at 95% confidential level using PROC GLM and LSMEANS, with statistical package

SAS (version 9.4 SAS Institute, Inc., Cary, NC, USA).

3. Results and discussion

3.1. Effect of initial pH on the cell growth and astaxanthin production

pH as one of the most important environmental parameters has been reported affecting the growth and astaxanthin production of H. pluvialis (Orosa et al., 2001; Sarada et al., 2002;

Zhang et al., 2003). In addition, the medium pH will strongly affect the dissolubility of ammonia gas in the liquid. Therefore, the initial study in this work is to evaluate the effect of the medium pH on the H. pluvialis culture so that ammonia gas feeding can be optimized. The pH commonly reported in the H. pluvialis culture is around 7 (Kobayashi et al., 1991; Kang et al., 2005). In this work, the pH effect was evaluated within a wide range from 4-8. As shown in Figure 1a, cells at

79 pH 4 and 8 exhibited a longer lag phase and lower cell density than the other cultures ( p < 0.05), while pH 6 and 7 resulted in a faster growth rate as well as the maximum cell density. The cells at pH 5 also had a longer lag phase, but eventually caught up the maximum cell density with the cultures at pH 6 and 7 ( p > 0.05). The specific growth rate and maximum cell density obtained from each culture were further summarized in Figure 1b. Overall, the trend of the cell growth with different pH is consistent to previous studies in which pH 7 was reported as the optimal pH for H. pluvialis cultures (Sarada et al., 2002; Zhang et al., 2003).

During the culture period, the culture was maintained as greenish color for most of time, and turned pinkish color at the very late stage (day 8 for pH7 and day 10 for pH6), indicating the accumulation of the astaxanthin by aging. As shown in Figure 1c, the culture at pH 4 resulted in higher astaxanthin content than other cultures. Orosa et al., (2001) also reported using pH 4 as one induction method for successfully inducing astaxanthin accumulation of H. pluvialis , compared to the culture at neutral pH. However, the astaxanthin content obtained in this study (2

– 5 mg/g DW or 0.2-0.5%, w/w) was lower than most other studies using two-stage method in which stressful conditions was applied to the second stage (1.1-3.6 % w/w, Wang et al., 2013).

To further enhance the astaxanthin production, appropriate induction methods need to be applied.

3.2. Cell growth and astaxanthin production of H. pluvialis under different N sources

In order to compare the nitrogen source effect on the cell growth and astaxanthin production, different nitrogen sources, ammonium, urea, and nitrate were tested. The effects of nitrogen source including ammonium, nitrate, and urea on H. pluvialis growth and astaxanthin production have been studied and nitrate is in general regarded as the better nitrogen source for the synthesis of the carotenoid including astaxanthin (Borowitzka et al., 1991; Sarada et al.,

2002). However, in order to provide a better way of using ammonia gas, the cell growth and astaxanthin accumulation using ammonia as nitrogen source need to be evaluated.

The ammonium, urea, nitrate were added to the medium at a same concentration (1.5 mM). The medium pH in these three cultures was adjusted to 7 an optimal level for algal growth

(Stross, 1963; Sarada et al., 2002; Zhang et al., 2003). Figure 2a shows that urea led to a higher growth rate than the other two nitrogen sources. While the culture using ammonium and nitrate had a similar growth rate, the maximum cell density obtained using ammonium was higher than that of nitrate based culture. Figure 2b summarizes the specific growth rate and the maximum cell density. The cell densities from the three nitrogen sources were significantly different ( p <

0.05). The specific growth rate of the urea based culture was significantly higher than the other cultures ( p < 0.05). The above growth and astaxanthin production performance with different nitrogen species were comparable to the previous report (Borowitzka et al. 1991), in which ammonium gave the best cell growth and nitrate gave the best carotenoid formation.

During the entire culture, the cells in the ammonium-containing medium retained a dark green color, indicating minimal astaxanthin synthesis. The other two cultures (nitrate and urea) turned pinkish at the later stage of culture, the astaxanthin contents of these two cultures were therefore determined. It was found that the astaxanthin content of the cells using urea and nitrate was not significantly different ( p > 0.05).

3.3. Cell growth and astaxanthin production using ammonia gas as nitrogen source

In the study of using H. pluvialis culture as ammonia gas scrubber, an acidic pH was preferred as it will increase the dissolubility ammonia. On the other hand, previous result also showed that the growth of the H. pluvialis cells was maintained at a higher level within the pH

81 range 5-7 (Figure 1b), while pH 4 resulted in the high astaxanthin accumulation (Figure 1c).

Therefore, pH 4 and 5 were used for the algal culture.

For most algal species including H. pluvialis, ammonia is an ideal nitrogen source if the concentration is controlled at an appropriate level (Natarajan, 1970; Abeliovich and Azov, 1976;

Azov and Goldman, 1982; Källqvist and Svenson, 2003). Therefore, in this work, the effect of ammonia gas loading rate on the H. pluvialis culture was first investigated. As shown in Figure

3a and 3b, the cell growth was low at 32.4 mg/L/day ammonia loading, and drastically increased when loading rate increased to 41.6 and 67.3 mg/L/day. When loading was further increased to

134.8 mg/L/day, the cell growth was significantly inhibited. The cultures at pH 4 and 5 exhibited a similar trend with the ammonia loading rate. The maximum cell density and specific growth rate of the H. pluvialis cultures are summarized in Figure 3c. Ammonia gas loading at

41.6 mg/L/day resulted in much higher cell density and growth rate than other loading rates

(p<0.05).

One unique feature using ammonia gas for H. pluvialis culture was that the culture was changing the color from green to pinkish at the later stage of cultivation, indicating the astaxanthin accumulation. Compared to Figure 2c where no astaxanthin in the ammonium- containing medium, the moderate astaxanthin accumulation of the cultures using ammonia gas indicates the unique feature of the H. pluvialis culture as ammonia gas scrubber. Quantitative analysis of the astaxanthin from the harvested biomass confirmed the observation. As shown in

Figure 3d, all the culture except the cases of 134.8 mg/L/day ammonia gas loading produced the astaxanthin. The astaxanthin content appeared the highest at 32.4 mg/L/day ammonia loading rate, and decreased as ammonia concentration increased ( p<0.05). At 134.8 mg/L/day ammonia

82 loading, analysis of cellular astaxanthin content was not available due to the complete inhibition of the cell growth.

Using ammonia gas a nitrogen source for H. pluvialis has never been attempted in the literature. Compared to the ammonium ions which need transporters (González-Ballester et al.,

2004; Antonenko et al., 1997), the ammonia can freely diffuse into the cells through partial difference between cell membrane and environment. Therefore, the algal growth is very sensitive to the ammonia gas loading applied to the photobioreactors. The diffusion of ammonia specie into the cell usually causes the raise of intracellular pH, which eventually inhibits the cell growth. Indeed, ammonia is known to induce oxidative stress to cells by deactivating antioxidant enzymes and increasing superoxide formation (Kosenko et al., 1999). Based on those facts, it was assumed that ammonia gas as a nitrogen source plays two roles in the H. pluvialis culture. On one hand, it will serve as a nitrogen source for the cell growth, although the cell growth performance was not as good as other nitrogen sources such as nitrate or urea. On the other hand, the stressful conditions caused by the ammonia inhibition induce the astaxanthin formation. This dual-effect of ammonia gas on the H. pluvialis culture was supported by the report by Wang et al., (2003), in which H. pluvialis cells can maintain their PSII function during the induction phase and red cyst cells maintain their structural integrity during the photosynthesis.

To balance the cell growth and the astaxanthin formation in the cells, different ammonia feeding strategies were used for feeding the photobioreactors (Table 1) to compromise the cell growth and astaxanthin synthesis.

3.4. Cell growth and astaxanthin production under different ammonia feeding strategies

Table 1 summarizes different ammonia gas fed schedule and corresponding culture color changes. Figure 4 shows that all the feeding schedules supported the cell growth. A summary of the growth performance is presented in Figure 5a. The data in Figure 5a is the cell density on the last three days of culture and does not represent in the steady-state. Cell densities of feeding strategy #2 at 32.4 and 41.6 mg/L/day were significantly lower ( p < 0.05) than other columns, most probably due to the initial growth inhibition by nitrogen starvation. In other feedings at 20 and 41.6 mg/L/day cases, the cell density was highest at feeding #4, following #3 and #1, which correlates with the ammonia gas input amount, meaning that the ammonia gas input has not been inhibitory to the cell growth. Unlike when the ammonia gas was fed continuously in Section 3.3., the cell density was highest at 67.3 mg/L/day ammonia concentration.

During the culture, the color change of each culture condition was also recorded as a preliminary screening of the astaxanthin production. As shown in Table 1, the strategy in

Column 2 of the 32.4 and 41.6 mg/L/day cases result in the color change on day 2 due to the initial nitrogen starvation which induced the carotenogenesis. The red color was maintained for the rest of culture at 32.4 mg/L/day case, while returning to green on day 5 and then brown on day 6 at 41.6 mg/L/day case. Other feeding strategies at 32.4 and 41.6 mg/L/day cases also resulted in the color change, but at a much later stage. When 67.3 mg/L/day ammonia containing gas was aerated into the reactor, however, all the cultures remained green, regardless of the different feeding strategies.

The astaxanthin contents obtained at 41.6 mg/L/day loading rate was further quantified.

Compared to the continuous supply of the ammonia gas (Section 3.3, Figure 3d), the astaxanthin accumulation through the intermittent ammonia gas feeding resulted in a mixed results, the

84 feeding strategies #2 and #4 resulted in the higher astaxanthin content, but the strategies #1 and

#3 are lower. Overall, the above results shows that intermittent ammonia feedings does not create a significant improvement of the astaxanthin content, but instead, the cell growth was reduced compared to the continuous feeding. Therefore, other strategies for inducing the astaxanthin content need to be explored.

3.5. Induction of astaxanthin from vegetative cells grown in ammonia gas culture

In the astaxanthin induction test, the vegetative cells grown in 67.3 mg/L/day ammonia load rate (Table 1) were subjected to treatment by different chemicals. As ammonium is the most reduced nitrogen form and cannot accept electrons, it was hypothesized that addition of an electron acceptor, astaxanthin can be induced.

In this study, ferric ion is used as an electron acceptor and compared with C/N ratio, which is regarded as one of the important factors in carotenogenesis; and H2O2, which generates

ROSs similar to high irradiance effect.

Figure 6a shows cell density and astaxanthin content for each case. Among four cases of

C/N ratio, cells became lysed and pale over time when C/N ratio was above 1. Although not apparent in Figure 6a, cells tended to aggregate and form clumps in all H2O2 tests. Therefore, only control, C/N 1, and ferric ion flasks looked healthy and red from naked-eye observation.

Figure 6b shows the quantitative data of all the induction methods. Overall, the cell density was significantly higher than control except FeCl 3 0.5 mg/L ( p < 0.05). Among the induction methods, however, only sodium acetate (C/N 1) significantly increased astaxanthin content compared to control ( p < 0.05).

Conclusion

In conclusion, H. pluvialis can be utilized as an ammonia scrubber system and increase its feasibility by producing astaxanthin. Among different pH levels, pH 5-7 showed the optimal growth performance but pH 4 showed the highest astaxanthin content. Among different ammonia gas loading rate, 41.6 mg/L/day showed the best cell density at both pH 4 and 5. In the real implementation, C/N ratio 1 can be applied to further increase the astaxanthin content.

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Table 1. Ammonia gas input and cell color change at different ammonia gas feeding strategy mode. * indicates days of ammonia gas supplied and blank indicates no ammonia gas fed. G, B, O, R, RG, RB and GB indicates H. pluvialis cell color, each corresponds to green, brown, orange, red, reddish green, reddish brown, and greenish brown. Shaded cell means the column was harvested.

20 ppm (32.4 mg/L/day) 41 ppm (41.6 mg/L/day) 82 ppm (67.3 mg/L/day)

Column 1 2 3 4 1 2 3 4 1 2 3 4

day1 * G G * G * G * G G * G * G * G G * G * G

day2 G * R * G * G G * R * G * G G * G * G * G

day3 * G R * G * G * G RG * G * G * G G * G * G

day4 G * R G * G G * RG G * G G * G G * G

day5 * G R G * G * G G G * G * G G G * G

day6 O * R O O B * B * B * G G * G G G

day7 * R R * R R * B B * B * GB * G G * G G

day8 R * R * R R B B * B GB G * G * G G

day9 RB RB GB * G G * G G

day10 RB G G G G

day11 G G G G

(a) 1.2E+06 1.0E+06 8.0E+05 pH 4 pH 5 6.0E+05 pH 6 4.0E+05 pH 7 2.0E+05 pH 8 Cell densityCell (cells/ml) 0.0E+00 0 1 2 3 4 5 6 7 8 9 10 11 12 day

1.2E+06 0.5 (b) cell density specific growth rate 1.0E+06 0.4 8.0E+05 0.3 -1 6.0E+05 0.2 day 4.0E+05 2.0E+05 0.1 Cell densityCell (cells/ml) 0.0E+00 0 4 5 6 7 8 pH

Figure 1. pH effect on Haematococcus pluvialis : cell growth (a), cell density and specific growth rate (b) and astaxanthin content of H. pluvialis culture (c). Medium contained both nitrate (1mM) and ammonium (0.5mM).

(a) 5.00E+06 4.00E+06

3.00E+06 urea 2.00E+06 NH4+ NO3-1 1.00E+06 Cell densityCell (cells/ml) 0.00E+00 0 1 2 3 4 5 6 7 8 9 10111213141516171819 day

(b) 5.00E+06 0.4 cell density specific growth rate 4.00E+06 0.3 3.00E+06 -1 0.2 2.00E+06 day 0.1 1.00E+06 Cell densityCell (cells/ml) 0.00E+00 0 NH4+ urea NO3-

(mg/g DW) (mg/g 1 Astaxanthin content content Astaxanthin 0 NH4+ urea NO3-

Figure 2. Cell growth (a), cell density and specific growth rate (b) and astaxanthin content of H. pluvialis culture (c) with ammonium, urea, and nitrate being used as the sole nitrogen source (with 1.5mM N equivalent) in MES-volvox medium. Initial medium pH was 7. Ammonium + (NH 4 ) column astaxanthin was not measured because the color remained pure green.

32.4 mg/L/day (pH4) (b) 32.4 mg/L/day (pH5) (a) 41.6 mg/L/day (pH4) 41.6 mg/L/day (pH5) 67.3 mg/L/day (pH4) 67.3 mg/L/day (pH5) 134.8 mg/L/day (pH4) 134.8 mg/L/day (pH5) 1.60E+06 1.40E+06

1.40E+06 1.20E+06 1.20E+06 1.00E+06 1.00E+06 8.00E+05 8.00E+05 6.00E+05 6.00E+05 4.00E+05

4.00E+05 Cell density (cells/ml) Cell density (cells/ml) 2.00E+05 2.00E+05 0.00E+00 0.00E+00 0 2 4 6 8 101214 0 2 4 6 8 10121416 Time (day) Time (day)

1.60E+06 0.6 (c) cell density (d) 4 1.40E+06 specific growth rate 0.5 3.5 1.20E+06 3 0.4 1.00E+06 2.5 8.00E+05 0.3 2 day-1 6.00E+05 1.5 0.2

Cell density (cells/ml) 4.00E+05 1 0.1 2.00E+05 0.5 Astaxanthin content (mg/g DW) 0.00E+00 0 0

Figure 3. Cell growth curve (a and b), cell density and specific growth rate (c) and astaxanthin content of H. pluvialis culture (d) with ammonia gas being used as the sole nitrogen source in MES-volvox medium. For each ammonia gas concentration, medium pH was adjusted at 4 and 5. No astaxanthin was determined at 134.8 mg/L/day (164 ppm) as the death of the cells.

(a) 32.4 mg/L/day (1) 32.4 mg/L/day (2) (b) 41.6 mg/L/day (1) 41.6 mg/L/day (2) 32.4 mg/L/day (3) 32.4 mg/L/day (4) 41.6 mg/L/day (3) 41.6 mg/L/day (4) 4.00E+05 8.00E+05

3.50E+05 7.00E+05

3.00E+05 6.00E+05

2.50E+05 5.00E+05

2.00E+05 4.00E+05 3.00E+05 1.50E+05

Cell density (cells/ml) 2.00E+05 1.00E+05 Cell density (cells/ml) 1.00E+05 5.00E+04 0.00E+00 0.00E+00 012345678910 0 1 2 3 4 5 6 7 8 Time (day) Time (day)

(c) 67.3 mg/L/day (1) 67.3 mg/L/day (2) 67.3 mg/L/day (3) 67.3 mg/L/day (4) 1.80E+06 1.60E+06 1.40E+06 1.20E+06 1.00E+06 8.00E+05 6.00E+05

Cell density (cells/ml) 4.00E+05 2.00E+05 0.00E+00 0 1 2 3 4 5 6 7 8 9101112 Time (day)

Figure 4. Growth curves of cells grown under different ammonia gas feeding strategies. Numbers in parenthesis indicates the column numbers corresponding to Table 1.

1.40E+06 0.6 cell density specific growth rate 1.20E+06 0.5 1.00E+06 0.4 8.00E+05 -1 0.3 6.00E+05 day 0.2 4.00E+05 Cell densityCell (cells/ml) 2.00E+05 0.1

0.00E+00 0

2.5

1.5

0.5 Aastaxanthin content (mg/g content DW) Aastaxanthin(mg/g 0 41.6 mg/L/day (1) 41.6 mg/L/day (2) 41.6 mg/L/day (3) 41.6 mg/L/day (4)

Figure 5. Cell density and specific growth rate (a) and astaxanthin content of H. pluvialis culture (b) with different ammonia gas feeding strategies. Numbers in parenthesis indicates the column numbers corresponding to Table 1.

(a) 6

2 Cell densityCell (g/L)

0 control C/N 1 H2O2 H2O2 H2O2 FeCl3 FeCl3 FeCl3 FeCl3 0.05 0.1 0.2 0.1 0.2 0.5 1

(b) 4 3.5

2.5

1.5

0.5 Astaxanthin content (mg/g content DW) Astaxanthin(mg/g 0 control C/N 1 H2O2 H2O2 H2O2 FeCl3 FeCl3 FeCl3 FeCl3 0.05 0.1 0.2 0.1 0.2 0.5 1

Figure 6. Cell density (a) and astaxanthin content (b) at different induction methods. The cell density was measured by weight at the end of the culture day.

CHAPTER 5. PHOSPHORUS REMOVAL DYNAMICS IN REVOLVING

MICROALGAL BIOFILM SYSTEMS TREATING WASTEWATER

Juhyon Kang and Zhiyou Wen

Abstract

Phosphorus removal in wastewater is important for water quality and environment protection. Microalgae have been suggested as an alternative advanced phosphorus treatment method for its simple, effective, and economic traits. In order to achieve phosphorus removal down to untraceable level, it is important to grow microalgae in an efficient cultivation system and to operate in optimal condition by understanding microalgal phosphorus removal kinetics.

To address these needs, microalgae was grown in revolving algal biofilm (RAB) systems with a wide range of initial phosphorus concentrations (synthetic BBM medium, wastewater streams A,

B, and C) using batch mode. The RAB system quickly removed phosphorus even at the highest phosphorus concentration (84.7 mg/L-TP) and produced high cell biomass. The phosphorus uptake kinetics was demonstrated by analyzing bound-P, acid-soluble P (ASP), and acid- insoluble P (AISP), and phosphorus fate was elucidated by calculating mass balance. In all types of wastewater, luxury phosphorus uptake was confirmed and the majority of liquid phosphorus was migrated into bound-P. Stream A was further investigated for continuous culture by differentiating harvest frequency and wastewater replacement rate. Higher cell density and faster phosphorus removal was observed in 5-day continuous culture than 2-day continuous culture

(p<0.0001) and it was correlated to the highest ASP on day 2 and lowest ASP on day 5 in batch culture of Stream A. Therefore, the research shows the importance of considering microalgal phosphorus removal kinetics in designing RAB wastewater treatment operation.

Keywords: wastewater treatment, phosphorus kinetics, mass balance, revolving algal biofilm reactors

1. Introduction

Phosphorus is one of the main targets in wastewater treatment concerning for eutrophication and water quality. Over-enrichment of phosphorus in receiving water, eutrophicaiton causes excessive growth of algae and cyanobacteria that forms wide layers on water surface. The high microorganism population causes low dissolved oxygen and the wide layers block light penetration that are necessary for other aquatic organisms (Correll, 1998;

Alonso-Rodrı́guez and Páez-Osuna, 2003). The excessive growth of microorganisms can lead to death of aquatic animals and toxic algal blooms, and cause odor problems that impairs quality of water use for domestic and industrial water supply and recreation (Jones and Lee, 1982; Sharpley,

1999).

Chemical dosage has been widely used to remove phosphorus in most wastewater treatment plant due to its simplicity (EPA, 2007), however, the increasing public attention to environment and the reinforced phosphorus regulations (Sharpley et al., 2003) require advanced phosphorus removal processes that can remove phosphorus down to untraceable level, such as membrane technology, electrodialysis, enhanced biological phosphorus removal (Zeng et al.,

2003; Sonune and Ghate, 2004; Mahmoud and Hoadley, 2012). These advanced methods show

99 excellent phosphorus removal efficiency but are expensive and complicated to operate and have issues of phosphorus recovery (Powell et al., 2008; Sengupta and Pandit, 2011). In this context, microalgae have been proposed as a simple, effective, and economic phosphorus removal agency

(Hoffmann, 1998; Wang et al., 2010; Park et al., 2011).

Employing microalgae to prevent eutrophication can be an irony but the well-controlled microalgal cultivation system can function better than any other advanced methods. The key for successful employment of microalgal phosphorus treatment is first, understanding microalgal phosphorus uptake mechanism and second, developing a sustainable and efficient microalgae cultivation system. Evolutionally, microalgae have developed a series of acclimation responses to phosphorus limitation, such as reduced photosynthesis, cell division and growth (Moseley and

Grossman, 2009; Tripathi et al., 2012), synthesis of phosphatase that has wide substrate specificity and synthesis of transporters that has high affinity to phosphate (Healey and Hendzel,

1975; Moseley and Grossman, 2009), and ‘overcompensation’ of phosphorus after starvation or

‘luxury uptake’ of phosphorus for provision (Kulaev and Vagabov, 1983). By using their developed ability to uptake more phosphorus than their need, phosphorus removal can be maximized.

Microalgal wastewater treatment system have been developed since 1950s (Oswald et al.,

1957) in different forms and can be classified as suspended and non-suspended culture.

Suspended culture includes raceway oxidation ponds (Oswald, 1988; Shen et al., 2009) or photobioreactors (López et al., 2006; Ruiz et al., 2011) whereas non-suspended culture includes immobilized polymers (de-Bashan and Bashan, 2010) or biofilm (Boelee et al., 2011; Posadas et al., 2013; Shi et al., 2014). Although each system has its own pros and cons, in general, non- suspended culture has advantages over suspended culture such as steep nutrient gradients and

100 high exchange rates (Hoffman, 1998), short hydraulic retention time, lower energy for mixing the broth, and easier and simpler harvest (Boelee, 2012).

Our lab has developed a unique vertical revolving algal biofilm (RAB) reactor based on a trough reservoir (Gross and Wen, 2014; Gross et al., 2015b). On top of the general advantages of non-suspended culture, our vertical RAB reactor can minimize land usage, which has been the biggest hindrance of biofilm implementation (Boelee, 2012), while maximize capture of sunlight and CO 2 because of its vertically upright configuration. By studying the phosphorus uptake dynamics in a wide range of initial phosphorus concentration in RAB system, we can learn cellular growth activities and use the information to optimize the microalgal wastewater treatment system.

2. Materials and methods

2.1. Algae cell strain and preparation

The microalgae Chlorella vulgaris (UTEX #265) was used as the initial seed culture for the revolving algal biofilm (RAB) due to its good attached growth performances (Gross et al.,

2013; Gross and Wen, 2014). To prepare the seed culture, the strain preserved on agar slant at

4°C was transferred to 250 mL Erlenmeyer flask containing 50 mL medium. The cells were grown in Bold’s Basal medium. The medium was autoclaved at 121 °C for 15 min before use.

The flasks were incubated at a rotatory shaker at 25 oC under continuous illumination (110 µmol s-1 m-2). The cells were stepwise transferred to a 20 L flat panel reactors. The exponential cells at the flat panel reactor was used to inoculate the revolving biofilm reactor (RAB) systems.

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2.2. RAB configuration and operation

The cell suspension in the flat panel reactor was added to the liquid reservoir of RAB systems for cells to attach on the surface of the attachment materials in the RAB systems. The detailed RAB reactor configuration has been described previously (Gross et al., 2015b). Briefly, a cotton duct belt spanning around shafts revolved continuously by motors at a linear velocity of

4cm∙sec -1. One shaft was submerged into a trough shape liquid reservoir and the other shaft was uplifted by a column so that the belt rotates vertically and the cells could alternatively rotate between liquid and gas phases. The surface area of the belt was 0.1-0.12 m 2 and liquid volume in the trough reservoir was 1.5 L. All the RAB systems were continuously illuminated at 110 m mol∙s -1m-2 at 25°C.

To establish the initial biofilm on the RAB system, the RAB system was run with distilled water to rinse any impurities on the belt for a day. After cleaning, the RAB system was inoculated with the cell suspension from the flat panel reactor and run for 10 days to establish a thick biofilm on the belt. During these 10 days operation, the cell suspension was periodically supplemented to the liquid reservoir to compensate water evaporation in the trough reservoir.

After 10 days of incubation, a thick biofilm was established at the attachment materials, then, this biofilm was harvested by scraping with a rigid plastic pan. The remaining cell colonies on the attachment materials surface served as a seed for the next growth cycle (re-growth). After additional two re-growth cycles (with the BBM medium), the biofilm was firmly stabilized. Then, the RAB systems were started for testing their nutrient removal capabilities by feeding the different streams as described as follows.

First, the artificial BBM medium with different phosphorus concentration was fed to the trough reservoir of the RAB systems (Table 1). The nitrogen of the medium was kept constant.

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The different N/P ratios of the influents were also presented in Table 1. For each case, the RAB system was rinsed with distilled water before running with the medium so that the interference from phosphorus soaked in the belt could be minimized. The biofilm was harvested from a certain surface area on daily basis. To compensate the liquid loss due to evaporation, the trough was supplemented with distilled water daily to make sure the liquid level in the tough was unchanged.

Three different wastewater streams were also tested including Supernatant from gravity thickening obtained from Chicago municipal wastewater (Stream A), Post digestion centrate obtained from Chicago municipal wastewater (Stream B), and effluent from a fermentation- based manufacturing plant (Stream C). The detailed characteristics of the streams are shown in

Table 2. All three streams were tested in batch and stream A was further tested in continuous mode. Similar to the test in the BBM medium, the RAB system was rinsed with distilled water to remove the interference of the residue nutrients soaked in the attachment materials. In batch culture, the wastewater was added to the trough, and the RAB systems were ran until the cells reach to the stationary phase. During the operation, distilled water was added to the trough daily to compensate the water evaporation. Cells were harvested and liquids were collected daily. In continuous culture, the liquid in the trough was replaced every two or five day at a certain percentage (25%, 50%, 75% and 100%) and the biomass was also harvested on the same time when liquid was replaced. During the two or five days, distilled water was added to make up for the evaporation. Stream A was used as an influent. The continuous operation was run until the cell biomass reach to the steady state.

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2.3. Analytical methods

Cellular phosphorus and liquid phosphorus were measured to determine the fate of the phosphorus during the operation of the RAB systems. The cellular phosphorus includes bound phosphorus, intracellular acid-soluble polyphosphate (ASP), and intracellular acid-insoluble polyphosphate. The method was modified from Aitchison and Butt (1973) and Kanai et al.

(1965). In summary, ~200 mg of freeze dried algae biomass was washed with 20 ml distilled water and centrifuged. This step was repeated twice and the combined supernatant was used to measure bound phosphorus (mg/L). The rinsed cell pellet was further extracted with 10 ml cold

10% trichloroacetic acid, sitting for 10 min at -20 °C and centrifuged. This step was repeated twice and the combined supernatant was neutralized with 3M KOH and measured as intracellular acid-soluble polyphosphate (ASP, mg/L). The ASP-extracted biomass residue was further extracted with 15 ml 2N KOH, sitting for 10 min at room temperature and centrifuged. This step was also repeated twice and the combined supernatant was neutralized with HCl and measured as acid-insoluble polyphosphate (AISP, mg/L). All the supernatants and liquid phosphorus were sampled and subjected for their total phosphorus measurement using a Hach total phosphorus kit

(TNT Reagent Set, Molybdovanadate method) with a spectrophotometer (Hach model DR 3900).

The phosphorus concentrations of the cellular phosphorus supernatants were then converted into biomass phosphorus content. The total nitrogen (Hach total nitrogen reagent set, persulfate digestion Test ‘N Tube method) and nitrate (Hach Nitrate TNT plus LR kit) were also measured using the same spectrophotometer and ammonia was measured by electrode (model 9512

HPBNWP Thermo Scientific, Beverly, MA) according to EPA-approved standard methods

4500-NH 3 E.

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2.4. Statistical analysis

The significant difference between data was analyzed by One-way analysis of variance

(ANOVA) at 95% confidential level using PROC GLM, with statistical package SAS (version

9.4 SAS Institute, Inc., Cary, NC, USA).

3. Result and discussions

3.1. Characterization of the different wastewater streams

As a benchmark of cell growth performance in the RAB system, synthetic BBM medium was firstly used for growing cells. N/P ratio has been one of the important parameters for microalgae growth (Rhee and Gotham, 1980; Sterner and Elser, 2002; Klausmeier et al., 2004) but the optimal N/P ratio in literature has varied widely depending on the growth conditions. In this study, the different N/P ratio for the RAB algae growth was investigated by setting the initial nitrogen concentration as the same level as that in the regular BBM medium while varying the initial phosphorus concentration (Table 1). It has been reported that the N/P ratio of secondary effluent in wastewater treatment process is about 4 (Ruiz et al., 2011) so the range of this study covers the real wastewater.

Table 2 shows features of three different wastewater streams. Stream A and B are municipal wastewater with low and high nutrient content while stream C is low in phosphorus but high in nitrogen. The wastewater was characterized in accordance with the parameters used by wastewater treatment plant. In Table 2, original TP measures total phosphorus of the wastewater containing all buoyant while filtered SP measures soluble phosphorus of the wastewater after filtering with 45 m m membrane filter. In theory, SP is usually lower than TP because many phosphorus compounds are not soluble in water (APHA, 2012). From the result, it

105 was confirmed that centrifuged TP was not much deviated from SP and total nitrogen alone can be a good parameter for nitrogen. Therefore, centrifuged TP and total nitrogen were used to characterize nutrient removal for the real wastewater study.

3.2. Cell growth of different wastewater streams

As shown in Figure 1a, cells grew better in higher initial phosphorus conditions (122.6,

53.6, and 12.4 mg/L) and the cell density at stationary phase was significantly higher than lower initial phosphorus conditions ( p<0.05). Similar results were shown in real wastewater (Figures

1b, 1c, and 1d). The cell density at stationary phase was higher in stream B than stream A and C

(p<0.05, F=590.35). Despite of ten times higher initial phosphorus, stream A had a slightly higher cell density than stream C ( p<0.05, F=4.99) probably due to ten times higher initial nitrogen in stream C.

In order to simulate a real wastewater treatment, running RAB system in continuous mode is important. In this paper, stream A was used to study the effect of harvest frequency and wastewater replacement rate. Figure 2a and 2b show 2-day harvest frequency and 5-day harvest frequency cell growth and nutrient removal, respectively. Both 2-day and 5-day harvest frequency were tested for 25, 50, 75, and 100% wastewater replacement rate. For example, 100% replacement rate means complete substitution of remaining liquid reservoir by wastewater.

Within 2-day harvest frequency, the cell density after day 12 was highest at 100% replacement rate, following 75, 25, and 50% ( p<0.05), which is proximately in the order of the replaced wastewater amount (Figure 1a). On the other hand, 5-day harvest frequency after day 30 showed close cell growth among all replacement rate ( p>0.05) and the cell density was higher

106 than 2-day harvest frequency ( p<0.0001) (Figure 1b), probably due to a longer solid retention time and wastewater retention time.

3.3. Nutrient removal of different wastewater streams

As shown in Figure 3a, the liquid phosphorus concentration of synthetic BBM medium is interesting because the liquid phosphorus removal was not effective in overall, probably due to the soaked phosphorus in the belt during the inoculation (Table 3). From this observation, the

RAB system trough was compensated with distilled water instead of cell suspension during inoculation process for real wastewater experiment. Figures 3b, 3c, and 3d show that liquid nutrient removal of wastewater was almost untraceable at the end of the treatment. The phosphorus and nitrogen removal efficiencies are summarized in Table 3. In all cases, nitrogen removal was quicker than phosphorus removal.

Liquid nutrient concentration is shown in Figure 4. In 2-day harvest frequency, 75 and

100% replacement rate showed immediate phosphorus removal, while 25 and 50% replacement rate took an extended length of time to remove phosphorus down to untraceable level (Figure 4a and 4b). In contrast, the phosphorus and nitrogen removal in 5-day harvest frequency was almost complete in earlier stage for all replacement rate ( p>0.05) (Figure 4c and 4d).

The result is comparable to other microalgal wastewater treatment systems. For example,

Ruiz et al. (2011) treated wastewater with different initial phosphorus (1.3-143.5 mg/L-P) and nitrogen (5.8-226.8 mg/L-N) using photobioreactors. When initial phosphorus was higher than

9.7 mg/L and initial nitrogen was higher than 48.2 mg/L, more than half of the initial substrate was still remained by day 9. Also, the cell growth was not significantly faster in higher initial phosphorus concentration. Another example, Wang et al. (2010) studied microalgal wastewater

107 treatment in flask culture. The microalgae could remove phosphorus down to untraceable level when the initial phosphorus was lower than 6.86 mg/L. However, when the initial phosphorus was 201.5 mg/L, about 25 mg/L of phosphorus was still left by day 9. The nitrogen removal was incomplete in all cases. Although cell growth was superior in higher nutrient wastewater, the faster cell growth alone was not sufficient for efficient nutrient removal. Powell et al. (2009) treated wastewater using algal stabilization pond. The phosphorus was removed completely when the initial phosphorus concentration was 5 and 15 mg/L-P, but significant amount of phosphorus still persisted after 24 days when the initial phosphorus was 30 mg/L-P. Biofilm- based wastewater has been studied mostly at low initial concentration (1.5-15 mg/L-P, 25-100 mg/L-N) and showed quick and efficient nutrient removal (Boelee et al., 2011; Posadas et al.,

2013; Shi et al., 2014).

The result implies positive application of RAB system in wastewater treatment because even the higher initial nutrient shows quick removal and higher cell density instead of slow removal and dragged cell growth. Enhancing growth factors for optimal cell growth and subsequent nutrient removal has been suggested, such as high light and temperature (Powell et al., 2009) and optimizing N/P ratio (Wang et al., 2010). However, considering that adjusting growth conditions in situ is unrealistic for real application, employing RAB system is a more suitable solution.

Considering real application of RAB system in the wastewater treatment plant, 5-day harvest frequency for Stream A will be adequate because it requires less labor and shows higher biomass production and nutrient removal performance. However, in places where large amount of wastewater should be treated, harvest frequency should be tested separately in regards of cellular nutrient uptake rate.

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3.4. Dynamics of luxury P uptake

Microalgal capability to uptake more phosphorus than the amount necessary for metabolism, so called luxury P uptake, has been recognized and quantified as different forms of intracellular polyphosphate (Baker and Schmidt, 1963; Miyachi et al., 1964; Kanai et al., 1965;

Aitchison and Butt, 1973). Luxury uptake is similar to overcompensation in that phosphate is excessively synthesized up to 20% of dry weight but is different because overcompensation occurs after depletion of phosphate while luxury uptake occurs in normal metabolism without previous phosphorus starvation (Fuhs and Chen, 1975; Kulaev and Vagabov, 1983; Eixler et al.,

2006). Microalgae assimilate phosphorus in the form of orthophosphate and accumulate phosphorus in the form of polyphosphate (Correll, 1998; Eixler et al., 2006). Typically, intracellular polyphosphates can be classified as acid-soluble polyphosphate (ASP) and acid- insoluble polyphosphate (AISP) according to their metabolic roles and extraction reagents

(Baker and Schmidt, 1963; Miyachi et al., 1964; Powell et al., 2008; Powell et al., 2009). In general, ASP measures phosphate that are actively involved in metabolism, such as orthophosphate and polyphosphate intermediates while AISP measures polyphosphate that are reserved for the case of phosphorus limitation (Miyachi et al., 1964; Kulaev and Vagabov, 1983;

Powell et al., 2009). In addition to these two types of phosphate, the algal biofilm also contains significant amount of bound P, which has been largely neglected in literature because significant amount of phosphorus would be attached on extracellular polymeric substance (EPS), a unique compounds in the biofilm systems (Tian et al., 2006; Gross et al., 2015a). By studying bound P,

ASP, and AISP, phosphorus migration flux and cellular metabolic activity can be illustrated.

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As shown in Figure 5a, AISP was constant in all phosphorus concentrations, implying that cells maintain their AISP and they were not under phosphorus limitation. The bound P was increasing in 122.6 and 53.6 mg/L-P while decreasing in 12.4 mg/L-P. Considering that bound P measures extracellular phosphorus, it is reasonable to see a decreasing bound P at lower initial phosphorus concentration. The initial peak of bound P on day 1 for 12.4 mg/L-P might be caused from soaked phosphorus on belt (Figure 5a).

The bound P, ASP, and AISP (mg P/mg dry weight) of the biomass obtained from different wastewater streams were compared (Figures 5b, 5c, and 5d). While bound P and ASP were in the order of Stream B, A, and C (p<0.0001), AISP was similar in Stream A and C, following Stream B ( p=0.0395). Similarly, Powell et al. (2009) also reported a higher ASP and

AISP of biomass when growing in medium containing 15 and 30 mg/L-P than that in 5 mg/L-P.

The authors concluded that high phosphate concentration can trigger luxury uptake. However, it should be noted that in their study, 15 and 30 mg/L-P showed similar ASP and AISP levels.

Similarly, even Stream B had higher ASP and AISP content in biomass, the numbers were not notably higher than other streams considering that Stream B contained more than 10 times of phosphorus. This observation leads to a conclusion that cells have upper limit on luxury phosphorus uptake and removing high phosphorus concentration using luxury uptake should be accompanied with high cell density. Aitchison and Butt (1973) also recognized that cells with high phosphate did not accumulate polyphosphate when the medium phosphate concentration was increased. In this regard, phosphorus mass balance for each stream will be discussed in next section.

In Stream A and C, ASP and AISP (mg P/mg dry weight) initially were highest and decreased then bounced back to a higher level (Figures 5b and 5d). The initial crest implies

110 luxury phosphorus uptake. The initial peak, decrease, and rebounding ASP and AISP has been observed in previous studies (Baker and Schmidt, 1963; Aitchison and Butt, 1973) but Powell et al. (2009) did not see this trend because they analyzed ASP and AISP infrequently, every four days. On the other hand, Stream B showed almost consistent ASP and AISP (mg P/mg dry weight) throughout the test. The consistently lowest AISP in Stream B supports that cells accumulate AISP in case of phosphorus limitation and further suggests that AISP accumulation is triggered by lower initial phosphorus concentration. Tripathi et al. (2012) also found that AISP increases when the cells are under phosphorus deficiency.

From the dynamics of ASP, harvest time in continuous mode can be adjusted to optimally utilize luxury uptake. For example, Stream A showed peak ASP on day 2 and lowest ASP on day

5 in batch culture (Figure 5b). This means that cells were actively growing and accumulating biomass on day 2 while finishing active growth stage on day 5. Therefore, if harvested every 2 days, cells could not consume all the phosphorus for growth and had to adjust to a new environment before reaching to their steady state. This interpretation can be confirmed by the higher cell density and quick phosphorus removal in 5-day continuous culture than 2-day continuous culture ( p<0.0001) (Figures 2 and 4). However, different parameters can be considered for Stream B because of consistently high ASP (Figure 5c) and quick liquid phosphorus removal (Figure 3c). Examples of consideration includes liquid nutrient removal performance or other on-site factors.

As shown in Figure 6, in continuous culture, 2-day harvest frequency had similar ASP and AISP content ( p>0.05) and had higher bound P than 5-day harvest frequency ( p<0.05). In all cases, AISP was higher than ASP, implying low cellular metabolic activities probably because the samples were collected at the end of the treatment process, at steady-state. In 25%

111 replacement rate of 5-day harvest frequency, AISP kept increasing and bound P and ASP were decreasing, indicating that cells were close to starvation.

3.5. P content in biomass

The phosphorus content in biomass, sum of bound P, ASP, and AISP, is shown in Figures

7 and 8. Considering that the typical microalgae biomass found in freshwater contains about 1-

1.3 % phosphorus (Kaplan et al., 1986; Oswald, 1988), the data in this study has quite higher range probably due to combined effect of the adsorbed bound-P and also luxuriously absorbed intracellular phosphorus. Except cells grown in BBM medium where significant amount of phosphorus still was untreated by the end of the course (Figure 7a), cellular phosphorus content

(%P of biomass) was in the order of the initial phosphorus concentration, i.e., highest at Stream

B, following Stream A and C ( p<0.0001). Similarly, Powell et al. (2008) showed higher phosphorus percentage in biomass at higher phosphate concentration. Also, the phosphorus content in biomass was increasing in Stream B (Figure 7c), meaning abundant phosphorus, but was decreasing in stream A and C (Figure 7b and 7d), meaning phosphorus limitation. Similarly,

Baker and Schmidt (1963) found that when excessive nutrient was provided, total cellular phosphorus was linearly increased. As shown in Figure 8, in continuous culture, 2-day harvest frequency had higher phosphorus content in biomass, corresponding to an efficient phosphorus removal in 5-day harvest frequency.

3.6. P mass balance

In order to demonstrate the fate of phosphorus in the RAB culture, mass balance was analyzed as follows.

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Ptotal (mg) = P-bound (mg) + ASP (mg) +AISP (mg) + P liquid (mg)

Phosphorus mass balance is summarized in Table 4 and 5 and visualized in Figures 7 and 8. As shown in Table 4 and 5, P total (mg) is well conserved in different days.

In characterizing phosphorus in different forms, different units can be used such as phosphorus content in biomass as each different forms (mg P/g DW) (Figures 5 and 6) and portions of each phosphorus forms in total phosphorus (%) (Figures 7 and 8). Although previous literature mostly presented data based on mg P/g DW (Miyachi et al., 1964; Kanai et al., 1965;

Aitchison and Butt, 1973; Healey and Hendzel, 1975; Eixler et al., 2006; Powell et al., 2009;

Tripathi et al., 2012), seeing phosphorus distribution among total phosphorus (%) gives an overall picture relates to the phosphorus mass balance.

In synthetic BBM medium culture (Figure 7a), portions of all phosphorus forms were increasing, meaning active cell adsorption and absoprtion in all cases, while liquid phosphorus portions were decreasing. As shown in Figures 7b, 7c, and 7d, ASP and AISP portion in Stream

A and C were relatively constant while ASP in Stream B was increasing. ASP portion in total phosphorus was highest in Stream B ( p<0.05) and similar in Streams A and C ( p>0.05), while

AISP portion was in the order of Stream C, A, and B ( p<0.0001). Therefore, the portions of ASP and AISP corresponds to the cellular metabolic activities and initial phosphorus concentrations.

In Stream B, the majority of phosphorus in liquid migrated to bound P and the portions of

ASP and AISP were almost consistent, meaning that extracellular adsorbed phosphorus plays an important roles in phosphorus removal under nutritious waste streams. In Stream A, liquid phosphorus still persisted, while Stream B and C showed quick liquid phosphorus removal. This is probably because Stream A had low cell biomass to treat liquid phosphorus while Stream B had good cell growth and Stream C had very low initial phosphorus concentration (1.6 mg/L-P).

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In continuous culture, bound P took the majority of portions in both 2-day and 5-day harvest frequency except 25% in 5-day (Figure 8). Except 25% in 5-day, ASP portion in 2-day harvest frequency was lower than that of 5-day harvest frequency ( p<0.001) while AISP portion was not significantly different ( p>0.05). This is in agreement with the higher cell density and quick phosphorus removal in 5-day continuous culture.

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Conclusion

In conclusion, vertical RAB system excels other microalgal wastewater treatment systems in cell growth and liquid nutrient removal. By studying wide range of initial phosphorus and nitrogen concentrations, microalgal phosphorus removal dynamics and mass balance in the

RAB system were demonstrated. Phosphorus content in biomass confirmed luxury uptake of phosphorus and cellular phosphorus distribution showed the kinetics of luxury phosphorus uptake. In phosphorus mass balance, phosphorus was conserved throughout the experiments and the majority of phosphorus in liquid migrated to bound-P. From continuous culture of Stream A, it was suggested to run 5-day harvest frequency in real application.

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Table 1 . Phosphorus and nitrogen concentration of BBM medium with modified phosphorus concentrations.

Reactors Total Nitrogen (mg/L-N) Total phosphorus (mg/L-P) N/P 1 41.18 122.6 0.34 2 41.18 53.6 0.77 3 41.18 12.4 3.33 4 41.18 7.19 5.73 5 41.18 3.25 12.7

Table 2 . Phosphorus and nitrogen content of different wastewater streams. Stream A:

Supernatant from gravity thickening obtained from municipal wastewater; Stream B: Post digestion centrate obtained from municipal wastewater, and Stream C: An effluent from a fermentation-based manufacturing plant.

Total Original TP a Centrifuged Filtered SP b Ammonia Nitrate Stream nitrogen (mg/L) TP (mg/L) (mg/L) (mg/L-N) (mg/L-N) (mg/L-N) A 7.0 ± 1.99 3.2 ± 0.64 3.1 ± 0.60 15.7 ± 6.46 14.2 ± 0.88 0.3 ± 0.07 B 84.7 ± 14.4 53.8 ± 1.8 45.6 ± 5.52 550 ± 26.5 541.8 ± 23.6 0.5 ± 0.00 C 0.63 0 0 169 126.8 2.03 a. TP, total phosphorus, including suspended matter, organic phosphorus, and reactive phosphorus (APHA, 2012) b. SP, soluble phosphorus is the phosphorus after filtering with 45 m m membrane filter.

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Table 3. Nutrient removal efficiency at the end of the treatment in batch culture

P removal efficiency (%) N removal efficiency (%)

122.6 mg/L 67.5 n/a 53.6 mg/L 47.4 n/a BBM medium 12.4 mg/L 61.6 n/a 7.19 mg/L 78.1 n/a 3.25 mg/L 95.4 n/a Stream A 71.6 94.9 Stream B 100 100 Stream C 100 96.7 * n/a: not available

Table 4. Phosphorus mass balance in batch culture

Bound P ASP AISP Liquid P P total day (mg) (mg) (mg) (mg) (mg) 1 4.75 2.55 1.69 106.3 115.3 122.6 3 11.1 9.36 3.15 75.7 99.3 mg/L 7 21.1 8.63 6.55 49.9 86.2 1 5.35 3.36 3.13 59.7 71.6 BBM 53.6 3 6.81 5.94 3.79 45.3 61.9 medium mg/L 7 9.19 7.05 4.38 40.5 61.1 1 5.26 3.00 1.96 47.8 58.0 12.4 3 7.51 4.24 3.09 28.7 43.6 mg/L 7 18.9 7.16 6.74 25.8 58.6 2 2.84 3.29 2.68 2.29 11.1 4 4.77 4.02 1.34 2.28 12.4 Stream A 5 9.48 3.22 2.00 1.41 16.1 6 9.11 3.91 2.54 1.67 17.2 7 7.55 4.38 2.66 1.24 15.8 2 8.62 7.31 1.78 42.1 59.8 4 18.7 14.4 2.49 17.7 53.2 7 41.5 27.0 1.91 0.29 70.7 Stream B 8 46.5 21.2 6.10 0 73.8 11 44.6 22.5 3.29 0 70.3 12 37.2 23.0 2.31 0 62.5 2 2.87 2.20 3.18 0 8.26 3 2.94 2.57 2.57 0 8.07 Stream C 4 4.00 1.23 1.83 0 7.06 5 4.96 2.46 2.68 0 10.1 7 3.19 2.24 2.73 0 8.15

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Table 5. Phosphorus mass balance in continuous culture

Harvest Replacement Bound P ASP AISP Liquid P P total day frequency rate (mg) (mg) (mg) (mg) (mg) 12 22.6 5.62 11.0 0.33 39.5 25% 16 20.8 3.56 13.0 0.18 37.5 20 17.8 2.93 7.79 0.15 28.7 12 11.1 7.26 11.5 0.62 30.5 50% 16 10.8 2.64 8.44 0.11 22.0 20 11.7 2.06 6.92 0.10 20.8 2-day 12 19.2 6.04 10.5 0.09 35.9 75% 16 16.4 2.59 6.74 0.21 25.9 20 26.3 4.59 11.4 0.09 42.4 12 30.5 3.49 10.7 0.18 44.8 100% 16 28.1 2.90 7.40 0.16 38.6 20 27.7 2.95 8.39 0.14 39.2 30 8.15 2.86 7.21 0.12 18.3 25% 40 11.6 5.85 26.4 0.05 43.9 50 4.0 1.43 22.3 0.13 27.9 30 42.0 9.23 18.4 0.29 69.9 50% 40 20.1 4.32 12.0 0.49 36.9 50 10.6 3.21 7.93 0.17 21.9 5-day 30 38.0 15.0 21.5 0.04 74.5 75% 40 31.6 6.45 13.2 0.19 51.4 50 34.8 6.73 14.8 0.08 56.4 30 32.9 5.36 13.0 0.16 51.3 100% 40 30.7 7.07 17.7 0.17 55.7 50 28.6 9.71 22.2 0.08 60.1

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20 8 122.6 mg/L 53.6 mg/L (b) 18 12.4 mg/L 7.19 mg/L 7 3.25 mg/L 16 6 14 (a) ) ) 2 2 5 12

10 4

8 3 cell density (g/m 6 Cell density (g/m 2 4

2 1

0 0 0 2 4 6 8 10 0 2 4 6 8 Time (day) Time (day)

30 7 (c) (d) 6 25

5 ) ) 2 20 2

4 15 3 Cell density (g/m 10 Cell density (g/m 2

5 1

0 0 0 1 2 3 4 5 6 7 8 91011121314 0 1 2 3 4 5 6 7 8 Time (day) Time (day)

Figure 1. Cell growth of different growth media in the batch culture of the RAB system, (a) synthetic BBM medium with different initial phosphorus concentration, (b) Stream A, (c) Stream B, and (d) Stream C.

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7 (a) 2d-25% 2d-50% 2d-75% 2d-100% 6 ) 2 5

3 Cell densityCell (g/m 2

0 2 4 6 8 10 12 14 16 18 20 Time (day)

14 (b) 12 5d-25% 5d-50% 5d-75% 5d-100% ) 2 10

4 Cell densityCell (g/m

0 5 10 15 20 25 30 35 40 45 50

Time (day)

Figure 2. Cell growth during the continuous culture feeding wastewater stream A (Table 1). Cell density of (a) every 2-day harvest frequency and (b) every 5-day harvest frequency.

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140 122.6 mg/L 18 (a) 53.6 mg/L (b) 120 12.4 mg/L 16 7.19 mg/L 3.25 mg/L 14 100 12 Total P 80 (mg/L-P) 10 Total N 60 8 (mg/L-N)

40 6 P concentration(mg/L)

P and N concentration(mg/L) 4 20 2 0 0 2 4 6 8 10 0 Time (day) 0 2 4 6 8 Time (day)

45 450 1.6 100 (c) (d) 40 400 1.4 90 35 350 80 1.2 70 30 Total P 300 Total P (mg/L-P) 1 60 25 250 Total N 0.8 Total N 50 20 (mg/L-N) 200 (mg/L-N) 0.6 40 15 150 30 P concentration(mg/L) P concentration(mg/L) N concentration(mg/L) N concentration(mg/L) 0.4 10 100 20 5 50 0.2 10 0 0 0 0 0 2 4 6 8 10 12 14 0 2 4 6 8 Time (day) Time (day)

Figure 3. Nutrient removal during batch culture of the RAB system. (a) Total phosphorus concentration in synthetic BBM media with different initial phosphorus concentration, (b) total phosphorus and nitrogen concentration in Stream A, (c) total phosphorus and nitrogen concentration in Stream B, and (d) total phosphorus and nitrogen concentration in Stream C.

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4 (a) 2d-25% 2d-50% 2d-75% 2d-100% 3

P concentration(mg/L) 0 0 2 4 6 8 10 12 14 16 18 20 Time (day) 15 (b) 2d-25% 2d-50% 2d-75% 2d-100%

5 N concentration(mg/L) 0 0 2 4 6 8Time 10 (day) 12 14 16 18 20 4 (c) 5d-25% 5d-50% 5d-75% 5d-100% 3

P concentration (mg/L-P) 0 0 5 10 15 20 25 30 35 40 45 50 Time (day) 20 (d) 5d-25% 5d-50% 5d-75% 5d-100% 15

N concentration(mg/-N) 0 0 5 10 15 20 25 30 35 40 45 50 Time (day)

Figure 4. Liquid nutrient concentration during the continuous culture feeding wastewater stream A (Table 2). (a) P and (b) N concentration on 2-day harvest frequency. (c) P and (b) N concentration on every 5-day harvest frequency.

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(a) 16 (b) 16

14 14

12 12

10 10 8 8 6 6 (mg (mg dryP/g weight)

4 (mg dryP/g weight) P distribution in biomass

P distribution in biomass 4 2 2 0 1 3 7 1 3 7 1 3 7 0 2 4 5 6 7 122.6 mg/L 53.6 mg/L 12.4 mg/L day P concentraiton (mg/L) Bound P ASP AISP Bound P ASP AISP

6 20 5

15 4

3 10 (mg (mg dryP/g weight) (mg (mg dryP/g weight) 2 P distribution in biomass P distribution in biomass 5 1

0 0 2 4 7 8 11 12 2 3 4 5 7 day day Bound P ASP AISP Bound P ASP AISP

Figure 5. Phosphorus mass distribution as different portions (bound P, ASP, and AISP) of batch culture in: (a) synthetic BBM media with different initial phosphorus concentrations, on day 1, 3, and 7 for the three highest phosphorus concentration cases, (b) Stream A, (c) Stream B, and (d) Stream C.

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(a) Bound P ASP AISP 12

(mg dry weight) (mg P/g 2 P in biomass distribution P

0 12 16 20 12 16 20 12 16 20 12 16 20 25% 50% 75% 100% Change of trough liquid

(b) Bound P ASP AISP 8 7 6 5 4 3

(mg dry weight) (mg P/g 2 P in biomass distribution P 1 0 30 40 50 30 40 50 30 40 50 30 40 50 25% 50% 75% 100% Change of trough liquid

Figure 6. Phosphorus mass distribution as different portions (bound P, ASP, and AISP) during continuous culture feeding wastewater stream A (Table 2). (a) Every 2-day harvest frequency and (b) 5-day harvest frequency.

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120 3 120 9 (a) (b) 100 2.5 8 100 7 80 2 80 6

60 1.5 5 60 4 %P oftotal P

40 1 %P ofbiomass %P oftotal P 40 3 %P ofbiomass 20 0.5 2 20 0 0 1 1 3 7 1 3 7 1 3 7 0 0 122.6 mg/L 53.6 mg/L 12.4 mg/L 2 4 5 6 7 P concentraiton (mg/L) day Liquid P (%) AISP (%) Liquid P (%) AISP (%) ASP (%) Bound P (%) ASP (%) Bound P (%) %P of biomass % P of biomass 120 10 120 4.5 (c) (d) 9 4 100 8 100 3.5 7 80 80 3 6 2.5 60 5 60 4 2 %P oftotal P %P ofbiomass %P oftotal P

40 %P ofbiomass 3 40 1.5

2 1 20 20 1 0.5

0 0 0 0 2 4 7 8 11 12 2 3 4 5 7 day day Liquid P (%) AISP (%) Liquid P (%) AISP (%) ASP (%) Bound P (%) ASP (%) Bound P (%) %P of biomass % P of biomass

Figure 7. Phosphorus mass percentage of each portion in total P and total P percentage of whole biomass in the batch culture system. Cells were grown in: (a) synthetic BBM media with different initial phosphorus concentration on day 1, 3, and 7 for the three highest phosphorus concentration cases, (b) Stream A, (c) Stream B, and (d) Stream C.

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120 18 (a) Liquid P (%) AISP (%) ASP (%) Bound P (%) %P of biomass 16 100 14 80 12 10 60 8

%P of total P total of %P 40 6 %P of biomass of %P 4 20 2 0 0 12 16 20 12 16 20 12 16 20 12 16 20 25% 50% 75% 100% Change of trough liquid

Bound P (%) ASP (%) AISP (%) (b) 120 18 Liquid P (%) %P of biomass 16 100 14 80 12 10 60 8

%P of total P total of %P 40 6 %P of biomass of %P 4 20 2 0 0 30 40 50 30 40 50 30 40 50 30 40 50 25% 50% 75% 100% Change of trough liquid

Figure 8. Phosphorus mass percentage of each portion in total P and total P percentage of whole biomass in the continuous culture feeding wastewater stream A (Table 2). (a) Every 2-day harvest frequency and (b) 5-day harvest frequency.

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CHAPTER 6. GENERAL CONCLUSION

The present body of study verifies the potential use of microalgae to treat different kinds of environmental pollutants, air and wastewater, as nutrients for cell growth. Different microalgae cultivation system was used for each pollutants to increase nutrient transfer to biomass: gas was bubbled through the reactors for air-born pollutants; and algal biofilm was submerged in trough liquid for water-born pollutants.

In culturing microalgae with ammonia gas and CO 2, the ammonia gas removal efficiency was consistently high (> 95%) and the majority (80‒90%) of the ammonia was removed by algal cell assimilation. The CO 2 removal rate and removal efficiency, however, highly depended on the CO 2 mass loading rate and it was explained in regards to the cellular metabolic activities. It was suggested that in comparing algal performance in CO 2 removal, it is important to take into account the CO 2 loading rate instead of concentration. The produced algal biomass contained high protein content with an amino acid profile close to the ideal protein compositions for different animal feeds, with a high ratio of essential amino acids.

As an effort to enhance the feasibility of microalgal ammonia scrubber system, a valuable byproduct, astaxanthin was produced via Haematococcus pluvialis. Ammonia was fed as both nitrogen source and astaxanthin inducer and different strategies were examined to enhance astaxanthin content, such as different fed-batch strategies and different chemicals to generate stressful condition were examined. From the data, it was concluded that H. pluvialis can utilize ammonia gas as a nitrogen source and astaxanthin induction but further astaxanthin induction strategies were only effective at C/N ratio 1.

Another type of pollutant, wastewater was treated with microalgae in RAB system. The

RAB system showed excellent cell growth and liquid phosphorus removal, even at high

131 phosphorus concentration (215.9 mg/L-TP). By analyzing intracellular phosphorus, cellular metabolic activities in regards to the initial phosphorus concentration was interpreted and luxury phosphorus uptake was proved. From the phosphorus mass balance, it was shown that the majority of liquid phosphorus migrated to the bound-P.

This thesis is very timely as Pope Francis calls public attention to environment protection in his encyclical on ecology and climate change, ‘Laudato Si’. Such efforts in this thesis will minimize anthropogenic impacts on Earth and future generations also can be gifted with the protected nature.

Future Research Recommendations

Much additional work should be made for furthering our findings realistic.

First, microalgal waste gas mitigation needs to be tested with the real outlet gas from

CAFOs as the present study only tested ammonia gas and carbon dioxide. The photobioreactor also needs to be tested in pilot-scale in the real CAFOs with different parameters, such as methods of collecting the outlet gas, different configurations of reactors, and outdoor or indoor environment.

Second, the potential use of microalgae biomass as animal feed additive should be validated for its bioavailability, possible toxicity, impact for animal performances, and its applicability. Especially after treating wastewater, the produced microalgae biomass might contain some amount of heavy metals and pharmaceutical chemicals. This concern should lead to a study of applicability of microalgae biomass as a fertilizer.

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Third, the low astaxanthin content in biomass in this study can be enhanced by higher light intensity or other growth parameters. For example, the cell growth performance in the bubble column is significantly influenced by the initial seed cell status. By inoculating different stages of cells, the cell growth and astaxanthin content can be enhanced. Also, feeding the real outlet gas from CAFOs to induce astaxanthin can be another future study topic.

Lastly, the RAB system should be run in continuous mode with different wastewater streams. As different wastewaters contain different nutrient compositions, studying various wastewater will give a clear picture in designing optimal RAB system operation.