1 Deoxygenation Treatment Strategy

Deoxygenation Treatment Strategy to Control Vampirovibrio chlorellavorus in Chlorella sorokiniana Cultures

Item Type text; Electronic Dissertation

Authors Attalah, Said

Publisher The University of Arizona.

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Download date 02/10/2021 21:48:38

Link to Item http://hdl.handle.net/10150/631467 1

DEOXYGENATION TREATMENT STRATEGY TO CONTROL VAMPIROVIBRIO CHLORELLAVORUS IN CHLORELLA SOROKINIANA CULTURES

Said Attalah

A Dissertation Submitted to the Faculty of the

DEPARTMENT OF BIOSYSTEMS ENGINEERING

In Partial Fulfillment of the Requirements

For the Degree of

DOCTOR OF PHILOSOPHY

In the Graduate College

THE UNIVERSITY OF ARIZONA

2018

STATEMENT BY AUTHOR

This dissertation has been submitted in partial fulfillment of the requirements for an advanced degree at the University of Arizona and is deposited in the University Library to be made available to borrowers under rules of the Library.

Brief quotations from this dissertation are allowable without special permission, provided that an accurate acknowledgement of the source is made. Requests for permission for extended quotation from or reproduction of this manuscript in whole or in part may be granted by the copyright holder.

SIGNED: Said Attalah

ACKNOWLEDGEMENTS

First, I would like to express my highest regards and sincere gratitude to my advisor Dr.

Peter Waller for his valuable guidance and continuous support during the course of this research. Without his guidance and dedication, I would not be able to achieve my PhD program. I am greatly thankful to my co-advisor Dr. Kevin Fitzsimmons for his support and advice. I was fortunate to take his classes and learn the essentials about algae. I express my profound gratitude to Dr. Kim Ogden for her advice and help to define and develop my research plan. Her insightful thoughts and pertinent comments were very valuable to my research project. I am honored to have her in my dissertation committee. My deep gratitude goes to Dr. Stephen Poe who accepted to be in my dissertation committee. His advice and encouragement helped me to be persistent and stay focused on my research goals.

My sincere acknowledgements are extended to Dr. Judith Brown, who helped me to develop my experimental design and allowed me to use her laboratory equipment. Her insightful comments and suggestions were crucial in the completion of this research.

I would like to thank S. Steichen, C. Brown, and C. N. Galvez who provided the stock culture and inoculum and helped with qPCR measurements. I would like to thank S. Gao for his help with nutrients preparation and ash free dry weight measurement.

My acknowledgements go also to Dr. M. Kacira, Dr. G. Ogden, Dr. F. Jia, G. Khawam, O.

Bertelsen, E. Leichtenberg, K. Lepley, I. Liang, M. Acedo, and Y. Mahdipoor, members of the RAFT team at the University of Arizona.

My exceptional thanks go to the BE department faculty and staff members for their continuous support. I am honored and proud to be a part of this great department.

I am grateful for the U.S. Department of Energy and Regional Algal Feedstock Testbed

(RAFT) project, University of Arizona, for supporting this study.

I am so thankful to my family for their love and support that gave me lot of courage and patience to achieve this project.

Thank you!

TABLE OF CONTENT

LIST OF FIGURES ...... 11

LIST OF TABLES ...... 16

ABSTRACT ...... 17

Chapter 1: Introduction ...... 19

1.1. Literature review ...... 19

1.1.1. Microalgae cultivation and biological contaminants issue ...... 19

1.1.2. Bacteria-algae interaction ...... 22

1.1.3. Chlorella sorokiniana characteristics ...... 23

1.1.4. Vampirovibrio chlorellavorus phenotypes ...... 24

1.1.5. Control strategies of biological contaminants ...... 25

1.2. Rationale ...... 26

1.3. Hypotheses ...... 27

1.3.1. Hypothesis 1 ...... 27

1.3.2. Hypothesis 2 ...... 27

1.4. Dissertation structure ...... 28

Chapter 2: Deoxygenation in algae cultivation systems ...... 31

2.1. Introduction ...... 31

2.2. Dissolved oxygen measurement methods ...... 31

2.3. Deoxygenation approaches ...... 32

2.4. Deoxygenation impact on microalgae and associated microorganisms ...... 33

Chapter 3: Technical evaluation and cost estimation of the deoxygenation process ...... 36

3.1. Introduction ...... 36

3.2. Dissolved oxygen removal ...... 37

3.3. Deoxygenation with nitrogen gas sparging ...... 38

3.4. Technical considerations and cost estimation of the deoxygenation process ...... 38

Conclusion ...... 40

Complete Dissertation References ...... 41

Appendix A: Application of deoxygenation-aeration cycling to control the predatory bacterium Vampirovibrio chlorellavorus in Chlorella sorokiniana cultures ...... 55

Abstract ...... 55

1. Introduction ...... 56

2. Materials and methods ...... 60

2.1. Pathogen-free cultures and co-cultures ...... 60

2.2. Deoxygenation-aeration cycling experiments ...... 61

2.3. Inoculum preparation and experimental media ...... 63

2.4. Algal suspension culture growth assessment ...... 63

2.5. Real time quantitative polymerase chain reaction for quantification of V. chlorellavorus

...... 65

2.6. Experimental reactor design and laboratory apparatus ...... 65

3. Results and discussion ...... 68

3.1. Effect of deoxygenation cycling on V. chlorellavorus -free C. sorokiniana ...... 68

3.2. Impact of deoxygenation-aeration cycling on V. chlorellavorus infection ...... 74

4. Conclusion ...... 83

5. Acknowledgements ...... 83

6. References ...... 83

Appendix B: Technoeconomic assessment of deoxygenation for control of Vampirovibrio chlorellavorus in Chlorella sorokiniana cultures ...... 92

Abstract ...... 92

1. Introduction ...... 93

2. Materials and methods ...... 95

2.1. Cultures and cultivation medium ...... 95

2.2. Laboratory and outdoor experimental protocols ...... 95

2.3. Experimental apparatus design and operating procedure ...... 98

2.3.1. Laboratory experimental setup ...... 98

2.3.2. Outdoor experimental setup ...... 100

2.4. Inoculum preparation and sampling approach ...... 101

2.5. Assessment of C. sorokiniana growth and V. chlorellavorus infection ...... 102

2.6. Technical assessment of nitrogen usage and cost estimation ...... 103

3. Results and discussion ...... 106

3.1. Pathogen-free C. sorokiniana monoculture and C. sorokiniana-V. chlorellavorus co-

cultures with continuous aeration ...... 106

3.2. Evaluation of the efficacy of short time deoxygenation on the growth of C. sorokiniana

infected by V. chlorellavorus ...... 108

3.3. Application of one-hour nitrogen gas sparging (deoxygenation) in C. sorokiniana -V.

chlorellavorus co-cultures associated with unidentified ciliates ...... 111

3.4. Nitrogen gas sparging and dissolved oxygen depletion assessment ...... 117

3.5. Nitrogen-usage evaluation and cost estimation ...... 122

4. Conclusion ...... 125

5. Acknowledgements ...... 125

6. References ...... 126

Appendix C: Data acquisition and dataloggers programing ...... 135

1. Purpose ...... 135

2. Campbell Scientific data logger CR300 for Laboratory experiments ...... 135

2.1. Description ...... 135

2.2. Operating program instructions ...... 136

3. Campbell Scientific data logger CR3000 for outdoor experiments ...... 137

3.1. Description ...... 137

3.2. Operating program instructions ...... 138

4. References ...... 140

Appendix D: Standard operating procedures ...... 141

1. Purpose ...... 141

2. Media recipe ...... 141

3. Standard curve for algae ash free dry weight (AFDW) and biomass measurement ...... 142

4. Vampirovibrio chlorellavorus filtration ...... 143

5. References ...... 144

Appendix E: Interaction between V. chlorellavorus , ciliates and C. sorokiniana ...... 146

1. Purpose ...... 146

2. Experiments and preliminary results ...... 146

3. Conclusion ...... 149

4. References ...... 150

LIST OF FIGURES

Fig. 1.1. Algae raceway integrated design (ARID) at the algae facility of the University of Arizona (Regional Algal Feedstock Testbed (RAFT) project, 2017). ARID prototype in operation with healthy algae circulating in open channels (upper left), healthy algae stored in ARID deep canal (upper right), collapsed Chlorella sorokiniana under the impact of Vampirovibrio chlorellavorus infection, in channels (bottom left) and in deep canal (bottom right)...... 21

Fig. 2.1. Dissolved oxygen removal with nitrogen gas sparging in Chlorella sorokiniana culture. Treated culture was subjected to 2-hr deoxygenation alternated with 2-hr aeration during the dark period. Control culture without aeration or deoxygenation. Growth measurement based on ash free dry weight (AFDW) biomass...... 33

Fig. 2.2. Dissolved Oxygen (DO) and ash free dry weight (AFDW) of Chlorella sorokiniana co- cultured with Vampirovibrio chlorellavorus. Treatment (TR) was subjected to deoxygenation with 1-hr nitrogen sparging during the dark period. Control (C) was continuously aerated during the dark period...... 35

Fig. A.1 . Experimental reactor design and apparatus. A) General view of the experimental apparatus arrangement. B) Experimental equipment and Campbell Scientific (CS) data logger for deoxygenation-aeration cycling management. C) Grow-light panels added to the incubator, and pH controllers used to regulate pH in the suspension culture. D) Light intensity measured with MP-200 Apogee silicon cell pyranometer placed at equidistance from one another to achieve consistent deoxygenation-aeration cycling during treatment 1-3 (TR1, TR2, and TR3), with the nonaerated positive control (C1), and aerated negative control (C2). E) Example of an experimental flask with tubing, sensors, and rubber stopper. F) Arrangement of the 1-gallon Erlenmeyer flasks containing the Chlorella sorokiniana suspension cultures placed inside the incubator...... 66

Fig. A.2. Daily ash free dry weight (AFDW) biomass (lines with markers), and biomass gain during the light period and biomass loss during the dark period (markers) in experiments 1-3 with Vampirovibrio chlorellavorus-free Chlorella sorokiniana: A-C) 4-, 6-, and 8-h deoxygenation treatments, respectively...... 71

Fig. A.3 . Comparison of the average ash free dry weight (AFDW) in experiments 1-3 with Vampirovibrio chlorellavorus-free Chlorella sorokiniana cultures, including nonaerated positive control (C1), aerated negative control (C2), and 4-, 6-, and 8-h deoxygenation treatments (TR). Error bars represent one standard deviation...... 72

Fig. A.4. Dissolved oxygen in experiments 1-3 with Vampirovibrio chlorellavorus-free Chlorella sorokiniana culture: nonaerated positive control (C1), aerated negative control (C2), and deoxygenation treatments (TR) with various deoxygenation times (4-, 6-, and 8-h) during dark period...... 73

Fig. A.5. Compared average dissolved oxygen between Vampirovibrio chlorellavorus -free Chlorella sorokiniana culture (Exp.1) and C. sorokiniana -V. chlorellavorus co-cultures (Exp. 4). During the 8-h dark periods, negative controls (C2) were continuously aerated, and treatments (TR) involved 2-h deoxygenation-2-h aeration cycles. The control C2 in Exp.4 represents an average of the aerated control duplicates, C2A and C2B...... 75

Fig. A.6 . Compared ash free dry weight (AFDW) and DNA ratios in aerated negative controls (C2A and C2B) and 2-h deoxygenation-aeration cycling treatment (TR1 and TR2) in three replicates (A-C) with Chlorella sorokiniana -Vampirovibrio chlorellavorus co-cultures (Exp.4). DNA ratios of V. chlorellavorus to C. sorokiniana are presented on log10 scale at the growth phase (1), stationary phase (2), and death phase (3) for the end of the 16-hr light periods (small markers) and end of the 8-h dark periods (large markers)...... 79

Fig. A.7 . Compared ash free dry weight (AFDW) in aerated negative control experiments between Vampirovibrio chlorellavorus -free Chlorella sorokiniana cultures (Exp. 1-3) and C. sorokiniana -V. chlorellavorus co-cultures (Exp. 4). (a) and (b) show the selected data for statistical t-test. Error bars represent one standard deviation...... 81

Fig. A.8 . Visual inspection and light microscopic observations (400x magnification) of Chlorella sorokiniana -Vampirovibrio chlorellavorus co-cultures. A) Treated co-cultures with deoxygenation-aeration cycles, left and middle flasks (TR) with brighter grass-green color, and aerated controls, the 2 flasks on the right (C) with foamy brown grime on the walls. B) Green single cells of C. sorokiniana in the treated co-cultures. C) Lysed and clumped cells of C. sorokiniana in the aerated controls...... 82

Fig. B.1 . Equipment for the laboratory experiments. A) General view of the Innova 4330 incubator, equipment for deoxygenation and aeration, and pH controllers. B) Grow-light panels were added to the incubator for culture illumination. C) Flasks containing Chlorella sorokiniana culture with sensors (dissolved oxygen and pH) and stone diffusers for nitrogen and air. In the present study, only the four flasks in the corners were considered; deoxygenation treatment replicates (TR1 and TR2) were on the left, and aerated control replicates (CA1 and CA2) on the right...... 99

Fig. B.2 . Equipment for the outdoor experiments. A) Polypropylene reactor containing Chlorella sorokiniana culture with dissolved oxygen probe (DO1, DO2, DO3) locations inside the reactor. B) Sealed lid with two small vents and connectors. C) Sensors and PVC-pipe structure inside the reactor. D) CO 2 cylinder and Campbell Scientific CR3000 data logger. E) Nitrogen cylinder and relays...... 101

Fig. B.3. Two-point calibration (saturation and zero) of the dissolved oxygen probes (DO1, DO2, DO3) measured with CR3000 data logger prior to the outdoor experiments with clean tap water (Exp. 6) and Chlorella sorokiniana culture (Exp.7)...... 103

Fig. B.4. ARID raceway prototype cultivated with Chlorella sorokiniana at the algae facility of the University of Arizona (Regional Algal Feedstock Testbed project, 2017)...... 105

Fig. B.5 . Average ash free dry weight (AFDW) biomass and dissolved oxygen (DO) with continuous aeration during the dark period. A) Pathogen-free Chlorella sorokiniana culture sustained growth period. B) Collapse of C. sorokiniana co-cultured with Vampirovibrio chlorellavorus indicating a high sensitivity of C. sorokiniana to V. chlorellavorus infection in a continuously aerated environment . Error bars represent one standard deviation ( n = 4 )...... 108

Fig. B.6 . Average ash free dry weight (AFDW) biomass and dissolved oxygen (DO) for Chlorella sorokiniana co-cultured with Vampirovibrio chlorellavorus . Aerated control (C) and deoxygenation treatment (TR) represent averages of the aerated control duplicates (CA1 and CA2) and the treatment duplicates (TR1 and TR2), respectively. During dark period, the controls were continuously aerated, and the treatments were subjected to 0.5-h deoxygenation at the beginning of the dark period (Exp. 3). Error bars denote one standard deviation. ... 109

Fig. B.7. Average ash free dry weight (AFDW) biomass and dissolved oxygen (DO) for Chlorella sorokiniana co-cultured with Vampirovibrio chlorellavorus . The aerated control (C) and deoxygenation treatment (TR) represent averages of the aerated control duplicates (CA1 and CA2) and the treatment duplicates (TR1 and TR2), respectively. During dark periods, the controls were continuously aerated, and the treatments were subjected to 1-h deoxygenation at the beginning of the dark period (Exp. 4). Error bars denote one standard deviation. ... 111

Fig. B.8. Ash free dry weight (AFDW) biomass and DNA ratio of Vampirovibrio chlorellavorus to Chlorella sorokiniana (log 10 scale) in the experiments with C. sorokiniana-V. chlorellavorus co-cultures contaminated by ciliates. Treatments subjected to 1-hr nitrogen sparging for deoxygenation during 8-hr dark period/day. A, B, and C are replicates...... 114

Fig. B.9. Average ash free dry weight (AFDW) biomass and dissolved oxygen for 3 replicates with Chlorella sorokiniana co-cultured with Vampirovibrio chlorellavorus in association with unidentified ciliates (Exp. 5). The treated culture (TR) was subjected to deoxygenation with 1-h nitrogen gas sparging at the beginning of the dark period, and the controls (C) were continuously aerated during the dark period. Error bars denote one standard deviation (n = 6). Points (a) and (b) represent the grouped data for a statistical comparison...... 115

Fig. B.10. Chlorella sorokiniana co-cultured with Vampirovibrio chlorellavorus contaminated by unidentified ciliates. The co-culture was subjected to deoxygenation treatment with 1-h nitrogen gas sparging at the beginning of the dark period. Visual symptoms of the collapsed culture in the aerated controls (upper left) and healthy treated culture (upper right). Light microscopy (400x magnification) observation of clumped and lysed C. sorokiniana cells (bottom left) and clean single cells (bottom right)...... 116

Fig. B.11. Compared dissolved oxygen (DO) concentrations in clean water and Chlorella sorokiniana culture under similar temperatures and deoxygenation treatment with 1-h nitrogen sparging at the beginning of the night. Logarithmic slope of the oxygen deficit vs time was used to calculate the oxygen depletion coefficient (K La20 )...... 118

Fig. B.12. Dissolved oxygen (DO) concentration with less than 1-h nitrogen sparging (deoxygenation treatment) in clean water (solid line) and Chlorella sorokiniana culture (dashed lines)...... 119

Fig. B.13. Dissolved oxygen (DO) fluctuations in clean water (solid line) and Chlorella sorokiniana culture (dashes) under similar temperatures over 24 hours with less than 1-h nitrogen sparging at the beginning of the night...... 120

Fig. B.14 . Chlorella sorokiniana growth indicators and environmental parameters in the outdoor reactor. A) Ash free dry weight (AFDW) biomass, and dissolved oxygen (DO) concentrations at various depths in the reactor: 6 cm (DO1), 30 cm (DO2), and 60 cm (DO3) below the culture surface. B) Culture temperature and solar intensity (measured outside the reactor)...... 121

Fig. C.1. Campbell Scientific data logger, CR300, installed in the laboratory for dissolved oxygen data acquisition...... 135

Fig. C.2. Campbell Scientific CR3000 datalogger components ...... 137

Fig. C.3. Campbell Scientific CR3000 wirings for the outdoor experiment (left). Nitrogen cylinder connected to a solenoid valve activated with a relay(right) through CR3000 program. .... 138

Fig. D.1. Linear correlation between the optical density at 750 nm wavelength (OD750) and the ash free dry weight (AFDW) for pathogen-free Chlorella sorokiniana monoculture...... 142

Fig. D.2. Vampirovibrio chlorellavorus filtration with a 2.0 µm pore size Whatman filter and vacuum flask apparatus (filtration level 1)...... 143

Fig. D.3. Vampirovibrio chlorellavorus filtration with a 0.22 µm pore size Whatman filter and vacuum flask apparatus (filtration level 2). Filtration process (left). Final filtrate (right) ready for use to infect Chlorella sorokiniana culture...... 144

Fig. E.1. Visual inspection of Chlorella sorokiniana in the presence of V ampirovibrio chlorellavorus and unidentified ciliates with deoxygenation treatment applying one-hour nitrogen gas sparging at the beginning of the dark period. Symptoms of the collapsed culture in the two aerated control flasks (C), from the right to the left with foamy brown grime on the walls, and healthy cultures in the three treated flasks (TR), from the left to the righ, with brighter grass-green color...... 147

Fig. E.2. Light microscopic observation showing Clumped and lysed Chlorella sorokiniana cells under the impact of Vampirovibrio chlorellavorus attack and contamination by unidentified ciliates (400x magnification)...... 148

Fig. E.3. Average DNA ratio of Vampirovibrio chlorellavorus to Chlorella sorokiniana (n = 6) in the experiments with C. sorokiniana-V. chlorellavorus co-cultures contaminated by ciliates. One-hour nitrogen sparging treatment (TR) for deoxygenation during 8-hr dark period/day. The controls (C) were continuously aerated...... 149

LIST OF TABLES

Table 1.1. Global market of selected microalgal high-value compounds (Markou & Nerantzis, 2013) ...... 19

Table A.1. Deoxygenation-aeration cycling experiments schedule...... 62

Table A.2. Light intensity measurements recorded for each flask during the experimental runs 68

Table B.1. Experimental plan ...... 97

Table B.2. Onsite nitrogen gas generator specifications (N-750-T, On-Site Gas Systems Inc., Newington, CT) ...... 123

Table B.3. Annual cost of deoxygenation process with an onsite nitrogen generator (N-750-T) and OSMAN SA100 air compressor in a commercial scale ARID raceway system with 24 raceways each covering 0.4 ha surface area ...... 125

Table C.1. CR300 datalogger program for data acquisition in the laboratory experiments ..... 136

Table C.2. CR3000 program instructions for data acquisition in outdoor experiment ...... 138

Table D.1. BG-11 media recipe (UTEX, 2018) ...... 141

ABSTRACT

Researchers and algae growers have shown deep interest in the cultivation of Chlorella microalgae because of its significant growth potential, high biomass quality, and broad range of applications.

Both small and large-scale cultivation systems (laboratory or outdoors) with Chlorella microalgae have been extensively studied, and promising results have been demonstrated in terms of cultivation procedures, growth media, growth parameters, harvesting, and processing; however, protection against biological contaminants such as the predatory Vampirovibrio chlorellavorus requires more investigation. Despite the high growth potential of Chlorella species, the impact of

V. chlorellavorus infection on the culture productivity is significantly damaging and cannot be counterbalanced naturally. Thus, a pressing need to develop appropriate preventative and/or curative strategies for an optimal control of this type of predator has become one of the priorities in the cultivation process of Chlorella microalgae.

The present research focusses on the management of dissolved oxygen (DO) during the nighttime (dark period) in co-cultures of Chlorella sorokiniana and V. chlorellavorus . We developed an unprecedented method to control the infection of C. sorokiniana by V. chlorellavorus . Because V. chlorellavorus is an obligate aerobe and C. sorokiniana is not, deoxygenating the culture assumably harms the predator and not the host. This research first examined the effect of deoxygenation on pathogen-free C. sorokiniana monoculture, and then on

C. sorokiniana -V. chlorellavorus co-cultures.

In the first section, initial experiments with pathogen-free C. sorokiniana cultures included different deoxygenation-aeration cycle times. Pure nitrogen gas was used to create anoxic conditions, and ambient air was used to reestablish aerobic conditions. In these initial experiments,

C. sorokiniana tolerated anoxic conditions for extended time intervals as long as 8 hours. In the experiments with infection by V. chlorellavorus , aerated controls collapsed while the deoxygenated cultures sustained a normal growth cycle. Visual observation showed a healthy green C. sorokiniana culture in the treated cultures and a brown slime and collapsed culture in the aerated controls.

In the second section, we evaluated the technical aspects of the deoxygenation practice and cost of nitrogen gas sparging as an appropriate method to create anoxic conditions in the cultivation system. In this experiment, DO concentrations were driven to low levels (0.2 ppm to 0.5 ppm) by sparging nitrogen gas for one hour at the beginning of the night (dark period), and then natural deoxygenation by dark respiration kept the oxygen concentration at a low level. The laboratory results showed that this method kept the DO levels low during the entire dark period and effectively controlled V. chlorellavorus infection in C. sorokiniana co-cultures.

The cost of the deoxygenation method was estimated in outdoor experiments with pathogen-free C. sorokiniana cultures. Nitrogen sparging for one-hour at the beginning of the dark period maintained dissolved oxygen concentrations at low levels (<0.5 ppm) throughout the night.

The total nitrogen injection per night per liter algae culture was then translated to annual commercial-scale raceway cost. Finally, the technical and economic feasibility of this process was evaluated for onsite nitrogen gas generators in commercial-scale reactors.

Keywords: Anoxic, biomass, infection, nitrogen, aeration, economic feasibility.

Chapter 1

Introduction

1.1. Literature review

1.1.1. Microalgae cultivation and biological contaminants issue

Microalgae are cultivated at small and commercial scales for various purposes such as nutraceuticals, pharmaceuticals, and cosmetics (Borowitzka, 2013; Markou & Nerantzis, 2013), fertilizer (Mulbry et al., 2005), feed (Madeira et al., 2017), and biofuel (Chisti, 2013; Safi et al.,

2014). Some high-value compounds are reported in Table 1.1 (Markou & Nerantzis, 2013).

Microalgae have also been used as a bioremediation method in wastewater treatment (Richmond,

2004; Park et al., 2011; Ramanna et al., 2014).

Table 1.1. Global market of selected microalgal high-value compounds (Markou & Nerantzis, 2013).

Global Market (million Production Added-value compound US$/annum) kt/annum Price $/kg Carotenoids 1200 - - β-Carotene 261 - 300-700 Lutein 233 - - Astaxanthin 240 - 2000-7000 Bioplastics - 64 - Fatty acids (omega-3) >700 85 0.88-3.8 Vitamins and supplement 68 - - Glycerol - 1995.5 0.3-1 Phycobilin >60 - - C-phycocyanin - 5 - B-phycoerythrin - - 50000

Considering the diversity of algae species, different cultivation modes are applied: autotrophic with light and inorganic carbon source (Yeesang & Cheirsilp, 2011), heterotrophic with an organic carbon source under dark conditions (Radmer & Parker, 1994), or mixotrophic under simultaneous autotrophic and heterotrophic conditions (Lowrey et al., 2015).

Among microalgae cultivation systems, open pond raceways have been commonly used since 1970s (Goldman, 1979). Their simplicity and low cost made them convenient for use in commercial microalgae cultivation (Chaumont 1993; Sheehan et al., 1998; Waltz 2009), especially for a low-value product like algal biofuel, which requires economically feasible techniques and environmentally friendly practices. Regardless of the cultivation system, proper management of algae cultivation systems including nutrients (macro and micro), vitamins, pH levels, temperature, salinity, mixing, and light should be considered to achieve an optimal production (Singh & Singh,

2015).

One of the undeniable challenges in algal cultivation raceways is biological contamination

(McBride et al., 2016). Lack of appropriate control of contaminants in algal cultivation systems usually leads to significant biomass loss (Wen-Li et al., 2011). In open cultivation raceways, exposure of algae culture to the open environment enhances contamination by grazers, predators, parasites, and competitors (Lincoln et al., 1983; Flynn et al., 2017) such as zooplankton (e.g. rotifers), phytoplankton (e.g. other microalgae), bacteria (e.g. Bdellovibrio sp .), and viruses (e.g.

Cyanophage ) (Lawrence, 2008; Wang et al., 2010). Contaminants affect algae growth potential and lead to frequent collapses which incur costly maintenance and decrease in productivity.

Contaminants can decrease algae productivity by 10 to 30% in open raceway ponds, and 5 to 10% in closed photobioreactors (Richardson et al., 2014). Bacterial predators are among the biological

contaminants which have significant impact on microalgae biomass. For instance, contamination of Chlorella species by the predatory Vampirovibrio chlorellavorus (Gromov & Mamkaeva, 1980) has been a significant issue in the paddlewheel raceways (Ganuza et al., 2016) and the Arid

Raceway Integrated Design (ARID) (Waller et al, 2012), especially during warm seasons (Li,

2015; Steichen et al., 2016). The impact of V. chlorellavorus infection on Chlorella cultures is so significant that the culture can collapse unexpectedly and quickly, within a day or two, when the attack occurs (Ganuza et al., 2016). Fig.1.1 illustrates some visual symptoms of a healthy C. sorokiniana culture and V. chlorellavorus infection in the ARID raceway during the Regional

Algal Feedstock Testbed (RAFT) research project (2017) at the algae facility of the University of

Arizona.

Fig. 1.1. Algae raceway integrated design (ARID) at the algae facility of the University of Arizona

(Regional Algal Feedstock Testbed (RAFT) project, 2017). ARID prototype in operation with healthy algae circulating in open channels (upper left), healthy algae stored in ARID deep canal

(upper right), collapsed Chlorella sorokiniana under the impact of Vampirovibrio chlorellavorus infection, in channels (bottom left) and in deep canal (bottom right).

Introduction and propagation of biological contaminants into the microalgae cultivation systems, whether in the closed bioreactors or open raceways, can be caused by water, air, nutrients, inoculum, dust, insects, birds, animals, or a cross-contamination via shared equipment. Although polycultures might be less affected than monocultures (McBride, 2016), preventative and/or curative control measures should be considered in both cases.

1.1.2. Bacteria-algae interaction

Understanding the interactions between bacteria and microalgae is a key to success in algae farming and algae-biotechnology. The coexistence of a specific bacterium and a specific alga in a specific environment clearly defines a special connection between the two organisms. The interaction mode could either be beneficial or detrimental to the algae culture, thus requires thorough investigation if appropriate management approaches of the phycosphere is intended.

Mutualism, commensalism, and parasitism (Ramanan et al., 2016) are the main modes of interaction that can occur in the phycosphere (Bell & Mitchell, 1972). Although bacteria-algae associations are necessary for the ecological balance, the impact of bacterial predation on algae culture can be devastating, especially in commercial-scale cultivation systems (Ganuza et al.,

2016).

Predatory bacteria are two types: obligate (e.g. Bdellovibrio spp .), or facultative (e.g.

Saprospira grandis ) (Pasternak et al., 2013). Obligate predators are the most damaging biological

contaminants because they cannot survive without killing their prey. For instance, the life cycle of the predatory V. chlorellavorus cannot be accomplished without decaying the host Chlorella species (Soo et al., 2016), which usually results in a quick collapse of Chlorella culture. Thus, an effective bacterial control strategy should be integrated into the cultivation process to prevent the culture loss.

Under optimal growth conditions, microorganisms undertake four growth stages: lag phase

(physiological acclimation), exponential phase (the fastest cell division stage), stationary phase

(unstable limited growth), and declining or death phase (continuous net loss) (Yates & Smotzer,

2007). However, unfavorable environmental conditions and/or contamination can interfere with the normal growth cycle and truncate the growth stages, or even cause an early death.

1.1.3. Chlorella sorokiniana characteristics

Chlorella sorokiniana is one of the most prominent microalgae species which have attracted many researchers and algae growers worldwide. The Chlorophyte C. sorokiniana belongs to the trebouxiophyceae class, chlorellales order, and chlorellaceae family (Shihira & Krauss,

1965). Chlorella microalgae are spherical unicellular species with a size ranging from 2 – 10 microns (Lizzul et al., 2018). Chlorella sorokiniana is qualified as a top producer based on its high growth potential, high biomass productivity (30 g m-2 d-1 in outdoor paddlewheel raceways), tolerates a wide range of temperatures (4 - 43 °C), and generates about 25% fatty acids (Neofotis et al., 2016). Huesemann et al., (2016) reported that C. sorokiniana species can reach a maximum growth rate of 5.9 d-1 at pH 6 and tolerate a wide range of pH (3 to 9) and up to 35 g L-1 (NaCl) salinity concentration. Chlorella sorokiniana is also viewed as a potential candidate for biofuel

production and can be cultivated in fresh or salt water for various purposes such as nutraceuticals, pharmaceuticals, cosmetics, and feed (Huesemann et al., 2016).

1.1.4. Vampirovibrio chlorellavorus phenotypes

Vampirovibrio chlorellavorus is a pleomorphic gram-negative bacterium with a size varying from 0.3 microns (vibrios) to 0.6 microns (cocci) and lives attached to the surface of

Chlorella species cells (Coder & Starr, 1978; Esteve et al., 1983; Coder & Goff, 1986).

Vampirovibrio chlorellavorus is nonculturable on agar media and requires co-culturing with

Chlorella species to survive (Coder & Starr, 1978). Symptoms of V. chlorellavorus infection on

Chlorella species were first observed by Mamkaeva in 1966, but without a detailed investigation.

In 1972, in an attempt to describe the predatory microorganism, Gromov and Mamkaeva misnamed the bacterium as Bdellovibrio chlorellavorus , but then they accurately renamed it in

1980 as V. chlorellavorus , arguing that Bdellovibrio species enter the periplasm of their hosts, and

Vampirovibrio attach to the surface of Chlorella species (Coder & Starr, 1978). Subsequent studies

(Esteve et al., 1983; Coder & Goff, 1986; Soo et al., 2015) with Chlorella and other algae species showed that V. Chlorellavorus only attacked Chlorella species ( C. vulgaris, C. sorokiniana, and

C. kessleri ) but not Scenedesmus, Ankistrodesmus, Scotiella, or Chlorococcum . Thus, V. chlorellavorus was identified as a Chlorella specific predator. Coder and Goff (1986) demonstrated that V. chlorellavorus connects to Chlorella species and ruptures their cell-wall, suggesting that such specificity might be related to the cell wall properties of the specific Chlorella species. Esteve et al., (1983) found that epibiont species like V. chlorellavorus attach to the hosts and decay their cells. Soo et al., (2015) characterized V. chlorellavorus as an obligate aerobic

epibiont predatory bacterium belonging to the cyanobacteria phylum, melainabacteria class, and vampirovibrionales order, with a life cycle including: prey location, attachment, development of secretion system and ingestion, binary division, and release. Steichen et al., (2016) reported that the fastest infection and death of C. sorokiniana by V. chlorellavorus occurred when the temperature exceeded 34 °C. Ganuza et al., (2016) found that V. chlorellavorus cannot tolerate a pH ≤ 3.5, thus can be controlled with pH-shock treatment. These characteristics are not exhaustive, but worth considering in the management of Chlorella culture cultivation.

1.1.5. Control strategies of biological contaminants

The presence of undesirable impurities or invaders such as biological contaminants in microalgae cultivation systems can cause substantial biomass losses. Thus, appropriate control strategies, preventative and/or curative, are crucial to reduce the impact of contamination and improve the algae productivity. The existing control procedures can be categorized into physical, chemical, biological, or integrated techniques. In general, the control approaches include selective growth conditions (Borowitzka, 2005), filtration and chemical pesticides (McBride et al., 2014), pulsed electric fields (Dempster, 2015), zooplanktivorous integration (Smith et al., 2010), salvage harvest (Carney and Lane 2014), chemical agents (Pouneva, 2006; Woo and Kamei, 2003; Fott &

Fott, 1967; Shurin et al., 2013; Benderliev et al., 1993; Webb et al., 2012; Bagwell et al., 2016), physical disruption and removal (Holm et al. 2008), biological control (Larkum et al 2012, Qin et al 2012), and genetic engineering (Loera‐Quezada et al, 2016). For instance, filtration was used to eliminate rotifers (Borowitzka, 2005), quinine was used to control ciliates (Moreno-Garrido &

canavate, 2001), and pH 3.5 shock-treatment was used to inhibit V. chlorellavorus (Ganuza et al.

2016).

Most of the previous approaches appear to successfully eradicate the targeted contaminants or at least limit their impact, but some treatments come with restrictions and are applied with caution to avoid the risk of affecting nontargeted species (Wang et al., 2010) or causing environment pollution.

1.2. Rationale

Chlorella sorokiniana is one of the prominent microalgae species with a high growth potential (Huesemann et al., 2016) and high lipid rate (Neofotis et al., 2016); however, it is highly sensitive to predation by the predatory bacterium V. chlorellavorus (Gromov & Mamkaeva, 1980).

Vampirovibrio chlorellavorus infection is usually severe and oftentimes leads to an unexpected and quick collapse of the culture (Gromov & Mamkaeva, 1980; Esteve et al., 1983; Coder & Goff,

1986; Soo et al., 2015; Bagwell et al., 2016; Ganuza et al., 2016). At a commercial scale production, the bacterial predation (e.g. V. chlorellavorus ) reduces the productivity significantly and makes the process costly and difficult to address (Ganuza et al., 2016). Most of the existing treatment methods apply chemical agents (Bagwell et al., 2016; Ganuza et al., 2016) which are proven effective at small scale, but might present risks such as pollution, prohibitive cost, and less feasibility, especially at large scale in open ponds. Thus, alternatives should be investigated to provide stakeholders with more effective options in terms of contaminants control strategies.

Vampirovibrio chlorellavorus is an obligate aerobic bacterium (Soo et al., 2016) and

Chlorella species tolerate anoxic conditions (Kobayashi et al., 1982). Based on the assumption

that a relationship between a specific bacterium and a specific alga can be controlled by modifying the environmental growth parameters, we suggest studying the impact of deoxygenation-aeration cycling on the growth of both V. Chlorellavorus and C. sorokiniana to assess whether deoxygenation could disrupt the bacterial predation process while keeping C. sorokiniana undergoing a normal growth cycle.

Deoxygenation of the culture can be achieved by sparging pure nitrogen gas (Butler et al.,

1994). This cost-effective and environmentally friendly practice (Butler, Schoonen & Rickard,

1994; Barnhart, 1995; Alatossava, Gursoy & Alatossava, 2010) is feasible in closed bioreactors and ARID raceway, which is a suitable reactor for the deoxygenation process because algae is stored in a deep canal at night.

The technical aspect and cost estimation of the suggested deoxygenation method were assessed for adequate application at different cultivation scales. An outdoor application was conducted in 150-L sealed translucent-polyethylene reactor, and then evaluated in a prototype

ARID raceway covering one-acre (0.4 ha) surface area.

1.3. Hypotheses

1.3.1. Hypothesis 1

Effective V. chlorellavorus pathogen population size could be manipulated by periodically deoxygenating the algal culture to abate the potential attack on C. sorokiniana and prevent the culture collapse.

1.3.2. Hypothesis 2

Vampirovibrio chlorellavorus pathogenicity can be prevented with short period deoxygenation by sparging nitrogen gas at the beginning of the night followed by natural deoxygenation for the rest of the night. This approach is technically and economically feasible at commercial scale with onsite nitrogen generators.

1.4. Dissertation structure

The present research focuses on the contamination of C. sorokiniana by V. chlorellavorus at various cultivation scales (laboratory and outdoors), aiming to develop a deoxygenation control strategy and evaluate the technical and financial aspects of the deoxygenation practice with nitrogen gas sparging.

Chapter 2 describes the commonly used deoxygenation methods and the impact of deoxygenation treatment on the algae culture and associated microorganisms. In relation to chapter

2, an unprecedented control strategy applying deoxygenation-aeration cycling to prevent or reduce

V. chlorellavorus predation while keeping C. sorokiniana culture within a normal growth cycle and harvestable biomass is developed in appendix A: “Application of deoxygenation-aeration cycling to control the predatory bacterium Vampirovibrio chlorellavorus in Chlorella sorokiniana cultures”.

The primary goal of this section (Appendix A) was to evaluate the control of V. chlorellavorus with deoxygenation-aeration cycling. The first objective was to determine the tolerance of pathogen-free C. sorokiniana to various deoxygenation cycle times. The second objective was to test and determine the efficacy of cyclical deoxygenation-oxygenation regimes to

limit the growth of V. chlorellavorus to sub-pathogenic levels in C. sorokiniana suspension cultures, based on the inability of V. chlorellavorus to grow anaerobically, whereas C. sorokiniana can thrive under either aerobic or anaerobic conditions. The final objective was to identify an appropriate deoxygenation cycling regime and evaluate the effect of deoxygenation on V. chlorellavorus infection of the algal test host, with respect to culture phenotypes, biomass production, and the extent of V. chlorellavorus infection, quantified by real time polymerase chain reaction (PCR) amplification. The hypothesis is that effective V. chlorellavorus population size, below a specific threshold where attack and infection occur, could be manipulated by periodically deoxygenating the culture. A quantitative PCR assay was used to monitor V. chlorellavorus levels in aerobic and deoxygenated co-cultures to determine a preliminary threshold apparently conducive to pathogenicity.

Chapter 3 discusses the technical aspects of deoxygenation by nitrogen sparging and cost assessment of the deoxygenation process in the algae cultivation systems. Gas transfer equations are presented, and benefits and drawbacks of each deoxygenation method are included. Appendix

B is a detailed investigation of “Technoeconomic assessment of deoxygenation for control of

Vampirovibrio chlorellavorus in Chlorella sorokiniana culture”. The first section in appendix B deals with the impact of continuous aeration on pathogen-free C. sorokiniana and co-cultures of

C. sorokiniana and V. chlorellavorus , the second section compares deoxygenation application times, and the third section assesses the technical feasibility and the cost of deoxygenation treatment with nitrogen gas sparging.

The overall purpose of the study in appendix B was to evaluate the cost and efficacy of a short deoxygenation cycle at the beginning of the night (dark period) followed by natural

deoxygenation for the rest of the night. The first objective was to apply this process in laboratory experiments with C. sorokiniana co-cultured with V. chlorellavorus . The second objective was to apply the deoxygenation process in outdoor reactor with pathogen-free C. sorokiniana monoculture to evaluate the technical feasibility of the deoxygenation method, calculate the required mass of nitrogen gas per unit volume of algae culture at small and commercial scales, and estimate the cost of the deoxygenation process.

Appendix C “Data acquisition and programming instructions” describes the data loggers and programs with instructions used to monitor dissolved oxygen and nitrogen sparging, in the laboratory and outdoor algae cultivation reactors.

Appendix D “Standard Operating Procedures” describes media recipe, algae growth measurement, and Vampirovibrio chlorellavorus filtration and suspension process.

Appendix E “Interaction between V. chlorellavorus , ciliates and C. sorokiniana ” is suggested as a potential topic for further investigation. Contamination by ciliates occurred in some experiments of the present research, and since contamination of C. sorokiniana by ciliates is very common, especially in open raceways, some preliminary data are included in Appendix E for future consideration.

Chapter 2

Deoxygenation in algae cultivation systems

2.1. Introduction

In algae cultivation systems, dissolved oxygen (DO) is generated naturally by algae photosynthesis as a byproduct, or artificially with aeration (air injection, mixing, direct contact with the atmosphere) and made available for respiration to algae and associated living organisms.

The DO is partially transferred back to the atmosphere, or trapped in the cultivation system.

Environmental parameters such as temperature, salinity, pH, and atmospheric pressure have a significant impact on DO fluctuations. Although high DO concentrations can be a good indicator of a healthy algae culture, DO excess in the system could inhibit algae growth and reduce productivity (Weissman, Goebel & Benemann, 1988). Nighttime algal biomass loss can be reduced by managing DO at low levels (Edmundson & Huesemann, 2015). However, low DO could be a growth limiting factor for other associated living organisms, thus appropriate management of DO levels is necessary.

2.2. Dissolved oxygen measurement methods

Various methods for monitoring DO levels in water or algae culture are available, including iodometry (Navoc et al., 1988), colorimetry, titrimetry, polarography (Hitchman, 1978; Clean

Water Team (CWT), 2004), amperometry (Jalukse et al., 2004), luminescent-based sensor (Jin et al., 2013), spectrophotometry (Labasque et al., 2004), and optical DO-sensors (Li et al., 2015).

Advantages, limits, and field of application of each method are described in the indicated

respective references. A suitable method will depend on the type of application in respect to laboratory or field operations.

2.3. Deoxygenation approaches

Elimination of DO from water can be achieved physically, chemically, or biologically (Ito et al., 1998; Moon et al., 1999; Li et al., 2001; Van der Vaart et al., 2006; Shao et al., 2008; Karimi et al., 2011). These approaches also apply to deoxygenation in algae cultivation systems.

Deoxygenation in closed algae reactors should not be difficult, especially during the nighttime

(dark period). However, during the daytime (light period) photosynthesis can impede the effect of deoxygenation and make the process strenuous.

Each deoxygenation method has specific advantages and drawbacks; therefore, appropriate selection criteria including practicality, cost-effective, and eco-friendly aspects should be considered. The use of nitrogen gas sparging to remove DO from water is proven cost-effective and environmentally friendly (Butler, Schoonen & Rickard, 1993; Barnhart, 1995; Ugwu, Aoyagi

& Uchiyama, 2007; Alatossava, Gursoy & Alatossava, 2010). Thus, this approach was applied in the present study to deoxygenate the algae culture, during dark periods. Fig. 2.1 illustrates the impact of nitrogen sparging on DO concentrations in C. sorokiniana culture with a replicated treatment compared to an untreated culture control. During the dark period, nitrogen gas sparging in the treatment decreased DO to zero, whereas in the control a part of DO was consumed by algae respiration, causing the concentrations to decline to near saturation level. During the light period,

DO concentrations increased due to photosynthesis, indicating normal algae growth which is also reflected in the increase of ash free dry weight (AFDW) biomass.

Fig. 2.1. Dissolved oxygen removal with nitrogen gas sparging in Chlorella sorokiniana culture.

Treated culture was subjected to 2-hr deoxygenation alternated with 2-hr aeration during the dark period. The control culture was left without aeration nor deoxygenation. Growth measurement based on ash free dry weight (AFDW) biomass.

2.4. Deoxygenation impact on microalgae and associated microorganisms

Dark respiration and photosynthesis are important activities for autotrophic microalgae to perform their metabolism and grow. During light period, microalgae perform photosynthesis to transform carbon dioxide into carbohydrates and provide oxygen as a byproduct (Garcia-Camacho et al., 2012; Masojídek, Torzillo & Koblížek, 2013). Microalgae are not the only source of DO in the cultivation system; artificial aeration or direct gas exchange with the atmosphere (e.g. mixing in open reactors) can create balanced DO levels in the algae cultivation systems. In addition to

outgassing from the cultivation system into the atmosphere, some of the DO is consumed by microalgae when dark respiration occurs, and by obligate or facultative aerobic microorganisms which thrive in association with microalgae. A partial or complete DO depletion in microalgae cultivation system can lead to some growth stress on both microalgae or other associated microorganisms, especially obligate aerobes. Therefore, deoxygenation practices should be carefully applied to avoid harmful consequences. For instance, the present study focuses mainly on the application of deoxygenation to control V. chlorellavorus in association with C. sorokiniana culture, which could be an issue for the nontargeted beneficial coexisting microorganisms.

However, C. sorokiniana can grow axenically (Ganuza et al., 2016) and tolerates low DO concentrations (Kobayashi et al.,1982), and V. chlorellavorus is an obligate aerobic bacterium, thus dark period deoxygenation is an appropriate control strategy. A deoxygenation example is presented in Fig. 2.2. Deoxygenation treatment (TR) was applied for 1-hr by sparging nitrogen gas during the dark period. The control (C) was continuously aerated during the dark period and did not receive any deoxygenation. In the treatment (TR), DO levels remained high during the light period and decreased to near zero in the dark with a slight decrease towards the end when the algae growth entered a stationary phase. However, DO never decreased to zero in the control (C) during the dark period, but dropped drastically during the light period at the end when the algae culture collapsed. These results show the damaging impact of V. chlorellavorus on C. sorokiniana under a continuous oxygenation (control), on one hand, and the efficacy of deoxygenation treatment in preventing C. sorokiniana from collapsing, on the other hand. The results imply that V. chlorellavorus attack was inhibited by low DO levels in the system.

Fig. 2.2. Dissolved Oxygen (DO) and ash free dry weight (AFDW) of Chlorella sorokiniana co- cultured with Vampirovibrio chlorellavorus . Treatment (TR) was subjected to deoxygenation with

1-hr nitrogen sparging during the dark period. Control (C) was continuously aerated during the dark period.

Chapter 3

Technical evaluation and cost estimation of the deoxygenation process

3.1. Introduction

Dissolved oxygen concentrations in aqueous systems can be controlled with aeration or deoxygenation depending on the intended purpose. These converse processes require assessment of the overall oxygen transfer coefficient (K La) to define appropriate designs and suitable equipment. In the case of aeration (e.g. wastewater treatment) (Taricska et al., 2009), KLa can be calculated with the following equations:

dC/dt = KLa (C ST – C) or (1)

ln [(C ST – C0) / (C ST – C)] = KLa Δt (2)

Plotting ln [(C ST – C0) / (C ST – C)] vs. time will result in a slope of KLa.

With: CST : Middepth oxygen saturation (ppm) at testing temperature T (°C), C 0: Oxygen concentration (ppm) at time t = 0, and C: Oxygen concentration (ppm) at a given time t.

-1 During the deoxygenation process, the oxygen depletion coefficient K La (min ) is calculated as the logarithmic slope of the oxygen deficit vs. time (Eq. 3)

KLa = [ln (Ci) – ln (Cf)]/Δt (3)

-1 with: K La: Oxygen depletion coefficient (min ), Ci: Initial oxygen concentrations (ppm) at testing temperature T (°C), Cf: Final oxygen concentration (ppm) at temperature T (°C), Δt: Time interval

(min),

In both cases, aeration or deoxygenation, KLa can be adjusted for standard temperature (Eq.

4):

20-T KLa20 = K La * (θ ) (4)

-1 with: KLa20 : Oxygen transfer/depletion coefficient (min ) at standard temperature (20 °C), and θ:

Temperature correction coefficient (1.024).

The standard oxygen transfer rate (SOTR) can be calculated with Eq. 5:

SOTR = KLa20 * CS20 * V (5)

-1 with: SOTR: Standard oxygen transfer rate (mg/min), KLa20 : oxygen transfer coefficient (min ),

CS20 : Dissolved oxygen concentration at saturation (mg/L) at standard conditions (20 °C and 1 atm), V = Volume of water (L).

3.2. Dissolved oxygen removal

Dissolved oxygen removal can be accomplished physically by heating, vacuuming, nitrogen gas sparging, microporous membranes permeation (Sinha & Li, 2000), chemically by adding sodium sulphite (Colombo, Berg, and Constant, 1998), or biologically by glucose oxidase

(Karimi et al., 2011). Considering drawbacks and limitations of each approach such as high energy use, risk of toxicity and pollution, prohibitive cost, and lack of practicality, careful selection of an appropriate method is crucial to make the process successful. for instance, using chemicals or heating might be suitable in water but not in algal cultures; however, nitrogen sparging appears to be an appropriate alternative to displace dissolved oxygen from both water and algae cultivation systems without adverse effects on the culture or the environment (Sawdon & Peng, 2014).

3.3. Deoxygenation with nitrogen gas sparging

Nitrogen (N 2) is an inert gas which can be used for multiple purposes such as food packaging (Kosiki & Itoh, 2002), storage and transportation (Alatossava, Gursoy, and Alatossava,

2009), freezing and cooling (Yanisko et al., 2011), and oxygen removal (Ivanova and Lewis,

2012), to name just a few. Sparging nitrogen gas to remove dissolved oxygen from water has been successfully applied (Butler et al., 1994; Barnhart, 1995; Sinha and Li, 2000; Sawdon & Peng,

2014). Nitrogen gas bubbling is also attractive because of its numerous benefits including low energy use, cost-effective, practicality, and environmentally friendly aspects. Thus, in the present study nitrogen gas sparging was applied to remove dissolved oxygen from C. sorokiniana culture and control V. chlorellavorus infection.

3.4. Technical considerations and cost estimation of the deoxygenation process

In general, the design of an optimized deoxygenation process with nitrogen gas sparging should consider appropriate injection flow rate, diffusion uniformity, reactor geometry, mixing intensity, and sparger location. Technically, deoxygenation is easier and faster with fine bubbles and pure nitrogen gas; however, it appears that the degree of purity of nitrogen gas does not show a significant difference in the remaining dissolved oxygen concentration in water treated with nitrogen gas containing less than 5 ppm oxygen, compared to nitrogen gas with less than 0.5 ppm oxygen (Butler et al., 1994). Higher nitrogen flow leads to higher deoxygenation rate (Barnhart,

1995), but long-time applications are not necessarily more effective (Butler et al., 1994).

The aforementioned general and technical aspects of the deoxygenation process with nitrogen sparging should be combined with a cost estimation and risk evaluation to include

investment (e.g. equipment, buildings, and infrastructure) and operational charges (e.g. energy, labor, delivery, and maintenance).

Nitrogen gas is abundantly available and thoroughly regulated for safe management

(Yanisko et al., 2011), which makes the nitrogen sparging approach reliable and attractive.

Nitrogen can be generated using multiple methods such as distillation, adsorption, or permeation, and stored in small volumes (containers), industrial volumes (large tanks), or provided directly from onsite generators (Ivanova et al., 2012).

Small nitrogen containers (e.g. 9in x 52in cylinders) might be convenient for small algae reactors cultivated for high value products, but in commercial reactors they are cumbersome and costly, thus not suitable. Instead, larger nitrogen tanks (e.g. 24in x 62in HPL-N2), or nitrogen generators (Myers & Froehlich, 2009) are more suitable because they are cost-effective and easy to manage.

Conclusion

Optimal algae productivity requires a continuous cultivation process, appropriate cultivation systems, and effective strategies to control biological contaminants. Chlorella sorokiniana is an important microalgae species, but highly sensitive to V. chlorellavorus predation under aerated conditions. The present research demonstrated that V. Chlorellavorus could be controlled using deoxygenation-aeration cycles.

One-hour deoxygenation treatment using nitrogen gas sparging at the beginning of the night followed by natural deoxygenation by algae dark respiration is suggested to be efficient to control V. chlorellavorus , and onsite nitrogen generators are suitable for deoxygenation of the algae culture at commercial scale in closed photobioreactors and ARID raceway, thus are herein recommended.

Further studies are needed to understand the specific timeframe, chemical signals and environmental conditions required by the predatory bacterium V. chlorellavorus to shift into a pathogenic mode, and develop resistance to anaerobic conditions. Also, more investigation on V. chlorellavorus "pathogenicity threshold" should be conducted to accurately assess the interaction of V. chlorellavorus- C. sorokiniana in the presence / absence of ciliates under the deoxygenation treatment.

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Appendix A

Paper submitted to Algal Research

Application of deoxygenation-aeration cycling to control the predatory bacterium

Vampirovibrio chlorellavorus in Chlorella sorokiniana cultures

S. Attalah a, * , P. Waller a, S. Steichen b, S. Gao c, C. C. Brown b, K. Ogden d, and J. K. Brown b a The University of Arizona, Department of Biosystems Engineering, Tucson, AZ 85721, USA. b The University of Arizona, School of Plant Sciences, Tucson, AZ 85721, USA. c Pacific Northwest National Laboratory, Richland, WA 99352, USA. d The University of Arizona, Department of Chemical and Environmental Engineering, Tucson, AZ 85721, USA.

* Corresponding author: [email protected] BE Department, Shantz Bldg. Rm. 403, Tucson, AZ 85721

Abstract : A previously untested approach was evaluated to enable management of the predatory bacterium, Vampirovibrio chlorellavorus , a pathogen of Chlorella sorokiniana , in suspension cultures grown in a laboratory test reactor. Because V. chlorellavorus is an obligate aerobic bacterium, whereas C. sorokiniana grows under aerobic and anaerobic conditions, deoxygenation of the culture was expected to be detrimental to the pathogen, but not to the algal host. The effect of deoxygenation on the uninfected (healthy) C. sorokiniana suspension cells, compared to the C. sorokiniana-V. chlorellavorus co-culture, was studied in relation to biomass, dissolved oxygen, ratio of C. sorokiniana to V. chlorellavorus DNA, and visual and light microscopic observations.

Preliminary experiments were conducted to test the effects of different deoxygenation-aeration cycling regimes on performance of V. chlorellavorus -free C. sorokiniana cultures. To an aerobic culture, pure nitrogen gas was introduced to create anoxic conditions, followed by the injection of ambient air to re-establish an aerobic environment. Under this repeated cycling regime, C.

sorokiniana was shown to tolerate the anoxic conditions for extended timespans that ranged from

2 to 8 hours over a 5-day test period. The analogous aerobic-anoxic cycling with the C. sorokiniana-V. chlorellavorus co-cultures resulted in ‘near-normal’ growth cycle and harvestable biomass, whereas the continuously-aerated (aerobic) co-cultures that were grown without the deoxygenation step in the cycle collapsed in 3 days. Visual and light microscopic observations revealed intact C. sorokiniana cells were present in the deoxygenated cultures, compared to the aerobically-grown, brown-colored algal cultures consisting of collapsed cells. Quantitative polymerase chain reaction analysis showed continuous increases in the ratio of V. chlorellavorus

(16S rDNA) to C. sorokiniana (18S rDNA) DNA in the aerated co-cultures, with greater increases during dark periods, while the pathogen-to-host DNA ratio in the deoxygenated co-cultures was relatively low and algal cells did not collapse, as would be expected following pathogen attack.

Keywords: anoxic, biomass, infection, nitrogen, quantitative polymerase chain reaction.

1. Introduction

Outdoor raceways expose algae cultures to contamination, which can cause cultures to collapse, reduce biomass productivity, and increase maintenance and re-inoculation costs [1].

Several strategies have been suggested to reduce contamination of algae cultures: salvage harvest

[2], pH shock treatment [3], use of chemical agents [4-9], iron limitation [10], physical disruption and removal [11], and biological control [12,13], with cost, side effects, and efficacy having determined the feasibility of each strategy.

Species of the chlorophyte genus, Chlorella , belong to the Trebouxiophyceae

(order, Chlorellales; family, Chlorellaceae), and are spherical or ellipsoidal unicellular species ranging from 2 to 15 microns in size [14]. A number of Chlorella species are cultivated for food, cosmetics, and vitamins, and some are of potential interest for producing biofuels because of their ability to produce 35 g/m 2 d-1 or more of biomass [15], tolerate a wide range of growth conditions

[16], can grow under both aerobic and anaerobic conditions [17], and have a specific growth rate as high as 5.9 d -1 [18]. However, several Chlorella species have been lucrative hosts of the predatory cyanobacterium, Vampirovibrio chlorellavorus , which parasitizes the algal host cells, leading to rapid death [19] and complete loss of harvestable biomass [20,21].

Vampirovibrio chlorellavorus [19,22,23] is a non-autotroph, obligate aerobic gram- negative cyanobacterium with a pleomorphic shape ranging from 0.3 microns (vibrios) to 0.6 microns (cocci) and lives attached to the surface of Chlorella spp . cells [19,24-26]. Under high temperatures, within several days after V. chlorellavorus presence has been detected in the

Chlorella culture [20], cells have been observed to clump, collapse and lyse [24], resulting in substantially reduced biomass [21,27]. With the recent increased interest in Chlorella cultivation for biofuel production, this predatory bacterial pathogen has become of a great concern to the industry [3]. In 1966, Mamkaeva [28] first reported an unidentified bacterium that attacked

Chlorella spp. In 1972, the bacterium was named Bdellovibrio chlorellavorus [22], and in 1980 the genus affiliation was amended to Vampirovibrio , resulting a name change to V. chlorellavorus

[23]. A hallmark of species within the genus, Bdellovibrio , is that they enter the periplasm of their hosts. However, V. chlorellavorus attaches to Chlorella microalgae and ruptures the cell wall, a tropism that might be associated with the particular cell wall properties of Chlorella host species

[24-26]. Predatory bacteria like V. chlorellavorus have been found to attach to their algal host using a (proposed) electron-dense pad, and to inject effectors, enzymes, and plasmid DNA into the algal host cell, leading to cell lysis and loss of the cellular content [25,26].

Experimental and natural host range studies have shown that V. chlorellavorus infects a number of Chlorella species, including C. kessleri , C. sorokiniana , and C. vulgaris , whereas species of Ankistrodesmus, Chlorococcum , Scenedesmus , and Scotiella were found to be non- hosts of V. chlorellavorus [19,20,25,26]. These findings are consistent with the observation that in experimental outdoor raceways, designed to establish optimal conditions for commercial cultivation of C. sorokiniana, the bacterial pathogen V. chlorellavorus has been associated with lysis of cells and death of C. sorokiniana [14,29] when the optical density was approximately ≥

1.0 at 750 nm [21,27].

Strategies have been developed to reduce losses in Chlorella algal reactors by limiting infection by the bacterial pathogen. An acid-induced drop in pH in reactors containing susceptible

Chlorella species co-cultured with V. chlorellavorus showed reduced rates of infection, extended duration of culture growth, and increased number of successive cultivation-runs [3]. Also, a treatment implemented to limit iron availability, resulted in decreased biomass ranging from zero to 9% in the iron-treated Chlorella cells infected with V. chlorellavorus , compared to cultures with no iron limitation, which showed a 72% decrease in biomass, indicating that a shortage of iron could limit pathogen attack [10].

The V. chlorellavorus is a non-autotroph obligate aerobe [26]; however, Chlorella species can grow under aerobic or anaerobic conditions [17]. Thus, deoxygenation was considered as an option for limiting the rates of V. chlorellavorus infection that would not significantly reduce the

growth of C. sorokiniana . However, although anoxic-oxic cycles might be expected to negatively affect V. chlorellavorus growth, such conditions could possibly alter the composition of beneficial microorganisms associated with the algal culture. Because C. sorokiniana is known to be capable of growing axenically [3] and can tolerate anaerobic conditions [17], the proposed hypothesis was that C. sorokiniana growth might not be significantly reduced when cultured in a partially- deoxygenated environment, and at the same time, such conditions might be detrimental to the mutualistic, non-pathogenic phycosphere bacterial community. To guard against possible negative effects of deoxygenation on the non-pathogenic bacteria, algal cultures were aerated with ambient air during the dark period after deoxygenation cycles.

Nitrogen gas sparging is cost-effective and environmentally-compatible [30-33] for deoxygenating water. Deoxygenation with the timed injection of nitrogen gas can be accomplished in closed bioreactors, and also in the deep canal of the testbed utilized here, referred to as the Algae

Raceway Integrated Design (ARID) [34,35], by covering the canal to enhance the effects of deoxygenation and/or aeration cycling.

The goal of this study was to identify the best cyclical deoxygenation-oxygenation regime in which the growth of V. chlorellavorus would be reduced to sub-pathogenic levels in suspension cultures of the algal host, C. sorokiniana , based on the inability of V. chlorellavorus to grow anaerobically, and ability of C. sorokiniana to thrive in aerobic or anaerobic environments. The first objective was to determine the tolerance of C. sorokiniana , DOE1412 [36], to various deoxygenation cycle times. The second objective was to define a deoxygenation cycling regime under which the effects of deoxygenation could be tested to evaluate its potential to abate V. chlorellavorus infection of C. sorokiniana , measured as algal culture phenotype, biomass

production, and V. chlorellavorus titer determined by quantitative polymerase chain reaction

(PCR) amplification. The rationale for the study was based on the hypothesis that the effective V. chlorellavorus pathogen population size, below a to-be-determined threshold, at which infection occurred, could be manipulated by periodically deoxygenating the algal culture. Finally, in this study, a quantitative PCR assay developed previously [27] was used to monitor V. chlorellavorus accumulation under aerobic and deoxygenated conditions, to determine the relative threshold conducive to pathogenicity.

2. Materials and Methods

2.1. Pathogen-free cultures and co-cultures

The C. sorokiniana monoculture was cultivated in modified BG-11 solid growth medium

-1 -1 -1 -1 - containing 0.1 g L urea, 0.012 g L MgSO 4, 0.035 g L NH 4H2PO 4, 0.175 g L KCl, 0.005 g L

1 FeCl 3 [37], with trace minerals [38]. Individual algal cells were selected and transferred from the solid media and cultivated in 50 mL, 250 mL, then in 20 L carboy reactors with continuous agitation by injection of ambient air. All cultures were grown under 200 µmole/m 2 s-1 light intensity with 16-h and 8-h light and dark cycles. The V. chlorellavorus culture used in this study was isolated from a co-culture of V. chlorellavorus and C. sorokiniana , DOE1412 [36], from the

ARID raceway [35] located at the University of Arizona, Tucson, AZ, USA. The bacterium was isolated by filtration through a 2 µm Whatman filter, and then inoculated to a laboratory culture of

C. sorokiniana to establish a fresh co-culture. Isolation, filtration, and co-culturing of V. chlorellavorus are described previously [27]. A one-liter co-culture was maintained by the weekly transfer of 200 mL of the co-culture to 800 mL fresh C. sorokiniana cells in modified BG-11

medium. The co-culture was grown in 2-L flasks in an incubator (I-36LL, Percival Inc, USA) at

200 µmole/m 2 s-1 light intensity with 16-h and 8-h light and dark cycles at 24 °C.

2.2. Deoxygenation-aeration cycling experiments

Four experiments were carried out to assess the effects of deoxygenation-aeration cycling impact on V. chlorellavorus -free C. sorokiniana and co-cultures of C. sorokiniana -V. chlorellavorus (Table A.1). Each experiment consisted of 16-h light period and 8-h dark period.

The first experiment (Exp. 1) with 4-h deoxygenation included 5 days with an alternating 2-h deoxygenation and 2-h aeration cycles, during the dark period, followed by alternating 4-h deoxygenation and 4-h aeration during the dark period for 4 days. Nutrients were replenished at the end of the 2-h deoxygenation-2-h aeration cycles experiment, in the dark period of day 5. The second and third experiments (Exp. 2 and 3), with 6-h and 8-h deoxygenation, respectively, were carried out for a duration of 5 days.

The deoxygenation tolerance of C. sorokiniana cultures was determined under conditions involving the absence of V. chlorellavorus (experiments 1-3). The experimental design included a nonaerated algal culture (C1, positive control), which was expected to have high algal biomass loss, an aerated negative control (C2), which was expected to have a normal growth, and a deoxygenation-aeration cycling treatment (TR). Because of the absence of aeration, the C1 control was subjected to a natural deoxygenation-oxygenation treatment due to C. sorokiniana dark- respiration and photosynthesis cycles; whereas, the C2 control consisted of a continuously-aerated culture and no deoxygenation, during dark period. The treatment TR had three replicates (TR1,

TR2, and TR3) in Exp. 1, and two (TR1 and TR2) in Exp. 2 and Exp. 3. The nonaerated C1 control with natural deoxygenation by dark respiration proved to be ineffective at reducing the dissolved

oxygen concentration to anoxic levels for experiments 1-3. As a result, no additional experiments were conducted with C1.

The fourth experiment (Exp. 4) with C. sorokiniana-V. chlorellavorus co-cultures and 2-h deoxygenation-2- h-aeration dark period cycling treatment considered three replicates, 4-1 to 4-3.

The deoxygenation-aeration treatment was replicated twice (TR1 and TR2), along with the aerated negative control, C2, which was replicated, and referred to as C2A and C2B.

Table A.1: Deoxygenation-aeration cycling experiments schedule.

Description Deoxygenation - aeration Schedule cycles 1- Application of deoxygenation- Exp. 1 aeration cycles, during the dark Alternating 2-h / 2-h From 2/23 to 2/27/17 period, with V. chlorellavorus -free Alternating 4-h / 4-h From 2/28 to 3/3/17 C. sorokiniana cultures. Exp. 2 Nonaerated positive control 6-h deoxygenation /2-h From 3/7 to 3/11/17 subjected to natural aeration deoxygenation-oxygenation process Exp. 3 (dark-respiration and 8-h deoxygenation From 3/17 to 3/21/17 photosynthesis), and continuously

aerated negative control during dark period.

2- Application of dark-period Replicate 4-1 deoxygenation-aeration cycles with Exp. 4 From 5/23 to 5/30/17 C. sorokiniana -V. chlorellavorus co- Alternating Replicate 4-2 cultures, and a continuously 2-h deoxygenation / 2-h From 6/15 to 6/20/17 aerated negative control. aeration Replicate 4-3 From 6/26 to 7/1/17

2.3. Inoculum preparation and experimental media

Prior to the initiation of experiments 1-3, the C. sorokiniana monoculture inoculum was diluted with distilled water, from a starting optical density of 5.0, at a wavelength of 750 nm

(OD750) measured with the Spectronic 200 spectrophotometer (Thermo Fisher Scientific, USA), to an OD750 of 1.0. Nutrients were replenished according to the methods (media) described above.

A volume of 3.5 L of the diluted C. sorokiniana culture was transferred into each flask at the beginning of each experimental replicate.

In Exp. 4, when the culture density in the carboy reached an OD750 of 1.0, the infected co- culture was first centrifuged (7000 rpm for 5 min) to pellet algal cells, and the supernatant was filtered (2-µm Whatman filter) to remove the C. sorokiniana cells and debris. Subsequently, 1 L of the V. chlorellavorus filtrate was added to 17 L of the V. chlorellavorus -free C. sorokiniana culture, to a final OD750 = 1.0. Nutrients were added to the diluted co-culture as described above, using the modified BG-11 recipe. As for experiments 1-3, a 3.5-L volume of the co-culture and nutrients were transferred to each flask (TR1, TR2, C2A, and C2B) to initiate the experiments.

2.4. Algal suspension culture growth assessment

Algal growth was determined based on the OD750 at the end of each light and dark period, and used to calculate the ash free dry weight (AFDW) in g L-1 (Eq. 1). The spectrophotometer was calibrated against the AFDW of C. sorokiniana culture at the beginning and end of Exp. 1, and a correlation between OD750 and AFDW was established. Algae samples were diluted five-fold, by the addition of 0 to 4 mL increments of distilled water to 1 mL of algae from the culture to achieve

1 to 5-fold dilutions, and cover the range of optical densities occurring from the early to the late

growth stages. The AFDW was measured by filtering algae samples on Whatman glass microfiber filters, drying at 75 ‐ overnight and ashing at 550 ‐ for 4 h. A linear correlation (Eq. 1) between

OD750 and AFDW was calculated as:

AFDW = 0.1617 * OD750 (1) with R 2 = 0.97.

Daily biomass gain and loss of C. sorokiniana were calculated [39,40] as the difference in biomass over each 16-h light period and 8-h dark period, respectively. A daily adjustment of water level in each flask was performed by adding an appropriate volume of distilled water. The student’s t-test, at α = 0.05 level of significance was carried out using the data analysis tool pack in Microsoft

Excel (2016) to determine the statistical significance of the differences observed in biomass concentration on specific days and for treatments compared to the experimental controls.

In Exp. 4, indicators of V. chlorellavorus infection resulted in a color change from green to yellow or brown, emittance of undesirable odor, the accumulation of foam on the culture surface, and presence of clumped or lysed cells, observed by light microscopy [24] at 400x magnification.

In addition, the DNA ratio of V. chlorellavorus (16S rDNA) to C. sorokiniana (18S rDNA) was used to determine the extent of accumulation of V. chlorellavorus . The estimated number of V. chlorellavorus cells present in total DNA isolated from the C. sorokiniana-V. chlorellavorus culture was estimated using the standard curve established previously for the real-time quantitative

PCR (qPCR) assay, using V. chlorellavorus -specific primers [21,27].

Throughout the study, the dissolved oxygen (DO) level was recorded at 5-min intervals using DO sensors (DO1200, Sensorex Inc., Garden Grove, CA), connected to the data logger

(CR300, Campbell Scientific, Logan, UT).

2.5. Real time quantitative polymerase chain reaction for quantification of V. chlorellavorus

The concentration of V. chlorellavorus cells was determined by qPCR analysis of the 16S ribosomal RNA gene (rDNA, single copy) as previously described [27]. For each time point, a 1- mL sample was collected, and pelleted using a microfuge, at 7500 x g for 10 min. The biomass was stored at -20 °C, prior to total DNA isolation using the cetyltrimethylammonium bromide

(CTAB) method [41]. The DNA was used as a template in the multiplex qPCR for quantification of V. chlorellavorus and C. sorokiniana 16S and 18S rDNA, respectively. The quantitation cycle

(Cq) values were converted to gene copy number using the linear regression equation, based on the standard curve values for 10-fold serially-diluted plasmid containing the target gene amplicon

[27]. The gene copy number for V. chlorellavorus and C. sorokiniana , respectively, were normalized [27], and expressed as the bacterial to algal cell ratio.

2.6. Experimental reactor design and laboratory apparatus

To investigate the effects of the deoxygenation-aeration cycling on C. sorokiniana and V. chlorellavorus , algal culture and co-culture (bacterium and alga) were grown in reactors consisting of five 4-L Erlenmeyer flasks, with incubation in a digital incubator-refrigerator-shaker

(Innova4330, New Brunswick Scientific Inc., Edison, NJ) (Fig. A.1). The flasks held 3.5-L volume of algal suspension culture to ensure that the sensors and diffusers could be housed inside the reactor vicinity. Pure nitrogen gas UN1066 was bubbled to deoxygenate the algal cultures and co- cultures [30-33], and ambient air was injected to re-establish the oxygenation. Clear tygon tubing equipped with check valves were used to connect aquarium air stones in the flasks to the air pumps and gas tanks containing nitrogen and carbon dioxide, respectively, through the vent located on

the side of the incubator. Rubber stoppers with holes drilled to allow entry of the tubes were added to seal the flasks, and to provide support for the sensors and diffusers. The Milwaukee pH controllers (MC110, Milwaukee Instruments Inc., Rocky Mount, NC) were used to regulate the pH of the culture at a pH ranging from 7.0 - 7.5, which was accomplished by triggering the injection of pure carbon dioxide at the high setpoint (pH 7.5). The cultures were mixed by shaking in the Innova4330 incubator at 80 rpm, and 24 °C. Two grow-light panels (45W LED, Watt Shine,

China) were attached to the incubator top to provide light to the culture. The heat produced by the lights caused the incubator temperature to fluctuate from 24 to 30 °C during the light and dark cycles.

Fig. A.1. Experimental reactor design and apparatus. A) General view of the experimental apparatus arrangement. B) Experimental equipment and Campbell Scientific (CS) data logger for

deoxygenation-aeration cycling management. C) Grow-light panels added to the incubator, and pH controllers used to regulate pH in the suspension culture. D) Light intensity measured with

MP-200 Apogee silicon cell pyranometer placed at equidistance from one another to achieve consistent deoxygenation-aeration cycling during treatment 1-3 (TR1, TR2, and TR3), with the nonaerated positive control (C1), and aerated negative control (C2). E) Example of an experimental flask with tubing, sensors, and rubber stopper. F) Arrangement of the 1-gallon

Erlenmeyer flasks containing the Chlorella sorokiniana suspension cultures placed inside the incubator.

The light intensity was measured for each of the flasks using a cell pyranometer (MP-200

Apogee silicon, Apogee instruments Inc., Logan, UT) at a height of 16 cm above the incubator platform, corresponding to the culture height in the flasks, in W m-2, converted to µmole/m 2 s-1

(Table A.2), assuming a sunlight radiation conversion [42,43]. Even though the center flask had a similar light intensity measurement, it received light from all sides, leading to a higher growth rate in Exp. 1. Consequently, in Exp. 2-4, the position of the grow-light panel was adjusted, and the light intensity was recorded for all of the experiments, confirming consistency between the experiments. Because the growth rate in the center flask, TR3, resulted in inconsistent biomass production, so the results from TR3 were only considered in Exp.1.

Table A.2: Light intensity measurements recorded for each flask during the experimental runs

Light Intensity

W m-12 (µmole/m 2 s-1)

Nonaerated Aerated Treatment Treatment Treatment

Experiments control control replicate 1 replicate 2 replicate 3

C1 C2 TR1 TR2 TR3

Exp. 1 36 (164) 36 (164) 37 (169) 34 (155) 41 (187)

Exp. 2 - 4 39 (178) 39 (178) 36 (164) 35 (160) 40 (183)

3. Results and Discussion

3.1. Effect of deoxygenation cycling on V. chlorellavorus -free C. sorokiniana

The effect of deoxygenation-aeration cycles on the growth of V. chlorellavorus -free C. sorokiniana was monitored in the treated cultures (TR1, TR2, and TR3) and compared to the nonaerated positive control C1, and the aerated negative control C2 (Exp. 1-3).

Based on the fluctuation in AFDW biomass for Exp. 1, grown with alternating 2- and 4-h deoxygenation-aeration cycle, the algal growth rate in the TR3 replicate was considered inconsistent with respect to biomass production in the other two replicates (Fig. A.2.A). This artifact occurred presumably because all sides of each of the middle flasks received greater exposure to light, compared to the outer two flask positions. Although the positions of the light panels were adjusted accordingly, for Exp. 2 (6-h deoxygenation), the TR3 replicate showed higher than expected biomass production, thus data for the TR3 replicate were excluded from the analyses. As with the 4-h deoxygenation treatment (Exp. 1), in Experiments 2 (6-h deoxygenation)

and 3 (8-h deoxygenation), the AFDW was slightly lower in TR1 and TR2 compared to the controls C1 and C2 (Figs. A.2-B and A.2-C, respectively).

The maximum AFDW was approximately 0.4 g/L over a four-day growth following inoculation with C. sorokiniana monoculture. The algae growth was probably hampered by light limitation and nutrient depletion [37, 44-47]. Throughout the three experiments (Exp. 1-3), macroscopic and light microscopic (400x magnification) viewing of the algal cultures in the flasks showed no evidence of stress such as emittance of undesirable odors, foam accumulation in the flasks, color alteration from green to yellow or brown, and clumping or lysed cells.

Fig. A.2. Daily ash free dry weight (AFDW) biomass (lines with markers), and biomass gain during the light period and biomass loss during the dark period (markers) in experiments 1-3 with

Vampirovibrio chlorellavorus -free Chlorella sorokiniana : A-C) 4-, 6-, and 8-h deoxygenation treatments, respectively.

The mean of biomass values for the replicated, nonaerated positive control (C1) and aerated negative control (C2) for Exp. 1-3, were calculated and compared statistically with the mean values for the treatments (TR) in Exp. 1-3 using the student’s t-test. The statistical differences were not significant (p > 0.05) between C1, C2, or any of the treatments (inset in Fig. A.3) at day 4 which marks the onset of the stationary phase, in the present experiments. Based on the standard deviation and error bars (inset in Fig. A.3), the variability in AFDW biomass (g/L) between experiments (1-

3) was trivial in C1 and C2, ranging from 0.389-0.394 with an average of 0.381 and 0.372-0.390 with an average of 0.376, respectively. However, in the replicated treatment (TR), the biomass varied moderately between experiments (1-3) at the beginning of the treatment (day 1 and day 2), but then the biomass difference decreased at the end of day 4, ranging from 0.362-0.374 with an average of 0.368; 0.330-0.354 with an average of 0.342; and 0.336-0.377 with an average of 0.356, in 4-, 6-, and 8-h treatments, respectively. Although there was no statistical difference between the controls and treatments, the 2x2 (4-h) treatment displayed a better trend (biomass variation was within the same range as the controls), compared to the 6 and 8-h treatments, which showed lower biomass and greater fluctuations between the dark and light periods. Thus, the 2x2 (4-h) deoxygenation-aeration appears to be the best optimized treatment as applied in the present study.

Fig. A.3. Comparison of the average ash free dry weight (AFDW) in experiments 1-3 with

Vampirovibrio chlorellavorus -free Chlorella sorokiniana cultures, including nonaerated positive control (C1), aerated negative control (C2), and 4-, 6-, and 8-h deoxygenation treatments (TR).

Error bars represent one standard deviation.

The dissolved oxygen concentrations (DO) in experiments 1-3 (Fig. A.4) increased consistently with aeration and photosynthesis. The negative control C2 received constant aeration during the dark period to maintain the DO level at above saturation, i.e. at 10 ppm, and as a result it reached a higher DO, compared to the other flasks exposed to the same light regime. The 4-h deoxygenation treatment in Exp. 1 and the positive control C1 had similar light period DO levels.

Both ended the dark period at approximately 6 ppm and reached almost 15 ppm during the light period. The 6-h and 8-h deoxygenation treatments (Exp. 2 and 3, respectively) decreased to zero

DO concentrations after 0.5 h of deoxygenation and reached approximately 10 and 12 ppm during the light period, respectively. There was a general decline in light period DO over the four days in

the 8-h treatment, which might be due to a continuous low DO levels during the dark periods; however this did not cause a significant difference in biomass production compared to the other treatments. Light microscopy observations made at the end of the experiments (1-3) showed that

C. sorokiniana cells were green and intact; however, it appeared that algae had not undergone cell division, indicating the possibility that light limitation and/or nutrient deficiency had occurred.

Fig. A.4. Dissolved oxygen in experiments 1-3 with Vampirovibrio chlorellavorus -free Chlorella sorokiniana culture: nonaerated positive control (C1), aerated negative control (C2), and deoxygenation treatments (TR) with various deoxygenation times (4-, 6-, and 8-h) during dark period.

In summary, the experiments reported here using V. chlorellavorus -free C. sorokiniana cultures show that C. sorokiniana was capable of tolerating anoxic conditions for 2x2-, 4-, 6-, and

8-h, respectively. This indicated that by imposing anoxic conditions of as much as 8-h in duration, growth and attack by the pathogenic bacterium, V. chlorellavorus , could be abated. However, considering the general trend in biomass production and the prospective high usage of the resources required for abatement, the shortest possible deoxygenation-aeration cycling of 2x2 appeared to be the most suitable, and so these conditions were examined in Exp. 4. Additional deoxygenation scenarios, perhaps with shorter application times, may be of interest for further optimizing the process.

3.2. Impact of deoxygenation-aeration cycling on V. chlorellavorus infection

The 2-h deoxygenation-2-h aeration cycling treatment was tested during dark periods in

Exp. 4 to control V. chlorellavorus infection in the C. sorokiniana-V. chlorellavorus co-cultures.

The algal growth and ratios of V. chlorellavorus to C. sorokiniana in the treated culture and the aerated control were compared. The experiment was carried out using a 16-h light period followed by 8-h dark period with 2x2-h deoxygenation-aeration cycles in replicated treatments (TR1 and

TR2), and aerated negative control, duplicated as C2A and C2B.

The average DO concentrations in Exp. 4 ( C. sorokiniana-V. chlorellavorus co-cultures ) and Exp 1 ( V. chlorellavorus -free C. sorokiniana culture) were compared. Even during the first

16-h light period, DO concentrations were significantly different in both treatment and control

(Fig. A.5). Nitrogen injection lowered the DO concentrations to zero for each 2-h deoxygenation treatment period, and air injection reestablished the aerobic conditions. Although both C2A and

C2B were continuously aerated, during the 8-h dark periods, they had lower average DO concentrations (6 ppm) during the dark period than in the aerated negative control (C2) in Exp. 1

(10 ppm), which was probably due to the consumption of oxygen by V. chlorellavorus . During the

16-h light periods, average DO levels in C2A and C2B (Exp. 4) were much lower (10 to 12 ppm) than in C2 (Exp. 1) which ranged from 20 to 25 ppm. Similarly, average DO concentrations in the treatments were much lower during the light periods in Exp. 4 than in Exp. 1. Even with lower oxygen in Exp. 4, the algal biomass during the growth phase was similar in all flasks in both Exp.

1 and 4. This result provided a further indication that the lower DO concentrations in Exp. 4 were probably due to the consumption of DO by V. chlorellavorus . At the end of the experiments, DO levels dropped significantly in the controls due to V. chlorellavorus infection and the death of C. sorokiniana (Fig. A.5). The dissolved oxygen levels also dropped somewhat in these treatments during the light period, but not by as much as in the controls.

Fig. A.5. Compared average dissolved oxygen between Vampirovibrio chlorellavorus -free

Chlorella sorokiniana culture (Exp.1) and C. sorokiniana -V. chlorellavorus co-cultures (Exp. 4).

During the 8-h dark periods, negative controls (C2) were continuously aerated, and treatments

(TR) involved 2-h deoxygenation-2-h aeration cycles. The control C2 in Exp.4 represents an average of the aerated control duplicates, C2A and C2B.

Biomass concentration and DNA ratios of V. chlorellavorus to C. sorokiniana were measured for three replicates (replicates 4-1 to 4-3) in Exp. 4 (Fig. 6-A-C), respectively. The 2-h deoxygenation-2-h aeration cycling treatment (TR1 and TR2) and aerated negative controls C2A and C2B had normal growth during the first three days, but after the fourth light period C2A and

C2B entered a death phase while TR1 and TR2 entered a stationary phase (Fig. 6-A). The death phase in the negative controls in replicates 4-2 and 4-3 began after the third light period (fig. A.6.B-

C), whereas the growth of the treated cultures continued to the stationary phase. Thus, the deoxygenation process limited the impact of V. chlorellavorus and kept C. sorokiniana growing for a longer time compared with the aerated controls.

Quantitative polymerase chain reaction (qPCR) measurements were taken after each light and dark cycle during the growth phase (1), the stationary phase (2), and death phase (3) (Fig. 6).

In the first replicate (4-1), Initial ratios after the first light period were in the range of 0.001 or lower. The ratios remained approximately the same after the first dark period (Fig. A.6.A-B). On day 2, both controls showed significantly increased ratios, whereas the treated co-cultures ratios remained relatively low. Ratios in C2A and C2B increased to a greater extent during the dark

(large markers) compared to light periods (small markers) on day 2 and 3 (Fig. A.6.A). DNA ratios over the dark period of day 2 increased sixfold in C2A, and fourfold in C2B, but remained almost the same in TR2, and below detection in TR1. Similarly, during the dark period of day 3, the ratios

increased sixfold in C2A, and fourfold in C2B, but only twofold in TR2, and they remained below detection in TR1.

Based on the biomass measurements, a quick collapse of the co-cultures was observed on day 4 in the replicate 4-1 controls, due to infection by V. chlorellavorus . During the dark period on day 4, C2A and C2B experienced much greater biomass loss than what is normal without infection, whereas the biomass loss in the treated cultures was within a normal range, in agreement with previous observations [40,48,49]. At the same time, corresponding with the initiation of the death phase in the controls, and stationary phase in the treatments, DNA ratios continued increasing exponentially in both controls, but were lower in TR2, and below detection in TR1.

Similar growth cycles and DNA ratios were observed in the replicated experiments 4-2 and 4-3, although fewer qPCR measurements were taken (Fig. A.6.B-C). The collapse of the controls, due to V. chlorellavorus , occurred when DNA ratios were greater than one; however, in TR2 there was not an associated major decline even with ratios in the range of 1 or greater, as shown in all three replicates (Fig. A.6.A-C). This may imply that the deoxygenation treatment prevented V. chlorellavorus from causing a collapse even when the V. chlorellavorus population was relatively high. Although the negative control samples had higher DNA ratios than the treated samples, the ratios observed for TR2 suggested that alternating 2-h deoxygenation-aeration cycles did not eradicate V. chlorellavorus .

The results of V. chlorellavorus quantification by qPCR indicated that deoxygenation- aeration cycling had a largely suppressive effect on the accumulation of the pathogenic cells. While some of the treated cultures were below the detection limit of 13 gene copies [27], other replicates in the experiment harbored detectable V. chlorellavorus cells. A whole V. chlorellavorus genome

sequence analysis predicts that this bacterium encodes a complete set of highly expressed genes required for aerobic respiration, but also encodes a lactate dehydrogenase gene facilitating fermentation of pyruvate for anaerobic survival [26]. Though the bacterium appears to be capable of surviving the alteration in oxygen cycles under most conditions tested here, evidence from other systems finds that pathogenic life stages can be closely regulated by sensing oxygen levels [50].

Because C. sorokiniana microalgae produce oxygen as they photosynthesize and grow, it is possible that the concentration of oxygen acts as a cue for V. chlorellavorus similarly by initiating expression of the Type 4 secretion system proteins. Further studies are needed to understand the specific timeframe, cues, and environmental conditions required by V. chlorellavorus to shift into a pathogenic mode, and likewise, to develop resistance to anaerobic conditions.

Fig. A.6. Compared ash free dry weight (AFDW) and DNA ratios in aerated negative controls

(C2A and C2B) and 2-h deoxygenation-aeration cycling treatment (TR1 and TR2) in three replicates (A-C) with Chlorella sorokiniana-Vampirovibrio chlorellavorus co-cultures (Exp.4).

DNA ratios of V. chlorellavorus to C. sorokiniana are presented on log 10 scale at the growth phase

(1), stationary phase (2), and death phase (3) for the end of the 16-hr light periods (small markers) and end of the 8-h dark periods (large markers).

A t-test assuming unequal variances of the biomass average of all replicates in Exp.4 including all the controls (n = 6) versus all the treatments (n = 6) showed no significant statistical difference (p > 0.05) after three days; however, the same group had a significant statistical difference (p = 0.02) after five days. The maximum biomass levels during the stationary phase in the treatments ranged from 0.35 to 0.45 g L-1, which is the same range as in the V. chlorellavorus - free cultures in Exp. 1-3. This indicates that the culture growth was limited by light or nutrients, or both, rather than by V. chlorellavorus, which had minimal, if any apparent impact on the biomass concentrations in the treatments.

The aerated negative control in experiments 1-3 was compared with the aerated controls in

Exp. 4 (Fig. A.7). At day 3 (a), where the infected cultures had not yet collapsed, the difference in biomass was not statistically significant (p > 0.05); however, after the infected C. sorokiniana cultures entered the death phase, on day 5 (b), the statistical difference was significant (p = 0.008).

Fig. A.7. Compared ash free dry weight (AFDW) in aerated negative control experiments between

Vampirovibrio chlorellavorus -free Chlorella sorokiniana cultures (Exp. 1-3) and C. sorokiniana -

V. chlorellavorus co-cultures (Exp. 4). (a) and (b) show the selected data for statistical t-test. Error bars represent one standard deviation.

Light microscopic observation for the co-cultures in the experimentally replicated flasks

(Exp. 4) showed characteristic signs of V. chlorellavorus infection, i.e., lysis and clumping C. sorokiniana cells, by one day post-inoculation. The co-culture in the negative control flasks became brown in color, and developed a foamy brown grime on the walls of the flasks (protein film on bubbles) as expected in a dying culture. During the death process/phase, the biomass density showed a sharp decline, and co-culture appeared increasingly brown and emitted an unpleasant odor (Fig. A.8.A). In contrast, the treated co-cultures showed no evidence of algal cell

collapse, and a light microscopic observation showed that the C. sorokiniana culture consisted of individually-growing green cells, indicative of algal health (Fig. A.8.B). Conversely, the untreated

C. sorokiniana co-cultures (negative control) contained lysed and clumped cells (Fig. A.8.C) characteristic of the appearance of C. sorokiniana cultures infected with V. chlorellavorus .

Fig. A.8. Visual inspection and light microscopic observations (400x magnification) of Chlorella sorokiniana-Vampirovibrio chlorellavorus co-cultures. A) Treated co-cultures with deoxygenation-aeration cycles, left and middle flasks (TR) with brighter grass-green color, and aerated controls, the 2 flasks on the right (C) with foamy brown grime on the walls. B) Green

single cells of C. sorokiniana in the treated co-cultures. C) Lysed and clumped cells of C. sorokiniana in the aerated controls.

4. Conclusion

The deoxygenation-aeration cycles did not significantly inhibit C. sorokiniana growth in the experiments with V. chlorellavorus -free C. sorokiniana cultures (Exp.1-3). In the C. sorokiniana-V. chlorellavorus co-cultures (Exp.4), the aerated controls collapsed; however, the deoxygenation treatment provided protection of C. sorokiniana cells against attack and predation by V. chlorellavorus . These results provide a robust demonstration that deoxygenation-aeration cycling can dramatically limit the negative effects of the V. chlorellavorus pathogen on its algal host, and offer great promise as an effective treatment for V. chlorellavorus management in indoor reactors, and likely for outdoor ponds as well.

5. Acknowledgements :

The authors are grateful for the U.S. Department of Energy and Regional Algal Feedstock Testbed

(RAFT) project, University of Arizona, for supporting this study.

6. References :

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Appendix B

Paper to be submitted to Algal Research

Technoeconomic assessment of deoxygenation for control of Vampirovibrio chlorellavorus in Chlorella sorokiniana cultures

S. Attalah a, P. Waller a* , S. Steichen b, C. C. Brown b, K. Ogden c, and J. K. Brown b a The University of Arizona, Department of Biosystems Engineering, Tucson, AZ 85721, USA. b The University of Arizona, School of Plant Sciences, Tucson, AZ 85721, USA. c The University of Arizona, Department of Chemical and Environmental Engineering, Tucson, AZ 85721, USA.

* Corresponding author: [email protected] BE Department, Shantz Bldg. Rm. 403, Tucson, AZ 85721

Abstract : Deoxygenation prevents Vampirovibrio chlorellavorus from attacking Chlorella sorokiniana . This study elaborates a practical approach to minimize the cost of the deoxygenation process in a C. sorokiniana cultivation system by sparging with nitrogen gas for one hour at the beginning of the night, and then utilizing natural deoxygenation by algae dark respiration to keep the oxygen concentration at a low level for the rest of the night. This method kept oxygen concentration low during the entire dark period and effectively controlled V. chlorellavorus infection in laboratory co-cultures of C. sorokiniana-V. chlorellavorus . The cost of this method was assessed in an outdoor experiment with pure water and pathogen-free C. sorokiniana cultures in a 150-L sealed translucent-polyethylene reactor. The outdoor reactor had a better seal than the laboratory experiments and maintained dissolved oxygen concentrations at near zero throughout the night in the C. sorokiniana culture. The total nitrogen sparging amount per night per liter of algae culture was evaluated for the 150-L reactor, then translated to large-scale per acre (0.4 ha) raceway cost. The present study suggests that onsite nitrogen generators are cost-effective and

appropriate for deoxygenating C. sorokiniana culture in closed bioreactors and algae raceways with tank storage at night, such as Algae Raceway Integrated Design (ARID).

Keywords : aeration, anoxic, algal biomass, bacterial infection, economic feasibility, nitrogen sparging.

1. Introduction

Bacterial infections have led to failures of many microalgae cultures in the Regional Algal

Feedstock Testbed (RAFT) project and other research and commercial projects [1]. Researchers have proposed many techniques to reduce the impact of infection/predation, including specific cultivation methods and strain selection [2,3], chemical [4-10], physical [11], and biological

[12,13] control.

Chlorella sorokiniana is highly sensitive to the predatory Vampirovibrio chlorellavorus

[1,10,14-17]. Cultivation of C. sorokiniana will continue to be hampered without an effective control strategy for V. chlorellavorus . Periodic deoxygenation of the algae culture controls V. chlorellavorus without harming C. sorokiniana , because C. sorokiniana can tolerate extended anoxic conditions [18,19] and V. chlorellavorus is aerobic [17]. Alternating 2-h deoxygenation and aeration cycle-times over 8-h dark periods prevented V. chlorellavorus from damaging C. sorokiniana cultures [19]. This technique is applicable to closed bioreactors [20] and the Algae

Raceway Integrated Design (ARID) reactor in which the culture is stored in a deep tank overnight

[21].

The deep tank of the ARID raceway is typically aerated to prevent anoxic conditions, but aeration might be unnecessary and even counterproductive. For instance, aeration can use more

energy than any other device in ARID [22]. In addition, the mixing caused by aeration during the night contributes to biomass loss [23,24]. Reduced mixing and low dissolved oxygen (DO) during the dark period could limit algae biomass loss at night [25]. Finally, daytime DO levels can reach a toxic range in intensive algae cultivation [26-29], and the daytime DO levels are reduced if the culture has a reduced DO at night [19].

Air injection (aeration) can prevent algae settling at night but may not be necessary. If mixing is required to suspend or resuspend the culture, alternatives to air injection include sump pumps [21], impellers and paddlewheels [30]. Regardless, Chlorella microalgae have a slow settling rate [31] so mixing is generally not required.

One of the concerns associated with prolonged periods of anoxic conditions in C. sorokiniana culture is that it might harm beneficial aerobes living in association with the culture.

However, because C. sorokiniana can grow axenically [1] and deoxygenation cycles, as long as 8- hours, do not reduce the culture productivity [19], deoxygenating the culture is suggested to be appropriate.

It is essential that algae biomass production for biofuel uses low-cost techniques to be economically feasible and compete with fossil fuels [32-34]. In addition to low cost, cultivation practices should be environmentally friendly and ideally simple to apply.

Physical, chemical, or biological [35-42] methods can deoxygenate water. Cost, labor, resource use, and practicality are crucial factors in the selection and design of an appropriate deoxygenation process. Nitrogen gas sparging has been used to deoxygenate water, and is a proven cost-effective, and environmentally friendly method [43-45]. Algae culture itself can become part of the deoxygenation process because it uses oxygen as part of the respiration and cell repair

process at night. However, just relying on natural deoxygenation by algae dark respiration does not decrease the DO concentration to a low enough level to reduce or prevent V. chlorellavorus pathogenicity/virulence [19].

The primary goal of this study was to develop a practical, cost-effective deoxygenation method based on a hypothesis that short cycle nitrogen gas sparging at the beginning of the night followed by natural deoxygenation by algae respiration for the rest of the dark period can prevent/reduce V. chlorellavorus pathogenicity. The first objective was to apply this treatment process in laboratory experiments with C. sorokiniana , DOE1412 [46], co-cultured with V. chlorellavorus . The second objective was to evaluate the cost of this deoxygenation method outdoor in a 150-L sealed translucent-polyethylene reactor with pathogen-free C. sorokiniana monoculture by calculating the required mass of nitrogen gas per unit volume of culture. Finally, cost and management aspects of deoxygenation by nitrogen sparging with an onsite nitrogen generator were assessed at a commercial-scale ARID raceway.

2. Materials and Methods

2.1. Cultures and cultivation medium

Chlorella sorokiniana [46] cells selection, V. chlorellavorus [14] isolation, and cultivation medium preparation (modified BG-11) [47,48] were conducted as described previously [19].

2.2. Laboratory and outdoor experimental protocols

This study included laboratory experiments with pathogen-free C. sorokiniana monoculture, C. sorokiniana -V. chlorellavorus co-cultures, and outdoor experiments with C. sorokiniana monoculture and clean water (Table B.1).

Five laboratory experiments were conducted in an incubator with four 4-L Erlenmeyer flasks subjected to 16-h light and 8-h dark cycles. The first experiment (Exp.1), with V. chlorellavorus -free C. sorokiniana monoculture, and the second experiment (Exp.2), with C. sorokiniana -V. chlorellavorus co-cultures , assessed C. sorokiniana growth and V. chlorellavorus infection potential, respectively, using continuous aeration with ambient air during the dark period in all experimental flasks. The third (Exp. 3) and fourth (Exp. 4) experiments were conducted with

C. sorokiniana -V. chlorellavorus co-cultures including aerated control with ambient air and deoxygenation treatments with pure nitrogen gas (UN1066) sparging for 0.5 and one hour at the beginning of the dark period, respectively, followed by natural deoxygenation by algae dark respiration to keep the oxygen concentration at a low level (<0.5 ppm). The fifth experiment (Exp.

5) used a 1-h deoxygenation treatment with pure nitrogen gas sparging in C. sorokiniana -V. chlorellavorus co-cultures associated with unidentified ciliates. This experiment (Exp. 5) was replicated three times (experiments 5-1 to 5-3).

In experiments 3-5, the experimental design included replicated deoxygenation treatment

(TR1, TR2) and replicated aerated control (CA1, CA2). The controls were continuously aerated with ambient air during the 8-h dark period.

The outdoor experiments (Exp. 6 and 7) were conducted using 113 L of clean tap water and C. sorokiniana monoculture, respectively, in a 150-L sealed translucent-polyethylene reactor, from 11/29/17 to 12/07/17 in Tucson, Arizona. In Exp. 6, mixing and aeration were applied during the day to increase DO to saturation in the clean water. Nitrogen sparging was applied for one hour at the beginning of the night to deoxygenate the water. In Exp. 7, no mixing was applied during the night. Nocturnal anoxic conditions were created by deoxygenating the culture with pure

nitrogen gas (UN1066) for one hour at the beginning of the night, followed by natural deoxygenation by algal dark respiration. During the day, photosynthesis naturally increased DO above saturation, and mixing of the algae culture was performed with a submersible water pump at a flow rate of 10 L min -1.

Table B.1. Experimental plan.

Experiments Description

Laboratory Experiments

Exp. 1: pathogen-free C. Continuous aeration in all flasks (No deoxygenation) sorokiniana . during dark period, with pathogen-free C. sorokiniana (No V. chlorellavorus ).

Exp. 2: Co-culture of C. Continuous aeration in all flasks (No deoxygenation) sorokiniana-V. chlorellavorus. during dark period, with C. sorokiniana infected by V. chlorellavorus .

Exp. 3: Co-culture of C. 0.5-h nitrogen sparging (deoxygenation treatment) and sorokiniana-V. chlorellavorus no aeration during dark period, with C. sorokiniana subjected to 0.5-h deoxygenation. infected by V. chlorellavorus . Negative control continuously aerated during dark period

Exp. 4: Co-culture of C. One-hour nitrogen sparging (deoxygenation treatment) sorokiniana-V. chlorellavorus and no aeration during dark period, with C. subjected to 1-h deoxygenation. sorokiniana infected by V. chlorellavorus . Negative control continuously aerated during dark period.

Exp. 5: C. sorokiniana -V. One-hour nitrogen sparging (deoxygenation treatment) chlorellavorus co-cultures associated and no aeration during dark period using C. with unidentified ciliates sorokiniana infected by V. chlorellavorus in association with unidentified ciliates. There were three replicates (Exp. 5-1 to 5-3). Negative control continuously aerated during dark period.

Outdoor experiments

Exp. 6: Clean water subjected to 1-h One-hour nitrogen sparging (deoxygenation treatment) deoxygenation. with clean water in a 150-L sealed translucent- polyethylene reactor during nighttime. Mixing and aeration applied only during daytime.

Exp. 7: pathogen-free C. One-hour nitrogen sparging (deoxygenation treatment) sorokiniana subjected to 1-h with C. sorokiniana monoculture in a 150-L sealed deoxygenation. translucent-polyethylene reactor during nighttime. No aeration. Mixing applied only during daytime.

2.3. Experimental apparatus design and operating procedure

2.3.1. Laboratory experimental setup

The laboratory apparatus included four 4-L Erlenmeyer flasks in a digital incubator- refrigerator-shaker (Innova4330, New Brunswick Scientific Inc., Edison, NJ), aquarium air pumps

(for ambient air injection in experiments 1 and 2), clear tygon-tubing, check valves, nitrogen and carbon dioxide gas cylinders, pH controllers (MC110, Milwaukee Instruments Inc., Rocky Mount,

NC), DO sensors (DO1200, Sensorex Inc., Garden Grove, CA), CR300 Campbell Scientific data logger (CR300, Campbell Scientific, Logan, UT), two grow-light panels (45W LED, Watt Shine,

China), and stone diffusers (Fig. B.1).

Rubber stoppers with holes for tubes were used to seal the flasks, and clear tygon-tubing connected the stone diffusers to the air pumps and gas cylinders. The culture pH was kept in the range of 7.0 - 7.5 by a pH controller (MC110, Milwaukee Instruments Inc., Rocky Mount, NC) and injection of pure carbon dioxide. Deoxygenation and aeration were achieved using pure nitrogen gas (UN1066) and ambient air, respectively. The culture was continuously agitated at a constant speed of 80 rpm, the DO levels were recorded at 5-minute intervals, and the incubator

temperature was set at 24 °C. During the light period, the incubator temperature increased to 30

°C because the two grow-light panels added heat. The light intensity at each flask’s location (TR1,

TR2, CA1, and CA2) inside the empty incubator was measured with a pyranometer (MP-200

Apogee silicon, Apogee instruments Inc., Logan, UT) at a height of 16 cm above the incubator platform, the culture height in the flasks. Light intensities ( I) were measured in W/m 2: I-CA1 = I-

CA2 = 39 W/m 2, I-TR1 = 36 W/m 2, and I-TR2 = 35 W/m 2.

Fig. B.1. Equipment for the laboratory experiments. A) General view of the Innova 4330 incubator, equipment for deoxygenation and aeration, and pH controllers. B) Grow-light panels were added to the incubator for culture illumination. C) Flasks containing Chlorella sorokiniana culture with sensors (dissolved oxygen and pH) and stone diffusers for nitrogen and air. In the present study, only the four flasks in the corners were considered; deoxygenation treatment replicates (TR1 and

TR2) were on the left, and aerated control replicates (CA1 and CA2) on the right.

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2.3.2. Outdoor experimental setup

The outdoor experimental setup included a 150-L sealed translucent-polyethylene reactor containing three identical DO probes (DO1, DO2, and DO3), pH probe, type-K thermocouple, stone diffuser, and submersible pump connected to a PVC-pipe structure (Fig. B.2).

The DO concentrations in the outdoor reactor were measured at 6 cm (DO1), 30 cm (DO2), and 60 cm (DO3) below the culture surface. The average DO data were recorded in a CR3000

Campbell Scientific data logger (CR3000, Campbell Scientific, Logan, UT) at 5-minute intervals.

The culture temperature was continuously recorded with the type-K thermocouple. The culture pH was kept in the range of 7.0 – 7.5 by a pH controller (MC110, Milwaukee Instruments Inc., Rocky

Mount, NC) and injection of pure CO 2 gas. A compressed pure nitrogen gas cylinder (2350 psi,

210cf, Class 2.2 UN1066 Non-Flammable, cryogenics and gas facility, the University of Arizona,

Tucson, AZ) was connected to a solenoid valve and a relay. The CR3000 datalogger interrupted nitrogen injection after one hour of deoxygenation, or sooner if the DO2 probe recorded near zero

DO level (0.2 ppm). Nitrogen gas was injected into the algae culture through a stone diffuser at a

-1 flow rate of 9 L min and 1-atm injection pressure (P out ). There was no mixing or aeration of the algae culture for the rest of the night. During the day, a submersible aquarium pump mixed the culture through a PVC-pipe structure at a flow rate of 10 L min -1. During the experiment with clean water, mixing and aeration were applied to re-establish DO saturation during the day. Solar intensity was measured with a silicon pyranometer (200SZ, Scientific sales Inc., Lawrenceville,

NJ) placed outside the reactor at 2 m elevation above the soil surface and connected to the CR3000 data logger.

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Fig. B.2. Equipment for the outdoor experiments. A) Polypropylene reactor containing Chlorella sorokiniana culture with dissolved oxygen probe (DO1, DO2, DO3) locations inside the reactor.

B) Sealed lid with two small vents and connectors. C) Sensors and PVC-pipe structure inside the reactor. D) CO 2 cylinder and Campbell Scientific CR3000 data logger. E) Nitrogen cylinder and relays.

2.4. Inoculum preparation and sampling approach

In the laboratory experiments (Exp. 1-5), C. sorokiniana inoculum, V. chlorellavorus filtrate, nutrients, and co-cultures were prepared as described in [19]: the pathogen-free C. sorokiniana culture and/or V. chlorellavorus -C. sorokiniana co-cultures were diluted to 0.16 g L -

1 biomass density. Modified BG-11 growth medium was added, then 3.5 L of the mixture was transferred into each experimental flask to initiate the experiments.

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In Exp. 5, an unexpected development was that ciliates infected the algae inoculum. As contamination by ciliates is common, especially in open ponds, the decision was made to proceed with the experiment and evaluate the effect of deoxygenation on the two contaminants (i.e. V. chlorellavorus and ciliates).

Experiment 6 was conducted with clean tap water and did not include algae culture. In the outdoor experiment with C. sorokiniana monoculture (Exp. 7), 113 L of C. sorokiniana culture were transferred to the experimental translucent-polyethylene reactor from an outdoor paddlewheel raceway used for the Regional Algae Feedstock Testbed (RAFT45) experiment at the

University of Arizona. The algae culture was transferred when the biomass density reached 0.2 g

L-1 AFDW, then nutrients were replenished using the modified BG-11 recipe, and the culture pH was adjusted to 7.0.

In the laboratory experiments (1-5), culture samples were taken daily from each flask at the end of the 16-h light period; whereas daily sampling in the outdoor experiment (Exp. 7) was performed at 9 am. The volume level in the reactors was adjusted with water after each sampling.

2.5. Assessment of C. sorokiniana growth and V. chlorellavorus infection

The algae growth was evaluated by measuring the DO concentrations and the culture optical density at 750 nm wavelength (OD750) which was then converted to AFDW biomass (g

L-1) as described previously [19]. The impact of V. chlorellavorus infection was assessed by

AFDW and DO measurements, light microscopic observation of lysed C. sorokiniana cells, and visual symptoms of V. chlorellavorus attack such as; algae discoloration from grass-green to brownish, release of repugnant smell, and froth-like accumulation on the culture surface and

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reactor walls. Additionally, in Exp. 5 the accumulation of V. chlorellavorus was reported by the ratio of V. chlorellavorus -16S rDNA- to C. sorokiniana -18S rDNA- as determined by a previously described quantitative polymerase chain reaction assay [19, 58]. The statistical significance of the difference in AFDW between the treated cultures and experimental controls was performed using the Student t-test at α = 0.05 level of significance (data analysis tool pack in

Microsoft Excel, 2016).

2.6. Technical assessment of nitrogen usage and cost estimation

Prior to the initiation of the outdoor experiments (Exp. 6 and 7), the DO probes were calibrated at the saturation point (saturated air) and the zero point (saturated solution of sodium sulfate) [50]. The probes accurately detected both saturation and zero DO levels (Fig. B.3).

Fig. B.3. Two-point calibration (saturation and zero) of the dissolved oxygen probes (DO1, DO2,

DO3) measured with CR3000 data logger prior to the outdoor experiments with clean tap water

(Exp. 6) and Chlorella sorokiniana culture (Exp.7).

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During deoxygenation, the oxygen depletion coefficient K La is calculated as the logarithmic slope of the oxygen deficit vs. time (Eq. 1) and then adjusted for temperature (Eq. 2)

[51]:

KLa = [ln (C i) – ln (C f)]/Δt (1)

20-T KLa20 = K La (θ ) (2)

-1 with: K La: Oxygen depletion coefficient (min ), C i: Initial oxygen concentration (ppm) at temperature T (°C), C f: Final oxygen concentration (ppm) at temperature T (°C), Δt: Time interval

-1 (min), K La20 : Oxygen depletion coefficient (min ) at standard temperature (20 °C), and

θ: Temperature correction coefficient (1.024).

The nitrogen tank lifetime (Eq. 3) and daily nitrogen gas injection volume (Eq. 4) were calculated [52] at a constant injection pressure (Pout = 1 atm)

Tt = P i / ΔP (3)

With: T t: Nitrogen tank lifetime (d), P i: Initial pressure (atm), and ΔP: Change in pressure (atm).

Vd = V i / T t (4)

With: V d: Daily nitrogen injected volume (L), V i: Nitrogen gas initial volume (L), and T t: Nitrogen gas total volume (L).

The economic assessment was made based on the ARID raceway, which is a suitable reactor for the deoxygenation process because algae culture is stored in a deep canal at night. The deep canal is not sealed from the atmosphere (Fig. B.4), but it would need to be sealed for the

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deoxygenation process to be effective. The volume in the tank is a function of cultivated area and culture depth.

Fig. B.4. ARID raceway prototype cultivated with Chlorella sorokiniana at the algae facility of the University of Arizona (Regional Algal Feedstock Testbed project, 2017).

The required amount of nitrogen gas and cost of the deoxygenation process were first calculated for the 150-L sealed translucent-polyethylene reactor, based on the cost of a compressed nitrogen gas cylinder. The volume and cost were then translated to a one acre (0.4 ha) commercial

ARID system with onsite nitrogen gas generator. The cost of nitrogen cylinders is based on cost rates at the cryogenics and gas facility of the University of Arizona (2017). The onsite nitrogen generator cost is estimated based on capital, power, and maintenance costs [53].

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3. Results and discussion

The first three sections focus on laboratory experiments. The last section includes an outdoor test and economic calculations. Section 3.1 presents the results of the initial laboratory experiments with continuous aeration and pathogen-free C. sorokiniana monoculture (Exp. 1), and

C. sorokiniana -V. chlorellavorus co-cultures (Exp. 2). Section 3.2 focuses on the impact of the

0.5-h and 1-h deoxygenation treatments on V. chlorellavorus pathogenicity. Section 3.3 evaluates the effect of deoxygenation on the combination of ciliates and V. chlorellavorus . Finally, section

3.4 includes an outdoor evaluation of a 1-h nitrogen injection cycle and economic calculations.

3.1. Pathogen-free C. sorokiniana monoculture and C. sorokiniana-V. chlorellavorus co-cultures with continuous aeration

In order to simulate typical management practice, pathogen-free C. sorokiniana culture

(Fig. B.5-A) was aerated during 8-h dark periods (Exp.1). Biomass concentration reached 0.54 g

L-1 AFDW after nine days. The consistent diurnal DO pattern indicated that the culture was healthy throughout the entire experiment. The DO concentrations ranged from 7 ppm during dark periods to 12 ppm during light periods. After nine days, healthy single green C. sorokiniana cells were observed in the microscope.

In Exp. 2, C. sorokiniana-V. chlorellavorus co-cultures were aerated during 8-h dark periods. Biomass concentration increased to 0.27 g L-1 AFDW, and then entered the death phase

(Fig. B.5-B). The DO concentrations ranged between 7 – 14 ppm during the first two days, and then ranged between 6 and 8 ppm during the death phase. Signs of bacterial predation began on the second day and included brown color of the culture, froth accumulation on the culture surface

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and the walls of the experimental reactors, repugnant odor, and clumped and lysed C. sorokiniana cells, all of which are typical symptoms of V. chlorellavorus infection [15].

As expected, aeration enhanced V. chlorellavorus virulence and pathogenicity and prompted lysis and decay of C. sorokiniana cells, which led to a rapid culture collapse.

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Fig. B.5. Average ash free dry weight (AFDW) biomass and dissolved oxygen (DO) with continuous aeration during the dark period. A) Pathogen-free Chlorella sorokiniana culture sustained growth period. B) Collapse of C. sorokiniana co-cultured with Vampirovibrio chlorellavorus indicating a high sensitivity of C. sorokiniana to V. chlorellavorus infection in a continuously aerated environment . Error bars represent one standard deviation ( n = 4 ).

3.2. Evaluation of the efficacy of short time deoxygenation on the growth of C. sorokiniana infected by V. chlorellavorus

Experiment 3 assessed the impact of 0.5-h deoxygenation at the beginning of the dark period and no aeration on C. sorokiniana-V. chlorellavorus co-cultures. The control was continuously aerated with ambient air during the entire dark period. Both treatment and control collapsed (Fig. B.6) and presented the typical signs of V. chlorellavorus infection, i.e., froth, unpleasant smell, clumping and lysed cells. The AFDW and DO measurements showed similar

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trends in the treatment and control. The daytime DO concentrations in the treatment decreased at the end of the experiment but were not as low as the DO concentrations in the control. There was reduction in infection impact in the treated cultures compared to the control; nevertheless, the visual signs of V. chlorellavorus pathogenicity indicated that 0.5-h nitrogen sparging was not an effective control method.

Fig. B.6. Average ash free dry weight (AFDW) biomass and dissolved oxygen (DO) for Chlorella sorokiniana co-cultured with Vampirovibrio chlorellavorus . Aerated control (C) and deoxygenation treatment (TR) represent averages of the aerated control duplicates (CA1 and CA2) and the treatment duplicates (TR1 and TR2), respectively. During dark period, the controls were continuously aerated, and the treatments were subjected to 0.5-h deoxygenation at the beginning of the dark period (Exp. 3). Error bars denote one standard deviation.

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Experiment 4 assessed the impact of 1-h deoxygenation at the beginning of the dark period and no aeration on C. sorokiniana-V. chlorellavorus co-cultures. The controls collapsed after three days of growth, whereas the deoxygenated culture with 1-h nitrogen sparging had sustained growth and no signs of V. chlorellavorus infection impact (Fig. B.7). During the first 3 days, it appears that the treatment slightly delayed C. sorokiniana growth as indicated by a lower biomass (0.3 g

L-1) compared to 0.35 g L-1 in the controls. However, the controls collapsed after three days of growth due to the infection by V. chlorellavorus , while the treated culture continued to grow for the entire experiment duration reaching 0.45 g L-1 AFDW. The efficacy of 1-h deoxygenation was confirmed by the significant difference in AFDW biomass between the treatment and control in all replicates after day 3 (Fig. B.7).

During the dark period, the 1-h deoxygenation treatment DO concentrations decreased to the anoxic range. The aerated control DO concentration was in the range of 6 to 7 ppm, but had a steady decrease in dark period DO concentration during the first three days. During the light period, DO concentrations remained in the range of 10-12 ppm in the treated culture for the entire run. However, in the controls, DO only increased to 11 ppm during the first two days and then declined sharply and remained below saturation, in the range of 4-5 ppm, for the rest of the run. In addition to the evidence of V. chlorellavorus predation from biomass and DO concentrations, light microscopy observation showed green individual cells in the treated culture while the untreated culture displayed the typical signs of V. chlorellavorus predation (brown culture, froth accumulation, and lysed cells).

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3.3. Application of one-hour nitrogen gas sparging (deoxygenation) in C. sorokiniana -V. chlorellavorus co-cultures associated with unidentified ciliates

The deoxygenation treatment with 1-h nitrogen sparging at the beginning of the dark period

(Exp. 4) was repeated three times in Exp. 5, which had C. sorokiniana -V. chlorellavorus co- cultures infected with unidentified and uncontrolled ciliates. The spontaneous emergence of ciliates was noticed two days after inoculation of C. sorokiniana and V. chlorellavorus co-cultures, and their presence persisted in all three replicates. In order to observe the efficacy of the treatment

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on a second contaminant, the decision was made to go ahead with the experiment. The aerated controls collapsed on the third day after inoculation in all three replicates while the deoxygenated cultures did not collapse; however, growth was minimal in the treated cultures after the third day, which is often observed with ciliate contamination. The treated cultures did not exhibit visual signs of V. chlorellavorus predation but had the reduced growth that is typically observed with grazing ciliates.

On the third day after inoculation of C. sorokiniana-V. chlorellavorus co-culture, the number of V. chlorellavorus per algal cell in the controls increased by approximately 100-fold relative to the treated cultures in each replicated experiment. After the third day, the trend was inconsistent (Fig. B.8), possibly due to the contamination by ciliates, which are known to graze on both bacteria and algae [54]. The delay in V. chlorellavorus accumulation under the treatment conditions reflects the dynamics of the physiologically similar attack phase Bdellovibrio spp. under anoxic conditions [55]. In the aerated controls algal populations declined rapidly upon passing a

10:1 ratio of V. chlorellavorus to C. sorokiniana . Observations of Bdellovibrio sp. grown in ambient air have indicated that predator to prey ratio can trigger massive prey lysis in a threshold dependent manner as well [56]. After the third day, treated cultures in this experiment did cross the 10:1 predator to prey ratio threshold, but did not exhibit immediate cell death, indicating that anoxic conditions disrupt the typical interaction dynamics.

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Student’s t-test, assuming unequal variances, n = 6, showed no statistical difference (p >

0.05) in AFDWs at the end of the growth phase (point a) but a highly significant statistical difference (p = 0.00001) at point b, the stationary phase for the treatment and the death phase for the control (Fig. B.9). As with Exp. 4, one-hour nitrogen sparging reduced DO concentrations in the treated culture (Exp. 5) to the anoxic range (< 0.5 ppm), while continuous aeration in the control maintained DO between 6 and 7 ppm (Fig. B.9). During the light period, DO concentrations remained in the range of 10-12 ppm in the treated culture for the entire run. However, in the controls, DO only increased up to 11 ppm during the first two days, then decreased sharply and remained below saturation, in the range of 4-5 ppm, for the rest of the experimental run.

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Fig. B.9. Average ash free dry weight (AFDW) biomass and dissolved oxygen for 3 replicates with Chlorella sorokiniana co-cultured with Vampirovibrio chlorellavorus in association with unidentified ciliates (Exp. 5). The treated culture (TR) was subjected to deoxygenation with 1-h nitrogen gas sparging at the beginning of the dark period, and the controls (C) were continuously aerated during the dark period. Error bars denote one standard deviation (n = 8). Points (a) and (b) represent the grouped data for a statistical comparison.

Light microscopy observation showed the presence of ciliates in all tested flasks. There were green individual C. sorokiniana cells in the treated flasks while the control flasks had clumping and lysed C. sorokiniana cells. In addition, the controls had brown color and unpleasant odor, typical signs of V. chlorellavorus predation (Fig. B.10). As suggested previously, ciliates graze on both bacteria and algae [54], which probably inhibited the algae biomass productivity in

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the treated culture even though it did not lead to a culture collapse. Thus, the deoxygenation treatment with 1-h nitrogen sparging effectively reduced the impact of V. chlorellavorus pathogenicity, but did not control ciliates. In contrast, the untreated co-cultures quickly collapsed due to infection by V. chlorellavorus and possibly also due to contamination by ciliates.

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Based on the combination of visual symptoms, light microscopy observation, and DNA ratios, it appears that the 1-h deoxygenation treatment prevented the culture from collapsing due to V. chlorellavorus predation.

3.4. Nitrogen gas sparging and dissolved oxygen depletion assessment

Experiments 6 and 7 were conducted outdoors and assessed nitrogen use and DO depletion during 1-h nitrogen sparging in a 150-L sealed translucent-polyethylene reactor. Exp. 6 was conducted with clean tap water, and Exp. 7 with C. sorokiniana monoculture free of V. chlorellavorus , at 0.28 g L-1 AFDW initial concentration. In both experiments, 1-h nitrogen sparging at the beginning of the night reduced DO to the anoxic range. In Exp. 6 with clean water,

DO decreased from 7 ppm to 0.5 ppm in 32 min whereas in Exp. 7 with algae culture, DO decreased from 16 ppm to 0.5 ppm in 40 min (Fig. B.11). The oxygen depletion coefficient K La at

30 °C was 0.109 min -1 in the clean water and 0.112 min -1 in the algae culture as calculated from the logarithmic slope of DO vs time (inset in Fig. B.11). Converting to standard temperature (20

-1 -1 °C) with Eq. 2, K La20 was 0.086 min , and 0.088 min , in the clean water and algae culture, respectively. The similar DO depletion coefficient calculated in both experiments indicates that the algae had minimal deoxygenation effect during the nitrogen sparging process. The rate of DO depletion slowed down at the end of the nitrogen sparging cycle in both cases, which means that the deoxygenation efficiency of the nitrogen sparging process decreased over time.

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Dissolved oxygen remained low for the entire night in the algae culture (Exp. 7) but increased to 0.6 ppm in pure water (Exp. 6), indicating that natural deoxygenation due to dark respiration contributes to the deoxygenation process and maintains low DO (Fig. B.12).

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Fig. B.12. Dissolved oxygen (DO) concentration with less than 1-h nitrogen sparging

(deoxygenation treatment) in clean water (solid line) and Chlorella sorokiniana culture (dashed lines).

During the day, aeration in the clean water (Exp. 6) increased DO concentrations up to saturation (8 ppm) in two hours, while photosynthesis in the algae culture (Exp.7) increased the

DO to saturation in 2.5 hours and to a maximum of 16 ppm, which is typical of a healthy algae culture (Fig. B.13).

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After sparging with nitrogen gas, the DO concentrations in the outdoor reactor remained much lower than in the indoor laboratory experiments with flasks. The outdoor reactor had a much better seal than the indoor reactors, which had gas exchange with the atmosphere due to the experimental design. This difference demonstrates the possible importance of sealing the reactor in the case of 1-h deoxygenation followed by natural dark respiration [19].

In Exp. 7 with C. sorokiniana monoculture, the deoxygenation process in the sealed reactor did not prevent algae growth. Deoxygenation might even have had a beneficial impact on the algae culture by keeping the DO low [25-28]. As usual, DO fluctuation correlated with temperature and

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solar radiation. The algae culture ceased growing on day 6 when clouds and low temperatures caused a biomass decline (Fig. B.14).

Fig. B.14. Chlorella sorokiniana growth indicators and environmental parameters in the outdoor reactor. A) Ash free dry weight (AFDW) biomass, and dissolved oxygen (DO) concentrations at

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various depths in the reactor: 6 cm (DO1), 30 cm (DO2), and 60 cm (DO3) below the culture surface. B) Culture temperature and solar intensity (measured outside the reactor).

3.5. Nitrogen-usage evaluation and cost estimation

In the sealed outdoor reactor with 113-L algae culture (Exp. 6), the daily pressure decrease in the nitrogen cylinder was 13.6 atm, and the discharge pressure (P out ) was 1 atm. The initial pressure in the nitrogen cylinder was 160 atm with a full initial gas volume of 5,946 L. The lifetime of the compressed nitrogen gas cylinder is calculated with Eq. 3 as T t = 160 atm/13.6 atm/day =

11 days, and the daily volume of nitrogen gas used is calculated with Eq. 4 as V d = 5,946 L/11 days = 540 L d-1 at standard temperature and pressure (STP) with diffusers supplying 9 L min -1.

The cost of one cylinder was $31, including $14 for the nitrogen gas and $17 for delivery. The applied volume of nitrogen gas per liter of algae culture was 540 L / 113 L ~ 5 L N gas / L culture.

Small nitrogen cylinders (9in x 52in) are only convenient for small reactors and high value products. For commercial-scale raceways, onsite nitrogen generators would be the most cost- effective source of nitrogen.

The estimation of total annual nitrogen consumption is based on the fact that C. sorokiniana is a warm season species, as is V. chlorellavorus [57]. In outdoor raceways in Arizona, V. chlorellavorus infection events occur during the warm season which runs from March to October.

A reasonable estimate based on previous outdoor experiments [58, 59] is that a system for control of V. chlorellavorus might be required for 16 infection events (100 days) per year during the warm season (March-October). In the prototype ARID raceway at the University of Arizona, the typical culturing process for C. sorokiniana was semi-continuous with 75% biomass harvest at OD750 =

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1.5, which generally occurred 14 days after inoculation of algae culture [58]. This approach is herein adopted for the economic analysis.

The price of nitrogen gas from generators is based on the nitrogen gas flow rate, pressure, purity, and equipment lifetime. In the present study, the pressure swing adsorption (PSA) nitrogen generator (N-750-T, On-Site Gas Systems Inc., Newington, CT) was used as the basis for economic calculation. The N-750-T generator specifications are described in table B.2.

Table B.2.

Onsite nitrogen gas generator specifications (N-750-T, On-Site Gas Systems Inc., Newington, CT)

N2 N2 N2 flow Outlet lifetime Power N2 tank source Purity rate pressure (yrs.) (W) capacity type (%) (L min -1) (atm) (L)

Onsite 95 8,885 5.44 20 17 9,691 generator (continuous PSA service) (N-750-T)

The prices of the generator, nitrogen receiving tank, and annual maintenance were provided by On-Site Gas Systems Inc.: $117,855, $13,670, and $970, respectively. Considering the expected

20-year lifetime of the generator under continuous service, the combined annual cost of the generator and tank is $6,576, neglecting rate of return.

The N-750-T generator provides 8,885 L min -1 (533,100 L h-1) nitrogen gas. An ARID raceway of one-acre (0.4 ha) surface area at 10 cm culture-depth would have a total volume of

400,000 L, which would be held in a sealed canal at night. Assuming that 75% of the culture is harvested when an infection takes place, the volume to be disinfected would be 100,000 L. This

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volume would require 100,000 L * 5 L N gas / L culture = 500,000 L N gas/application/raceway.

Therefore, one N-750-T generator would be enough to treat the whole culture volume with one- hour nitrogen gas sparging/application. For a maximum profitability, the nitrogen generator should produce nitrogen continuously for 24 h d -1. If storage is available or the raceways could be treated sequentially, then 24 ARID raceways (each covering 0.4 ha surface area) could be treated per day with one-hour nitrogen sparging/raceway.

The electricity consumption for the N-750-T generator is only 17 W; thus, the electrical usage per one-hour application per day is 0.017 kWh. Therefore, for 100 days of operation/year, the total energy requirement would be 4,080 kW. At an energy cost of $0.10 kWh-1, the generator energy cost is $408 yr -1. Per raceway cost would be $17 yr -1. It should be noted that the N-750-T nitrogen generator requires high flow compressed air to provide high purity ( ≥ 95%) of nitrogen gas. An OSMAN SA100-air compressor (Shenzhen Osman air compressor manufacture CO.,

LTD, Guangdong, Mainland, China) would provide 12,800 L min -1 at 75 kW. The price of the

OSMAN SA100-air compressor is $6,470, and the cost of energy use is $18,000 yr -1 at an energy cost of $0.10 kWh -1. Thus, per raceway energy cost would be $750 yr -1.

The estimated annual cost of the nitrogen generator (N-750-T) and the air compressor

(OSMAN SA100) with one-hour nitrogen injection and 24 ARID raceways is reported in table

B.3.

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Table B.3. Annual cost of deoxygenation process with an onsite nitrogen generator (N-750-T) and

OSMAN SA100 air compressor in a commercial scale ARID raceway system with 24 raceways each covering 0.4 ha surface area

Generator / raceway costs $/yr $/m 2/yr $/m 2/night Annualized capital cost* 287.50 0.07 0.0007 Electricity** 767 0.19 0.0019 Maintenance 40.41 0.01 0.0001 Total 1,094.91 0.27 0.0027 *Added cost of nitrogen generator, receiving tank, and air compressor spread over 20 years lifetime and service **Electricity cost for both generator and air compressor

This approach with the aforementioned nitrogen sparging management and equipment showed that the deoxygenation process will cost only $0.27/m 2/yr.

4. Conclusion :

The infection process of V. chlorellavorus on C. sorokiniana culture can be effectively reduced or prevented with a deoxygenation control strategy with one-hour nitrogen gas sparging at the beginning of the dark period. Natural dark respiration maintains the oxygen concentration in the anoxic range for the rest of the dark period. In commercial-scale ARID raceways, the cost of deoxygenation treatment using onsite nitrogen generators with 1-h nitrogen sparging would be only $0.27/m 2/yr.

5. Acknowledgements :

The authors are grateful for the U.S. Department of Energy and Regional Algal Feedstock Testbed

(RAFT) project, University of Arizona, for supporting this study.

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